Structural characterisation of the fungal Pmt4 homodimer

Domain: physics
Published: 2025-12-14
Editorial outcome: minor_revision

Abstract
Protein O-mannosyltransferases (PMTs) are conserved endoplasmic reticulum membrane-embedded enzymes responsible for the transfer of mannose from dolichol phosphate-mannose (Dol-P-Man) to serine/threonine-rich protein substrates or unfolded proteins. PMTs from three subfamilies form obligate dimers with different substrate specificities and require the concerted action of their transmembrane domains (TMDs) and a luminal MIR domain for catalysis. Here, we present structures, native mass spectrometry, and structure-based mutagenesis of the fungal Pmt4 homodimer. The core fold of the TMDs and MIR domain is conserved with the Pmt1-Pmt2 heterodimer, indicating a shared catalytic mechanism. Distinct from Pmt4, the MIR domain interacts in cis with the TMDs of the same subunit and has a β-hairpin insertion required for O-mannosylation of substrates. We further identify a cytosolic binding site for substrate Dol-P-Man within the Pmt4 TMDs, which is conserved amongst PMTs and important for in vivo activity. Thus, we provide a framework to understand the substrate specificity and regulation of the Pmt4 homodimer.

Introduction
Protein O-mannosylation is an evolutionarily conserved and essential post-translational modification that occurs in the endoplasmic reticulum (ER) of eukaryotes1. In yeast, the majority of proteins entering the secretory pathway (including 90% of all cell wall proteins) are O-mannosylated2 and unfolded protein O-mannosylation (UPOM) is important for ER protein quality control3, whilst impairment of O-mannosylation in humans leads to congenital muscular dystrophies4. The transfer of mannose from dolichol monophosphate-activated mannose (Dol-P-Man) to the hydroxyl group of serine/threonine residues of substrate proteins is catalysed by membrane-embedded glycosyltransferases of the GT-C superfamily5. The most widely conserved family is the protein O-mannosyltransferases (PMT in yeast, POMT in mammals), which target largely unstructured regions within a range of substrates1,2. More recently, mammalian TMCT1-4 and TMEM260 enzymes were also found to O-mannosylate specific folded domains6.

In Saccharomyces cerevisiae, six active PMT proteins are classified into three families on the basis of sequence similarity (Supplementary Fig. 1): PMT1 (Pmt1 and Pmt5), PMT2 (Pmt2, Pmt3 and Pmt6) and PMT4 (Pmt4)7. These proteins form obligate dimers within or across families8 and, despite a degree of functional redundancy, a Pmt1-Pmt2 heterodimer and Pmt4-Pmt4 homodimer account for the majority of O-mannosylation9. In contrast, a heterodimer comprising POMT1 and POMT2 (PMT4 and PMT2 family members, respectively) is the sole PMT-like O-mannosyltransferase present in mammals10.

The cryo-EM structure of the S. cerevisiae (sc) Pmt1-Pmt2 heterodimer, involved in both UPOM and O-mannosylation of bona fide S/T-rich substrates of the secretory pathway, revealed each subunit to comprise 11 transmembrane domains (TMDs) that share a fold with the catalytic subunits of oligosaccharyltransferases11 and tryptophan C-mannosyltransferases12. A further structure of the scPmt1-Pmt2 complex bound to the donor Dol-P and acceptor peptide confirmed that the active site is located in a luminal pocket of each subunit, where an invariant DE motif within the loop between TMD1 and TMD2 catalyses transfer of the mannose by an inverting mechanism. Aside from the transmembrane region, PMT proteins contain a luminal MIR domain between TMD7 and TMD8, so-called due to their presence in mannosyltransferases, inositol triphosphate receptors and ryanodine receptors13. Within the scPmt1-Pmt2 structure, the scPmt1-MIR domain interacts in trans with the TMDs of scPmt2 next to the active site and the scPmt2-MIR domain is disengaged11, although the dynamic nature of these interactions cannot be inferred. Furthermore, the characteristic β-trefoil fold of PMT2-MIR domains was shown to comprise mannose-specific binding sites, which are in part conserved with carbohydrate-binding modules (CBMs) and important for O-mannosylation of S/T-rich peptides14. This led to a model whereby the MIR domain binds to a mannosylated peptide product to keep it away from the adjoining PMT active site and ensure enzyme processivity.

Currently, there is no structural information available for the PMT4 family of O-mannosyltransferases and it cannot be inferred from the scPmt1-Pmt2 structure how the MIR domains would interact with the TMDs within the Pmt4-Pmt4 homodimer. In contrast to the recognition of both soluble and membrane proteins by Pmt1-Pmt2 and its additional involvement in UPOM15, Pmt4 predominantly mannosylates S/T-rich regions in membrane protein substrates9,16,17, however, the molecular basis of this substrate specificity is also unknown. In addition, yeast Pmt4 provides a powerful model for understanding the significant group of congenital muscular dystrophies linked to reduced O-mannosylation of α-dystroglycan (αDG) by POMT1-POMT2, as it mannosylates αDG-derived peptides and is catalytically impaired by the insertion of pathogenic POMT1 mutations18.

Here, we use a combination of structural biology, native mass spectrometry and in vivo functional assays to characterise the fungal Pmt4 homodimer. The MIR domain of Pmt4 interacts in cis with the TMDs of the same subunit and contains a unique β-hairpin insertion important for O-mannosylation of substrates. The structure of Dol-P bound within the Pmt4 luminal active site highlights a strictly conserved catalytic mechanism between PMT families. Surprisingly, our data reveal an additional binding site for Dol-P-Man on the cytosolic side of the membrane, which we show to be important for the O-mannosyltransferase activity of Pmt4.

Results
The structure of the Pmt4 homodimer
To investigate the structure of Pmt4, we took advantage of its thermostable homologue from the thermophilic fungus Chaetomium thermophilum, which has 48% sequence identity to yeast Pmt4 (Supplementary Fig. 1). C. thermophilum Pmt4 (ctPmt4) was overexpressed in S. cerevisiae and purified via a C-terminal deca-histidine tag in detergent lauryl maltose neopentyl glycol (LMNG) supplemented with cholesteryl hemisuccinate (CHS) (Supplementary Fig. 2A). Structural analysis by single-particle cryo-electron microscopy (cryo-EM) confirmed ctPmt4 is a homodimer (Fig. 1A), allowing processing of the data with either no symmetry (C1) or two-fold rotational symmetry (C2) imposed (Supplementary Fig. 3A, B and Table 1). Within the membrane region, the 3.2 Å C2 map showed the same features as the 3.4 Å C1 map but at higher resolution (Supplementary Fig. 3C), and was therefore used to build a two-fold symmetric model for the ctPmt4 transmembrane domains (TMDs) (Supplementary Fig. 4A). In contrast, the luminal MIR domains were arranged asymmetrically and only resolved in the C1 reconstruction at a resolution that precluded direct model building (Supplementary Fig. 3C). Therefore, we purified the ctPmt4-MIR domain from Escherichia coli via a N-terminal hexa-histidine tag (Supplementary Fig. 2B) and solved the crystal structure at 1.2 Å resolution by molecular replacement (Table 2) (Supplementary Fig. 4B). Our final composite model for the ctPmt4 homodimer (Fig. 1A, B) was derived from rigid body fitting these high-resolution structures for the TMDs and MIR domain into the C1 reconstruction and directly building models for their connecting linkers into the remaining density.

The 11 TMDs of ctPmt4 have a highly similar arrangement to those of scPmt1 and scPmt211, superimposing with a root-mean-square deviation (RMSD) of 1.6 Å and 1.4 Å respectively (Fig. 1C). Moreover, the ctPmt4 homodimer adopts the same overall morphology within the membrane region as the scPmt1-Pmt2 heterodimer, with a rhombic cavity separating the dimer halves (Fig. 1B). As in scPmt1-Pmt2, residues at the cytosolic ends of TMD6 (W263 and L266) and TMD8 (R629 and R630) form the major dimer interface, bounding the rhombic cavity on two sides at the cytosolic side of the membrane (Fig. 1D). However, the luminal ends of TMD9 from each dimer half are tilted further towards each other, resulting in an additional single constriction point at F662 and V663 and a smaller rhombic cavity than for scPmt1-Pmt2 (Fig. 1B, D). Interestingly, the residues at these dimer interfaces are largely conserved (Supplementary Fig. 1) and thus do not provide a satisfactory explanation for highly specific Pmt4-Pmt4 or Pmt1-Pmt2 dimer formation. Therefore, the N-terminus of scPmt1 and the interfacial lipid, which additionally contribute to the scPmt1-Pmt2 dimer interface11 and are not resolved in the ctPmt4 map, could be key elements that drive exclusive heterodimer formation.

The luminal MIR domains of ctPmt4 adopt a strikingly different arrangement with respect to the TMDs, compared to within the scPmt1-Pmt2 heterodimer; whilst the scPmt1- and scPmt2-MIR domains are positioned to interact in trans with the TMDs of the opposing dimer subunit, the first ctPmt4-MIR domain is rotated by 30o relative to the membrane and consequently interacts in cis with the TMDs from the same dimer subunit (Fig. 1B, C). Interestingly, whilst the second ctPmt4-MIR domain is also positioned in cis, it is rotated a further 30o so that it no longer interacts with the TMDs, akin to the scPmt2-MIR domain. Therefore, despite major differences in the cis/trans nature of their interactions, the MIR domains of both the ctPmt4 homodimer and scPmt1-Pmt2 heterodimer have a similar asymmetric arrangement, whereby one MIR domain is attached to the TMDs and the other is detached from them (Fig. 1B). As a result of this cis arrangement of the ctPmt4-MIR domains, the homodimer is maintained solely by the interactions defining the rhombic membrane cavity and perhaps explain why an additional luminal contact is present (Fig. 1D).

Pmt4-MIR domains have a functionally important β-hairpin insertion
Previously, high-resolution crystal structures were solved and functionally annotated for MIR domains from S. cerevisiae PMT2 family members14. Therefore, in order to be able to compare PMT2 and PMT4 family MIR domains from the same species, we additionally purified the scPmt4-MIR domain (Supplementary Fig. 2C) and solved the crystal structure to 1.3 Å resolution (Table 2) (Supplementary Fig. 4B). Aside from minor conformational differences in the β4−β5 and β8−β9 loops, the scPmt4- and ctPmt4-MIR domain structures are highly similar (RMSD 0.87 Å over 194 Cα-atoms) (Fig. 2A) and comprise the characteristic peculiar β-trefoil fold i.e. six β-hairpins arranged within a pseudo-threefold symmetry (three MIR-motifs: MIRm1, 2, 3), altogether forming a β-barrel and a triangular cap (Fig. 2B).

Overall, the scPmt4-MIR domain closely resembles scPmt2 in its core fold (RMSD 1.38 Å over 168 Cα-atoms) (Fig. 2C) and accordingly putative binding sites for O-mannosylated peptides could also be located in each MIR-motif (α, β, γ and δ) (Fig. 2B). The highly conserved LH(S/T)H and DxNN (x: any residue) fingerprints in MIRm1 and MIRm2 define the α and β sites respectively (Supplementary Fig. 5), which are likely to form mannose-specific binding sites similar to other CBMs19. However, the fungal Pmt4 structures have a short three-residue insertion (ct: E449-Y451, sc: E442-D444) in MIRm2 relative to scPmt2 that is part of a unique α-helix (αMIR) of unknown function (Fig. 2B and Supplementary Fig. 5). Unlike MIRm1 and MIRm2, MIRm3 displays higher sequence divergence across PMT families and deviates from scPmt2 in exposed loop regions (Fig. 2C and Supplementary Fig. 5). Interestingly, whilst in the scPmt2-MIR the sugar-binding γ site of MIRm3 is sterically blocked, the γ site within the scPmt4-MIR is open and comprises a LWTH motif more similar to the α and β sites (Supplementary Fig. 5). In addition, the PMT4 family MIR domains (including the homologous human POMT1-MIR) do not contain the PMT2-MIRm3 δ site insertion14. Instead, three exposed loops, β1−β2, β8−β9 and β11−β12, are positioned around a central cavity that corresponds to the PMT2-MIRm3 δ site, which we altogether define as a distinct Pmt4-MIR δ site (Fig. 2B and Supplementary Fig. 5).

Characteristically, the Pmt4-MIR domain structures reveal a unique β-hairpin insertion (β-ins) of seven residues (RY(D/E)(D/N) GRΦ) in β2−β3 of MIRm1 (Fig. 2B, C and Supplementary Fig. 5). β-ins is extensively stabilised by internal hydrogen-bonding and a central salt-bridge between the conserved 4th and 6th residue (ct: D370:R372, sc: D364:R366) (Fig. 2D). It is fixed on top of the cap scaffold by hydrogen-bonding of the tyrosine (ct: Y368, sc: Y362) to a glutamate (ct: E441, sc: E434), and of the centrally-fixed arginine to the main chain around a proline (ct: P434, sc: P427) within MIRm2. The tyrosine also stacks onto this proline and is part of an aromatic cluster within the cap (including ct: Y364 and W501, sc: Y358 and W496).

To assess the importance of β-ins on the O-mannosyltransferase activity of Pmt4 in vivo, yeast strains were created that express either FLAG-tagged wild-type or mutated scPmt4 (scPmt4-FLAG), specifically with a deletion of β-ins (Δ362-368) or alanine substitutions of residues crucial for stabilising its fold (Y362A/D364A/R366A (YxDxR), P427A/P430A (PP) and W496A) (Fig. 2D). Notably, using nano differential scanning fluorimetry (nanoDSF), we found that the isolated scPmt4-MIR domain with the most severe Δ362-368 mutation was folded, albeit with a slightly reduced stability relative to the wild-type domain (Supplementary Fig. 6A). All full-length scPmt4-FLAG variants were subsequently shown to be integrated in the membrane at the same level as wild-type scPmt4-FLAG (Supplementary Fig. 6B), allowing them to be tested using two assay systems. Firstly, we took advantage of the synthetic temperature sensitivity of the double deletion strain Δpmt1pmt4, which only grows at 37 °C when complemented with enzymatically active scPmt4 (or scPmt1)20. Whilst wild-type and PP scPmt4-FLAG complement the temperature-sensitive phenotype, none of the other mutants restore thermal tolerance to Δpmt1pmt4 (Supplementary Fig. 6C), indicating that these scPmt4 variants have little or no mannosyltransferase activity. Secondly, we used the cell wall protein Ccw5 as a reporter, as it shows a markedly altered glycosylation pattern when scPmt4 activity is impaired16. In Δpmt4-CCW5 cells expressing wild-type scPmt4, O-mannosylation in a S/T-rich region prevents N-glycosylation of residue N114 and Ccw5 has an apparent molecular mass of ~40 kDa. In contrast, lack of scPmt4-based O-mannosylation allows N-glycosylation of Ccw5, resulting in a shift in apparent molecular mass to ~90–120 kDa. Consistent with the temperature-sensitive growth phenotype, the glycosylation pattern of Ccw5 isolated from yeast strains expressing scPmt4 β-ins mutant proteins Δ362-368, YxDxR and W496A is very similar to that of the null mutant pmt4 (Fig. 2E), demonstrating that the enzyme function is largely lost in vivo. Therefore, β-ins is generally required for O-mannosylation of substrate proteins by Pmt4.

As observed previously in the PMT2-MIR crystal structures14, the Pmt4-MIR domains are extensively bound to small polar ligands present in the crystallisation solutions (acetate or HEPES) or cryoprotectant buffers (glycerol or ethylene glycol) (Fig. 2B and Supplementary Fig. 7A). In particular, the α and β sites, which were demonstrated for scPmt2 to bind O-mannosylated peptides14, are again occupied by ligands in ctPmt4-MIR and scPmt4-MIR respectively, in line with the expectation that they are also sugar binding sites. Given that both the open γ site and β-ins of ctPmt4-MIR are also bound to acetate ligands (Fig. 2B, D and Supplementary Fig. 7A), these too may be mannose binding sites, which would indeed provide an explanation for the importance of β-ins in O-mannosylation by Pmt4. Furthermore, the ctPmt4- and scPmt4-MIR domains have an asymmetric surface charge distribution with both a highly positively and negatively charged patch (Supplementary Fig. 7A). Strikingly, the bound ligands are aligned in a linear trace starting from the positively charged surface at the δ and γ sites, extending through β-ins and up to the α and β sites and the rather negatively charged surface (Supplementary Fig. 7A). This suggests a defined substrate/product peptide route, in line with the model that PMT-MIR domains coordinate a mannosylated peptide to ensure a processive enzyme reaction14.

The active site of PMTs is located in a deep cavity on the ER luminal side of the TMDs, and it has been postulated that the TMD-MIR interface plays a role in guiding substrates to or products away from the active site11,14. As in the scPmt1-Pmt2 heterodimer, the δ site of the ctPmt4-MIR domain is positioned at the TMD-MIR interface, opposing the active site (Supplementary Fig. 7B). In the attached ctPmt4 TMD-MIR interface, the active site is completely covered by this δ site, whilst in the detached conformation, the MIR domain is rolled aside and the active site is fully accessible (Supplementary Fig. 7C). However, PMT2-specific residue insertions in the δ and δ’ sites, which in the scPmt1-Pmt2 heterodimer are primarily responsible for TMD-MIR contacts, are absent in PMT4 sequences11,14. Instead, the attached MIR domain directly contacts the TMDs via the β8−β9 loop, whereby exposed histidine and charged residues (E465, H466, K468) form a relatively weak interface with aromatic residues of the TMDs (F319, H561, W562) (Supplementary Fig. 7D). The rolling movement between the attached and detached state of the MIR domain is likely to be facilitated both by having this weak δ site interaction and the flexible ~20 residue linkers connecting the MIR domains and TMDs, which are arranged asymmetrically in our structure (Fig. 1A). Indeed, the poor resolution of the ctPmt4-MIR domains in the C1 cryo-EM reconstruction demonstrates that they are rather dynamically associated with the TMDs (Supplementary Fig. 3C). Although in the scPmt1-Pmt2 structure the MIR domains act in trans, their general orientation and surface properties are corresponding11. Thus, whilst the full trajectories and conformational space of the Pmt4-MIR domains are yet to be clarified, the alternate association with their TMDs likely acts as a general mechanism to regulate access or catalysis at the active site.

The active site is highly conserved across PMT families
Our cryo-EM reconstructions revealed two single-chain lipid species bound within each membrane domain of the ctPmt4 homodimer, arranged so that the hydrocarbon tails protrude into the central rhombic cavity (Fig. 3A and Supplementary Fig. 8A). The head group of one lipid species is positioned in the luminal membrane leaflet in the active site and was thus modelled as the reaction product Dol-P, as in the scPmt1 structure11. Similarly, only the first seven isoprene repeats of the dolichol chain are resolved in the density and these are accommodated within the membrane in a hydrophobic groove formed by residues from TMD5 (I241), TMD6 (Y245, L248, F251) and TMD7 (F284, I292, F296, Y299) (Supplementary Fig. 8B). The Dol-P phosphate group is also stabilised by the conserved residues H105, K244, R666 and H672 (Supplementary Figs. 1 and 8C), however is retracted 4.7 Å from the acceptor threonine binding site compared to within scPmt1 (Fig. 3B). Accordingly, the R666 side chain is rotated 60o to stabilise this alternative position of the Dol-P phosphate, similar to its position in scPmt2 where Dol-P was not bound (Supplementary Fig. 8C). Therefore, the dolichol product of the O-mannosylation reaction appears to be positioned closer to its putative release pathway. This implicit flexibility likely also explains why the density for the Dol-P phosphate group is consistently weaker than for the dolichol tail (Supplementary Fig. 8A)11.

Superimposition of ctPmt4 with both scPmt1 and scPmt2 indicates that the remaining residues for binding the donor dolichol and those expected to bind the acceptor threonine (F83, D84, H87 and F669) are conserved in their positions in the active site (Fig. 3B and Supplementary Fig. 8C). In addition, comparison with the bacterial oligosaccharyltransferase PglB21 indicates that D84 and D197 are positioned appropriately for the coordination of a Mn2+ ion (Supplementary Fig. 8D), however, like in the scPmt1-Pmt2 structure11, we did not observe clear density for a catalytic ion. Taken together, the structural similarities indicate that the catalytic mechanism is strictly conserved across PMT families.

Aside from the conserved interactions with the acceptor threonine residue, it was previously observed that scPmt2 does not specifically interact with the remainder of the bound PYT substrate peptide11. Regardless, the equivalent substrate binding pocket of ctPmt4 does show some differences in peptide-proximal residues that could contribute to substrate recognition (Fig. 3C). Most notably, the ctPmt4 δ’ site within the MIR linker does not contain the four residue insertion specific to the PMT2 family14 and, as such, adopts a distinct conformation that brings it much closer to the expected position of the substrate peptide. However, the ctPmt4 peptide-proximal residues and δ’ site are virtually identical to those within scPmt1 (Supplementary Fig. 8E). This suggests that these features are unlikely to be strong determinants of substrate specificity, as would be expected given PMTs do not recognise their substrates via defined sequence motifs2.

Interestingly, the attached MIR domains in the ctPmt4 homodimer and scPmt1-Pmt2 heterodimer occupy significantly different positions relative to the TMDs, with the ctPmt4-MIR domain sitting much closer to the substrate peptide binding site (Fig. 3C). It is therefore tempting to speculate a role for the Pmt4-MIR domain in selecting membrane protein substrates17, either through direct substrate recognition or by sterically favouring membrane protein access to the active site.

Pmt4 binds Dol-P-Man on the cytosolic side of the membrane
The head group of a second single-chain lipid is bound within a positively charged pocket adjacent to the cytosolic end of TMD9 (Fig. 3A). To identify the lipid species, ctPmt4 was purified in dodecyl maltoside (DDM) detergent (Supplementary Fig. 2A) and subjected to native mass spectrometry (native MS). The resulting spectrum confirms that ctPmt4 also forms a homodimer in this detergent, and shows that both monomer and dimer are associated with adduct peaks corresponding to bound ~1100 Da and ~1700 Da species (Fig. 4A and Supplementary Table 1). Subsequent lipidomics analysis revealed that, in addition to a mixture of the common <1000 Da phospholipids present in the S. cerevisiae ER membrane (Supplementary Fig. 9A, B), >1000 Da Dol-P-Man and cardiolipin are associated with the purified protein (Fig. 4B and Supplementary Fig. 9C). Further dissociation of bound lipid species during native MS indicated that the ~1100 Da adduct peaks correspond to Dol-P-Man with an average of 13 isoprene units (Supplementary Fig. 9D). As observed previously22, the mitochondrial lipid cardiolipin (~1400 Da) is likely present as an artefact of protein purification from total cellular membranes, suggesting that detergent extraction can to some extent alter native lipid composition. Intriguingly, cardiolipin (~1400 Da) is not large enough to fully account for the ~1700 Da adduct peaks observed in native MS; these are therefore likely to correspond to a detergent adduct with a lipid, for example, cardiolipin or the Dol-P present in the active site.

Strikingly, we found that Dol-P-Man is a good fit for the lipid density in the cytosolic binding pocket, with the phosphate group being stabilised by positively charged residues (R218, R638 and R642) and the mannose group by histidine side chains (H219 and H637) (Fig. 3D and Supplementary Fig. 8A). This coordination is highly analogous to the binding of Dol-P product, and ergo Dol-P-Man substrate, in the active site of PMT proteins (Fig. 3B and Supplementary Fig. 8C). Similarly, the density for the head group is weaker than for the dolichol tail, implying some flexibility within the Dol-P-Man binding site. To further verify this Dol-P-Man interaction, alanine mutants of the coordinating arginine and histidine residues in the ctPmt4 cytosolic pocket were purified and were confirmed using nanoDSF to have the same thermal stability in DDM as the wild-type protein (Supplementary Fig. 10A). Subsequently native MS showed that, whilst single arginine or double H219A/H637A mutations were insufficient to prevent Dol-P-Man binding (Supplementary Fig. 11), double R218A/R642A or triple R218A/R638A/R642A (RRR) mutations abolished the interaction (Fig. 4C and Supplementary Fig. 10), confirming that Dol-P-Man is the single-chain lipid species bound within the cytosolic pocket of ctPmt4.

Only the first six isoprene units of Dol-P-Man are resolved in the density, whilst lipidomics analysis shows that species with up to 17 units are associated with ctPmt4 (Fig. 4B and Supplementary Fig. 9C). This suggests that more than half of the dolichol tail is disordered within the central rhombic cavity, as for Dol-P within the active site. Of the ordered portion, the first two isoprene repeats are accommodated in a hydrophobic groove along the membrane normal between TMD4, TMD6 and TMD9 (Fig. 3A and Supplementary Fig. 8B). However, this trajectory is interrupted by the side chain F649, which sits in the groove and forces the dolichol tail to kink 120o (Fig. 3E). The remaining resolved isoprene repeats then adopt an alternative path along the membrane plane, which crosses hydrophobic surfaces of TMD8 and TMD9 (Fig. 3A, E and Supplementary Fig. 8B). In fact, the dolichol tail passes so close to G648 within TMD9 that we reasoned a larger side chain in this position would sterically impede Dol-P-Man binding. Indeed, native MS showed that Dol-P-Man was no longer bound to a G648L mutant of ctPmt4, despite it exhibiting the same thermal stability as the wild-type protein (Fig. 4D and Supplementary Fig. 10A, C). Therefore, the Dol-P-Man interaction requires both stabilisation of its dolichol tail in the observed conformation and its phosphate-mannose headgroup in the cytosolic binding pocket.

Dol-P-Man binding to a cytosolic pocket is functionally important for Pmt4
We next wanted to assess whether the observed binding of Dol-P-Man to the cytosolic pocket of ctPmt4 is important for its function. First, the O-mannosyltransferase activity of ctPmt4 in S. cerevisiae was demonstrated through complementation of the Δpmt1pmt4 mutant with ctPmt4 expressed from a constitutive ADH1 promoter. The in vivo functionality of ctPmt4 was similar to scPmt4, restoring normal growth at elevated temperatures (Supplementary Fig. 12A) and alleviating the clumping phenotype observed in the S. cerevisiae Δpmt1pmt4 mutant (Supplementary Fig. 12B). HA-tagged ctPmt4 (ctPmt4-HA) was then shown to mannosylate the previously described chimeric scPmt4 substrate FUS-Axl2TMZZ in Δpmt4 cells17 (Fig. 5A), providing an assay system for in vivo ctPmt4 activity. We verified the stability and membrane integration of RRR, G648L and H219A/H637A ctPmt4-HA variants in the Δpmt4 strain (Supplementary Fig. 12C) and found that cells expressing RRR or G648L mutants of ctPmt4-HA, which cannot bind Dol-P-Man at the cytosolic pocket, accumulated hypo-mannosylated FUS-Axl2TMZZ at ~70 kDa compared to those expressing wild-type or H219A/H637A ctPmt4-HA (Fig. 5A). Whilst constitutive expression of RRR and G648L ctPmt4-HA did not impair the growth of Δpmt4 (Supplementary Fig. 12D), overexpression of these mutants from a GAL promoter had a dominant negative effect, leading to impaired cell growth (Fig. 5B) and increased cell clumping (Supplementary Fig. 12E) relative to overexpression of wild-type His-tagged protein (ctPmt4-His). However, FUS-Axl2TMZZ is hypo-mannosylated only in the presence of overexpressed G648L ctPmt4-His, implying that increased protein amounts compensate for the mannosylation defect of the RRR ctPmt4-His variant (Supplementary Fig. 12F). Nevertheless, altogether these results show that Dol-P-Man binding to the cytosolic pocket of ctPmt4 impacts its O-mannosyltransferase activity.

We were intrigued whether cytosolic Dol-P-Man binding is a general feature important for protein O-mannosylation by PMTs, given that residues involved in Dol-P-Man binding to ctPmt4 are not strictly conserved (Supplementary Fig. 1). Strikingly, a comparison with the AlphaFold DB23,24 or cryo-EM models for yeast and human PMT4 (scPmt4, hsPOMT1), PMT1 (scPmt1) and PMT2 (scPmt2, hsPOMT2) family members revealed a common positively charged cytosolic cavity in the same position as the Dol-P-Man binding site in ctPmt4, particularly in the vicinity of the phosphate group (Fig. 5C and Supplementary Fig. 13A). Furthermore, a similar feature also appears to be present within the distinct O-mannosyltransferases TMTC1-4 and TMEM260 found in mammals6 (Supplementary Fig. 13B). To assess whether this cavity is important for the enzymatic activity of scPmt4, alanine substitution mutants of the basic residues K212, R216 and K635 were expressed in a Δpmt4-CCW5 reporter strain. Whilst the stability and membrane integration of these variants in vivo was unchanged relative to wild-type scPmt4-FLAG (Supplementary Fig. 12G), expression of K635A and K212A/K635A scPmt4-FLAG lead to an increase in N-glycosylated Ccw5, demonstrating that these variants are impaired for protein O-mannosylation (Fig. 5D). These results provide a compelling argument for the conserved cytosolic cavity being functionally important at least within fungal Pmt4 and potentially beyond, although its role in binding Dol-P-Man needs to be verified for other PMTs. Interestingly, the cytosolic N-termini, which in our ctPmt4 homodimer structure are not resolved and likely flexible, at least partially occupy the cytosolic cavity within scPmt1 and scPmt4 models (Fig. 5C and Supplementary Fig. 13A) and thus could act as an alternative natural ligand to Dol-P-Man.

Methods
Construct design
All plasmids used in this study are shown in Supplementary Table 2. For recombinant protein expression, wild-type full-length ctPmt4 was cloned with a C-terminal 3C-H tag downstream of the bidirectional GAL1/GAL10 promoter in the pMT929 vector22 (pMT929 ctPmt4). The scPmt4-MIR domain (residues 330-540), scPmt4-MIR Δ362-368 variant and ctPmt4-MIR domain (residues 337-537) were cloned into a pET24d vector (Novagen) with an N-terminal MKH PM tag. pMT929 ctPmt4 point mutations were subsequently created using the Quikchange Lightning site-directed mutagenesis kit (Stratagene).

For in vivo functional studies of scPmt4, K212A, R216A, K635A and K212A/K635A point mutations were introduced into plasmid pJK4-B18 using Quikchange Lightning site-directed mutagenesis kit, taking care to culture all subsequent E. coli transformants at 20 °C. Recombinant PCR31 was used to introduce Δ362-368, Y362A/D364A/R366A, P427A/P430A and W496A mutations into pJK4-B1, with the resulting DNA fragments being cloned into NcoI/EcoNI linearised plasmid via homologous recombination in yeast. For constitutive expression in yeast strains, ctPmt1 and ctPmt4 were amplified from cDNA and cloned with a C-terminal HA tag into the pREP3-adh vector digested with NotI/XhoI or NotI/BamHI, respectively. Point mutations were introduced into pREP3-adh ctPmt4-HA using Quikchange Lightning site-directed mutagenesis. All constructs were routinely validated through sequence analysis.

S. cerevisiae strain construction
All S. cerevisiae strains used in this study are shown in Supplementary Table 3. To create Δpmt4-CCW5, strain pmt4::TRP132 was transformed33 with plasmid pML7825. To create strain pmt4-G, the PMT4 deletion cassette was amplified from strain pmt4::TRP132 and transformed into DS1-2b34. Transformants were selected on synthetic complete (SC) medium-Trp and verified by PCR analyses. Strain pmt4-G was subsequently transformed with plasmids pML7825 and pJH117 to create the reporter strains pmt4-G/CCW5 and pmt4-G/AXL, respectively.

Growth of strains
For recombinant protein expression, pMT929 ctPmt4 variants were transformed into DS1-2b34 and plated on SC-Trp plates for 48 h, before replating on SC-Leu plates for 24 h. A stationary SC-Leu pre-culture containing raffinose as the carbon source was diluted 12.5-fold into YP medium containing 1% galactose and incubated at 30 °C for 24 h. The Pmt4-MIR domains were expressed in E. coli Rosetta 2 (DE3) (Novagen) by autoinduction35 overnight at 18 oC. All cells were harvested by centrifugation and stored at −20 oC until further use.

Yeast strain CFY38 was transformed with pREP3-adh plasmids (scPmt1-HA, scPmt4-HA, ctPmt1-HA, ctPmt4-HA) and grown at 25 °C in SC -Leu medium until OD ~1. From these cultures, light micrographs were taken, then a total of 105 cells and 10-fold serial dilutions thereof were spotted onto YPD plates, which were incubated for 3 days at 25 °C and 35 °C, respectively. pREP3-adh ctPmt4-HA variants or pMT929 ctPmt4 variants were transformed into yeast strains pmt4-G or pmt4-G/AXL and selected on SC-Leu or SC-Leu/-Ura media respectively. Verified transformants were grown at 30 °C in SC selective medium supplemented with 2% glucose (pREP3-adh ctPmt4-HA variants) or 2% galactose (pMT929 ctPmt4 variants). For growth curves of pmt4-G strains, the OD was measured at 60 min intervals and light micrographs were taken during logarithmic growth.

Yeast strains Δpmt1pmt4, Δpmt4-CCW5 or pmt4-G/CCW5 were transformed with pRS423 empty vector, pJK4-B1 plasmid or mutated variants and selected on SC-His medium. Three individual colonies of each Δpmt1pmt4 transformant were grown at 25 °C and 37 °C on YPAD plates for 3 days.

Preparation of cell extracts and western blotting
Whole cell extracts and crude membranes were prepared for analysis of Pmt4 expression as previously described15. Cell wall extracts were prepared for analysis of Ccw5 glycosylation as previously described16. The Pmt4 substrate protein FUS-Axl2TMZZ was analysed according to the protocol from Hutzler and coworkers17. Western blot analysis was performed as detailed by Bausewein and coworkers18. Monoclonal mouse anti-FLAG M2 (#F3165, Sigma-Aldrich) and anti-HA (#2362, Cell Signalling) antibodies were used at a dilution of 1:5000 and ZZ tag peroxidase-coupled anti-rabbit-IgG antibody from goat (#A6154, Sigma-Aldrich) was used at a dilution of 1:10,000. GAPDH-directed antibodies (#A9521, Sigma-Aldrich) and Sec61-directed antibodies (gift from K. Römisch) derived from rabbit were used at a dilution of 1:5000 and 1:2500, respectively. As secondary antibodies, peroxidase-coupled anti-rabbit-IgG antibody from goat (#A6154, Sigma-Aldrich) and anti-mouse antibody from rabbit (#A9044, Sigma-Aldrich) were used at a dilution of 1:10000 and 1:5000 respectively.

Protein purification
For the purification of full-length ctPmt4 variants, S. cerevisiae cell pellets were lysed in 50 mM HEPES (pH 7.5), 500 mM NaCl, 10% (v/v) glycerol, 1 mM EDTA by beating with glass beads at 500 rpm for 3 × 4 min. The lysate was cleared by centrifugation at 1000 x g for 25 min and the cell membranes were harvested and washed once by centrifugation at 160,000 x g for 30 min. The membranes were solubilised for 30 min in solubilisation buffer (50 mM HEPES (pH 7.5), 500 mM NaCl) supplemented with either 0.5% (w/v) LMNG (Anatrace) or 0.5% (w/v) DDM (Anatrace) for cryo-EM or native MS applications, respectively. These detergents were present in all subsequent buffers at a concentration closer to the critical micelle concentration: either a 0.01% LMNG/0.001% (w/v) CHS mixture (Anatrace) for cryo-EM or 0.03% (w/v) DDM for native MS. Solubilised membranes were centrifuged at 160,000 x g for 30 min and the supernatant was incubated with Ni-NTA agarose (Qiagen) for 30 min at 4 °C. This resin was washed with 40 column volumes (CV) wash buffer (solubilisation buffer with 30 mM imidazole) and ctPmt4 was eluted by on-column cleavage via incubation with 5 mg His6-tagged HRV-3C Protease in 5 CV wash buffer for 1 h at 4 °C. Size exclusion chromatography (SEC) was performed with a Superdex 200 increase 10/30 GL column (Cytiva) equilibrated with 20 mM HEPES (pH 7.5), 200 mM NaCl. Protein-containing fractions were concentrated with a VivaSpin Protein Concentrator MWCO 100,000 (Cytiva) to 20 μM for native MS.

For the purification of Pmt4-MIR domains, E. coli cell pellets were lysed in lysis buffer (50 mM HEPES (pH 7.5), 300 mM NaCl, 30 mM imidazole) using a M-110L Microfluidiser (Microfluidics). The lysates were cleared by centrifugation at 48,000 x g for 25 min, and the supernatant was applied to a 1 ml HisTrap HP column (Cytiva). The column was washed with 20 CV 50 mM HEPES (pH 7.5), 1 M NaCl, 30 mM imidazole, followed by 5 CV lysis buffer. The MIR domains were eluted with 50 mM HEPES (pH 7.5), 300 mM NaCl, 250 mM imidazole and applied to a HiLoad 16/60 Superdex 75 pg (Cytiva) column equilibrated in SEC buffer. The SEC buffer for scPmt4-MIR was 50 mM MES (pH 6.0), 400 mM NaCl for x-ray crystallography and 20 mM HEPES (pH 7.5), 150 mM NaCl for nanoDSF, whilst the SEC buffer for ctPmt4-MIR was 20 mM HEPES (pH 7.5), 150 mM NaCl. Protein-containing fractions were concentrated with a 10 kDa Amicon-Ultra centrifugal filter (Merck Millipore).

X-ray structure determination
All crystallisation was performed via the sitting drop vapour diffusion technique at 18 °C. The scPmt4-MIR domain was concentrated to 20.4 mg/ml and used to set 400 nl drops in a 1:1 ratio with 0.1 M HEPES (pH 7.5), 20% (w/v) PEG 8000. Crystals were cryo-protected by soaking in mother liquor supplemented with 20% (v/v) glycerol and data were collected at beamline ID30A-1 at the European Synchrotron Radiation Facility (ESRF, Grenoble). Data were integrated with XDS36 and scaled with AIMLESS37. Molecular replacement was performed in Phaser-MR38 using a single molecule of Arabidopsis thaliana SDF2-like protein (PDB 3MAL) as a search model. Coot39 and phenix.refine40 of the Phenix package (version 1.15.2)41 were used for iterative model building and refinement cycles.

The ctPmt4-MIR domain was concentrated to 40 mg/ml and used to set 300 nl drops in a 1:1 ratio with 1.4 M sodium acetate (pH 7.0), 0.1 M sodium cacodylate (pH 6.5). The crystal was cryo-protected by soaking in mother liquor supplemented with 20% (v/v) ethylene glycol and data were collected at beamline ID30A-3 at the ESRF. Data were integrated with XDS and scaled with AIMLESS and STARANISO within the autoPROC toolbox42. Molecular replacement was performed in Phaser-MR of the Phenix package using the scPmt4-MIR domain as a search model, then model building and refinement as for the scPmt4-MIR structure. Crystallographic data, refinement and model statistics are summarised in Table 2.

Cryo-EM grid preparation and data collection
ctPmt4 was used at 1.7 mg/ml after SEC without further concentration and 3 μl sample was applied to a glow-discharged holey carbon-coated grid (Quantifoil 300 mesh, Cu 1.2/1.3), before plunge-freezing in liquid ethane using a Vitrobot Mark IV (Thermo Fisher). Freezing was performed at 100% humidity and 6 °C with 15 s adsorption, 5 s blot time, 10 blot force and 1 s drain time and grids were stored in liquid N2 until data collection. Data were collected at beamline CM01 at the ESRF on a 300 kV Titan Krios transmission electron microscope on a K2 detector (Gatan). 9879 images were collected at 0.827 Å/pixel and a magnification of 165,000, with a total dose of 42.7 e-/Å2 over 40 frames. EPU (Thermo Fisher) was used to setup and monitor data collection.

Image processing
Motion correction was carried out with dose-weighting using the Relion 3.143 implementation of MotionCor244 and CTF estimation of unweighted micrographs was performed via GCTF45. 2,199,173 particles were picked using WARP46 and imported into cisTEM 1.0 beta47 for 2D classification followed by 3D auto-refinement of the selected classes into a single class. These centred particles were imported into Relion 3.1 for all subsequent processing. 3D classification (with image alignment) and 3D auto-refinement were followed by another round of 3D classification (without image alignment) and 3D auto-refinement. For the map without applied symmetry (C1), Bayesian polishing, 3D auto-refinement and post-processing yielded a 3.4 Å reconstruction. With C2 symmetry applied, Bayesian polishing, 3D auto-refinement, 3D classification without image alignment, 3D auto-refinement and post-processing yielded a 3.2 Å reconstruction. Both reconstructions were low-pass filtered to 6.0 Å to yield more continuous density for flexible, low-resolution regions. Local density sharpening of the C1 map was performed with LocScale48 in order to represent each region at its local resolution. The data processing pipeline is summarised in Supplementary Fig. 3A.

Model building and refinement
A SWISS-MODEL homology model49 for the ctPmt4 TMDs was generated from the scPmt2 TMDs (PDB 6P25) and sequence alignment in Supplementary Fig. 1, then jiggle-fit50 into one subunit within the C2 reconstruction in Coot. Loop and helical deviations were manually repositioned to fit the density and the L566-P571 loop was built within the C2 map low-pass filtered to 6.0 Å. Models and geometry restraint files for Dol-P and Dol-P-Man were generated using eLBOW51 of the Phenix package (version 1.15.2) then also fitted into the C2 density within Coot (version 0.8.9.2). The complete resulting model for one ctPmt4 subunit was then duplicated and jiggle-fit into the density for the second subunit within the C2 reconstruction. Real-space refinement in Phenix40 with secondary structure, geometry and C2 symmetry restraints produced the final C2 model for the ctPmt4 TMDs (Table 1). Subsequently, this TMD model and two copies of the ctPmt4-MIR domain structure (with ligand, water molecules and flexible termini removed) were jiggle-fit into their corresponding densities within the C1 reconstruction in Coot. Finally, the linkers connecting the MIR domains and TMDs were manually built and real-space refined in Coot within the C1 map low-pass filtered to 6.0 Å. This produced the final composite model for the ctPmt4 homodimer, which was validated in Phenix52 against the C1 reconstruction (Table 1). Structural figures were prepared in UCSF ChimeraX (version 1.10)53 and PyMOL (version 2.5; Schrödinger, LLC).

nanoDSF
Full-length ctPmt4 variants purified in 20 mM HEPES (pH 7.5), 200 mM NaCl, 0.03% (w/v) DDM for native MS were diluted to 0.3 mg/ml and two repeats for each variant were loaded into Prometheus standard capillaries (NanoTemper Technologies). The Prometheus NT.48 (NanoTemper Technologies) was used to monitor the temperature-dependent change in tryptophan fluorescence at 330 nm and 350 nm emission wavelengths, with a 1.5 °C/min temperature gradient from 20 °C to 90 °C. The melting temperature (Tm) was determined by detecting the maximum of the first derivative of the fluorescence intensity ratio at 350 nm and 330 nm. scPmt4-MIR domain variants purified in 20 mM HEPES (pH 7.5), 150 mM NaCl were diluted to 1 mg/ml and three repeats for each variant were measured as for ctPmt4, except the Prometheus Panta (NanoTemper Technologies) was used with a 1 °C/min temperature gradient from 25 °C to 90 °C.

Native MS
Protein samples were buffer exchanged into 200 mM ammonium acetate (pH 8.0) supplemented with 2x CMC of the respective detergent using Micro Bio-Spin 6 columns (BioRad) prior to MS analyses. These samples were directly introduced into the mass spectrometer using gold-coated capillary needles prepared in-house54. Data were collected on a Q Exactive UHMR Hybrid Quadrupole-Orbitrap mass spectrometer (Thermo Fisher) in negative polarity. The instrument parameters used for data collection were: capillary voltage 0.9 kV, S-lens RF 100%, quadrupole selection from 400 to 20,000 m/z range. The HCD collision energy was 300 V for all acquired spectra, with the exception of the mass spectrum shown in Supplementary Fig. 10E where it was 200 V. Source fragmentation was 0 V for all acquired spectra, with the exception of the mass spectrum shown in Supplementary Fig. 9D where it was 150 V. In-source trapping was 250 V for ctPmt4 wild-type mass spectrum in Fig. 4A and 300 V for all other ctPmt4 spectra. The ion transfer optics in negative mode was set as follows: injection flatapole −5 V, inter-flatapole lens −4 V, bent flatapole −2 V, transfer multipole 0 V. The resolution of the instrument was 8,750 at m/z = 200 (transient time of 64 ms) for all acquired spectra, with the exception of the mass spectrum shown in Supplementary Fig. 9D where it was 140,000 at m/z = 200. Argon pressure in the HCD cell was maintained at approximately 5 × 10−10 mbar and source temperature was kept at 250 °C. The noise level was set at 3 rather than the default value of 4.64. Calibration of the instruments was performed using 10 mg/ml solution of caesium iodide in water. Where required, baseline subtraction was performed to achieve a better-quality mass spectrum. Data were analysed using the Xcalibur 3.0 (Thermo Scientific), NaViA55 and UniDec56 software packages.

Lipidomics analysis
Co-purified lipids have been identified in a similar manner as described previously57. Briefly, proteins were digested with trypsin (1:20 units) for 15 h at 37 °C in a thermomixer (Eppendorf). The digest was lyophilized and re-dissolved in 68% solution A (ACN:H2O 60:40, 10 mM ammonium formate and 0.1% formic acid) and 32% solution B (IPA:ACN 90:10, 10 mM ammonium formate and 0.1% formic acid) for analysis by reverse phase liquid chromatography tandem MS (RPLC-MS/MS). LC-MS/MS was performed using a Dionex UltiMate 3000 RSLC Nano system coupled to an LTQ Orbitrap XL hybrid mass spectrometer (Thermo Fisher). The tryptic digest mixture was loaded onto a pre-equilibrated C18 trap (Acclaim PepMap m-Precolumn 300 mm, 5 mm) followed by a C18 column (Acclaim PepMap 100, C18, 75 mm 3 15 cm; Thermo Fisher) and separated by using the following step gradient of solution A and B at 300 nl/min and 40 °C: 0-10 min with 32% solvent B, solvent B was ramped to 65% over 1 min, then the concentration of solvent B was increased to 80% buffer over 6 min and held for 10 min, then ramped to 99% B over 6 min and this concentration was held for 7 min. The nanoflow reversed-phase liquid chromatography was directly coupled to the mass spectrometer via a dynamic nanospray source. Typical MS conditions were spray voltage of 1.6 kV and capillary temperature of 275 °C. The LTQ-Orbitrap XL was set-up in negative ion mode and in data-dependent acquisition mode to perform five MS/MS scans per MS scan. Survey full-scan MS spectra were acquired in the Orbitrap (m/z 350−2000) with a resolution of 60,000 at m/z = 200. Collision-induced dissociation (CID) fragmentation in the linear ion trap was performed for the five most intense ions at an automatic gain control target of 30,000 and a normalized collision energy of 38% at an activation of q = 0.25 and an activation time of 30 ms. Data were analysed using Xcalibur 3.0 (Thermo Fisher).

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The manuscript describes the structural and biochemical characterization of fungal Pmt4. The authors determined the homodimeric enzyme's structure by cryo-EM and its poorly defined in cryo-EM maps MIR domain by X-ray crystallography. The overall structure is similar to a previously solved structure of a heterocomplex Ptm1-Ptm2 from the same family. The new observations include cis rather than in trans contacts between MIR and trans-membrane domain, differences in the structure of MIR domain, specifically the presence of  -hairpin insertion, and observation of plausible cytoplasmic Dol-P-Man binding site. Using native mass spectroscopy, it is shown that Dol-P-Man co-purified with wild-type Pmt4 is depleted in mutants.\n\nThe important questions related to the difference in specificity are not addressed in the study, which makes it somewhat less impactful. Also, I could not find sufficient information in the manuscript about the catalytic site and substrate-binding site. Their detailed comparison between Pmt1-Pmt2 and Ptm4 would be a valuable addition to the manuscript.\n\nSome mutagenesis studies require better controls. The mutation of the plausible Dol-P-Man binding site includes multiple positively charged residue substitutions by alanine residues. Do the mutations preserve the correct insertion of Ptm4 into the membrane? Are the mutants folded correctly?\n\nInspection of the cryo-EM density shows that Dol-P and Dol-P-Man densities can be modeled as such but there are ambiguities. The density modeled as Dol-P is fragmented. Coordination of the phosphate group by positively charged residues should have made its density well defined, yet, the density of strong electron scattering phosphate is weaker than that of the isoprenoid chain. This ambiguity must be explicitly mentioned.\n\nOn the cytoplasmic side, similarly, the density of Dol-P-Man is not well defined and the sugar ring only partially fits the density. The cryo-EM-based interpretation of the density as Dol-P-Man is plausible but not certain. Again, the phosphate group is not well defined. Given that the density is fragmented, it could also be interpreted as a lipid. The reconstruction shows densities for other aliphatic chains that are not bridged to allow interpreting them as lipids.\n\nThe study provides new experimental insights into Ptm4 structure and function but at the same time, leaves an impression of missing important fundamental insights into the subtype-specific substrate selection and catalytic mechanism.\n\nIn the discussion section, a plausible flippase function of Pmt4 is proposed. Do structural features support this suggestion? Is there any indication that Pmt4 may facilitate the negatively charged phosphate group of Dol-P-Man crossing the membrane?\n\nBelow are several additional specific comments:\n(1) Page numbers are missing.\n(2) What is the identity and homology between Pmt family members?\n(3) From line 161. How do you know if the mutated MIR domain remained folded? The anti-FLAG western blots indicate the expression of the protein but not its correct insertion or folding.\nLine 197: It would be helpful if the active site and substrate binding site could be explicitly shown in one of the main figures.\nLine 201. I could not understand what it means: “… the active site is completely triangulated …”\nLine 217: All the aliphatic chain densities resolved in the map appear to be single-channel lipids. How are densities modeled as Dol-P or Dol-P-Man different from the others?\nTable 1: Model resolution should be reported at 0.5 FSC cutoff. Cutoff 0.143 is used for two independently refined half-maps merged into one consensus map.", "reviewer_2": "The authors have combined crystallography, cryo-EM, native MS, and lipidomics to study the structure and composition of the Pmt4 homodimer and compare this to the known structure of the Pmt1-Pmt2 heterodimer. They also used mutagenesis, in particular to study the effect of specific mutations on the capacity of the dimer to bind Dol-P-Man. In my review, I focused on the mass spectrometry, as this most aligns with my own expertise. Overall, the MS work is solid, although I have a few suggestions that could potentially increase the level of confidence in some of the identifications.\n\n*In native MS, particularly of large proteins and complexes, peak broadening due to adduct formation and incomplete desolvation is typical, and this is observed in Figure 4 as expected. However, this makes it difficult to identify small bound species such as lipids just from the mass difference between the bound and unbound state of the protein. Indeed, the authors write on line 243 that the mass differences are ~1100 Da and ~1700 Da, and in Table S1, the observed mass increases due to cardiolipin or Dol-P-Man binding fluctuate between proteins. The authors also state on lines 249-250 that the ~1700 Da mass increase cannot be due to cardiolipin alone, but speculate that this might be due to simultaneous binding of a lipid and a detergent molecule (indeed, they refer to this assignment as “putative” several times). The identification of cardiolipin and Dol-P-Man in the lipidomics experiments lends credibility to the authors’ assignments. However, given the importance of this result, I wonder whether the authors could consider performing a tandem MS experiment to dissociate the bound species from the dimer and measure their accurate mass in the low-mass region of the spectrum, and potentially even perform further characterization using MSn. This approach has been described by one of the groups involved in the current work; see for example Gault et al., Nat Methods, 2020, or Wu et al., Nat Protoc, 2025.\n*On lines 259-260, the authors write that “single arginine or double H219A/H637A mutations were insufficient to prevent Dol-P-Man binding” and refer to Figure S10. With the aforementioned caveat about the lipid identification, this statement seems fine. However, in the figure caption, they write twice that these mutations “do not affect binding” of the lipid. This seems like a much stronger statement to me and implies that there is no effect on affinity. I suggest moderating the figure caption and using a similar phrasing to that in the main text.\n*In the native MS methods, the HCD energy range is given as 150-300 V, and source fragmentation as 0-100 V. These are significant ranges, so could the authors specify which spectra were acquired with which parameters?\n*Also in the native MS methods, on line 676, the authors describe settings for positive mode; however, the native spectra in Figures 4, S9, and S10 all seem to have been acquired in negative mode. Can the authors clarify what work was carried out in positive mode, or is there a typo?\n\nMinor points:\n*Line 232: “we do not observed” --> “we did not observe”\n*Line 261: “R218/R638/R642” --> “R218A/R638A/R642A”\n*There is some artifact in the PDF file (possibly due to file conversion) affecting the phrases “GAPDH-directed antibodies” (starting on line 435) and “prepared in UCSF Chimera” (starting on line 612).\n*“m/z” should be italicized\n*Line 703: Can the authors specify the m/z value at which the resolution was defined?\n*Data availability statement: The authors have uploaded their MS data to the PRIDE repository (and should be commended for this) so the dataset identifier should be added here.", "reviewer_3": "McDowell et al. provide the first structural insight into PMT4 homodimers which act as biosynthetic enzymes for fungal protein O-mannosylation. The authors have resolved the structure of Chaetomium thermophilum Protein O-mannosyltransferase 4 (Pmt4) homodimers by single-particle cryo-electron microscopy (CryoEM) and resolved high-resolution crystal structures of the fungal PMT4-MIR domains that protrude into the ER-lumen. Using native mass spectrometry and lipidomic analysis, the authors provide further insight into PTM4 homodimer assembly and lipid composition to identify co-purified Dol-P-Man donor substrate. Guided by their structures, the authors map key residues in the PMT4 homodimer and PMT4-MIR domains which are further validated by mutations and deletions in functional studies. Remarkably, the authors identify two distinct sites with bound Dol-p and Dol-p-Man in the PMT4 homodimer, which opens for new hypotheses on how GT-C type enzymes may utilize donor substrates in catalytic cycles. These findings are truly novel and exciting as they potentially suggest an intrinsic “flippase” mechanism for positioning Dol-p-Man from the cytosolic side towards the ER-lumen. Further experiments and structural studies are clearly needed to test this hypothesis; however these findings will likely change current views and have boarder impact on the understanding of the mammalian POMT1/POMT2 orthologs, and by extension, also other related enzymes (DPY19L1-4, TMTC1-4 and TMEM260).\n\nOverall, the manuscript is well written, the experiments skillfully executed, the results are detailed and clear. The discussion is balanced and aligned with the results, and the authors provide insightful conclusions. I have minor comments and suggestions for the authors (below) but my enthusiasm for the work is high and I would argue that the manuscript should be considered favorably.\n\nGeneral suggestions:\nLine 27: Consider rephrasing the sentence. Reading the abstract, I misinterpreted the sentence to believe that CryoEM was resolved for both Chaetomium thermophilum and Saccharomyces cerevisiae Pmt4 homodimers.\nLine 41: In addition to POMTs, mammalian O-mannosylation is also initiated by TMTCs and the recently described TMEM260 enzyme. Moreover, transfer of mannose to distinct protein substrates also includes the actions of DPY19Ls (C-Man), and all these enzymes are collectively classified as GT-C enzymes together with PMTs. The readership may benefit if this was clarified in the introduction to provide a broader scope/justification for this study.\nLine 88: The introduction or result section may benefit from a paragraph explaining the evolutionary relationship between ctPmt4 and scPmt4, with a brief statement on why the ctPmt4 dimer was chosen for CryoEM studies.\nLine 109: Are the TMD6/8 residues in the dimer interface in Pmt1-2 and Pmt4 present in other Pmts? Considering that Pmts don’t randomly dimerize, does the Pmt4 dimer interface provide insight into the predominant dimerization events among Pmts? Is this a general feature also conserved in mammalian POMT dimers? These questions can enrich the discussion paragraph.\nLine 142: Regardless of outcome, it would be useful to know if the authors have attempted to capture a potential interaction between scPmt4-MIR and O-mannosylated peptides, either in co-crystallization experiments or other biochemical assays. As is now, the statement on mannose binding/CBMs is rather speculative.\nLine 190-196: Without experimental evidence for the “substrate/product peptide route”, this paragraph is rather speculative. I would suggest that the authors consider rephrasing “highly suggestive” with softer language.\nLine 240: DDM extraction may, or may not, preserve the native lipid content of the assembly. This should be briefly stated for the readership.\nLine 244: Why was the identification of cardiolipin interesting? Was this expected or unexpected?\nLine 287-300: The result paragraph for the functional assay with the FUS-Axl2TMZZ substrate is not perfectly clear to me. What was the rational? How many technical/biological replicates were done for Figure 5A? Why is there no loading control? These questions are rooted in my difficulties in interpreting the results in Figure 5A. If the assay based on assessing the absence/accumulation of the indicated band at ~68 kDa, why is there a difference in intensity between EV and WT and why is there no major accumulation in RRR and H219A/H637A (compared to EV)? The authors should consider clarifying.\nLine 314-315: Does this argument also apply to other GT-C enzymes, e.g. DPY19Ls or TMTCs?\nLine 320: The discussion should perhaps briefly state that one current limitation is lack of insight into the Pmt4 dimer-substrate peptide complex, which precludes further insight into the catalytic cycle of the reaction.\nLine 1045: Figure S1: Size exclusion elusion profiles of ctPmt4 appear to migrate as monomers? It may be beneficial to indicate the migration pattern of molecular standards on the Superdex 200 Increase SEC. It may be further beneficial to indicate which elution fractions were used for CryoEM. It is unclear why ctPmt4 migrates at ~100 kDa (LMNG-CHS) and ~70 kDa (DDM) in the following SDS-PAGE analysis?"}, "author_response": "Response to reviewers\nFirst of all, we would like to thank all reviewers for their thoughtful comments and\nconstructive criticism, which have helped improve the quality of our manuscript.\nReviewer #1 (Remarks to the Author):\nThe manuscript describes the structural and biochemical characterization of fungal Pmt4. The\nauthors determined the homodimeric enzyme's structure by cryo-EM and its poorly defined in\ncryo-EM maps MIR domain by X-ray crystallography. The overall structure is similar to a previously\nsolved structure of a heterocomplex Ptm1-Ptm2 from the same family. The new observations\ninclude cis rather than in trans contacts between MIR and trans-membrane domain, differences in\nthe structure of MIR domain, specifically the presence of b-hairpin insertion, and observation of\nplausible cytoplasmic Dol-P-Man binding site. Using native mass spectroscopy, it is shown that\nDol-P-Man co-purified with wild-type Pmt4 is depleted in mutants.\nThe important questions related to the difference in specificity are not addressed in the study,\nwhich makes it somewhat less impactful. Also, I could not find sufficient information in the\nmanuscript about the catalytic site and substrate-binding site. Their detailed comparison between\nPmt1-Pmt2 and Ptm4 would be a valuable addition to the manuscript.\nAlthough Pmt1-Pmt2 and Pmt4 act on distinct proteins, the determinants of their\nsubstrate specificity are very poorly defined in the literature. Indeed, there are no strict\nsequence motifs for substrate recognition, rather a vague propensity for Ser/Thr rich\nregions in hydrophilic, intrinsically disordered domains (Neubert et al., 2016). In terms\nof specificity, Pmt1-Pmt2 acts on soluble and membrane proteins, whilst Pmt4 seems to\npreferentially mannosylate membrane proteins (Hutzler et al., 2007). Some substrates\n(e.g. CCW5, Ecker et al., 2003) are mannosylated by both Pmt1-Pmt2 and Pmt4.\nAltogether, the substrate specificity is broad, overlapping and ill-defined. Therefore, it is\nto be expected that the structures do not show clear-cut differences that can be easily\ncorrelated with specificity, rather actually there is likely to be quite some overlap in\ntheir features.\nIn the manuscript, we compare the active site of Pmt4 with those of Pmt1 and Pmt2\n(Figures 3B and S7C) and find no difference in the residue arrangement, aside from the\n60° rotation in R666 that we describe in the text. We therefore conclude that the\ncatalytic mechanism is conserved across the PMT families and, as we cannot extend\nmechanistic insights beyond Bai et al., 2019, do not discuss this in much more detail.\nTo rationalize differences in substrate specificity in terms of structure, we rely on\ndifferences between the structures of Pmt1-Pmt2 and the Pmt4 homodimer, with the\ncaveat that our current structure does not contain bound substrate peptide. Within the\nTMDs, these structures have many more similarities than they have differences, whilst in\nthe Pmt1-Pmt2 structure, the only specific contacts between Pmt2 and the bound PYT\npeptide are with the acceptor threonine residue (Bai et al., 2019). Nevertheless, we now\nadd a comparison of the equivalent substrate binding region in Pmt4, Pmt1 and Pmt2\n(Figure 3C and S7E). We find some differences in the peptide-proximal residues and δ’\nsite between Pmt4 and Pmt2, however these features are identical in Pmt1 indicating\nthey do not hugely contribute to differences in substrate specificity between these\nfamilies. Another notable difference is in the positioning of the attached MIR domains\nwith respect to the TMDs, where the Pmt4 MIR domain sits much closer to the active site\nand could thereby also contribute to substrate selection. We have now added these\nobservations to the results section.\nGiven that there is some evidence that PMTs are found in a complex with the Sec61\ntranslocon and OST (Loibl et al., 2014), it could also be that Pmt1/Pmt2 and Pmt4\nachieve substrate specificity by interacting with different translocon-associated\ncomplexes e.g. in response to the translocation of particular substrates. This more\nspeculative point has been added to the discussion.\nSome mutagenesis studies require better controls. The mutation of the plausible Dol-P-Man\nbinding site includes multiple positively charged residue substitutions by alanine residues. Do the\nmutations preserve the correct insertion of Ptm4 into the membrane? Are the mutants folded\ncorrectly?\nAll ctPmt4 mutants used for both native MS and in vivo assays were purified from\nmembranes and subjected to nanoDSF, showing no change in their unfolding\ntemperature relative to the wild type protein (Figure S9A). This strongly indicates that\nthe wild type and mutant ctPmt4 variants have the same fold. This is further supported\nby native MS, which shows that dimer formation is not disrupted and that the putative\ncardiolipin adduct is still able to bind (Figure 4). These mutants were also present at\nequivalent levels to the wild type protein in the membrane (Figure S11C). We assert that\ntogether this already provides sufficient evidence that the mutants are correctly\ninserted in the membrane and properly folded.\nUnfortunately, it is not possible to purify the equivalent mutants of scPmt4 in sufficient\namount and purity for the same analysis. However, these are also present at the same\nlevel as the wild type protein in the membrane (Figure S11G), indicating correct\ninsertion and, as incorrect folding of the TMDs would likely lead to impaired membrane\nintegration or membrane extraction, likely correct folding.\nInspection of the cryo-EM density shows that Dol-P and Dol-P-Man densities can be modeled as\nsuch but there are ambiguities. The density modeled as Dol-P is fragmented. Coordination of the\nphosphate group by positively charged residues should have made its density well defined, yet,\nthe density of strong electron scattering phosphate is weaker than that of the isoprenoid chain.\nThis ambiguity must be explicitly mentioned.\nOn the cytoplasmic side, similarly, the density of Dol-P-Man is not well defined and the sugar ring\nonly partially fits the density. The cryo-EM-based interpretation of the density as Dol-P-Man is\nplausible but not certain. Again, the phosphate group is not well defined. Given that the density is\nfragmented, it could also be interpreted as a lipid. The reconstruction shows densities for other\naliphatic chains that are not bridged to allow interpreting them as lipids.\nWe agree with the reviewer that the densities for Dol-P and Dol-P-Man are not as well\nresolved as for the surrounding TMDs. In both cases, this is likely due to two reasons:\n1. The headgroups exhibit flexibility with respect to the isoprenoid chain, explaining\nwhy the densities for the phosphate groups are weaker than expected.\n2. These lipids only have partial occupancy in the structure. From the relative\nabundance of the species resolved in the native mass spectrum in Figure 4A, we\ncan estimate that ~40% Pmt4 dimers have at least one molecule of Dol-P-Man\nbound. Dol-P species are not unambiguously resolved by native MS, so their\noccupancy is likely to be even lower, in-line with the weaker, more fragmented\ndensity relative to Dol-P-Man.\nWe have added some clarification of these points in the text.\nThe modelling of the active site density as Dol-P is in full agreement with the binding\nand coordination of this species previously observed in the Pmt1-Pmt2 structure (Bai et\nal., 2019). Indeed, inspection of the Dol-P density in the Pmt1-Pmt2 map confirms that\nthe density for the phosphate group here is also weaker than for the isoprenoid chain.\nThe phosphate group occupies a slightly different position compared to the Pmt1-Pmt2\nmodel, which is supported by the clear density of the rotated R666 side chain. The\nrepositioning of Dol-P adds further weight to the argument that the headgroup has some\nflexibility within the active site; indeed, as the product of the reaction one would expect\nthat the lipid should not be stably bound.\nThe modelling of Dol-P-Man in the cytosolic site is of course novel and required more\ncareful justification. Our rationale was as follows: 1) the density unambiguously\ncorresponds to a single chain and is not fragmented. 2) the coordination of the\nheadgroup by His and Arg residues closely mimics the binding of Dol-P-(Man) in the\nactive site. The sugar likely has some flexibility given that the capping cytoplasmic loop\n(residues 695-728) is also not resolved. 3) we clearly see Dol-P-Man in the native MS\nand do not see density to assign Dol-P-Man in the active site. To further verify the\ncytoplasmic lipid is Dol-P-Man, we made mutants in the binding site and confirmed they\nare still membrane integrated and folded (see response to previous point). Dol-P-Man\nbinding was abolished in several of these mutants (but notably not binding of the\ncardiolipin adduct), which in our view confirms our modelling of Dol-P-Man.\nThe study provides new experimental insights into Ptm4 structure and function but at the same\ntime, leaves an impression of missing important fundamental insights into the subtype-specific\nsubstrate selection and catalytic mechanism.\nSubstrate specificity has been discussed more extensively in response to the first point,\nbut we have added Figure 3C, Figure S7E and corresponding text to address this. With\nregards to the catalytic mechanism, the Pmt4 and Pmt1-Pmt2 catalytic sites are virtually\nidentical, strongly indicating a conserved “inversion reaction mechanism for mannosyl\ntransfer”. As we do not resolve a bound metal ion, substrate peptide or Dol-P-Man in the\nactive site, we cannot extend the description of the catalytic mechanism beyond what is\nalready presented in Bai et al., 2019. Therefore, we focused our manuscript more\nextensively on the novel features of Pmt4 with respect to Pmt1-Pmt2.\nIn the discussion section, a plausible flippase function of Pmt4 is proposed. Do structural features\nsupport this suggestion? Is there any indication that Pmt4 may facilitate the negatively charged\nphosphate group of Dol-P-Man crossing the membrane?\nH657 points into the TMD groove connecting the cytosolic and luminal Dol-P-Man\nbinding sites and is part of a highly conserved HYΦP motif in TMD9. Interestingly, this\nmotif is also found in the active site, where H672 interacts with the Dol-P(-Man)\nheadgroup. We therefore think this motif forms a plausible intermediate binding site for\nthe lipid headgroup as it is flipped. We have now added this point to the discussion and\nindicate the HYΦP motif below the sequence alignment in Figure S1.\nBelow are several additional specific comments:\n(1) Page numbers are missing.\nWe apologise for this omission – we believe there was an error during the PDF\nconversion.\n(2) What is the identity and homology between Pmt family members?\nThis is variable, but identity within a PMT family is typically >30% and between PMT\nfamilies 25-30%. ctPmt4 and scPmt4 have a sequence identity of 48%. To illustrate this,\na percent identity matrix including all S. cerevisiae homologues is now shown below the\nPMT sequence alignment (now Figure S1).\n(3) From line 161. How do you know if the mutated MIR domain remained folded? The anti-FLAG\nwestern blots indicate the expression of the protein but not its correct insertion or folding.\nThe western blot in Figure S5B has now been obtained with samples derived from total\nmembranes rather than whole cell lysate, demonstrating that all the MIR domain\nmutants are present in the membrane at the same level as the wild type protein. In\naddition, we created the most severe b-ins mutation, the b-ins deletion (Δ362-368), in\nour scPmt4 MIR domain construct. We found that this variant could be purified to the\nsame yield and homogeneity as the wild type construct, and that the domain was folded\nwhen assessed by nanoDSF, albeit with a slightly reduced stability. These points have\nbeen added to the text and the comparison of nanoDSF traces for the wild type and\nΔ362-368 variants are shown as Figure S5A.\nLine 197: It would be helpful if the active site and substrate binding site could be explicitly shown\nin one of the main figures.\nThe panel in Figure 3B shows the active site of ctPmt4 along with the position of dol-P\nand the acceptor threonine derived from the scPmt2 structure. It is superimposed with\nscPmt1 to demonstrate the highly conserved arrangement of residues in the active site.\nThe equivalent superimposition with scPmt2 is shown in Figure S7C.\nWe have now added panels as Figure 3C and Figure S7E plus a corresponding description\nin the text, which show the substrate peptide binding sites of scPmt2 or scPmt1\nrespectively superimposed with ctPmt4.\nLine 201. I could not understand what it means: “… the active site is completely triangulated …”\nWe changed the wording to “the active site is completely covered by this δ site” to\nimprove clarity.\nLine 217: All the aliphatic chain densities resolved in the map appear to be single-channel lipids.\nHow are densities modeled as Dol-P or Dol-P-Man different from the others?\nWe agree there are other densities, particularly in the C1 map, that could correspond to\nlipids, but they are generally more fragmented compared to the assigned Dol-P-Man\ndensity. We further cannot rule out that these additional densities also correspond to\nDol-P species, but we have no evidence to assign them as such. Our rationale for\nmodelling Dol-P and Dol-P-Man in their respective densities was primarily dependent on\nthe chemistry of their binding sites and experimental validation, as outlined in more\ndetail above in response to the previous point regarding these lipids.\nTable 1: Model resolution should be reported at 0.5 FSC cutoff. Cutoff 0.143 is used for two\nindependently refined half-maps merged into one consensus map.\nModel resolution at 0.5 FSC cutoff are now given instead in Table 1.\nReviewer #2 (Remarks to the Author):\nThe authors have combined crystallography, cryo-EM, native MS, and lipidomics to study the\nstructure and composition of the Pmt4 homodimer and compare this to the known structure of the\nPmt1-Pmt2 heterodimer. They also used mutagenesis, in particular to study the effect of specific\nmutations on the capacity of the dimer to bind Dol-P-Man. In my review, I focused on the mass\nspectrometry, as this most aligns with my own expertise. Overall, the MS work is solid, although I\nhave a few suggestions that could potentially increase the level of confidence in some of the\nidentifications.\n*In native MS, particularly of large proteins and complexes, peak broadening due to adduct\nformation and incomplete desolvation is typical, and this is observed in Figure 4 as expected.\nHowever, this makes it difficult to identify small bound species such as lipids just from the mass\ndifference between the bound and unbound state of the protein. Indeed, the authors write on line\n243 that the mass differences are ~1100 Da and ~1700 Da, and in Table S1, the observed mass\nincreases due to cardiolipin or Dol-P-Man binding fluctuate between proteins. The authors also\nstate on lines 249-250 that the ~1700 Da mass increase cannot be due to cardiolipin alone, but\nspeculate that this might be due to simultaneous binding of a lipid and a detergent molecule\n(indeed, they refer to this assignment as “putative” several times). The identification of cardiolipin\nand Dol-P-Man in the lipidomics experiments lends credibility to the authors’ assignments.\nHowever, given the importance of this result, I wonder whether the authors could consider\nperforming a tandem MS experiment to dissociate the bound species from the dimer and measure\ntheir accurate mass in the low-mass region of the spectrum, and potentially even perform further\ncharacterization using MSn. This approach has been described by one of the groups involved in the\ncurrent work; see for example Gault et al., Nat Methods, 2020, or Wu et al., Nat Protoc, 2025.\nWe thank the reviewer for this insightful suggestion. Indeed, we attempted to perform\ntandem MS experiments using both the Q Exactive UHMR (the instrument used for this\nstudy) and the Orbitrap Eclipse to dissociate and analyse the bound species in the lower\nm/z region. However, under our experimental conditions, the observation of the protein\ncomplex necessitates the application of collision voltages in the HCD cell downstream of\nthe quadrupole for the Q Exactive UHMR. This step is essential because application of in-\nsource trapping voltages could not effectively liberate the protein from the surrounding\nmicelles. As a result, we are unable to isolate a micelle-free protein complex within the\nquadrupole for subsequent selection and activation steps.\nThis limitation precludes the isolation of a specific charge state of the protein-lipid\ncomplex prior to activation, which is a prerequisite for the type of tandem MS (and\npotential MSⁿ) analysis suggested by the reviewer. We fully agree that such an approach\nwould offer valuable structural insights, and we are actively exploring alternative\nexperimental strategies that might enable this type of analysis in future work. As noted,\nwe also attempted native MS experiments using the Orbitrap Eclipse, but were unable to\nobtain sufficiently resolved spectra. The challenges encountered may stem from the\nheterogeneity of the protein system, which involves a monomer/dimer equilibrium and\nthe presence of multiple bound lipid species, all contributing to a high degree of spectral\ncomplexity.\nTo further probe the identity of the bound species, we conducted an additional\nexperiment by increasing the activation energies to the maximum levels permitted by\nthe instrument (300 V HCD, 150 V source fragmentation, and 300 V in-source trapping)\nand increasing the resolution to 140,000 at m/z = 200. In parallel, we also optimised\nthe mass spectrometry parameters to enhance transmission of low-mass ions,\nfacilitating the detection of smaller dissociated species. Under these optimised\nconditions, we were able to dissociate the bound Dol-P-Man and observe its accurate\nmass in the low-mass region. The resulting peaks showed excellent agreement with the\nDol-P-Man species identified in our lipidomics analyses, lending further support to our\nassignment. This experiment is now included as Figure S8D. Unfortunately, the 1700 Da\nspecies could not be resolved any further using the same approach.\n*On lines 259-260, the authors write that “single arginine or double H219A/H637A mutations were\ninsufficient to prevent Dol-P-Man binding” and refer to Figure S10. With the aforementioned caveat\nabout the lipid identification, this statement seems fine. However, in the figure caption, they write\ntwice that these mutations “do not affect binding” of the lipid. This seems like a much stronger\nstatement to me and implies that there is no effect on affinity. I suggest moderating the figure\ncaption and using a similar phrasing to that in the main text.\nWe thank the reviewer for pointing this out. We have now revised the figure caption in\nline with the suggestion, replacing the wording “do not affect binding” to “do not\nsubstantially impair binding” to better reflect the interpretation presented in the main\ntext.\n*In the native MS methods, the HCD energy range is given as 150-300 V, and source\nfragmentation as 0-100 V. These are significant ranges, so could the authors specify which spectra\nwere acquired with which parameters?\n*Also in the native MS methods, on line 676, the authors describe settings for positive mode;\nhowever, the native spectra in Figures 4, S9, and S10 all seem to have been acquired in negative\nmode. Can the authors clarify what work was carried out in positive mode, or is there a typo?\nWe thank the reviewer for highlighting these unclear points. We have now clarified in\nthe Methods section the specific voltages used for each spectrum shown. Regarding the\npolarity, the reviewer is correct, this was a typo. All native MS spectra, including those\nin Figures 4, S9 and S10, were acquired in negative ion mode, and we have corrected the\ntext accordingly.\nMinor points:\n*Line 232: “we do not observed” --> “we did not observe”\nCorrected.\n*Line 261: “R218/R638/R642” --> “R218A/R638A/R642A”\nCorrected.\n*There is some artifact in the PDF file (possibly due to file conversion) affecting the phrases\n“GAPDH-directed antibodies” (starting on line 435) and “prepared in UCSF Chimera” (starting on\nline 612).\nWe thank the reviewer for pointing out that. It seems to be an artifact from the PDF\nconversion during the submission process.\n*“m/z” should be italicized\nThis has been corrected in the text and Figure 4.\n*Line 703: Can the authors specify the m/z value at which the resolution was defined?\nThe resolution was defined at m/z = 200. This has now been included in the text.\n*Data availability statement: The authors have uploaded their MS data to the PRIDE repository\n(and should be commended for this) so the dataset identifier should be added here.\nWe have now added the relevant statement along with the dataset identifier to the data\navailability statement.\nReviewer #3 (Remarks to the Author):\nMcDowell et al. provide the first structural insight into PMT4 homodimers which act as biosynthetic\nenzymes for fungal protein O-mannosylation. The authors have resolved the structure of\nChaetomium thermophilum Protein O-mannosyltransferase 4 (Pmt4) homodimers by single-\nparticle cryo-electron microscopy (CryoEM) and resolved high-resolution crystal structures of the\nfungal PMT4-MIR domains that protrude into the ER-lumen. Using native mass spectrometry and\nlipidomic analysis, the authors provide further insight into PTM4 homodimer assembly and lipid\ncomposition to identify co-purified Dol-P-Man donor substrate. Guided by their structures, the\nauthors map key residues in the PMT4 homodimer and PMT4-MIR domains which are further\nvalidated by mutations and deletions in functional studies. Remarkably, the authors identify two\ndistinct sites with bound Dol-p and Dol-p-Man in the PMT4 homodimer, which opens for new\nhypotheses on how GT-C type enzymes may utilize donor substrates in catalytic cycles. These\nfindings are truly novel and exciting as they potentially suggest an intrinsic “flippase” mechanism\nfor positioning Dol-p-Man from the cytosolic side towards the ER-lumen. Further experiments and\nstructural studies are clearly needed to test this hypothesis; however these findings will likely\nchange current views and have boarder impact on the understanding of the mammalian\nPOMT1/POMT2 orthologs, and by extension, also other related enzymes (DPY19L1-4, TMTC1-4 and\nTMEM260).\nOverall, the manuscript is well written, the experiments skillfully executed, the results are detailed\nand clear. The discussion is balanced and aligned with the results, and the authors provide\ninsightful conclusions. I have minor comments and suggestions for the authors (below) but my\nenthusiasm for the work is high and I would argue that the manuscript should be considered\nfavorably.\nWe thank the reviewer for these positive comments and have addressed their specific\nsuggestions below.\nGeneral suggestions:\nLine 27: Consider rephrasing the sentence. Reading the abstract, I misinterpreted the sentence to\nbelieve that CryoEM was resolved for both Chaetomium thermophilum and Saccharomyces\ncerevisiae Pmt4 homodimers.\nWe have rephrased the abstract.\nLine 41: In addition to POMTs, mammalian O-mannosylation is also initiated by TMTCs and the\nrecently described TMEM260 enzyme. Moreover, transfer of mannose to distinct protein substrates\nalso includes the actions of DPY19Ls (C-Man), and all these enzymes are collectively classified as\nGT-C enzymes together with PMTs. The readership may benefit if this was clarified in the\nintroduction to provide a broader scope/justification for this study.\nWe thank the reviewer for this suggestion. The introduction has now been updated to\nput the POMTs in the context of these other enzymes involved in protein mannosylation.\nLine 88: The introduction or result section may benefit from a paragraph explaining the\nevolutionary relationship between ctPmt4 and scPmt4, with a brief statement on why the ctPmt4\ndimer was chosen for CryoEM studies.\nIn our hands, scPmt4 has proved largely recalcitrant to purification and structural\nstudies, whilst generally the recombinant production of C. thermophilum thermostable\nproteins has consistently yielded high resolution structures. We now start the results\nsection with “to investigate the structure of Pmt4, we took advantage of its\nthermostable homologue from the thermophilic fungus Chaetomium thermophilum,\nwhich has 48 % sequence identity to yeast Pmt4” and include a percent identity matrix\nwith the PMT sequence alignment (now Figure S1).\nLine 109: Are the TMD6/8 residues in the dimer interface in Pmt1-2 and Pmt4 present in other\nPmts? Considering that Pmts don’t randomly dimerize, does the Pmt4 dimer interface provide\ninsight into the predominant dimerization events among Pmts? Is this a general feature also\nconserved in mammalian POMT dimers? These questions can enrich the discussion paragraph.\nThis is an intriguing question, as the residues at the TMD6/8 dimer interface are\nrelatively few and largely conserved across PMTs and POMTs (Figure S1), therefore\ndon’t provide a satisfactory explanation for the formation of distinct dimer species.\nSimilarly, the TMD9 residues at the luminal constriction point in the Pmt4 dimer are also\nconserved. In the Pmt1-Pmt2 structure, the Pmt1 N-terminus, the Pmt1 MIR domain and\na lipid additionally contribute to the dimer interface and could promote preferential\nheterodimer formation by these species. We have added this point to the text. This is\npresumably less important for POMTs where the formation of only one dimer species is\npossible.\nLine 142: Regardless of outcome, it would be useful to know if the authors have attempted to\ncapture a potential interaction between scPmt4-MIR and O-mannosylated peptides, either in co-\ncrystallization experiments or other biochemical assays. As is now, the statement on mannose\nbinding/CBMs is rather speculative.\nThis is something we are actively working on both for the MIR domains and full-length\nprotein, but due to the time-frame and complexity of the experiments will likely yield\ndata for a future manuscript.\nLine 190-196: Without experimental evidence for the “substrate/product peptide route”, this\nparagraph is rather speculative. I would suggest that the authors consider rephrasing “highly\nsuggestive” with softer language.\nWe have removed “highly” from this statement.\nLine 240: DDM extraction may, or may not, preserve the native lipid content of the assembly. This\nshould be briefly stated for the readership.\nThis is now stated in the context of the observed cardiolipin binding (see below).\nLine 244: Why was the identification of cardiolipin interesting? Was this expected or unexpected?\nAs a mitochondrial lipid, we do not expect Pmt4 to be associated with cardiolipin in vivo\nand think it is more likely to be present in our sample as an artifact of protein\npurification from total cellular membranes. Indeed, this has been observed previously\nfor purified ER membrane proteins via the same lipidomics/native MS approach\n(McDowell et al., 2020). We have now clarified this point in the text.\nLine 287-300: The result paragraph for the functional assay with the FUS-Axl2TMZZ substrate is\nnot perfectly clear to me. What was the rational? How many technical/biological replicates were\ndone for Figure 5A? Why is there no loading control? These questions are rooted in my difficulties\nin interpreting the results in Figure 5A. If the assay based on assessing the absence/accumulation\nof the indicated band at ~68 kDa, why is there a difference in intensity between EV and WT and\nwhy is there no major accumulation in RRR and H219A/H637A (compared to EV)? The authors\nshould consider clarifying.\nAs we found ctPmt4 expressed in S. cerevisiae has an altered substrate specificity\nrelative to scPmt4, we tried several different substrate mannosylation assays and\ndetermined FUS-Axl2TMZZ to have the clearest readout. We performed three biological\nreplicates and agree the chosen representative western blot was rather ambiguous; we\nhave now replaced the western blot in Figure 5A with a replicate that shows these\neffects more clearly. In the absence of O-mannosylation (i.e. with the EV control), cells\naccumulate hypo-mannosylated FUS-Axl2TMZZ that runs on SDS-PAGE at ~70 kDa. We\nsee that the wild type and H219A/H637A (which native MS showed is not impaired for\nDol-P-Man binding) do not accumulate this species, whilst the RRR and G648L variants\n(which are impaired for Dol-P-Man binding) do accumulate hypo-mannosylated FUS-\nAxl2TMZZ, as per the EV control. The western blot of total membranes was repeated and\nanalysed with respect to the amounts of ctPmt4-HA and Sec61 as loading control (Figure\nS12C).\nLine 314-315: Does this argument also apply to other GT-C enzymes, e.g. DPY19Ls or TMTCs?\nTo answer this question, we superimposed our structure of ctPmt4 with structural\nmodels for other GT-C enzymes that use Dol-P-Man as a substrate. Strikingly, we found\nthat human TMTCs and TMEM208 appear to also have a positively charged cavity in the\nposition where we observe cytosolic Dol-P-Man binding, suggesting it is a general\nfeature of O-mannosyltransferases. We have now added this observation to the text and\nFigure S12. According to these comparisons, DPY19, PIG and ALG enzymes do not\nappear to present a positively charged cavity in this exact position; indeed these (largely\npredicted) models generally differ in the length/arrangement at the cytosolic ends of\nthe contributing TMDs.\nLine 320: The discussion should perhaps briefly state that one current limitation is lack of insight\ninto the Pmt4 dimer-substrate peptide complex, which precludes further insight into the catalytic\ncycle of the reaction.\nWe added this sentence to the discussion: “However, more detailed insights into the\ncatalytic mechanism, substrate specificity and regulation of PMTs requires\ncharacterisation of the PMT4 homodimer bound to specific substrates, and more\ngenerally, the conformational landscape and putative higher order assemblies of PMTs.”\nLine 1045: Figure S1: Size exclusion elusion profiles of ctPmt4 appear to migrate as monomers? It\nmay be beneficial to indicate the migration pattern of molecular standards on the Superdex 200\nIncrease SEC. It may be further beneficial to indicate which elution fractions were used for\nCryoEM. It is unclear why ctPmt4 migrates at ~100 kDa (LMNG-CHS) and ~70 kDa (DDM) in the\nfollowing SDS-PAGE analysis?\nThe manual for the Superdex 200 increase 10/30 GL column (Cytiva) shows that a 158\nkDa standard elutes at 12.5 ml, similar to the volume for ctPmt4 (92 kDa). Nevertheless,\nwe would argue that the soluble standard proteins used for calibration do not provide\ngood markers for a membrane protein, whose hydrodynamic radius and elution volume\nare altered by the associated detergent micelle. In addition, proteins eluted between 10\nand 15 ml are expected to cover the range ~44-440 kDa, indicating that species of quite\ndifferent molecular weights migrate closely together. We think that the cryo-EM and\nnative-MS together provide much better evidence for the oligomeric state of purified\nctPmt4. At the reviewer’s suggestion, we have now indicated the fraction(s) used for\nthese experiments on the SEC profiles in Figure S1A.\nWe thank the reviewer for noticing the discrepancy in the gels in Figure S1A. Upon\ncloser inspection, we realised the uncropped gel for ctPmt4 in LMNG-CHS was not\nperfectly straight in the image, which led to an offset of the markers. This has now been\ncorrected showing that ctPmt4 runs consistently in both detergents just above the 70\nkDa marker. The uncropped gels are provided in the Source Data file."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors addressed all my comments adequately and the manuscript can be published.", "reviewer_3": "The authors have addressed my comments."}, "author_response": ""}]

Decision letter