Interplay of SLC33A1-dependent and -independent Golgi sialic acid O-acetylation in CASD1 catalysis

Domain: biology
Published: 2026-04-01
Editorial outcome: major_revision

Abstract
Sialic acid O-acetylation is implicated in the modulation of sialoglycan recognition and ganglioside biology. The sugar modification is catalyzed by CASD1, a Golgi membrane protein that encompasses a luminal catalytic domain and a multipass transmembrane domain. The mechanism of how acetyl-CoA is provided to the Golgi remains poorly understood. Here, we show that the acetyl-CoA transporter SLC33A1 provides acetyl-CoA to the luminal domain of CASD1 and that patient-derived SLC33A1 variants linked to inherited neurodevelopmental and neurodegenerative disorders impair ganglioside 9-O-acetylation. Under conditions that enable the formation of 7,9-di-O-acetylated sialoglycans, genetic inactivation of SLC33A1 impaired di-O-acetylation, but unexpectedly, still enabled mono-O-acetylation. Structure prediction and site-directed mutagenesis revealed a second active site in CASD1 that shares striking similarities with the catalytic acetyl-CoA binding transmembrane tunnel of the lysosomal acetyltransferase HGSNAT. Together, our data provide strong evidence that CASD1 has dual functionalities and catalyzes 7,9-di-O-acetylation through SLC33A1-dependent luminal acetylation and SLC33A1-independent transmembrane acetylation.

Introduction
Sialic acids (Sia), a diverse family of nine-carbon α-keto acid sugars, are versatile glycan building blocks in vertebrates. Members of this sugar class are prominently found at the distal ends of N- and O-glycans of glycoproteins and are an integral component of the glycan moiety of glycosphingolipids, the gangliosides1,2. Sialoglycans are involved in numerous molecular interactions, fulfill critical functions in development and immunity, and are frequent targets for pathogens to promote host cell entry1,3,4. The major biosynthetic route of Sia starts from N-acetylmannosamine and provides 5-N-acetylneuraminic acid (Neu5Ac), the most frequent Sia type. Size, shape, and functionality of Neu5Ac is further modulated by a variety of post-synthetic modifications5. In humans, the most prevalent modification is O-acetylation at the C7 and/or C9 hydroxyl groups of Sia, which typically yields mono-O-acetylated Neu5,9Ac (9-O-Ac Sia) and di-O-acetylated Neu5,7,9Ac (7,9-O-Ac Sia)6. Not only can 9-O-acetylation mask recognition sites7–10, it can also create novel sites, which are exploited by viruses that possess specialized Sia-binding proteins for the specific attachment to 9-O-acetylated sialoglycans, such as influenza C virus and the human coronaviruses OC43 and HKU111–13. Furthermore, 9-O-acetylation also impacts ganglioside biology. For example, the addition of a single acetyl group to the C9 hydroxyl group of the terminal α2,8-linked Sia residue of the ganglioside GD3 counteracts the pro-apoptotic activity of the parental molecule and promotes the survival of various cancer cells14–16. Also, in the nervous system, 9-O-acetylated gangliosides are implicated in developmental, regenerative, and neuroprotective processes17–19.

The key enzyme in the biosynthesis of 9-O-acetylated sialosides is the sialate O-acetyltransferase (SOAT) CAS1 domain containing 1 (CASD1), a Golgi-resident multipass transmembrane (TM) protein composed of a luminal-oriented catalytic domain (LCD) and a C-terminal transmembrane region (CTMR) of unknown function20,21. In vitro studies with the isolated LCD demonstrated transfer of acetyl groups from acetyl-coenzyme A (acetyl-CoA) to cytidine 5′-monophosphate (CMP)-activated Sia (CMP-Sia)20, the donor substrate of the Golgi-resident sialyltransferases. This indicates that O-acetylation occurs prior to sialylation and that the LCD depends on luminal acetyl-CoA. Since acetyl-CoA is membrane impermeable and not known to be produced within the Golgi, a transport system is required that provides acetyl-CoA to the Golgi lumen22.

In 1997, a putative acetyl-CoA transporter cDNA was identified by its ability to increase ganglioside 9-O-acetylation upon transfection-mediated expression23. The encoded protein, solute carrier family 33 member A1 (SLC33A1; also referred to as AT-1 or ACATN), shares sequence similarity with transporters of the major facilitator superfamily (MFS)24. In its solubilized state, the transporter exists as a dimer and exhibits functional characteristics consistent with an acetyl-CoA/CoA antiporter25,26. SLC33A1 was initially found in the ER23,25 and it remained unclear whether it contributes to Golgi acetylation only upon overexpression. Subsequent studies led to the discovery of ER-based Nε-lysine acetylation, implicated in protein quality control and regulation of autophagy27–29, and a potential Golgi function of SLC33A1 was not further pursued. Gene silencing indicated that SLC33A1 is important for cell viability25, which may explain why knockout (KO) cells to dissect SLC33A1-dependent processes are still largely lacking. However, a comprehensive understanding of the SLC33A1 acetylation network is of medical relevance, given that genetic dysfunction of SLC33A1 in humans is associated with neurodevelopmental and neurodegenerative disorders. The most devastating type is the autosomal recessive Huppke-Brendel syndrome (HBS), a childhood-onset disorder characterized by profound psychomotor retardation, congenital cataracts, hearing loss, cerebellar atrophy, hypomyelination, and premature death30,31. SLC33A1 mutations acquired through dominant inheritance have been identified in patients affected by spastic paraplegia type 42 (SPG42)32 and late-onset cerebellar ataxia (ATX)33. The effect of disease-causing SLC33A1 mutations on Sia O-acetylation is not known.

To date, the precise mechanism by which acetyl groups are delivered to the Golgi lumen and the specific contribution of SLC33A1 for Golgi Sia O-acetylation remain elusive. By generating a panel of KO cells, we show here that SLC33A1 is essential for the formation of 9-O-acetylated GD3 (9-O-Ac-GD3) and that HBS-, SPG42- and ATX-derived SLC33A1 mutations impair ganglioside 9-O-acetylation. Moreover, we demonstrate that the O-glycan-specific sialyltransferase ST8SIA6 drives the formation of 7,9-O-acetylated sialoglycans in a process that depends on CASD1 but relies only partially on SLC33A1. By combining structure prediction, site-directed mutagenesis and molecular dynamics simulations, we show that the CTMR of CASD1 functions as a TMSOAT with direct access to cytosolic acetyl-CoA. Taken together, we establish a two-catalytic sites model for di-O-acetylation involving SLC33A1-dependent luminal Sia O-acetylation by the LCD of CASD1 and SLC33A1-independent TM Sia O-acetylation by the CTMR of CASD1.

Results
SLC33A1 is required for the formation of 9-O-acetylated GD3 in HAP1 cells
To define the role of the acetyl-CoA transporter SLC33A1 in Sia 9-O-acetylation, we generated SLC33A1-deficient cells (ΔSLC33A1) by CRISPR/Cas9. Initially, we targeted SLC33A1 in the near-haploid human cell line HAP1 by introducing a microdeletion in exon 1, leading to a premature stop codon and a massive truncation of the translation product (Supplementary Fig. 1). As a functional readout for Sia 9-O-acetylation, we assessed the formation of 9-O-Ac-GD3 by immunofluorescence staining with two antibodies that allow discrimination between the 9-O- and the non-O-acetylated form of GD3 (Fig. 1a, b). The synthesis of GD3 from the precursor ganglioside GM3 is catalyzed by the sialyltransferase ST8SIA1, which extends the glycan moiety of GM3 by an α2,8-linked Sia residue that can carry an O-acetyl group at position C9 (Fig. 1c). When expressed in parental HAP1 cells (HAP1-WT), ST8SIA1 induced the de novo display of both GD3 and 9-O-Ac-GD3, as shown previously20 and in Fig. 1a. Expression of ST8SIA1 in HAP1-ΔSLC33A1 cells yielded GD3, but not its 9-O-acetylated counterpart (Fig. 1b; left panel). The defect was rescued by the expression of human SLC33A1 (Fig. 1b, top right image), providing evidence that SLC33A1 is a crucial component of the Golgi-resident O-acetylation machinery. Albeit ST8SIA1 expression in HAP1 cells resulted in clusters of GD3- and 9-O-Ac-GD3 positive cells as shown in Fig. 1a, their overall yields were surprisingly low, as highlighted by flow cytometry (Supplementary Fig. 2a). Using glycosphingolipid profiling by multiplexed capillary gel electrophoresis with laser-induced fluorescence detection (xCGE-LIF; see schematic representation in Fig. 1d), we identified globotriaosylceramide (Gb3Cer) as the major glycosphingolipid of HAP1 cells and uncovered that GM3 was present only as a minor component (Fig. 1e). This indicates that HAP1 cells convert lactosylceramide, the shared precursor of GM3 and Gb3Cer, predominantly to Gb3Cer, which leads to the observed shortage in the GD3 precursor GM3. Profiling of Chinese hamster ovary (CHO) cells, another cell type known to produce 9-O-Ac-GD3 upon ST8SIA1 expression21,34, revealed that they convert lactosylceramide exclusively to GM3 (Fig. 1f). Consistent with this, transient expression of ST8SIA1 in CHO cells yielded over 80% GD3-positive and 46% 9-O-Ac-GD3-positive cells (Supplementary Fig. 2b), which prompted us to generate Slc33a1-deficient CHO cells as an alternative model to study SLC33A1 functions in a cellular context.

Loss of Slc33a1 abolishes 9-O-acetylation of GD3 and GD2 in CHO cells
Human and hamster SLC33A1 share 95% protein sequence identity (Supplementary Fig. 3) and the hamster Slc33a1 gene has exon-intron boundaries essentially identical to the human gene (schematically depicted in Fig. 2a). To inactivate Slc33a1 in CHO cells, we used three independent CRISPR/Cas9 approaches. Two single guide RNA (gRNA) approaches yielded mutant clones that harbor frameshift-inducing microindels in either exon 1 (Ex1fs) or exon 3 (Ex3fs) of the Slc33a1 gene, while co-application of two gRNAs resulted in an 11.8 kb-deletion spanning exon 1 through 5 (Ex1-5 del). Full details on the mutations are provided in Supplementary Fig. 4. Using total ganglioside extracts of CHO-WT and mutant cells, we examined the ST8SIA1-induced conversion of GM3 to GD3 with concomitant formation of 9-O-acetylated GD3. d Schematic work-flow of GSL-derived glycan profiling by xCGE-LIF. Profiles of GSL-derived glycans of HAP1 (e) and CHO cells (f). Migration times of defined glycan structures were used to assign peaks65. APTS 8-aminopyrene-1,3,6-trisulfonic acid trisodium, xCGE-LIF multiplexed capillary gel electrophoresis with laser-induced fluorescence detection, DAPI 4′,6-diamidino-2-phenylindole, dg derived glycan, GSL glycosphingolipid.
corresponding 9-O-acetylated form, as shown exemplarily in Fig. 2c for the CHO-ΔSlc33a1 clone “Ex1-5 del”. The O-acetylation defect was rescued by additional expression of SLC33A1 (Fig. 2c, far right lane of the top TLC plate). Thus, while genetic inactivation of Slc33a1 did not affect the synthesis of GD3 and GD2 per se, it fully abolished their modification by 9-O-acetylation.

To rule out indirect effects of Slc33a1-deficiency on cell homeostasis, we assessed the metabolic activity of WT and mutant cells in a tetrazolium-reduction assay (Fig. 2e). CHO-ΔCasd1 cells, which lack Sia 9-O-acetylation but harbor a functional Slc33a1 gene, were included as an additional control. Over a period of 72 h, we observed no obvious differences between WT and Slc33a1 KO cells, and after 96 h, only one out of three Slc33a1-deficient clones, namely “Ex1fs”, showed a statistically significant reduction in metabolic activity when compared to the WT control. This argues for a clonal difference rather than a specific effect due to Slc33a1-deficiency, and we selected clone “Ex1-5 del” for further analyses. Quantification of acetyl-CoA by liquid chromatography–mass spectrometry (LC-MS) revealed elevated levels of this central metabolite in CHO-ΔSlc33a1 cells compared to control cells (Fig. 2f), which excluded an overall shortage of cellular acetyl-CoA as a possible cause for impaired 9-O-acetylation in Slc33a1-KO cells.

As summarized schematically in Fig. 2g, our data obtained so far suggest that SLC33A1 forms a functional entity with CASD1. To examine whether the two proteins also target to the same compartment, we investigated their subcellular localization by indirect immunofluorescence analysis upon expression of V5-tagged constructs in CHO cells. As shown previously20, CASD1 colocalizes well with the Golgi marker α-mannosidase II (Fig. 2h, upper panel). Although SLC33A1 displays a more diffuse perinuclear distribution, it nonetheless shows substantial colocalization with the marker protein (Fig. 2h, lower panel), indicating that a significant fraction of the SLC33A1 pool resides in the Golgi apparatus. Our finding contrasts with the strict ER localization observed by subcellular fractionation of CHO cells25, but is consistent with immunofluorescence microscopy data demonstrating colocalization of SLC33A1 with the Golgi marker proteins giantin and syntaxin 6 in MCF-7 cells30.

Patient-derived mutations in SLC33A1 impact the formation of 9-O-Ac-GD3. After the successful generation of Slc33a1-KO cells, we established a complementation approach to evaluate the functional consequences of patient-derived SLC33A1 variants. As a proof-of-concept, we analyzed p.A110P and p.Y366* underlying childhood onset HBS30, p.S113R causing SPG4232, and p.G509S, which has been identified in a patient with late-onset cerebellar ATX33 (see Supplementary Table 1 for details on all patient-derived SLC33A1 mutations known so far). Each variant was co-expressed with ST8SIA1 in CHO-ΔSlc33a1 cells, and their capacity to restore GD3 9-O-acetylation was assessed by immuno-TLC. As seen before, expression of SLC33A1-WT readily complemented the ΔSlc33a1 cells, as indicated by the appearance of a prominent 9-O-Ac-GD3 signal (Fig. 3a, upper panel, +WT/gray box). In contrast, hardly any rescue was achieved with the two HBS variants p.A110P and p.Y366* (blue boxes), indicating that the underlying mutations severely impair SLC33A1 function. The SPG42 variant p.S113R (red box) allowed limited formation of 9-O-Ac-GD3, and the highest complementation capacity was seen for the ATX variant p.G509S (yellow box). For all set-ups, prominent GD3 signals were observed (Fig. 3a, lower panel), validating the functional expression of ST8SIA1 and excluding limitations in the general ganglioside biosynthesis. Detection of 9-O-Ac-GD3 by flow cytometry revealed that each of the analyzed patient-derived variants had a statistically significant lower complementation capacity compared to SLC33A1-WT (Fig. 3b, upper panel; see Supplementary Fig. 5 for gating strategy). Notably, the HBS variants p.A110P and p.Y366* reached only 11.2% and 3.5%, respectively, of the complementation capacity of SLC33A1-WT, whereas a higher restoration capacity of 32% and 57% was seen for p.S113R and p.G509S, respectively. As confirmed by Western blot analysis, all transporter variants were expressed, including the C-terminally truncated nonsense variant p.Y366*, which migrated faster in the SDS-PAGE compared to full-length SLC33A1 (Fig. 3b, lower panel), as shown previously30. The anti-Flag antibody used showed two non-specific bands (marked by asterisks in Fig. 3b), with the lower band migrating close to the truncated variant p.Y366*. For clearer visualization, we analyzed additionally V5-tagged constructs, as shown in Supplementary Fig. 6. Together, our data indicate that each of the analyzed mutations significantly impaired SLC33A1 function, with the HBS-derived mutations having the most severe effect.

Structural analysis of pathogenic SLC33A1 variants
During the revision of this manuscript, Zhou and co-workers solved a cryo-electron microscopy (cryo-EM) structure of human SLC33A1 in complex with acetyl-CoA36. The structure revealed a canonical MFS-fold with 12 transmembrane helices (TMH) arranged as two distinct N- and C-terminal 6-TMH bundles (see Fig. 3c, f for illustration). Among the known patient-derived mutations, Zhou et al. analyzed only the S113R variant, revealing that residue S113 lies in TMH2 near the substrate cavity and suggesting that substitution of S113 with a bulky arginine residue may interfere with acetyl-CoA binding36. In contrast to an earlier report that described the p.S113R variant as inactive24, Zhou et al. reported substantial residual transport activity36, a finding that matches well with the partial complementation activity observed for p.S113R in our cellular approach (Fig. 3b).

To obtain structural insights into the other patient-derived mutations analyzed in our study, we mapped the A110P, G509S, and Y366* variants onto the SLC33A1 structure. Residue A110 locates to TMH2 and forms an intrahelical hydrogen bond with Y106 (Fig. 3c, d). Since proline lacks the backbone amide proton required for such bonding, substituting A110 by proline likely induces a helix kink that affects the positioning of Y106. The resolved cryo-EM structure of SLC33A1 adopts an inward-facing conformation in which the extra-cytosolic site is sealed by coordinated packing of TMH1–2 and TMH7–8, reinforced by a lateral gate formed by TMH2 and TMH1136. In this configuration, Y106 occludes the luminal face of the substrate cavity (Supplementary Fig. 7) and links the luminal ends of TMH2 and TMH11 through a backbone-side chain hydrogen bond with D472 (Fig. 3d). The A110P substitution likely perturbs the helical geometry of TMH2, displacing Y106 and disrupting extra-cytosolic closure—an effect consistent with the pronounced loss of complementation activity observed for the HBS variant p.A110P (Fig. 3b).

The ATX mutation G509S affects a highly conserved residue (Supplementary Fig. 8) that locates to the luminal end of TMH12, which closely approaches the tip of TMH11 (Fig. 3e). As the smallest amino acid, a glycine permits short interhelical distances and the formation of interhelical backbone-to-backbone Cα-H…O=C hydrogen bonds37. Inspection of the cryo-EM structure revealed that the two Cα-H atoms of G509 are in hydrogen bond distance to the backbone carbonyl oxygen atoms of V471 (3.0 Å) and L474 (2.6 Å) of TMH12 (Fig. 3e). Substitution of G509 by a larger serine residue may disrupt this tight arrangement but whether this could affect the architecture of the adjacent luminal loop remains unclear, as this loop is not resolved in the cryo-EM structure36. Finally, we located the truncation site of the HBS variant p.Y366* to the C-terminal end of TMH8 (see Fig. 3f for schematic representation). The resulting loss of two-thirds of the C-terminal 6-TMH bundle likely render this variant non-functional, consistent with its failure to rescue the SLC33A1 knockout (Fig. 3a, b).

Loss of SLC33A1 impairs 7,9-di-O-acetylation of Golgi-localized sialoglycoproteins. Previously, we and others found that many mammalian cell lines display O-acetylated sialoglycans in the Golgi even if significant cell surface O-acetylation is missing20,38. For CHO cells, we demonstrated that the Golgi-confined O-Ac-Sia residues are predominantly 7,9-di-O-acetylated and attached to N- and O-glycans of so far uncharacterized glycoproteins20,39. Having CHO-ΔSlc33a1 cells in hand, we examined whether SLC33A1-deficiency prevents the O-acetylation of Golgi-confined sialoglycans. As analytical tools, we used virolectins, which are based on the hemagglutinin-esterase (HE) proteins of viruses that use 9-O-acetylated sialoglycans as host cell receptors. Specifically, we used HEs derived from influenza C virus (ICV) and bovine coronavirus (BCoV), which were produced as soluble Fc-chimera of HE variants with a genetically inactivated esterase active site (HE0-Fc)20,39. The resulting ICV-HE0-Fc construct recognizes 9-O-Ac-Sia, whereas the BCoV-derived probe largely prefers 7,9-O-Ac-Sia and shows only limited binding affinity toward 9-O-acetylated ligands11,39. In line with our previous data39, staining of permeabilized CHO-WT cells with BCoV-HE0-Fc yielded bright perinuclear signals in virtually all cells, whereas the signals obtained with ICV-HE0-Fc were less frequent and mostly of weaker intensity (Fig. 4a, CHO-WT panel). Golgi localization of the intracellular virolectin ligands was confirmed by co-staining with the Golgi marker α-mannosidase II (Supplementary Fig. 9). No virolectin binding was detected in CHO-ΔCasd1 cells, which were used as a negative control (Fig. 4a, right panel). Intriguingly, all three Slc33a1-KO clones showed an intermediate staining pattern. Compared to CHO-WT cells, we observed a severe loss in BCoV-HE0-Fc binding, while the respective ICV-HE0-Fc binding remained unchanged (Fig. 4a, middle panel). This finding implies that SLC33A1 critically contributes to 7,9-di-O-acetylation of Golgi-confined sialoglycans and, unexpectedly, uncovers that a basal level of mono-O-acetylated sialoglycans prevails even in the absence of the acetyl-CoA transporter.

SLC33A1 is essential but not sufficient for the ST8SIA6-driven formation of 7,9-di-O-acetylated sialoglycans
In an attempt to dissect SLC33A1-dependent and -independent forms of Sia O-acetylation, we searched for a sialyltransferase that is capable of producing 7,9-di-O-acetylated sialoglycans destined for cell surface display, as this would facilitate our analysis. Currently, our knowledge on 7,9-di-O-acetylated sialoglycans is sparse, and sialyltransferases responsible for their biosynthesis have not been identified. Based on literature reports, we considered ST8SIA6 as a promising candidate, as it produces an α2,8-linked disialyl motif on O-glycans40,41, a glycan structure for which both mono- and di-O-acetylated forms have been reported42,43. When we expressed ST8SIA6 in CHO-WT cells, we observed a striking increase in cell surface ligands for both ICV- and BCoV-HE0-Fc (Fig. 4b, c), indicating that ST8SIA6 induced the de novo display of 9-O- and 7,9-di-O-acetylated sialoglycans on the cell surface. The virolectin binding was sensitive to de-O-acetylation by the sialyl-7,9-O-acetylesterase NanS from Tannerella forsythia39, thereby validating the specificity of the used detection tools (Fig. 4c, solid vs. dashed green tracing).

Mock-transfected CHO-WT cells showed only weak surface binding of the two virolectins. The resulting signals were barely visible by immunofluorescence microscopy (Fig. 4b, left panel), but clearly evident in the flow cytometry histograms (Fig. 4c) by a slight, NanS-sensitive shift (solid vs. dashed black traces) when compared to the secondary antibody control (gray fill). Using a gating strategy that included these weakly stained cells (Supplementary Fig. 10), we observed virolectin binding on up to 30% of the mock transfected CHO-WT cells (Fig. 4d; mock), but on none of the CHO-ΔSlc33a1 cells (Fig. 4f; mock). When we challenged the Slc33a1-deficient cells by expressing ST8SIA6, we detected significant ICV-HE0-Fc binding by both fluorescence microscopy (Fig. 4e, lower middle image) and flow cytometry (Fig. 4f, striped dark green column). This indicated that the formation of 9-O-acetylated sialoglycans occurred in an ST8SIA6-dependent but SLC33A1-independent manner. Compared to ST8SIA6-expressing CHO-WT cells, however, we observed a profound loss in BCoV-HE0-Fc binding (Fig. 4e, top middle image), indicating diminished 7,9-di-O-acetylation due to Slc33a1-deficiency. Exogenous expression of human SLC33A1 restored BCoV-HE0-Fc binding (Fig. 4e; top right image). The complementation led to a ratio of BCoV- to ICV-HE0-Fc-positive cells of 1 to 1 (Fig. 4f; solid vs. striped light green columns), whereas without complementation, the ratio was only 0.55 to 1 (solid vs. striped dark green columns). In the latter setting, the actual underlying ratio of 7,9-di-O- to 9-O-acetylated Sia ligands was probably even lower, considering that BCoV-HE0-Fc is not fully selective for 7,9-di-O-acetylated Sia and can bind, albeit with low affinity, to 9-O-acetylated ligands11,39.

In Casd1-KO cells, the expression of ST8SIA6 alone was not sufficient to yield O-acetylated sialoglycans (Fig. 4g, h). Co-expression of ST8SIA6 with human CASD1, however, restored Sia O-acetylation (Fig. 4g, right panel) and yielded an almost equal number of BCoV- and ICV-HE0-Fc positive cells (Fig. 4h; solid vs. striped light green columns). Western Blot analysis confirmed the expression of ST8SIA6 in WT and mutant cells (Supplementary Fig. 11), ruling out the possibility that Slc33a1- or Casd1-deficiency impaired ST8SIA6 expression. The expression and subcellular localization of all transfected constructs were verified by immunofluorescence microscopy (Supplementary Fig. 12). Virolectin stains of permeabilized mock- and ST8SIA6-transfected cells are provided in Supplementary Fig. 13. Collectively, our data showed that the ST8SIA6-driven formation of O-acetylated sialoglycans requires CASD1 but relies only partially on SLC33A1.

Selective inactivation of the luminal domain of CASD1 recapitulates the phenotype of ΔSlc33a1 cells
As shown schematically in Fig. 5a, our previous in vitro analysis of the isolated LCD of CASD1 revealed that the luminal domain transfers acetyl groups from acetyl-CoA to CMP-Neu5Ac via a covalent acetyl-enzyme intermediate that involves the hydroxyl group of the catalytic triad nucleophile S9420. Introduction of an S94A mutation prevented the formation of the acetyl-enzyme intermediate and thus abolished the SOAT activity of the LCD20. The function of the CTMR of CASD1, however, has remained largely unexplored. Based on the AlphaFold2 (AF2) model of CASD1, the CTMR encompasses 14 TMHs that are arranged around a central pore (Fig. 5b). This raises the possibility that the CTMR functions as a membrane SOAT with direct access to cytosolic acetyl groups. If CASD1 comprises two SOAT domains, with only the LCD being dependent on SLC33A1-mediated acetyl-CoA influx, we can expect that inactivation of the LCD and SLC33A1-deficiency will lead to similar outcomes with respect to Sia O-acetylation. To study this experimentally, we complemented CHO-ΔCasd1 cells with either CASD1-WT or CASD1-S94A. As already shown in Fig. 4f, complementation of CHO-ΔCasd1 cells with CASD1-WT enables, in the presence of ST8SIA6, the formation of 7,9-di-O- and 9-O-acetylated sialosides (Fig. 5c, left panel). When we repeated this experiment with CASD1-S94A instead of CASD1-WT, we detected a substantial amount of 9-O-acetylated sialosides (Fig. 5c, middle right image), but hardly any 7,9-di-O-acetylated sialosides (Fig. 5c, top right image). This outcome closely resembles the phenotype of ST8SIA6-expressing CHO-ΔSlc33a1 cells (Fig. 4e, middle panel). Furthermore, complementation of CHO-ΔCasd1 cells with CASD1-WT, but not CASD1-S94A, enabled the formation of 9-O-Ac-GD3 in the presence of the GD3 synthase ST8SIA1 (Fig. 5d and Supplementary Fig. 14). Thus, selective inactivation of the LCD by introducing an S94A mutation into full-length CASD1 recapitulated both the “impaired 7,9-di-O-acetylation” phenotype (Fig. 4e, middle panel) and the “loss of GD3 9-O-acetylation” phenotype (Fig. 2b) of CHO-ΔSlc33a1 cells that became apparent in the presence of ST8SIA6 and ST8SIA1, respectively.

The CTMR of CASD1 functions as transmembrane SOAT
In the Pfam database44, the CTMR of CASD1 has been assigned to the AT3 clan (CL0316) since an iterative searching strategy indicated distant sequence similarity to bacterial sugar acyltransferases of the acyltransferase-3 (acyl_transf_3; AT3) family45. As shown in Supplementary Table 2, the AT3 clan consists of ten protein families, including the HGSNAT_cat family that comprises the catalytic multi-TM domain of the heparan-α-glucosamide-N-acetyltransferase (HGSNAT). HGSNAT is part of the lysosomal heparan sulfate (HS) degradation machinery and transfers acetyl groups from cytosolic acetyl-CoA to terminal glucosamine residues of luminal HS to enable their subsequent hydrolysis46. The recently solved structures of apo- and ligand-bound states of HGSNAT provided detailed insight into the catalyzed TM acyl-group transfer reaction47–49. A unique 9-TMH core architecture has been identified that comprises a “scaffold core” and a “catalytic core” (see Fig. 6a, b for schematic representation)47. The latter binds cytosolic acetyl-CoA in an extended conformation with the acetyl group pointing towards the luminal-oriented residue H297, which catalyzes the transfer reaction via a ternary complex mechanism49. An overlay of the HGSNAT structure with the AF2 model of CASD1 uncovered that TMH5-13 of CASD1 share the 9-TMH core architecture of HGSNAT (Fig. 6b, upper panel), whereas the remaining protein parts lack structural homology (see Supplementary Fig. 15 for superposition of the full-length proteins). In HGSNAT, the catalytic residue H297 is positioned in a luminal loop (connecting TMH2 and TMH3)47–49, whereas the equivalent loop in CASD1 (connecting TMH5 and TMH6) lacks a histidine. However, we identified CASD1-H446, located at the luminal end of TMH5, as potential functional equivalent (Fig. 6c, upper panel). This was further substantiated by comparing CASD1 with an AT3 domain-containing O-acetyltransferase. To date, there are no experimental structures of AT3 domains, but computational studies on the Salmonella O-antigen modifying acetyltransferase OafB suggested that its AT3 domain adopts a novel fold as framework for TM acyl-group transfer50. Our inspection of the AF2 model of OafB revealed a TMH arrangement that is very similar to the 9-TMH core of HGSNAT (Fig. 6b; bottom panel). This includes the splitting of the last “catalytic core”-TMH into two kinked half-helices, a characteristic feature of HGSNAT47–49 that appears to be conserved in both CASD1 and OafB (Supplementary Fig. 16). By superimposing the AF2 models of CASD1 and OafB, we identified OafB-H25 and CASD1-H446 as positional equivalents (Fig. 6c, bottom panel). Notably, H25 is essential for OafB activity and strictly conserved across all bacterial AT3 domain-containing proteins analyzed so far51. Likewise, the residue H446 is conserved not only across CASD1 orthologues of the sialylation competent vertebrate lineage, but also in Cas1p, a Golgi O-acetyltransferase that modifies the capsular polysaccharide glucuronoxylomannan of the pathogenic fungus Cryptococcus neoformans52, as well as in the reduced wall acetylation (RWA) proteins, four Golgi-localized CTMR domain-only proteins that play vital roles in cell wall O-acetylation in Arabidopsis53 (Supplementary Fig. 17).

Closer examination of the CASD1 model revealed that H446 is situated at the bottom of a luminal pocket that could serve as acceptor binding site. A structure-guided approach was then used to derive an initial CTMR–CMP-Neu5Ac complex, in which we sought to test the stability and orientation preference of the acceptor substrate, using molecular dynamics (MD) simulations. The initial complex, embedded in a 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) lipid bilayer model, was used to perform three independent MD simulations that commenced from a different configuration and different initial velocity distributions (see Supplementary Fig. 18 for details). The converged structure snapshot after 2000 ns MD simulation is shown in Fig. 6d. From this simulation, we observed that CMP-Neu5Ac occupies the luminal binding pocket in an orientation that positions the 9′-hydroxyl group of CMP-Neu5Ac within hydrogen bonding distance to H446 (see further detail in Supplementary Fig. 18d). This simple MD model supports the notion that the CTMR of CASD1 can accommodate CMP-Neu5Ac in an orientation that allows O-acetylation via H446, possibly by activating the 9′-hydroxyl group for a nucleophilic attack on the carbonyl group of acetyl-CoA. Collectively, these data indicate that the CTMR of CASD1 functions as an SLC33A1-independent TM SOAT with direct access to cytosolic acetyl-CoA.

To directly demonstrate that the CTMR of CASD1 functions as an O-acetyltransferase, we performed an on-bead enzyme assay using detergent-solubilized, Myc-V5-tagged CASD1 immobilized on Myc nanobody-coated beads as enzyme source (Fig. 6e). After incubation with CMP-Neu5Ac and acetyl-CoA, we quantified the acetyltransferase reaction product HS-CoA using a thiol-sensitive dye, as described recently for HGSNAT48. Under the used conditions, CASD1-S94A retained over 40% of the WT activity (Fig. 6f). Since this mutant lacks a functional LCD, the remaining SOAT activity can be attributed to the CTMR. Although only the double mutant S94A;H446A—harboring defects in both LCD and CTMR—was expected to be inactive, the single mutant CASD1-H446A likewise showed no detectable activity. Effective enzyme loading was verified by Western blot analysis, as shown exemplarily for one experiment in Fig. 6f and for all biological replicates in Supplementary Fig. 19.

To circumvent potential destabilizing effects of detergent solubilization, we employed cellular complementation using CHO-ΔCasd1 cells to further assess the functional impact of CASD1 mutations. The ability of CASD1 variants to restore 9-O-acetylated sialosides upon co-expression with ST8SIA6 was analyzed by flow cytometry using the ICV-derived virolectin (Fig. 6g; with dot plots and expression controls shown in Supplementary Figs. 20 and 21, respectively). In the cellular setting, CASD1-S94A retained >60% of the WT complementation activity. To definitively exclude any contribution from the mutated LCD, we replaced the entire domain with a FLAG epitope. The resulting deletion mutant, CASD1-ΔLCD, exhibited complementation activity comparable to CASD1-S94A, indicating that the CTMR alone accounts for the residual activity. CASD1-H446A, like the double mutant CASD1-S94A;H446A, reached no significant activity over the mock control. Likewise, substitution of H446 with alternative residues (H446Y/F/Q/S) invariably abolished complementation activity (Supplementary Fig. 22). In HGSNAT, alanine replacement of N286, positionally equivalent to CASD1-H446 by structural superposition (see Fig. 6h), abolished enzymatic activity and caused abnormal oligomerization48, highlighting a critical role of this position in protein folding and stability. Moreover, HGSNAT-N286 plays a pivotal functional role by stabilizing the acetyl group of acetyl-CoA within the active site47–49. Based on the structural overlay, we identified CASD1-Q714 as putative functional equivalent of HGSNAT-N286, even though the two residues lie in distinct yet adjacent TMHs of the 9-TMH core (Fig. 6h). Q714 is strictly conserved across phylogenetic distant CASD1 orthologues (Supplementary Fig. 17), and its replacement by alanine reduced complementation activity by ~35% (Fig. 6g). This loss increased to >90% when Q714A was combined with the LCD-inactivating S94A mutation (Fig. 6g; CASD1-S94A;Q714A), indicating a functional role of Q714 for CTMR-mediated O-acetylation.

Furthermore, as shown in Fig. 6i, we found that the four CASD1 residues K435, R428, R489 and R496 align structurally with the four positively charged HGSNAT residues that bind the adenosine diphosphate moiety of acetyl-CoA47–49. In HGSNAT, these sites are essential (R275 and R267) or contributes significantly (R345 and K341) to enzymatic activity48. Consistent with this, alanine substitution of the corresponding CASD1 residues severely impaired complementation activity when combined with the LCD-inactivating S94A mutation (Fig. 6g). In the presence of a functional LCD, however, the loss-of-function effect of the K435A, R428A, R489A and R496A mutations was largely masked by the activity of the LCD. Notably, the four positively charged residues identified as critical for CASD1’s CTMR function are not only structurally congruent with the basic acetyl-CoA-binding residues of HGSNAT (Fig. 6i), but also with a quartet of basic residues in the OafB model (Supplementary Fig. 23), further reinforcing the structural homology of the 9-TMH cores across HGSNAT, CASD1 and OafB.

Methods
All plasmids generated in this study, together with the sequences of the primers used for their construction, are provided in Supplementary Data 1. This Excel file contains multiple worksheets listing the primers used for the construction of SLC33A1, sialyltransferase and CASD1 plasmids, as well as the oligonucleotides employed for the insertion of guide RNAs into CRISPR/Cas plasmids. All primers and oligonucleotides were synthesized by Sigma-Aldrich. Newly generated plasmids described in this study are available from the corresponding author upon reasonable request.

Generation of expression plasmids
For the generation of the plasmid pcDNA3-Flag-SLC33A1-Myc, which encodes human SLC33A1 with an N-terminal Flag- and a C-terminal Myc-tag, we first modified pcDNA3 (Invitrogen) by inserting Flag- and Myc-tag coding sequences into the HindIII/KpnI and XhoI/XbaI restriction sites, respectively, through adapter ligation of the oligonucleotide pairs FLAG-HK-F/FLAG-HK-R and MYC-F/MYC-R. The coding region of human SLC33A1 (accession no. NM_004733.4) was amplified by RT-PCR from total RNA of SK-MEL-28 cells (CLS Cell Lines Service) using the gene-specific primers MG15 and MG16. The resulting PCR product was then ligated into the KpnI/XhoI sites of the modified pcDNA3 vector. Mutations underlying the SLC33A1 variants p.A110P, p.Y366*, p.S113R and p.G509S were introduced into pcDNA3-Flag-SLC33A1-Myc by fusion PCR using the primers MG15 and MG16 in combination with the following mutagenesis primer pairs: MG17/MG18 for generating pcDNA3-Flag-SLC33A1(A110P)-Myc, MG21/MG22 for generating pcDNA3-Flag-SLC33A1(Y366*), MG19/MG20 for generating pcDNA3-Flag-SLC33A1(S113R)-Myc, and MG23/MG24 for generating pcDNA3-Flag-SLC33A1(G509S)-Myc. The plasmid pcDNA3-V5-SLC33A1-Myc was generated from pcDNA3-Flag-SLC33A1-Myc by replacing the Flag-tag coding HindIII/KpnI-fragment by the V5-tag encoding oligonucleotide pair MH178/MH179.

For the generation of pcDNA3-Myc-CASD1, the coding region of human CASD1 was amplified by PCR using the primer pair MaA146/MaA147 and pcDNA3-V5-CASD1-Myc20 as template. The resulting PCR product was ligated into the BamHI/XhoI sites of pcDNA3 (Invitrogen). Prior to this ligation step, the Myc-tag coding sequence was inserted into the HindIII/BamHI sites using the oligonucleotides Myc7s and Myc7as. The plasmid pcDNA3-Myc-CASD1(S94A) was generated from pcDNA3-Myc-CASD1 by replacing the BamHI/EcoRI fragment by the corresponding fragment excised from pcDNA3-V5-CASD1(S94A)-Myc20. The plasmid pcDNA3-Myc-V5-CASD1 was generated by insertion of the V5-tag encoding oligonucleotides MH258/MH259 into the BamHI site of pcDNA3-Myc-CASD1. The plasmid pcDNA3-Myc-V5-CASD1(S94A) was generated from pcDNA3-Myc-V5-CASD1 by replacing the SacII/EcoRI fragment by the corresponding fragment excised from pcDNA3-Myc-CASD1(S94A). The plasmid pcDNA3-Myc-V5-CASD1(S94A;H446A) was generated by fusion PCR using pcDNA3-Myc-V5-CASD1(S94A) as template and the mutagenesis primer pair LyB8/LyB7 in combination with the flanking primers LyB6 and AMB46. The resulting PCR fragment was ligated into the SacII/XbaI sites of pcDNA3-Myc-V5-CASD1, thereby replacing the original SacII/XbaI fragment. All other amino acid exchanges within the CTMR of CASD1 were introduced accordingly using either pcDNA3-Myc-V5-CASD1 or pcDNA3-Myc-CASD1(S94A) as template and the mutagenesis primers listed in Supplementary Data 1. For the generation of the plasmid pcDNA3-Myc-V5-CASD1-ΔLCD, the sequence encoding the LCD of CASD1 (amino acid residues 61-300) was substituted by an in-frame insertion of a sequence stretch that encodes the FLAG epitope MDYKDDDD. In a first step, the sequence stretches flanking the deletion site were amplified with the primers LyB6/AMB47 and AMB48/AMB46, of which AMB47 and AMB46 contain complementary sequence stretches that encode the FLAG epitope. In a second step, the obtained amplification products were fused in a PCR reaction with the primer pair LyB6/AMB46, and the final product was ligated into the SacII/XbaI sites of pcDNA3-Myc-V5-CASD1, replacing the original SacII/XbaI fragment.

The plasmid pcDNA3-V5-ST8SIA1 was generated by cloning the HindIII/NotI fragment excised from pcDNA3.1-zeo-V5-ST8SIA120 into the HindIII/NotI sites of the pcDNA3 (Invitrogen). The bi-cistronic construct pcDNA3-ST8SIA1-2A-B4GALNT1 carries the sequence encoding human ST8SIA1 (NM_003034.3) without stop-codon followed in-frame by the sequence encoding the 2A “self-cleaving” peptide from equine rhinitis A virus (QCTNYALLKLAGDVESNPGP) and the sequence encoding human B4GALNT1 (NM_001478.5). This tripartite coding sequence was generated by PCR in three steps: (i) amplification of a fragment encoding ST8SIA1 fused to the 2A peptide using the primer pair LS58/LS67 and pcDNA3-V5-ST8SIA1 as template, (ii) amplification of a fragment encoding the 2A peptide fused to B4GALNT1 using the primer pair LS66/LS62 and pTWIST-Amp-High-Copy-hB4GALNT1 as template (Twist Biosciences, insert generated by custom DNA synthesis), and (iii) fusion PCR with the flanking primers LS58 and LS62. The resulting PCR fragment was ligated into the NotI/XbaI sites of pcDNA3. The plasmid pcDNA3-V5-ST8SIA6 was generated by the amplification of the coding sequence of human ST8SIA6 (accession no. NM_001004470.2) using the primer pair MaA90/MaA93 and pcDNA3.1*/C-(K)-DYK-ST8SIA6 (GeneScript, cat. OHu03577) as template. The resulting PCR product was ligated into the BamHI/XbaI restriction sites of pcDNA3-V5-CASD1-Myc, thereby replacing the original CASD1-Myc encoding BamHI/XbaI fragment by the ST8SIA6 encoding sequence. The identity of all plasmids was confirmed by Sanger sequencing. Sequences of the PCR primers are listed in Supplementary Data 1.

Cell culture
HAP1 cells (Haplogen Genomics, Austria; RRID: CVCL_Y019) were grown in Iscove’s Modified Dulbecco’s Medium (Gibco) supplemented with 5% fetal calf serum (FCS) and 1 mM sodium pyruvate. Chinese Hamster Ovary K1 (CHO-K1) cells were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM)/Ham’s F12 1:1 (PAN-Biotech) supplemented with 5 % FCS. Cultures were maintained at 37°C with a humidified atmosphere of 5% CO2.

CRISPR/Cas9-mediated genome editing
HAP1-ΔSLC33A1 cells (HAP1_SLC33A1_36819-02) were custom-made by Haplogen Genomics (Austria), using the Streptococcus pyogenes nuclease Cas9 (SpCas9) together with a guide RNA containing the gene-specific sequence 5’-GTAGACCGCATCAACCAACG-3’ for targeting exon 1 of human SLC33A1. The obtained HAP1-ΔSLC33A1 clone carries a 2-bp frameshift deletion in exon 1 of SLC33A1 as shown in Supplementary Fig. 1.

CHO-ΔCasd1 cells were established previously21 by introducing a frameshift mutation in exon 2 of Casd1 using the single guide RNA strategy described below. CHO-ΔSlc33a1 cells were generated by using the pX330-U6-Chimeric_BB-CBh-hSpCas9 plasmid (Addgene plasmid # 42230, a gift from Feng Zhang)63 with the gene-specific target sequence inserted into the BsbI sites of the guide RNA scaffold. CCTop-CRISPR/Cas9 target online predictor64 was used for the identification of target sites. Information on the selected sites and the respective oligonucleotide sequences used for the generation of single guide RNAs targeting exon(Ex) 1, 3 or 5 of Slc33a1 are provided in Supplementary Data 1. The resulting plasmids pX330-Slc33a1-Ex1, pX330-Slc33a1-Ex3, and pX330-Slc33a1-Ex5 were introduced into CHO cells, either individually for single guide RNA approaches to induce frameshift mutations or as a combination of pX330-Slc33a1-Ex1 and pX330-Slc33a1-Ex5 to introduce a multiple exon-spanning deletion. To facilitate the selection of transfected cells, an enhanced green fluorescent protein (EGFP) reporter plasmid (pEGFP-C1, Clontech) was co-transfected in a 4:1 ratio of pX330-based plasmid(s) to pEGFP-C1. Single green fluorescent colonies were selected and genotyped by PCR and Sanger sequencing. Clones with homozygous or heterozygous frameshift mutations or multiple-exon spanning deletions were subcloned and genotyped on genomic and transcript level. For the analysis of genomic DNA, 2×105 cells were lysed in 20 µL of lysis buffer (10 mM Tris-HCl pH 7.6, 50 mM NaCl, 6.25 mM MgCl2, 0.045% NP-40, 0.45% Tween-20) containing 1 mg mL−1 proteinase K. After an incubation for 1 h at 56 °C and heat-inactivation of the proteinase for 15 min at 95 °C, an aliquot of 1 µL of lysate was analyzed by PCR using the following primer combinations: 5’-CCGGGGATCTGGCTCTGTGC-3’ and 5’-GGATGCTCCCTGCCAGGCC-3’ for amplification of the Ex1 target site, 5’-CAGTTCTACCAATATTGCAT-3’ and 5’-GTGCACAATACTGAACACA-3’ for amplification of the Ex3 target site, 5’-TTCCACCATTC-CAGTGTTCTCT-3’ and 5’-ATTTCTAAGCAGTTCAGCAACTCTC-3’ for amplification of the Ex5 target site, and 5’-ATGTCACCGACCATCTCC-3’ and 5’-ATTTCTAAGCAGTTCAGCAACTCTC-3’ for the analysis of deletions spanning exon 1 to 5. The first primer of each combination was also used for Sanger sequencing of the obtained PCR products. Mutations were confirmed on transcript level by Sanger sequencing of RT-PCR products, which were obtained from total RNA using the following primer pairs: 5′-ATGTCACCGACCATCTCC-3′ (5′-end of exon 1) and 5′-GAGGGACCATTGGAACAGCA-3′ (exon 3), 5′-AACGTGGGC-TATGCTTCCAC-3′ (3’-end of exon 1) and 5′-AGAGCTGTCACA-CATGAGCC-3′ (exon 6), and 5′-ATGTCACCGACCATCTCC-3′ (5′-end of exon 1) and 5′-AGAGCTGTCACACATGAGCC-3′ (exon 6).

Transfection of mammalian cells
If not stated otherwise, transfections were performed with Polyethylenimine ‘Max’ (PEI MAX, Polysciences). Cells were grown to 70–80% confluency in 6-well plates containing 2 mL of culture medium per well or, alternatively, on 10 cm dishes containing 10 mL of culture medium. Per well of a 6-well plate, 2 µL PEI MAX were diluted in 100 µL of OptiMEM (Gibco) and mixed with 100 µL of OptiMEM containing 2 µg of plasmid DNA. After an incubation for 15–20 min at room temperature, the PEI MAX-DNA mixture was added to the culture medium. Cells grown in 10 cm dishes were transfected accordingly with reagent amounts scaled by a multiplication factor of 5. Transfections were stopped after 6 h by replacing the PEI MAX-containing medium by fresh culture medium. Cells were harvested 48 h after transfection for ganglioside analyses and after 24 h for immunofluorescence microscopy. For the subcellular localization analysis shown in Fig. 2h, transfections were performed with Metafectene (Biontex) instead of PEI MAX. All steps were performed in 6-well plates as described above, except that 3 µL of Metafectene and 0.5 µg of plasmid DNA were used per well.

Immunofluorescence analysis
Cells were seeded on glass coverslips placed in 6-well plates and transfected as described above. Adherent cells were washed with PBS, fixed with 4% paraformaldehyde in PBS for 15 min at room temperature, and washed again with PBS. For the detection of Golgi localized proteins, cells were permeabilized with 0.2% (v/v) Triton X-100 in PBS for 10 min at 4 °C. After blocking for 30 min with 1% bovine serum albumin (BSA) in PBS, cells were stained for 1 h at room temperature with the following primary antibodies or virolectins diluted in 1% BSA in PBS: mouse anti-9-O-Ac-GD3 mAb (1 µg mL−1; Santa Cruz cat. Sc-32269, clone UM4D4), mouse anti-GD3 mAb R24 (2.5 µg mL−1; purified by protein A affinity chromatography from cell culture supernatant of hybridoma cells ATCC HB-8445), mouse anti-Flag mAb (5 µg mL−1; Sigma-Aldrich cat. F1804, clone M2), rabbit anti-Flag pAb (DYKDDDDK-Tag antibody; 0.28 µg mL−1; Cell Signaling cat. 2368), mouse anti-Myc mAb (10 µg mL−1; Thermo Fisher, MA1-980, clone 9E10), mouse anti-V5 mAb (1 µg mL−1; Acris cat. SM1691PS, clone SV5-PK1), chicken anti-V5 pAb (2 µg mL−1; Abcam cat. ab9113), rabbit anti-α-ManII pAb (1:10,000; kindly provided by Kelley Moremen), BCoV-HE0-Fc (40 µg mL−1), and ICV-HE0-Fc (10 µg mL−1). Cells were washed and incubated with secondary antibody (diluted in 1% BSA in PBS) for 1 h at room temperature: goat anti-chicken IgY Alexa Fluor 488-conjugate (1:1000; Invitrogen cat. A-11039), donkey anti-human IgG Dylight 550-conjugate (1:1000; Invitrogen cat. SA5-10127), goat anti-mouse IgG Alexa Fluor 488-conjugate (1:500; Invitrogen cat. A-11029), sheep anti-mouse IgG Cy3-conjugate (1:1000; Sigma-Aldrich cat. C2181), donkey anti-mouse IgG Alexa 555 (1:500; Invitrogen cat. A-31570), goat anti-mouse IgG2a Alexa Fluor 488-conjugate (1:500; Invitrogen cat. A-21131) rabbit anti-mouse IgG3 DyLight 549-conjugate (1:5000; Rockland cat. 610-442-043), goat anti-mouse IgM Alexa Fluor 568-conjugate (1:500; Invitrogen cat. A-21043), goat anti-rabbit IgG Alexa Fluor 488-conjugate (1:500; Invitrogen cat. A-11008), sheep anti-rabbit IgG Cy3-conjugate (1:1000; Sigma-Aldrich cat. C2306). After washing with PBS and H2O, air-dried cells on cover slips were mounted in 3 µL Vectashield mounting medium containing 4′,6-diamidino-2-phenylindole (DAPI) (Vector Laboratories). Stained cells were analyzed using an Axiovert 200M microscope equipped with an ApoTome module and an AxioCam MRm digital camera (Zeiss). Images were processed using the software ZEN 2012 (Zeiss).

Glycan profiling
Profiling of GSL-derived glycans by multiplexed capillary gel electrophoresis coupled to laser-induced fluorescence detection (xCGE-LIF) was performed as described65. Briefly, total GSLs extracted from 107 cells were digested with 1 µL of LudgerZyme Ceramide Glycanase (Ludger) in a total volume of 6.5 µL LudgerZyme Ceramide Glycanase RXN Buffer for 24 h at 37 °C. Samples were dried and mixed with 2 µL of 8-aminopyrene-1,3,6-trisulfonic acid trisodium salt (APTS, Sigma-Aldrich; 20 nM in 3.5 M citric acid, 2 µL of 2-picoline borane complex (Sigma-Aldrich; 2 M in DMSO) and 2 µL of H2O. The reaction mixture was incubated at 37 °C in the dark and stopped after 16.5 h by the addition of 100 µL of acetonitrile/H2O (80:20, v/v). APTS-labeled glycans were purified by hydrophilic interaction chromatography-solid phase extraction (HILIC-SPE) and analyzed by xCGE-LIF using a remodeled ABI PRISM 3100-Avant genetic Analyzer (Applied Biosystems). Peaks were assigned according to the migration time of structurally defined glycan standards.

Ganglioside extraction and immuno-thin-layer chromatography
Total gangliosides were extracted with chloroform/methanol (1:2, v/v; 3 mL per 107 cells) in conjunction with ultrasonic dispersion, followed by solid-phase extraction on Chromabond C18 columns (Macherey-Nagel)21,39. For TLC, total gangliosides of an equivalent of 2×106 cells were spotted on Nano-DURASIL-20 HPTLC plates coated with 0.2 mm silica gel 60 (Macherey-Nagel). Plates were chromatographed in chloroform/methanol/H2O (50:40:10, v/v/v) containing 0.05% calcium chloride and dried. Plates were then chromatographed twice in 0.5% isobutyl methacrylate polymer (TCI, cat. M0086), dried and incubated overnight in PBS at 37 °C. After blocking with 2% BSA (w/v) in PBS for 1 h at room temperature, plates were washed with PBS and stained for 1 h at room temperature with the following primary antibodies diluted in PBS: mouse IgM anti-9-O-Ac-GD3 mAb (5 µg mL−1; Ancell, ANC-212-820, clone UM4D4), mouse IgM anti-9-O-Ac-GD3 mAb (1:40; Thermo Scientific; cat. MA1-34707, clone M-T6004), mouse IgG3 anti-GD3 mAb R24 (10 µg mL−1; affinity purified from cell culture supernatant of hybridoma cells ATCC HB-8445)20, mouse IgG3 anti-9-O-Ac-GD2 mAb 8B6 (10 µg mL−1; kindly provided by OGD2 Pharma, Nantes), or mouse IgG2a anti-GD2 mAb (15 µg mL−1; Kerafast EWI023, clone ME361). Plates were washed with PBS and incubated with the following secondary antibodies for 1 h at room temperature: goat anti-mouse IgG IRDye 800CW-conjugate (1:10,000; LI-COR Biosciences, cat. 926-32210) or goat anti-mouse IgM IRDye 800CW-conjugate (1:20,000; LI-COR Biosciences, cat. 926-32280). TLC plates were washed and signals were recorded using an infrared imaging system (Odyssey Imaging System, LI-COR Biosciences). Purified GD3 (Sigma-Aldrich, cat. 345752) and GD2 (Sigma-Aldrich, cat. 345743) were used as standards (0.2 µg per lane). The corresponding 9-O-acetylated forms were generated by in vitro O-acetylation of purified GD2 and GD3 using the Campylobacter jejuni O-acetyltransferase NeuD, recombinantly expressed as maltose-binding protein (MBP) fusion protein in Escherichia coli21,39. In a total volume of 20 µl, 27.5 µM ganglioside were incubated in the presence of 10 µg MBP-NeuD, 1.5 mM acetyl-CoA (Sigma-Aldrich), 0.05% sodium cholate, 10 mM MgCl2, 1 mM dithiothreitol and 50 mM MES buffer pH 7.2. After 6 h at 37 °C, the reaction was stopped by adding an equal volume of methanol. Gangliosides were purified by solid-phase extraction on Chromabond C18 columns (Macherey-Nagel) and dissolved in chloroform/methanol (1:2, v/v). Uncropped images of the TLC plates are available in the Source Data file.

Cell viability assay
Cell viability and proliferation was measured in a tetrazolium dye assay using water-soluble tetrazolium reagent-1 (WST-1) according to the manufacturer’s protocol (Roche). CHO cells were seeded at a density of 500 cells per 100 µL culture medium per well of a 96-well culture plate and cultivated for 24 h, 48 h, 72 h or 96 h. The medium was changed, followed by the addition of 10 µL of WST-1 assay reagent. After 2 h of incubation at 37 °C and 5% CO2, absorbance at 450 nm was measured against a reference wavelength at 690 nm using a microplate reader (PowerWave 340, Bio-Tek). Cell-free wells were used as blank control.

Acetyl-CoA measurements
CHO cells were seeded at a density of 3×106 cells per 10 cm dish and allowed to adhere for 24 h at 37 °C and 5% CO2. Acetyl-CoA was extracted from cells by chloroform-methanol liquid-liquid extraction66. Adherent cells were washed three times with ice-cold PBS and cellular metabolism was quenched by the addition of 500 µL of methanol pre-cooled to -20 °C. Isotopically U-13C-labeled metabolite yeast extract (Isotopic solutions, cat. G83-14-07-2021) was dissolved in water and used as internal standard. An aliquot corresponding to 150 µg of yeast dry cell weight in a total volume of 1 mL dest. H2O was added to each dish and cells were scraped and transferred to a microfuge tube containing 500 µL of chloroform pre-cooled to -20 °C. Cell extraction was performed for 20 min at 4 °C in an Eppendorf mixer at 1400 rpm. After phase separation by centrifugation (5 min at 16,100×g, 4 °C), the interphase was retained for protein extraction as described below and acetyl-CoA was isolated from the polar phase by 2-(2-pyridyl)ethyl silica gel based solid-phase extraction (SPE)67. SPE columns packed with functionalized (2-pyridyl)ethyl silica gel (100 mg, Merck, cat. 54127-U) were equilibrated with 1 mL of equilibration buffer (45% ACN, 20% H2O, 20% acetic acid, 15% isopropanol (v/v), pH 3). After equilibration, the samples (dissolved in the polar methanol phase) were loaded on the SPE column and washed with 1 mL of the equilibration buffer. Acetyl-CoA was eluted from the column using 2 mL of methanol/250 mM ammonium formate 4:1 (v/v, pH 7). The solvent was evaporated under a nitrogen stream while samples were kept on ice and dried samples were stored at -80 °C. Prior to analysis, samples were reconstituted with 100 µL 50% ACN in dest. H2O. Acetyl-CoA levels were quantified by hydrophilic interaction chromatography (HILIC) coupled to mass spectrometry (HILIC-MS)68. An aliquot of 10 µL was injected into an UPLC system (Vanquish Flex, Thermo Scientific) equipped with a zwitterionic HILIC column (Atlantis Premier BEH z-HILIC VanGuard FIT, 1.7 µm, 2.1×150 mm, Waters). Analytes were loaded for 1 min at 90% B, and separated using a linear gradient ranging from 90 to 65% B in 5 min with a flow rate of 500 µL min−1 (A: 15 mM ammonium bicarbonate (AmBiCa) in H2O, pH 9; B: 90% ACN, 15 mM AmBiCa in H2O, pH 9). The UPLC system was coupled via an electrospray-ionization (ESI) source to a tribrid orbitrap mass spectrometer (Orbitrap Fusion Lumos, Thermo Scientific). Single ion monitoring (SIM)-MS analysis of acetyl-CoA was performed in positive ion mode to quantify acetyl-CoA. Parameters were set as follows: orbitrap resolution: 60k (at m/z 200), maximum ion injection time: 118 ms, normalized AGC target: 100%, RF lens: 50%, isolation windows: 12C-acetyl-CoA: 810.1330±5 m/z, U-13C-acetyl-CoA: 833.2102±5 m/z, retention time: 2.8–3.8 min. Samples were measured in a randomized sequence. An external standard row with 7 standards ranging from 0.19 ng on column to 12.5 ng was measured before and after the samples to enable quantification.

LC-MS raw data were processed using TraceFinder (version 5.0.889.0, Thermo Fisher Scientific) with the following parameters: detection algorithm: Genesis, peak detection strategy: highest peak, peak area: 1%, smoothing: 5, min peak height S/N: 2. Manual integration of peaks was performed when necessary. Acetyl-CoA data from cell extracts were normalized to U-13C-labeled acetyl-CoA as internal standard to compensate for any possible variations during sample processing and to total protein content as a surrogate for cell number. For protein determination, the interphase of each cell extract was washed with methanol, pelleted by centrifugation (10 min at 16,100×g, 4 °C) and air dried at 35 °C for 15 min. Proteins were dissolved in 500 µL of 8 M urea, assisted by sonication (Branson Ultrasonic Sonifier S-450 equipped with a cup horn). The bicinchoninic acid (BCA) protein assay was performed on serially diluted samples according to the manufacturer’s protocol (Pierce).

Western Blot analysis of whole cell lysates
For the detection of Flag-tagged SLC33A1 variants, cell lysates were prepared using n-dodecyl-β-D-maltopyranoside (DDM; Anatrace) as detergent. A total number of 1×106 transfected CHO cells were lysed in 70 µL of DDM-lysis buffer (50 mM Tris-HCl pH 8.0, 150 mM NaCl, 5 mM EDTA, 1% DDM, 200 U mL−1 aprotinin, 10 µg mL−1 leupeptin, 2 mM PMSF) for 15 min on ice. After the removal of cell debris by centrifugation at 13,000×g for 15 min at 4 °C, lysates were mixed with an equal volume of 2-fold concentrated Laemmli sample buffer. After an incubation for 10 min at 65 °C, samples of an equivalent cell number (9×104 cells per lane) were separated by 10% SDS-PAGE, and proteins were transferred to polyvinylidene difluoride (PVDF) membrane by semi-dry blotting. Membranes were blocked in 3% BSA in PBS and incubated with rabbit anti-Flag pAb (DYKDDDDK-Tag antibody; 0.02 µg mL−1; Cell Signaling cat. 2368) overnight at 4 °C. Membranes were washed and incubated for 1 h at room temperature with goat anti-rabbit IgG horseradish peroxidase (POD)-conjugate (1:20,000; Sigma-Aldrich cat. A6154), followed by enhanced chemiluminescence (ECL) detection using an Amersham Imager 680 (Cytiva). Bound antibodies were stripped by incubating the membranes in stripping buffer containing 2% SDS and 0.8% β-mercaptoethanol in 62 mM Tris-HCl (pH 6.8) for 30 min at 50 °C. After washing with H2O, membranes were blocked and stained with mouse anti-actin mAb (1:100,000; Merck Millipore cat. MAB1501, clone C4) and goat anti-mouse IgG POD-conjugate (1:20,000; SouthernBiotech cat. 1010-05) following the same protocol as described above. All antibodies were diluted in blocking solution (3% BSA in PBS). All antibodies were diluted in blocking solution (3% BSA in PBS).

For the detection of V5-tagged ST8SIA6 (Supplementary Fig. 11), 1×106 cells were lysed in 70 µL of Triton-lysis buffer (50 mM Tris-HCl pH 8.0, 150 mM NaCl, 5 mM EDTA, 2% Triton X-100 (v/v), 0.5% sodium deoxycholate (w/v), 200 U mL−1 aprotinin, 10 µg mL−1 leupeptin, 2 mM PMSF). Samples were processed as described above and an equivalent of 7×104 cells was applied per lane. After blotting, PVDF membranes were blocked with Odyssey blocking buffer (LI-COR Biosciences) diluted 1:2 with PBS. Antibody staining was performed in the same buffer system using mouse anti-V5 mAb (0.1 µg mL−1; Acris cat. SM1691PS, clone SV5-PK1) followed by goat anti-mouse IgG IRDye 800CW-conjugate (1:20,000; LI-COR Biosciences cat. 926-32210) and rabbit anti-Actin pAb (1:1000; Sigma-Aldrich cat. A2066) followed by anti-rabbit IgG IRDye 800CW-conjugate (1:20,000; LI-COR Biosciences cat. 926-32211). Signals were detected by an infrared imaging system (Odyssey Imaging System, LI-COR Biosciences). Uncropped images of the blots are available in the Source Data file.

Structure prediction and analyses
The Protein Data Bank (PDB) coordinate files of the AF2 models of human CASD1 and Salmonella OafB were downloaded from the AlphaFold Protein Structure Database69,70 with the accession AF-Q96PB1-F1 and AF-A0A0H2WM30-F1, respectively. The PyMOL Molecular Graphics System (DeLano Scientific) was used for visual inspection of the structural models and for producing graphical images. Superpositions of structures were obtained by employing the super command implemented in PyMOL.

Expression and purification of virolectins
The virolectins BCoV-HE0-Fc and ICV-HE0-Fc were produced as secreted soluble Fc-chimera in baculoviral infected Trichoplusia ni (High5) insect cells and were purified from the cell culture supernatant by protein A affinity chromatography39. The expressed proteins encompass an enzymatically inactive HE ectodomain (amino acid residues 19-388 of BCoV-LUN HE carrying an S40A exchange or residues 15-631 of influenza C/California/78 HE carrying an S71A exchange) followed by the Fc part of human IgG120,39.

Flow cytometry
Cells were washed with excess of FACS buffer (PBS containing 1% BSA and 1 mM EDTA) and adjusted to 2.5×105 cells per 100 µL of FACS buffer. Samples were stained for 1 h at 4 °C with the following primary antibodies or virolectins in 100 µL FACS buffer: anti-9-O-Ac-GD3 mAb (1:20; mouse IgM, Thermo Scientific, cat. MA1-34707, clone M-T6004), BCoV-HE0-Fc (40 µg mL−1), or ICV-HE0-Fc (10 µg mL−1). Cells were washed twice and incubated with secondary antibody for 30 min at 4 °C using rat anti-mouse IgM PE-conjugate (1:80; BD cat. 553409) or goat anti-human IgG R-Phycoerythrin (R-PE)-conjugate (0.5 µg mL−1; Jackson ImmunoResearch, cat. 109-115-098). Cells were washed and suspended in 1 mL of FACS buffer. 15,000 events per sample were measured on a CyFlow ML flow cytometer (Sysmex Partec) equipped with a 488 nm laser. Data were analyzed using the FlowJo software Version 7.6.

Enzymatic de-O-acetylation
To validate specific virolectin staining, O-acetyl groups were removed by the sialate-7,9-O-acetylesterase NanS from Tannerella forsythia. Recombinant NanS carrying a C-terminal hexa-histidine-tag was expressed in Escherichia coli and purified by affinity chromatography39. Cells were adjusted to 2.5×105 cells per 100 µL of FACS buffer and incubated in the presence of 5 µg of NanS for 1 h at room temperature. Afterwards, cells were stained and analyzed by flow cytometry as described above.

Computational methods
All molecular dynamics (MD) simulations were performed using the GPU version of Gromacs 202271,72. The Gromos 54A773,74 force field was used to model protein as well as the model lipid bilayer (DPPC: 1,2-dipalmitoyl-sn-glycero-3-phosphocholine) membrane75,76. The force field parameters for the ligand CMP-Neu5Ac were obtained from the GROLIGFF77 web server. A model structure of the human CASD1 TM domain spanning P307-H796 was generated by AlphaFold2 using the ColabFold platform with default parameters69,78. The CMP-Neu5Ac ligand was positioned in the binding pocket by using structure-guided manual docking based on the co-crystal structures of Heparan α-glucosamine N-acetyltransferase (PDB: 8JKV, 8JL1, 8JL3, 8JL4, 8TU9, 8VKJ, 8VLG, 8VLI, 8VLU, 8VLY). The protonation state of titratable groups was chosen appropriate to pH 7.0. Each ligand-protein complex was embedded into a model membrane, composed of 512 DPPC molecules, using the GROMACS program g_embed. The complex was resized in the XY directions (corresponding to the plane of the membrane) to 10% of their original size and to insert the α-helices any overlapping lipid and water molecules were deleted. The complex was then grown stepwise to the original size. The final simulations rectangular box had approximate dimensions of 12.9 nm × 12.9 nm × 8.9 nm and contained ~27,000 water molecules and overall ~111,000 atoms (see Supplementary Fig. 18a).

Each system was simulated under periodic boundary conditions in a rectangular box. The pressure was maintained at 1 bar by weakly coupling the system to a semi-isotropic pressure bath using isothermal compressibility of 4.6×10−5 bar−1 and a coupling constant τP = 1 ps in the XY and Z directions, corresponding to the plane and normal directions of the bilayer, respectively79. The temperature of the system was maintained at 298 K by independently coupling the protein-ligand complex, lipids and water to an external temperature bath with a coupling constant τT = 0.1 ps using a Berendsen thermostat79. A nonbonded interaction cut-off of 0.14 nm was used. Long-range electrostatics were treated with the Reaction Field80 method. All bond lengths within the protein and lipids were constrained using the LINCS81 algorithm. The simple-point charge (SPC)82 water model was used and constrained using the SETTLE83 algorithm.

Each system was energy minimized for 1000 steps using steepest descent method followed by a position-restrained MD simulation where all heavy atoms of protein-ligand complex were restrained to their original position using 1000 kJ/mol/nm2 allowing lipid and water molecules to equilibrate. The restraints were removed and the whole system was allowed to equilibrate for 5 ns. The production MD simulations were performed for 2000 ns (in triplicate – two starting with a different initial velocity distribution and one starting from a different configuration) for each system. Supplementary Fig. 18 shows that substrate relaxation and binding orientation are reached within the simulated timeframe, consistent with the process being accessible on the brute-force MD timescale used here. All coordinates, velocities, forces, and energies were saved every 10,000 steps for analysis. The stability of the protein and protein-ligand complexes were evaluated by measuring the root mean square deviation (RMSD) of protein backbone as well as ligand atoms by fitting the backbone atoms of protein. The molecular dynamics checklist can be found in Supplementary Table 3. The molecular dynamics simulation data (starting structure, force field parameter files, molecular dynamics parameter, and trajectories) generated in this study have been deposited in the Zenodo database (https://doi.org/10.5281/zenodo.18795789)84.

On-bead O-acetyltransferase assay
To measure CASD1-mediated SOAT activity, we performed an on-bead enzyme reaction with affinity-captured CASD1 as enzyme source, and quantified the acetyltransferase reaction product HS-CoA via fluorescence-based thiol detection. Briefly, we expressed WT and mutant forms of N-terminally Myc-V5-tagged CASD1 in CHO-ΔCasd1 cells using the PEI transfection protocol described above. For each enzyme reaction, Myc-V5-CASD1 was affinity captured from the detergent lysate of 1.5×10⁷ cells harvested 48 h after transfection. Lysis was performed in 600 µL DDM-lysis buffer (50 mM Tris pH 8.0, 150 mM NaCl, 5 mM EDTA, 0.5% DDM, 200 U mL−1 aprotinin, 10 µg mL−1 leupeptin) for 1 h at 4 °C, and the obtained lysates were clarified by centrifugation at 16,100×g for 15 min at 4 °C. Anti-Myc nanobody-coupled magnetic agarose beads (Myc-Trap, ChromoTek) were added to the cleared lysates using 40 µL of the Myc-Trap suspension, and the affinity capture was performed for 2 h at 4 °C on a rotary wheel. Beads were washed once with 1 mL of wash buffer (10 mM MES pH 6.5, 1 mM MnCl₂, 150 mM NaCl) with 0.05% DDM and twice without detergent, followed by two washes with reaction buffer (10 mM MES pH 6.5, 1 mM MnCl₂). Affinity beads incubated with the detergent lysate of mock-transfected cells were used as a non-enzyme control.

The on-bead enzyme reaction was performed in a total volume of 60 µL reaction buffer containing 1 mM acetyl-CoA trisodium salt (Sigma-Aldrich) and 1 mM CMP-Neu5Ac sodium salt (Roche) with the affinity-captured material as enzyme source. After a 3-h incubation at 37 °C with continuous rotary motion, beads were removed by magnetic separation. The bound protein was subsequently analyzed by Western blotting with an anti-V5 antibody as described above for the detection of V5-tagged ST8SIA6. The reaction mixture was passed through a 50 kD cut-off filter. Ten microliters of the ultrafiltrate were combined with 100 µL of Measure-iT Thiol Assay Reagent (Thermo Fisher Scientific) and the fluorescence intensity was measured using an Infinite 200 Pro microplate reader (Tecan) with λEX/λEM = 485/520 nm. For the calculation of relative fluorescence intensities, the signal from the non-enzyme control was defined as 0% and the signal from the WT control was set to 100%. Statistical significance was determined by one-way ANOVA using GraphPad Prism v8.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This manuscript by Albers et al. focuses on a membrane protein sialate O-acetyltransferase CAS1 domain-containing 1 (CASD1). The primary finding—that CASD1 possesses two active sites, one dependent on SLC33A1 (acetyl-CoA transporter) and one independent—is exciting and significantly advances our understanding of this enzyme and Golgi acetylation processes. The manuscript is well-structured, with a clear presentation of results, and this reviewer identifies no major concerns. I strongly support its publication in Nature Communications, with some minor concerns detailed below.\n1. In Figure 2f, only the Ex 1-5 del clone is presented. Why were the other two Ex clones omitted? Including all three would allow readers to better assess clonal variation in the data. Additionally, could the authors clarify when acetyl-CoA levels were measured and whether these measurements align with the time points in Figure 2e?\n2. In Figure 3b, the anti-Flag western blot shows an impurity band near the Flag-SLC33A1-Y336* truncated protein. Could the authors clarify its identity or source? The band’s intensity appears correlated with ST8SIA1 expression.\n3. The MD simulation details should be included in the main methods section rather than relegated to the supplementary material. Given the complexity of the simulation system, the energy minimization and equilibration steps seem brief. Furthermore, the production phase of only 100 ns is relatively short; with current computational resources, extending it to 1 µs would be feasible and likely provide more robust insights.", "reviewer_2": "This manuscript makes some interesting observations regarding the mechanisms of sialic acid acetylation in mammals. This is an active area of research in the glycobiology field with important ramifications on our understanding of genetic disorders, cell signalling and host-pathogen interactions.\nThe main claims of this manuscript are:\n1) SLC33A1 is a Golgi acetyl-CoA antiporter that provides most of the acetyl-CoA substrate to the luminal catalytic domain of CASD1 to acetylate CMP-sialic acid.\n2) CASD1 has a catalytic transmembrane domain that enables SLC33A1-independent CMP-Sia acetylation using cytosolic pools of acetyl-CoA.\nThe first claim is well supported by the data presented. My main critique of this part of the manuscript is that it leans a little too heavily on AlphaFold2 models to interpret the consequences of disease-causing mutations on protein functon(L227-274, Fig 3d, S8), especially given that the protein has been modeled as a monomer when it is claimed to exist as a homodimer within the membrane. Have the authors considered examining these mutations in the context of the dimer AF2/3 model? Regardless, this mutation analysis is conjectural and should be substantially trimmed down.\nI am less convinced by the conclusions of the second part of the manuscript, which arguably makes the more interesting claim. The main findings in this section hinge upon data generated using two ‘virolectins’: BCoV-He0-Fc and ICVHe0-Fc. The latter is claimed to be specific for 9-O-Ac-Sia, while the former is claimed to recognize 7,9-di-O-Ac-Sia and, to a lesser extent, 9-O-Ac-Sia. The imperfect specificity of BCoV-He0-Fc makes interpreting this data difficult. It is unclear if weak signals observed with this lectin are due to weak recognition of 9-O-Ac-Sia, strong binding to a small amount of 7,9-di-O-Ac-Sia, or binding to some other uncharacterized glycoform (e.g. 7-O-Ac-Sia). An orthogonal means of detecting and quantifying the glycoforms in these cells that is independent of the virolectins is required to provide greater confidence in the authors’ interpretations of this data. MS-based glycomics seems well-suited to this task.\nFurther to this, stronger evidence is required before concluding that the CASD1 transmembrane domain has acetyltransferase activity. The claim is currently supported by partial structural homology to HGSNAT and OafB (as determined using AlphaFold2 models and without absolute conservation of active site residues), an MD simulation, and some virolectin data (which has its limitations, as discussed above). Direct detection and structural assignment of the products generated by CASD1 and its mutants (e.g. WT vs S49A vs H446A) in vitro would be far more convincing.\nRegarding the MD simulation, it is not clear to me if the Sia O-9 is in close proximity to the acetyl group of acetyl-CoA, or not. If it is, this should be illustrated more clearly. At present I don't think it is. Indeed, when looking at Figure 6f, I wondered if this site might actually be catalyzing acetyl migration from O-9 to O-7, rather than performing an acetylation with Ac-CoA. Have the authors considered the possibility that the TMD of CASD1 is actually a ‘migratase’ rather than a transferase?\nI think this is an interesting and important study and my hope is that the authors might be given the chance to further support their conclusions through the acquisition of more data using orthogonal techniques.\nSome further comments/questions regarding figures:\nFigure S5. Please quantitate and plot the degree of colocalization of these constructs over a larger number of cells. It appears to be imperfect, even for differently tagged versions of SLC33A1. Also, is B4GALT1 really in the same compartments as SLC33A1? It doesn’t look like it is to me… and if not, is it really a useful control for PLA experiment?\nFigure 3. What is the extra unassigned band in the western blot of Fig 3b?\nFigure 7. In my view, there is no strong evidence that SLC33A1 forms a complex with CASD1. Co-localisation and PLA certainly aren’t sufficient to make that claim. As such, it’d be best if they weren’t represented this was in the concluding cartoon.\nFigure 6d. WT control and H446A mutant needed.", "reviewer_3": "The manuscript by Malena Albers et al., “Interplay of SLC33A1-dependent and -independent Golgi sialic acid O-acetylation 1 in CASD1 catalysis”, describes a novel biosynthetic pathway for generation of mono and di-O-acetylated sialic acids and sialoglycoconjugates containing these molecules. The authors demonstrate that the putative AcCoA transporter SLC33A1 is essential for the synthesis of GD3 ganglioside containing a N-acetyl-neuraminic acid residues bearing 9-O-acetylation catalyzed by the sialyltransferase ST8SIA1. The authors also demonstrate that the mutant SLC33A1 variants identified in patients with two rare neurological genetic disorders are inactive or only partially active. They further show that the acetylation reaction in the Golgi lumen is catalyzed by a luminal (LCD) domain of a multi-spanning transmembrane protein CASD1 with previously unknown function. The authors further show that this reaction requires a catalytic triad with a nucleophile Ser94, however, the Ser94Ala mutation that abolishes catalytic activity at the LCD site, does not block synthesis of mono- 9-O-acetylated sialosides. The authors explain this paradox by identifying the second active site (CTMR) situated in a transmembrane channel formed by the TM domains 5,6,7 and 13 of CASD1. Using structural modeling and dynamic stimulations, they further demonstrate that the CTMR catalytic channel has a structural and a functional similarity with the catalytic channel of lysosomal N-acetyltransferase HGSNAT that catalyses transmembrane acetylation of lysosomal heparan sulfate using the cytoplasmic pool of AcCoA. Similar function and a structure has a catalytic channel of a bacterial O-acetyltransferase OafB. This reaction involves a nucleophile His residue essential for synthesis of di-7,9-O-acetylated sialosides but not of mono- 9-O-acetylated sialosides.\nOverall, the results show that transmembrane acetylation is a common physiological process in multiple species including humans and that a synthesis of O-sialylated sialosides is a complex highly regulated process. The experiments are well designed and support main conclusion of the authors. The manuscript describes a novel biological phenomenon and is of potential interest for the broad readership of the journal, however, there are several points that need to be addressed.\nFirst, most (if not all) experiments were conducted with immortalised cell lines overexpressing proteins of interests, thus it remains to be shown if the identified pathways are conserved in specific cells and tissues, especially in the neurons, where the gangliosides play essential roles. I suggest that the authors acknowledge this limitation in the end of the Discussion.\nSecond, the catalytic activities of LCD and CTMR domains need to be directly demonstrated in vitro using, for example, homogenates of CHO delta-Casd1 cells overexpressing WT and Ser/His CASD1 mutans, complemented with AcCoA and different sialosides as acceptors. Such assay is easily conducted for HGSNAT and has been essential to prove its enzymatic activity. Besides, such experiment would help to define the structural selectivity of the two active sites and whether the acetylation of the first site (presumably at 9-O position) by CTMR affects the acetylation at the O-7 position by LCD.\nFinally, the authors postulate in the Discussion and show on the scheme in Fig.7 that SLC33A1 facilitates the transport of CoA back to the cytoplasm. My impression is that this function has not been proven in the current work and remains to be a hypothesis. If so, this needs to be emphasized in the Discussion.\nI also have some specific suggestions:\nFigure 2h, PLA experiment. The images are small and do not allow to conclude in which cellular compartment the signal was detected. The signal is located in the perinuclear puncta which could be either Golgi or catabolic compartments such as lysosomes or autophagosomes. The cells need to be co-labeled for Goldi either using the same slides or separate slides co-labeled with a Golgi marker anti-Flag anti-V5 antibodies like in the figure S5\nFigure 3b. What is the identity of a ~30 kDa band on the panel? Is it related (fragment) to SLC33A1 or just an unspecifically reacting protein? This needs to be explained in the figure legend.\nFigure 4a. As in the Figure 2, Golgi localisation of proteins carrying mono and di-O-acetylated sialic acids needs to be confirmed by co-labeling with a Golgi marker. Moreover it would be interesting to compare images of permeabilized and non-permeabilized cells in panels b-d to show whether the proteins carrying 7,9-O-acetyl sialic acid are also present intracellularly.\nThe Figure 7 could be edited to include the synthesis of O-Ac-GD3 ganglioside by ST8Sia1.\nSome readers would benefit from the explanation that viral lectins cannot recognize and bind mono and di-acetylated sialic acids in gangliosides and are, thus, specific for sialylated glycoproteins."}, "author_response": "We would like to sincerely thank the reviewers for their careful reading of our manuscript and for providing thoughtful and constructive comments. We greatly appreciate the time and effort invested in reviewing our work, which has helped us improve both the clarity and quality of the manuscript. We have carefully considered each comment and made revisions to address the concerns raised. In the following, we provide detailed, point-by-point responses to all comments, highlighting the changes made in the revised manuscript where appropriate.\nPoint-by-point response:\nReviewer #1 (Remarks to the Author):\nThis manuscript by Albers et al. focuses on a membrane protein sialate O-acetyltransferase CAS1 domain-containing 1 (CASD1). The primary finding—that CASD1 possesses two active sites, one dependent on SLC33A1 (acetyl-CoA transporter) and one independent—is exciting and significantly advances our understanding of this enzyme and Golgi acetylation processes. The manuscript is well-structured, with a clear presentation of results, and this reviewer identifies no major concerns. I strongly support its publication in Nature Communications, with some minor concerns detailed below.\nWe are grateful to Reviewer 1 for the encouraging comments.\n1. In Figure 2f, only the Ex 1-5 del clone is presented. Why were the other two Ex clones omitted? Including all three would allow readers to better assess clonal variation in the data. Additionally, could the authors clarify when acetyl-CoA levels were measured and whether these measurements align with the time points in Figure 2e?\nThe acetyl-CoA levels were measured 24 hours after cell seeding, which corresponds to the first time point presented in Fig. 2e. Information on the timing is described in the Methods section and is now also provided in the legend of Fig. 2 for clarity.\nThe choice of the Ex 1-5 del clone for the acetyl-CoA measurement was guided by the chronological sequence of our study and the rationale underlying the generation of this particular clone. Upon identifying residual Sia O-acetylation in our initial ΔSlc33a1 clones, Ex1 and Ex3, we wondered whether exon skipping or the presence of an unannotated alternative upstream exon enabled the formation of functional translation products. To rigorously exclude this possibility, we generated the Ex 1-5 del clone, characterized by a large deletion encompassing most of the Slc33a1 locus. Given this, the majority of our subsequent experiments were performed with the Ex 1-5 del clone. The acetyl-CoA measurements were conducted at a later stage of our study and at that timepoint, we determined that analysing the Ex 1-5 del clone would be sufficient to exclude acetyl-CoA shortage as a potential cause of the observed loss-of-function phenotype.\n2. In Figure 3b, the anti-Flag western blot shows an impurity band near the Flag-SLC33A1-Y336* truncated protein. Could the authors clarify its identity or source? The band’s intensity appears correlated with ST8SIA1 expression.\nWe sincerely thank all three Reviewers for identifying the omitted labelling of nonspecific bands in the anti-Flag Western blot shown in Fig. 3b. We observed, indeed, two non-specific bands at approximately 30 kDa and 50 kDa, which appeared consistently across all lanes, including the empty vector control.\nIn the Results section, we added the following paragraph:\n“The anti-Flag antibody used showed two non-specific bands (marked by asterisks in Fig. 3b), with the lower band migrating close to the truncated variant p.Y366*. For clearer visualization, we analyzed additionally V5-tagged constructs, as shown in Supplementary Fig. 6.”\n3. The MD simulation details should be included in the main methods section rather than relegated to the supplementary material. Given the complexity of the simulation system, the energy minimization and equilibration steps seem brief. Furthermore, the production phase of only 100 ns is relatively short; with current computational resources, extending it to 1 µs would be feasible and likely provide more robust insights.\nWe thank Reviewer 1 for this valuable suggestion. We have now completed 2000 ns of MD calculations in triplicate. Overall, we conclude the same outcome. The new data set is shown in the revised version of Supplementary Fig. 17. A converged structure snapshot after 2000 ns MD simulation is shown in the revised version of Fig. 6d. The description of the MD simulation details have been transferred to the main methods section as suggested. The paragraph describing the MD simulation in the Results section has been revised and now reads as follows.\n“Closer examination of the CASD1 model revealed that H446 is situated at the bottom of a luminal pocket that could serve as acceptor binding site. A structure-guided approach was then used to derive an initial CTMR–CMP-Neu5Ac complex, in which we sought to test the stability and orientation preference of the acceptor substrate, using molecular dynamics (MD) simulations. The initial complex, embedded in a 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) lipid bilayer model, was used to perform three independent MD simulations that commenced from a different configuration and different initial velocity distributions (see Supplementary Figure 17 for details). The converged structure snapshot after 2000 ns MD simulation is shown in Fig. 6d. From this simulation, we observed that CMP-Neu5Ac occupies the luminal binding pocket in an orientation that positions the 9´-hydroxyl group of CMP-Neu5Ac within hydrogen bonding distance to H446 (see further detail in Supplementary Figure 17d). This simple MD model supports the notion that the CTMR of CASD1 can accommodate CMP-Neu5Ac in an orientation that allows O-acetylation via H446, possibly by activating the 9´-hydroxyl group for a nucleophilic attack on the carbonyl group of acetyl-CoA.”\nReviewer #2 (Remarks to the Author):\nThis manuscript makes some interesting observations regarding the mechanisms of sialic acid acetylation in mammals. This is an active area of research in the glycobiology field with important ramifications on our understanding of genetic disorders, cell signalling and host-pathogen interactions.\n1) SLC33A1 is a Golgi acetyl-CoA antiporter that provides most of the acetyl-CoA substrate to the luminal catalytic domain of CASD1 to acetylate CMP-sialic acid.\n2) CASD1 has a catalytic transmembrane domain that enables SLC33A1-independent CMP-Sia acetylation using cytosolic pools of acetyl-CoA.\nThe first claim is well supported by the data presented. My main critique of this part of the manuscript is that it leans a little too heavily on AlphaFold2 models to interpret the consequences of disease-causing mutations on protein functon(L227-274, Fig 3d, S8), especially given that the protein has been modeled as a monomer when it is claimed to exist as a homodimer within the membrane. Have the authors considered examining these mutations in the context of the dimer AF2/3 model? Regardless, this mutation analysis is conjectural and should be substantially trimmed down.\nDuring the revision of our manuscript, a first cryo-EM structure of human SLC33A1 in complex with acetyl-CoA has been solved (PDB ID 9M0S). The corresponding paper includes an analysis of the SPG42 mutation S113R, while other patient-derived mutations are not mentioned (Zhou et al. 2025 Cell Discov. 11:36; doi: 10.1038/s41421-025-00793-1).\nInstead of relying on AlphaFold models, we now map the mutations A110P, G509S and Y366* onto the cryo-EM structure of SLC33A1 and discuss their potential consequences on the basis of the experimental structure. With regard to the S113R mutation, we refer to Zhou et al. 2025. The original paragraph in the Results section has been replaced with a newly written version (see page 11 of the marked-up version). In the revised Fig. 3, we now show a cartoon representation of the cryo-EM structure of SLC33A1 instead of an AF2 model. The former Supplementary Fig. 8 (AF2 models of SLC33A1 variants) has been deleted. Furthermore, we have removed the former Supplementary Fig. 7, which provided evidence for the cytosolic orientation of the N- and C-termini of SLC33A1 through selective membrane permeabilization, as this information is now comprehensively established by the recently published cryo-EM structure of SLC33A1.\nWith regard to the oligomerization state of SLC33A1, the following points can be stated. Using analytical ultracentrifugation of solubilized SLC33A1, Peng and co-workers observed SLC33A1 homodimers (Peng et al. 2014 J. Neurosci. 34:6772; doi: 10.1523/JNEUROSCI.0077-14.2014), while the cryo-EM map of SLC33A1, like its AF2 model, shows a monomer. A similar discrepancy has been observed for the human dopamine transporter SLC6A3 (PDB ID 9EO4; Das et al. 2019 JBC 294:P5632; doi: 10.1074/jbc.RA118.006178). As shown by Chadda et al., transporter dimerization can be driven solely by differences in lipid solvation energetics between monomeric and dimeric states, without requiring specific protein-protein interactions (Chadda et al 2021 ELife 10:e63288; doi: 10.7554/eLife.63288). In such cases, the solubilization environment might become critical in determining whether the transporter is observed as a monomer or a dimer.\nI am less convinced by the conclusions of the second part of the manuscript, which arguably makes the more interesting claim. The main findings in this section hinge upon data generated using two ‘virolectins’: BCoV-He0-Fc and ICVHe0-Fc. The latter is claimed to be specific for 9-O-Ac-Sia, while the former is claimed to recognize 7,9-di-O-Ac-Sia and, to a lesser extent, 9-O-Ac-Sia. The imperfect specificity of BCoV-He0-Fc makes interpreting this data difficult. It is unclear if weak signals observed with this lectin are due to weak recognition of 9-O-Ac-Sia, strong binding to a small amount of 7,9-di-O-Ac-Sia, or binding to some other uncharacterized glycoform (e.g. 7-O-Ac-Sia). An orthogonal means of detecting and quantifying the glycoforms in these cells that is independent of the virolectins is required to provide greater confidence in the authors’ interpretations of this data. MS-based glycomics seems well-suited to this task.\nWe also thank the reviewer for the suggestion to employ MS-based glycomics to further characterize the O-acetylated glycoforms in our system. While we would have been eager to pursue this approach, the proposed analysis presents substantial technical challenges that extend well beyond the scope of the current study. Specifically, our work focuses on the enzymatic formation of 9- and 7,9-O-acetylated sialosides in the context of an O-glycan-specific sialyltransferase. This would necessitate the analysis of O-acetylated O-linked glycans attached to as-yet-unidentified carrier proteins—a task that demands specialized glycomics workflows not routinely available in standard laboratories.\nAfter discussing with experts in the field of MS-based glycan analytics, it became clear that such an analysis would require (i) non-standard glycomics, (ii) extensive method development to ensure sensitivity and specificity; and (iii) timely data interpretation by a specialist in glycomics. Moreover, even with these efforts, the inherent instability of O-acetyl groups poses a fundamental obstacle. These groups are highly susceptible to spontaneous loss, migration, and hydrolysis during sample handling, chromatography, and MS/MS fragmentation (Refs. 6, 56). As a result, O-acetylated species are frequently underrepresented or misassigned, and positional isomers—such as 7-O-acetyl versus 9-O-acetyl—cannot be reliably distinguished using conventional fragmentation techniques.\nThis challenge is further exacerbated by the conditions typically used in glycomics workflows. For instance, O-glycan release via base-mediated β-elimination—commonly employed in glycomics—inevitably leads to the hydrolysis of acetyl esters. Even when employing advanced fragmentation methods such as EThcD for glycopeptide analysis, only a small number of O-acetylated O-glycan structures have been confidently assigned to date (see Ref. 42, 43; Pap et al. 2020 Mol. Omics, 16:156). Notably, these structures have been identified primarily on proteins for which O-glycosylated peptides can be obtained that are particularly amenable to MS analysis—namely, those with high ionization efficiency and only a single O-glycosylation site—representing an additional constraint.\nWhile new analytical methods are emerging, they have been successfully applied primarily to well-characterized, highly O-acetylated model proteins—such as bovine submandibular mucin (Vos et al., 2023, Nat. Commun. 14:6795). This underscores the broader difficulty of analyzing O-acetylated O-glycans in complex biological systems, where these glycoforms are less abundant. Taken together, MS-based detection and structural characterization of O-acetylated sialic acids—both in general and specifically on O-glycans—remain significant and unresolved analytical challenges.\nFurther to this, stronger evidence is required before concluding that the CASD1 transmembrane domain has acetyltransferase activity. The claim is currently supported by partial structural homology to HGSNAT and OafB (as determined using AlphaFold2 models and without absolute conservation of active site residues), an MD simulation, and some virolectin data (which has its limitations, as discussed above). Direct detection and structural assignment of the products generated by CASD1 and its mutants (e.g. WT vs S49A vs H446A) in vitro would be far more convincing.\nWe thank Reviewer 2 for this insightful comment. In response, we have now provided (i) direct biochemical evidence for the acetyltransferase activity of the C-terminal transmembrane region (CTMR) of CASD1 through an in vitro assay, and (ii) additional functional validation of the catalytic transmembrane tunnel via an extensive mutational analysis of newly generated single and double mutants.\n(i) To directly assess acetyltransferase activity, we adapted a thiol-based assay to quantify the release of free CoA (HS-CoA) from acetyl-CoA during the transfer of an acetyl group to CMP-Neu5Ac. This approach was inspired by a recently published method for HGSNAT (Zhao et al., 2024, Nat Commun 15:5388) and proved robust and reliable under our experimental conditions. In contrast, an alternative HPLC-based method intended to detect the O-acetylated product, CMP-Neu5,9Ac₂, failed to yield sensitive results. Although chromatographic separation of the standards (CMP-Neu5Ac and CMP-Neu5,9Ac₂) was successful, the method lacked sufficient sensitivity to detect the low micromolar concentrations of product generated—levels that were confidently quantified using the thiol assay. Prolonging reaction times to enhance product accumulation was not feasible, as CMP-Neu5Ac is chemically labile under the assay conditions (pH 6.5, 37°C), undergoing self-degradation via a phosphate–carboxylate anhydride intermediate (Kajihara et al., Chem. Eur. J. 2011, 17:7645). Under the conditions used, a substantial amount of the input CMP-Neu5Ac (500 µM) hydrolyzed to CMP and Neu5Ac within 3 hours.\n(ii) Guided by structural superposition of our CASD1 model with the cryo-EM structure of HGSNAT, we targeted conserved residues lining the predicted transmembrane catalytic tunnel. We generated a panel of single and double alanine mutants, including combinations with the S94A mutation to inactivate the luminal catalytic domain (LCD), thereby isolating the contribution of the CTMR. As shown in the new Figure 6g, five residues within the CTMR were found to be essential for 9-O-acetylation of sialic acid—phenotypes that were only fully apparent in the absence of functional LCD activity. These findings provide strong functional support for the structural and mechanistic parallels between CASD1 and HGSNAT, and collectively reinforce the conclusion that the CTMR of CASD1 functions as a transmembrane acetyltransferase.\nTogether, these new data offer compelling biochemical and genetic evidence for the catalytic role of the CASD1 CTMR and further validate the proposed architecture of its transmembrane tunnel. The data are presented in the new Figs. 6e-i and Supplementary Figs. 18-21, and are described in the Result section (see pages 20-22 of the marked-up manuscript) as follows:\n“To directly demonstrate that the CTMR of CASD1 functions as an O-acetyltransferase, we performed an on-bead enzyme assay using detergent-solubilized, Myc-V5-tagged CASD1 immobilized on Myc nanobody-coated beads as enzyme source (Fig. 6e). After incubation with CMP-Neu5Ac and acetyl-CoA, we quantified the acetyltransferase reaction product HS-CoA using a thiol-sensitive dye, as described recently for HGSNAT48. Under the used conditions, CASD1-S94A retained over 40% of the WT activity (Fig. 6f). Since this mutant lacks a functional LCD, the remaining SOAT activity can be attributed to the CTMR. Although only the double mutant S94A;H446A—harbouring defects in both LCD and CTMR—was expected to be inactive, the single mutant CASD1-H446A likewise showed no detectable activity. Effective enzyme loading was verified by Western blot analysis, as shown exemplarily for one experiment in Fig. 6f and for all biological replicates in Supplementary Fig. 18.\nTo circumvent potential destabilizing effects of detergent solubilization, we employed cellular complementation using CHO-ΔCasd1 cells to further assess the functional impact of CASD1 mutations. The ability of CASD1 variants to restore 9-O-acetylated sialosides upon co-expression with ST8SIA6 was analysed by flow cytometry using the ICV-derived virolectin (Fig. 6g; with dot plots and expression controls shown in Supplementary Figs. 19 and 20, respectively). In the cellular setting, CASD1-S94A retained >60% of the WT complementation activity. To definitively exclude any contribution from the mutated LCD, we replaced the entire domain with a FLAG epitope. The resulting deletion mutant, CASD1-ΔLCD, exhibited complementation activity comparable to CASD1-S94A, indicating that the CTMR alone accounts for the residual activity. CASD1-H446A, like the double mutant CASD1-S94A;H446A, reached no significant activity over the mock control. Likewise, substitution of H446 with alternative residues (H446Y/F/Q/S) invariably abolished complementation activity (Supplementary Fig. 21). In HGSNAT, alanine replacement of N286, positionally equivalent to CASD1-H446 by structural superposition (see Fig. 6h), abolished enzymatic activity and caused abnormal oligomerization48, highlighting a critical role of this position in protein folding and stability. Moreover, HGSNAT-N286 plays a pivotal functional role by stabilizing the acetyl group of acetyl-CoA within the active site47, 48, 49. Based on the structural overlay, we identified CASD1-Q714 as putative functional equivalent of HGSNAT-N286, even though the two residues lie in distinct yet adjacent TMHs of the 9-TMH core (Fig. 6h). Q714 is strictly conserved across phylogenetic distant CASD1 orthologues (Supplementary Figure 16), and its replacement by alanine reduced complementation activity by ~35% (Fig. 6g). This loss increased to >90% when Q714A was combined with the LCD-inactivating S94A mutation (Fig. 6g; CASD1-S94A;Q714A), indicating a functional role of Q714 for CTMR-mediated O-acetylation.\nFurthermore, as shown in Fig. 6i, we found that the four CASD1 residues K435, R428, R489 and R496 align structurally with the four positively charged HGSNAT residues that bind the adenosine diphosphate moiety of acetyl-CoA47, 48, 49. In HGSNAT, these sites are essential (R275 and R267) or contribute significantly (R345 and K341) to enzymatic activity48. Consistent with this, alanine substitution of the corresponding CASD1 residues severely impaired complementation activity when combined with the LCD-inactivating S94A mutation (Fig. 6g). In the presence of a functional LCD, however, the loss-of-function effect of the K435A, R428A, R489A and R496A mutations was largely masked by the activity of the LCD. Notably, the four positively charged residues identified as critical for CASD1’s CTMR function are not only structurally congruent with the basic acetyl-CoA-binding residues of HGSNAT (Fig. 6i), but also with a quartet of basic residues in the OafB model (Supplementary Figure 22), further reinforcing the structural homology of the 9-TMH cores across HGSNAT, CASD1 and OafB.”\nRegarding the MD simulation, it is not clear to me if the Sia O-9 is in close proximity to the acetyl group of acetyl-CoA, or not. If it is, this should be illustrated more clearly. At present I don't think it is.\nIndeed, when looking at Figure 6f, I wondered if this site might actually be catalyzing acetyl migration from O-9 to O-7, rather than performing an acetylation with Ac-CoA. Have the authors considered the possibility that the TMD of CASD1 is actually a ‘migratase’ rather than a transferase?\nWe thank Reviewer 2 for this interesting thought. We cannot formally exclude the possibility that the transmembrane catalytic site facilitates acetyl migration from O-9 to O-7, a step that occurs at low frequency already non-enzymatically (Ref. 56). The structural similarity with HGSNAT, however, suggests that a role as transmembrane acetyltransferase remains a plausible function, in particular in light of the new data presented in Fig. 6g, as outlined in more detail above.\nI think this is an interesting and important study and my hope is that the authors might be given the chance to further support their conclusions through the acquisition of more data using orthogonal techniques.\nWe sincerely appreciate Reviewer 2’s encouraging feedback and hope that our new data satisfactorily resolve the reviewer’s concern.\nSome further comments/questions regarding figures:\nFigure S5. Please quantitate and plot the degree of colocalization of these constructs over a larger number of cells. It appears to be imperfect, even for differently tagged versions of SLC33A1. Also, is B4GALT1 really in the same compartments as SLC33A1? It doesn’t look like it is to me… and if not, is it really a useful control for PLA experiment?\nWe acknowledge that under transient overexpression conditions, certain constructs exhibited incomplete Golgi localization, accompanied by significant ER retention. To address this issue, we aimed at establishing stable co-transfectants. Despite extensive optimization of transfection conditions, antibiotic selection protocols, and clonal expansion, the co-expression of the plasmids—particularly those encoding the multi-pass transmembrane proteins SLC33A1 and CASD1—proved highly inefficient. This was reflected in the eventual undetectable expression levels of the multi-pass constructs in stable lines. In contrast, the single-pass transmembrane construct B4GALT1 was consistently amenable to stable integration and selection, yielding high-level, uniform expression in 80–100% of cells, with exclusive localization to the Golgi apparatus. These findings highlight the inherent difficulties in maintaining stable co-expression of multi-pass transmembrane proteins, due to their hydrophobic nature, complex folding requirements and low natural abundance.\nIn response to the reviewer’s comment, we acknowledge that we were unable to fully resolve the concerns regarding the PLA experiment. As a result, we have removed the PLA data from the manuscript. To address the question of subcellular localization, we have now included new data demonstrating the colocalization of transiently expressed SLC33A1 and CASD1 with a validated Golgi marker (Fig. 2h), which provides clear and consistent evidence of their shared Golgi localization. We have revised the Result section and the Discussion section accordingly (see below).\nResult section (page 10 of the marked-up manuscript):\n7\n“To examine whether the two proteins also target to the same compartment, we investigated their subcellular localization by indirect immunofluorescence analysis upon expression of V5-tagged constructs in CHO cells. As shown previously20, CASD1 colocalizes well with the Golgi marker α-mannosidase II (Fig. 2h, upper panel). Although SLC33A1 displays a more diffuse perinuclear distribution, it nonetheless shows substantial colocalization with the marker protein (Fig. 2h, lower panel), indicating that a significant fraction of the SLC33A1 pool resides in the Golgi apparatus. Our finding contrasts with the strict ER localization observed by subcellular fractionation of CHO cells25, but is consistent with immunofluorescence microscopy data demonstrating colocalization of SLC33A1 with the Golgi marker proteins giantin and syntaxin 6 in MCF-7 cells30.”\nDiscussion section (page 23 of the marked-up manuscript)\n“In contrast to the previous assumption that acetyl-CoA reaches the Golgi via vesicular transport from the ER54, the Golgi localization of SLC33A1 and CASD1 observed in the present study, consistent with findings by Huppke et al.30, suggest that acetyl-CoA is translocated directly from the cytosol to the Golgi lumen. The positioning of the acetyl-CoA transporter and the O-acetyltransferase within the same compartment ensures the effective utilization of imported acetyl-CoA by the LCD of CASD1.”\nFigure 3. What is the extra unassigned band in the western blot of Fig 3b?\nWe appreciate the reviewer’s careful inspection of Fig. 3b. This point has been addressed in the revised version of our manuscript as outlined in our response to Reviewer 1 (see point 2).\nFigure 7. In my view, there is no strong evidence that SLC33A1 forms a complex with CASD1. Co-localisation and PLA certainly aren’t sufficient to make that claim. As such, it’d be best if they weren’t represented this was in the concluding cartoon.\nWe appreciate this comment and have changed the concluding cartoon accordingly.\nFigure 6d. WT control and H446A mutant needed.\nAs outlined above, we have expanded our analysis by generating a larger panel of CASD1 mutants. In the revised manuscript, the expanded panel is shown in Fig. 6g and includes WT control and H446A mutant.\nReviewer #3 (Remarks to the Author):\nThe manuscript by Malena Albers et al., “Interplay of SLC33A1-dependent and -independent Golgi sialic acid O-acetylation 1 in CASD1 catalysis”, describes a novel biosynthetic pathway for generation of mono and di-O-acetylated sialic acids and sialoglycoconjugates containing these molecules. The authors demonstrate that the putative AcCoA transporter SLC33A1 is essential for the synthesis of GD3 ganglioside containing a N-acetyl-neuraminic acid residues bearing 9-O-acetylation catalyzed by the sialyltransferase ST8SIA1. The authors also demonstrate that the mutant SLC33A1 variants identified in patients with two rare neurological genetic disorders are inactive or only partially active. They further show that the acetylation reaction in the Golgi lumen is catalyzed by a luminal (LCD) domain of a multi-spanning transmembrane protein CASD1 with previously unknown function. The authors further show that this reaction requires a catalytic triad with a nucleophile Ser94, however, the\nOverall, the results show that transmembrane acetylation is a common physiological process in multiple species including humans and that a synthesis of O-sialylated sialosides is a complex highly regulated process. The experiments are well designed and support main conclusion of the authors. The manuscript describes a novel biological phenomenon and is of potential interest for the broad readership of the journal, however, there are several points that need to be addressed.\nFirst, most (if not all) experiments were conducted with immortalised cell lines overexpressing proteins of interests, thus it remains to be shown if the identified pathways are conserved in specific cells and tissues, especially in the neurons, where the gangliosides play essential roles. I suggest that the authors acknowledge this limitation in the end of the Discussion.\nWe thank Reviewer 3 for this suggestion and have now added the following statement to the Discussion (see page 25 of the marked-up manuscript):\n“Future work will require validation in more physiologically relevant models, such as primary cells or animal systems, as the current study relies on cell lines and protein overexpression.”\nSecond, the catalytic activities of LCD and CTMR domains need to be directly demonstrated in vitro using, for example, homogenates of CHO delta-Casd1 cells overexpressing WT and Ser/His CASD1 mutans, complemented with AcCoA and different sialosides as acceptors. Such assay is easily conducted for HGSNAT and has been essential to prove its enzymatic activity. Besides, such experiment would help to define the structural selectivity of the two active sites and whether the acetylation of the first site (presumably at 9-O position) by CTMR affects the acetylation at the O-7 position by LCD.\nThe catalytic activity of the luminal catalytic domain (LCD) has been directly demonstrated in our previous study using an in vitro assay, employing the isolated LCD as the enzyme source (Baumann et al. 2015 Nat. Commun. 6:7673). As detailed in our response to Reviewer 2, we now provide direct experimental evidence for O-acetyltransferase activity associated with the C-terminal transmembrane region (CTMR) of CASD1. In accordance with Reviewer 3’s suggestion, we performed an on-bead enzyme assay using detergent-solubilized, Myc-V5-tagged CASD1 immobilized on Myc nanobody-coated beads as the enzyme source (see schematic in new Fig. 6e). Following incubation with CMP-Neu5Ac and acetyl-CoA, the reaction product, HS-CoA, was quantified using a thiol-sensitive dye, as recently established for HGSNAT (Ref. 49). Under these conditions, CASD1-S94A, which lacks a functional LCD, retained over 40% of the WT activity (Fig. 6f). This residual activity therefore unambiguously demonstrates that the CTMR possesses intrinsic O-acetyltransferase function.\nHowever, determining the precise regioselectivity of acetylation by the two catalytic domains remains a significant analytical challenge. The O-acetyl group on sialic acid is inherently labile and prone to spontaneous migration along the glycerol side chain during sample preparation—particularly under\n9\nalkaline and acidic conditions or during derivatization steps. In our earlier work (Ref. 20), we established that the LCD transfers an acetyl group from acetyl-CoA to CMP-Neu5Ac, yielding CMP-Neu5,9Ac . This product was analyzed by DMB-HPLC, a method that involves acid-induced release of 2 Sia from CMP-Sia and subsequent DMB-derivatization of the free reducing end—processes that are known to promote acetyl group loss and migration. Importantly, because acetyl-group migration can mask the original site at which the acetyl group is introduced, this phenomenon inherently limits our ability to determine the regioselectivity of the two catalytic domains of CASD1 with complete certainty.\nTo address this limitation in our current study, we have tried to incorporate a more direct approach for the analysis of the CTMR of CASD1. [editorial note: confidential, unpublished information redacted]\nWe employed an HPLC-based assay to monitor O-acetylated CMP-Sia directly. While this method enabled adequate separation of the standards CMP-Neu5Ac and CMP-Neu5,9Ac , it lacked the 2 sensitivity required to detect the low micromolar levels of product generated under our assay conditions—levels that were readily quantifiable via the thiol-based assay for HS-CoA. Extending reaction times to increase product accumulation was not feasible, as CMP-Neu5Ac is chemically unstable under the reaction conditions (pH 6.5, 37°C), undergoing self-degradation via a phosphate–carboxylate anhydride intermediate (Kajihara et al., Chem. Eur. J. 2011, 17:7645). Under these conditions, a substantial proportion of the input CMP-Neu5Ac (500 µM) hydrolyzed to CMP and Neu5Ac within 3 hours.\nFinally, the authors postulate in the Discussion and show on the scheme in Fig.7 that SLC33A1 facilitates the transport of CoA back to the cytoplasm. My impression is that this function has not been proven in the current work and remains to be a hypothesis. If so, this needs to be emphasized in the Discussion.\nWe thank Reviewer 3 for raising this point, which helped us identify and address missing information in the manuscript. SLC33A1’s function as an antiporter has been demonstrated previously by the group of Luigi Puglielli (see Ref. 25 and 26). We have now added this information in the Introduction section (see page 5 of the marked-up version) as outlined below.\n“In its solubilized state, the transporter exists as a dimer and exhibits functional characteristics consistent with an acetyl-CoA/CoA antiporter25, 26.\nI also have some specific suggestions:\nFigure 2h, PLA experiment. The images are small and do not allow to conclude in which cellular compartment the signal was detected. The signal is located in the perinuclear puncta which could be either Golgi or catabolic compartments such as lysosomes or autophagosomes. The cells need to be co-labeled for Goldi either using the same slides or separate slides co-labeled with a Golgi marker anti-Flag anti-V5 antibodies like in the figure S5\nWe thank Reviewer 3 for this comment. As explained in our response to Reviewer 2, we have removed the PLA data from the manuscript. To address the question of subcellular localization, we have now included new data demonstrating the colocalization of transiently expressed SLC33A1 and CASD1 with a validated Golgi marker (Fig. 2h), which provides clear and consistent evidence of their shared Golgi localization.\nWe have addressed this point as explained in our response to point 2 of Reviewer 1.\nFigure 4a. As in the Figure 2, Golgi localisation of proteins carrying mono and di-O-acetylated sialic acids needs to be confirmed by co-labeling with a Golgi marker.\nWe are grateful for this suggestion, which helps us to improve our manuscript. Golgi localization of mono-and di-O-acetylated sialoglycans is now demonstrated by co-localization with the Golgi marker α-mannosidase II. The new data set is presented in the new Supplementary Figure 9. In the Results section on page 14 of the marked-up manuscript, we added the following sentence:\n“Golgi localization of the intracellular virolectin ligands was confirmed by co-staining with the Golgi marker α-mannosidase II (Supplementary Fig. 9).”\nMoreover it would be interesting to compare images of permeabilized and non-permeabilized cells in panels b-d to show whether the proteins carrying 7,9-O-acetyl sialic acid are also present intracellularly.\nFor non-transfected CHO-WT cells, we have shown the presence of intracellular 7,9-O-acetylated sialosides in Fig. 4a (see upper left image). However, since the comment of Reviewer 3 may relate to comparing permeabilized and non-permeabilized cells after ST8SIA6 expression, we extended our intracellular analysis. The new data set is shown in the new Supplementary Figure 13 and includes virolectin stains of CHO cells of all three genotypes (WT, ΔSlc33a1 and ΔCasd1) both before and after ST8SIA6 expression. The overall outcome corresponds to that of the cell surface staining shown in Figs. 4b-h, with the notable difference that intracellularly, CHO-WT cells—and to a certain extent also CHO-ΔSlc33a1 cells—exhibit virolectin ligands already prior to ST8SIA6 expression. These ligands contribute to the overall ligand profile observed after ST8SIA6 expression, thereby complicating the interpretation of intracellular analyses relative to that of cell surface analyses. Identification of the sialyltransferase that drives the formation of the intracellular O-acetylated sialosides in non-transfected CHO-WT cells requires future work and was beyond the scope of the current study.\nIn the Results section (see page 16 of the marked-up version), we have added the following sentence:\n“Virolectin stains of permeabilized mock- and ST8SIA6-transfected cells are provided in Supplementary Fig. 13.”\nMoreover, we have added the following explanatory note to the legend of Supplementary Fig. 13 for clarity:\n“Please note that intracellular virolectin ligands present before ST8SIA6 expression (mock) contribute to the total staining signal detected following ST8SIA6 expression (+ ST8SIA6).”\nThe Figure 7 could be edited to include the synthesis of O-Ac-GD3 ganglioside by ST8Sia1.\nWe thank Reviewer 3 for this suggestion. However, including an additional scheme depicting the synthesis of O-Ac-GD3 in Fig. 7 would largely duplicate the information already presented in Fig. 2g. To\n11\navoid redundancy while still facilitating comparison with Fig. 7, we have instead adapted Fig. 2g so that it now displays the luminal catalytic domain of CASD1 in the same format used for Fig. 7.\nSome readers would benefit from the explanation that viral lectins cannot recognize and bind mono and di-acetylated sialic acids in gangliosides and are, thus, specific for sialylated glycoproteins.\nAs shown by Zimmer and co-workers, influenza C virus (ICV) binds to 9-O-acetylated sialic acids on both glycoproteins and gangliosides (Zimmer et al. 1992 Eur. J. Biochem. 204:209). Using the ICV-derived virolectin, we have obtained similar results in our own experiments (unpublished), further supporting the published data.\n12"}, {"round": 2, "reviewer_comments": {"reviewer_1": "My concerns have been addressed in this revision and I do not have any further comments.", "reviewer_2": "I would like to thank the authors for their significant efforts in addressing the concerns I raised in my initial review. The revised manuscript is greatly improved and I think it makes a significant contribution to the glycobiology field.\nI have a couple of comments:\n1) It was unclear to me why disruption of Slc33a1 or the CASD1 LCD abrogates 9-O-Ac GD3 biosynthesis (as detected by TLC with an antibody to 9-O-Ac GD3) but does not disrupt 9-O-NeuAc (as detected by immunofluorescent microscopy with the virolectin ICV-HE0-185). Does this apparent discrepancy result from the different detection reagents involved (mAb vs virolectin)? Is there a reason why the TLCs are never developed using the virolectins? Using the same detection reagent in both assays could address this apparent contradiction.\n2) The CoA detected with the in vitro assay in Fig 6f could be a result of hydrolysis rather than acetyl transfer. I accept that it is difficult to quantitate the acetylated CMP-NeuNAc but it would be helpful (and feasible) to show by MS that this species is forming in an enzyme-dependent manner within the samples. This in vitro demonstration of enzyme activity would strengthen the most significant claim of the manuscript (i.e. that CASD1 is a bifunctional enzyme).", "reviewer_3": "The authors have been very responsive in addressing the reviewers comments and suggestions and the revised manuscript has been greatly improved compared to the original draft. This reviewer also appreciates the difficulty of analyzing O-acetylated O-glycans in complex biological systems due to their extreme lability, which makes some experiments suggested by the reviewers unfeasible with the currently available Glycomics methodology. However, the recently released cryo-EM structure of SLC33A1 provides now additional support for the main conclusions of the manuscript. I do not have additional comments or concerns."}, "author_response": "REVIEWER COMMENTS\nWe would like to sincerely thank the Editor and the Reviewers for their careful evaluation of our revised manuscript and for the constructive comments and suggestions. We greatly appreciate the time and effort invested in reviewing our work.\nWe are pleased that Reviewer 1 and 3 find that the revised version has satisfactorily addressed their comments. We are grateful for their positive feedback and thoughtful suggestions, which have helped us to improve the clarity and quality of the manuscript.\nWe sincerely thank Reviewer 2 for the additional comments. We have carefully considered these points and provide detailed, point-by-point responses below.\nPoint-by-point response:\nReviewer #1 (Remarks to the Author):\nMy concerns have been addressed in this revision and I do not have any further comments.\nWe thank the reviewer for confirming that the revised manuscript addresses all of their concerns.\nReviewer #2 (Remarks to the Author):\nI would like to thank the authors for their significant efforts in addressing the concerns I raised in my initial review. The revised manuscript is greatly improved and I think it makes a significant contribution to the glycobiology field.\nWe thank the reviewer for their positive assessment of the revised manuscript and for recognizing its contribution to the field.\nI have a couple of comments:\n1) It was unclear to me why disruption of Slc33a1 or the CASD1 LCD abrogates 9-O-Ac GD3 biosynthesis (as detected by TLC with an antibody to 9-O-Ac GD3) but does not disrupt 9-O-NeuAc (as detected by immunofluorescent microscopy with the virolectin ICV-HE0-185). Does this apparent discrepancy result from the different detection reagents involved (mAb vs virolectin)? Is there a reason why the TLCs are never developed using the virolectins? Using the same detection reagent in both assays could address this apparent contradiction.\nWe sincerely thank the reviewer for this important comment, which allowed us to clarify this point. Notably, the two assays employ distinct sialyltransferase, ST8SIA1 and ST8SIA6, reflecting different glycan biosynthetic pathways. We address the concern in two parts: firstly, the detection reagents; and secondly, the functional implications of the two sialyltransferases.\n1\n1. Detection reagents\nAt the outset of the study, we focused on ST8SIA1-dependent synthesis of 9-O-acetylated GD3 (9-O-Ac-GD3), for which we used the well-characterized, commercially available monoclonal antibody UM4D4, which specifically recognizes 9-O-Ac-GD3. In contrast, virolectins such as ICV-HE0-Fc are not commercially available and were developed in-house during the course of the study. Notably, ICV-HE0-Fc, which recognizes 9-O-acetylated sialic acids in both α2,3- and α2,8-linkages, was not available at the initial stage and was introduced later to detect 9-O-acetylated sialoglycoproteins produced by the O-glycan-specific ST8SIA6.\nIn response to the reviewer’s suggestion, we have repeated the experiment shown in Fig. 5d using both UM4D4 and ICV-HE0-Fc in parallel, ensuring methodological consistency. The results confirm that the luminal catalytic domain (LCD) of CASD1 is essential for ST8SIA1-dependent 9-O-Ac-GD3 synthesis, as evidenced by the loss of the 9-O-Ac-GD3 signal with the CASD1-S94A mutant. This is now shown by staining with UM4D4 and ICV-HE0-Fc (see Fig. A). Importantly, ICV-HE0-Fc detected not only 9-O-Ac-GD3 but also additional 9-O-acetylated ganglioside species (marked by asterisks), indicating broader reactivity. Given that the identity of these additional ganglioside species remains to be fully characterized, and considering that their biosynthesis may involve distinct sialyltransferases, we have chosen to present this supplementary data exclusively in the rebuttal letter. This allows us to address the reviewer’s concern while preserving the focus of the manuscript.\nFig. A: Immuno-TLC of total\ngangliosides extracted from CHO-\nΔCasd1 cells complemented with\nCASD1-WT or CASD1-S94A. CHO-\nΔCasd1 cells were transiently\ntransfected with a plasmid encoding\nST8SIA1 in combination with a\nplasmid encoding either CASD1-WT\nor CASD1-S94A. Extracted\ngangliosides were separated by TLC\nand plates were stained with anti-\nGD3 antibody R24, anti-9-O-Ac-GD3\nantibody UM4D4 or the virolectin\nICV-HE0-Fc. Bands detected by ICV0-\nHE-Fc but not UM4D4 are marked by\nasterisks\n2\n2. Sialyltransferases\nWe propose that the apparent discrepancy in the observed product profiles stem from distinct substrate preferences of ST8SIA1 and ST8SIA6. ST8SIA6 generates 9-O-acetylated and 7,9-di-O-acetylated sialosides. It may thus preferentially utilize 9-O-acetylated and 7,9-di-O-acetylated CMP-sialic acid (CMP-Sia), produced by the CTMR of CASD1 and the concerted action of LCD and CTMR, respectively. According to this model, disruption of Slc33a1 or the CASD1 LCD would abolish 7,9-di-O-acetylated sialoside formation but preserve ST8SIA6-mediated 9-O-acetylation—which is consistent with our experimental data.\nIn light of prior evidence for the cellular occurrence of 7-O-acetylated GD3 (7-O-Ac-GD3) and the previously proposed CASD1/ST8SIA1-mediated formation of 7-O-Ac-GD3 (Erdmann et al. 2006 Glycoconj. J. 23:23; Arming et al. 2011 Glycobiology 21:553), it is plausible that ST8SIA1, in contrast, utilizes 7-O-acetylated CMP-Sia—generated by the LCD of CASD1—as a substrate to synthesize 7-O-Ac-GD3. This product can undergo spontaneous ester migration to form 9-O-acetylated GD3, a non-enzymatic process that occurs efficiently even at physiological pH (Zhang et al. 2026 Angew. Chem. Int. Ed. 65:e17989). According to this model, loss of Slc33a1 or the CASD1 LCD would disrupt the initial 7-O-acetylation step, thereby abrogating the synthesis of 7-O-Ac-GD3 and, consequently, its downstream conversion to 9-O-Ac-GD3—which is consistent with the experimentally observed loss of 9-O-Ac-GD3.\nCollectively, this conceptual model—based on differential substrate utilization by ST8SIA1 and ST8SIA6, coupled with the proposed regioselective functions of CASD1’s catalytic domains—provides a coherent and mechanistically plausible explanation for the observed phenotypic differences.\nFor clarity, we have revised the corresponding paragraph in the Discussion Section (page 23 of the revised main manuscript) and included a schematic of the proposed pathway for the two-step generation of 9-O-acetylated GD3 as new Supplementary Figure 23 (see below).\nThe revised paragraph in the Discussion Section now reads as follows.\n“Our MD simulations on the CTMR model show binding of CMP-Neu5Ac in an orientation that would allow 9-O-, but not 7-O-acetylation, suggesting that the CTMR is selective for the C9 hydroxyl group, while the LCD mediates 7-O-acetylation. At first glance, this appears puzzling, since our data also demonstrated that the LCD, together with SLC33A1 and ST8SIA1, mediates the formation of 9-O-acetylated gangliosides. However, in light of prior evidence for the cellular occurrence of 7-O-acetylated GD3 (7-O-Ac-GD3)56, 57, it is plausible that ST8SIA1 uses 7-O-acetylated CMP-Sia—produced by the LCD of CASD1—as a donor substrate, thereby initially forming 7-O-Ac-GD3. This product can subsequently undergo non-enzymatic conversion to 9-O-Ac-GD3, as the 7-O-acetyl group is prone to\n3\nmigrate to C9, yielding a thermodynamically more stable primary ester bond58,59 (see Supplementary Fig. 23 for a schematic of the proposed pathway). ”\nSupplementary Figure 23.\nProposed Model for the generation of 9-O-acetylated GD3. Based on prior evidence for 7-O-acetylated GD3 (7-O-Ac-GD3)17,18, we propose a model, in which ST8SIA1 preferentially utilizes 7-O-acetylated CMP-Sia—generated by the LCD of CASD1—as a substrate, producing 7-O-Ac-GD3. Since the 7-O-acetyl group is prone to spontaneous migration to C9 of Sia, 7-O-Ac-GD3 is converted to 9-O-acetylated GD3 (9-O-Ac-GD3) over time19. This non-enzymatic process may take place within the Golgi apparatus or after transport to the plasma membrane (PM). According to this model, loss of SLC33A1 or inactivation of the CASD1 LCD through the S94A mutation disrupts the initial 7-O-acetylation step, thereby preventing the downstream production of 9-O-acetylated GD3.\n2) The CoA detected with the in vitro assay in Fig 6f could be a result of hydrolysis rather than acetyl transfer. I accept that it is difficult to quantitate the acetylated CMP-NeuNAc but it would be helpful (and feasible) to show by MS that this species is forming in an enzyme-dependent manner within the samples. This in vitro demonstration of enzyme activity would strengthen the most significant claim of the manuscript (i.e. that CASD1 is a bifunctional enzyme).\nWe sincerely appreciate the reviewer’s insightful suggestion. While we do agree that direct detection of O-acetylated CMP-Neu5Ac as a reaction product would be ideal, we believe that such a demonstration remains technically challenging under current experimental conditions, due to several interconnected factors:\n1. Low enzyme activity resulting from limited protein yield following immunoprecipitation of full-length CASD1 from detergent-solubilized lysates, which restricts the amount of active enzyme available for in vitro assays.\n2. The inherent instability of CMP-sialic acid, which undergoes rapid non-enzymatic hydrolysis to CMP and free sialic acid under the assay conditions (pH 6.5), as previously documented.\n3. The extreme lability of the O-acetyl group under standard mass spectrometric conditions, which further complicates the detection of the product, particularly given the low amounts.\nFollowing the Editor’s guidance that that demonstrating the formation of O-acetylated CMP Neu5Ac in vitro was not necessary for this revision, we have added the following statement to the Discussion Section (page23) to acknowledge it as an important direction for future work.\n“Further studies are needed to directly demonstrate the in vitro formation of O-acetylated CMP-Sia by the CTMR of CASD1 and to unambiguously define the regioselectivity of the two catalytic domains of CASD1. Because acetyl group migration can mask the initial acetylation site, real-time methods such as nuclear magnetic resonance spectroscopy may be required.”\nWe thank the reviewer for the constructive feedback and remain committed to advancing this line of research in future work.\nReviewer #3 (Remarks to the Author):\nThe authors have been very responsive in addressing the reviewers comments and suggestions and the revised manuscript has been greatly improved compared to the original draft. This reviewer also appreciates the difficulty of analyzing O-acetylated O-glycans in complex biological systems due to their extreme lability, which makes some experiments suggested by the reviewers unfeasible with the currently available Glycomics methodology. However, the recently released cryo-EM structure of SLC33A1 provides now additional support for the main conclusions of the manuscript. I do not have additional comments or concerns.\nWe thank the reviewer for their constructive comments and are pleased that the revised version satisfactorily addresses their concerns.\n5"}, {"round": 3, "reviewer_comments": {"reviewer_2": "The authors are to be commended for repeating the experiment in Fig. 5d with both UM4D4 and ICV-HE-Fc to ensure methodological consistency. This demonstrates key differences in what the detection reagents recognize. Given how important these detection reagents are in formulating the conclusions of the paper, I don't understand why the authors would want to keep it in the rebuttal only. Please add it to the SI or manuscript so the reader can appreciation the differences between these critical reagents.\nRegarding the in vitro assay, the authors state: \"We believe that a definitive in vitro demonstration would require substantial optimization—including purification of highly active recombinant CASD1, refinement of reaction conditions (e.g., pH buffering, stabilizing agents), and the use of advanced analytical platforms.\" I don't disagree with this statement but isn't the point of this work to provide a definitive demonstration of the enzyme activity rather than just suggestive data?"}, "author_response": "REVIEWERS' COMMENTS\nPoint-by-point response\nWe sincerely thank Reviewer 2 for the careful evaluation of our revised manuscript. We deeply appreciate the time and effort they devoted to reviewing our manuscript.\nReviewer #2 (Remarks to the Author)\nThe authors are to be commended for repeating the experiment in Fig. 5d with both\nUM4D4 and ICV-HE-Fc to ensure methodological consistency. This demonstrates key differences in what the detection reagents recognize. Given how important these detection reagents are in formulating the conclusions of the paper, I don't understand why the authors would want to keep it in the rebuttal only. Please add it to the SI or manuscript so the reader can appreciation the differences between these critical reagents.\nWe thank the reviewer for this helpful comment. The additional TLC experiment has now been included as new Supplementary Figure 14.\nRegarding the in vitro assay, the authors state: \"We believe that a definitive in vitro demonstration would require substantial optimization—including purification of highly active recombinant CASD1, refinement of reaction conditions (e.g., pH buffering, stabilizing agents), and the use of advanced analytical platforms.\" I don't disagree with this statement but isn't the point of this work to provide a definitive demonstration of the enzyme activity rather than just suggestive data?\nWe thank the Reviewer for this feedback. Regarding the in vitro assay, we followed the Editor’s guidance that demonstrating the formation of O-acetylated CMP Neu5Ac in vitro was not necessary for the current revision. We aim to address this point in future work."}]

Decision letter