# Structural basis for human chondroitin sulfate chain polymerization

Poushalee Dutta, Rosa Lorizolla Cordeiro, Mélanie Friedel-Arboleas, Marie Bourgeais, Sylvain Vallet, Margot Weber, Margaux Molinas, Elisabetta Boeri-Erba, Rebekka Wild

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Poushalee Dutta, Rosa Lorizolla Cordeiro, Mélanie Friedel-Arboleas, Marie Bourgeais, Sylvain Vallet, et al. Structural basis for human chondroitin sulfate chain polymerization. 2025. hal-05381182

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Structural basis for human chondroitin sulfate chain polymerization

Author information

Poushalee Dutta¹, Rosa Lorizolla Cordeiro¹, Mélanie Friedel-Arboleas¹, Marie Bourgeais¹, Sylvain D. Vallet¹, Margot Weber¹, Margaux Molinas¹, Elisabetta Boeri-Erba¹, Rebekka Wild¹,#

¹Institut de Biologie Structurale, UMR 5075, University Grenoble Alpes, CNRS, CEA, 38000 Grenoble, France.

#Correspondence: rebekka.wild@ibs.fr

Abstract

Chondroitin sulfates are complex polysaccharide chains that regulate various biological processes at
the cell surface and within the extracellular matrix. Here, we identify four heterodimeric complexes
responsible for chondroitin sulfate chain polymerization in humans: CHSY1-CHPF, CHSY1-CHPF2,
CHSY3-CHPF and CHSY3-CHPF2. Using a novel *in vitro* glycosylation assay based on chemo-
enzymatically synthesized fluorescent substrates, we demonstrate that all four complexes exhibit chain
polymerization activity. The cryo-EM structure of the CHSY3-CHPF complex provides, for the first
time, molecular insights into the chondroitin sulfate chain polymerization reaction. The architecture of
the catalytic sites suggests that CHSY1 and CHSY3 are enzymatically active, while CHPF and CHPF2
primarily play a stabilizing role. Mutational analysis of purified enzyme complexes, combined with an
*in cellulo* complementation assay, confirms that only CHSY1 and CHSY3 have bifunctional
glycosyltransferase activities. Based on the spatial arrangement of the catalytic sites, we propose that
chondroitin sulfates chain polymerization follows a non-processive, disruptive mechanism.
**Introduction**

Chondroitin sulfates (CS) are linear, negatively charged, and complex polysaccharide chains
ubiquitously expressed on the cell surface and within the extracellular matrix of almost all higher
eukaryotes. The polysaccharide backbone consists of repeating disaccharide units of N-acetyl-D-
galactosamine (GalNAc) and D-glucuronic acid (GlcA), and carries sulfate groups at various
positions. CS chains are covalently bound to specific serine residues of core proteins, together forming
chondroitin sulfate proteoglycans (CSPGs)¹,². CSPGs play a key role in regulating a broad range of
cellular processes, including cellular signaling, morphogenesis, extracellular matrix remodeling and
neuronal plasticity³⁻⁶. This extraordinary functional versatility is associated with the structural
diversity of CS chains, triggering interactions with a wide range of bio-active molecules⁷,⁸. Defects in
the CS biosynthesis machinery have been linked to osteoarthritis, inflammation, pathogen infections,
and various cancers⁹⁻¹².

Biosynthesis of CS occurs within the Golgi-lumen and is initiated by the addition of the
tetrasaccharide linker (glucuronic acid-galactose-galactose-xylose, GlcAβ1-3Galβ1-3Galβ1-
4Xyl)¹³,¹⁴. The subsequent addition of a first GalNAc moiety to the non-reducing end of the tetra-
saccharide linker is followed by alternating additions of GlcA and GalNAc moieties, forming the
chondroitin sulfate backbone¹,¹⁵⁻¹⁷, which undergoes further modifications, including 4-O and 6-O
sulfation of GalNAc and 2-O sulfation of GlcA, to complete CS biosynthesis¹⁸,¹⁹.

Four human genes – *CHSY1* (Q86X52), *CHSY3* (Q70JA7), *CHPF* (Q8IZ52) and *CHPF2* (Q9P2E5) – have been associated with CS chain polymerization. *CHSY1* and *CHSY3* (abbreviations for Chondroitin Sulfate Synthase-1 and -3) were previously reported to exhibit both *GlcA*-transferase-II and *GalNAc*-transferase-II activities²⁰,²¹. In contrast, *CHPF* (Chondroitin Polymerizing Factor), formerly known as *ChPF*²² or *CSS2* (Chondroitin sulfate synthase 2)²³, has been suggested to enhance *CHSY1* activity rather than catalyzing chain elongation by itself. Last, there is *CHPF2* (Chondroitin Polymerization Factor-2), which was previously also called *CSGlcA-T*²⁴,²⁵. Sequence analysis suggests that these four proteins, hereafter referred to as CS synthases, are type II membrane proteins with a shared topology: a short N-terminal cytoplasmic tail, followed by a single transmembrane helix, a flexible stem loop, and a large Golgi-luminal catalytic domain²⁶. Analogous to the heterodimeric heparan sulfate polymerase EXT1-EXT2²⁷,²⁸, previous studies suggested that co-expression of CS synthases – *CHSY1-CHPF*, *CHSY1-CHSY3* and *CHSY1-CHPF2* – enhances CS biosynthesis²⁵,²⁹. These findings suggest that CS synthases may also form heterodimeric complexes in the cell. The functional necessity of having four distinct CS synthases remains unclear.

In this study, we investigate the formation of heterodimeric CS polymerase complexes by combining
co-expression experiments with *in silico* analysis. Cryo-electron microscopy (cryo-EM) structure
determination reveals the architecture of the CHSY3-CHPF CS polymerase complex, which consists
of four glycosyltransferase (GT) domains. Trough extensive *in vitro* and *in cellulo* mutagenesis
studies, we provide detailed molecular insight into CS chain polymerization, unveiling the functional
role of the individual CS synthases.

**Results**

*CS synthases assemble into four distinct CS polymerase complexes:*

We set out to investigate whether CS synthases can assemble into homo- and/or heterodimeric
complexes, in order to identify the functional CS polymerase unit in human cells. Co-expression
studies were performed in human embryonic kidney Freestyle 293-F cells. Single transfections did
not result in detectable amounts of proteins, indicating that the proteins were not expressed and
secreted in monomeric or homodimeric forms (Fig. 1a). Interestingly, protein expression was
observed for four out of the six possible heterodimeric protein combinations: CHSY1-CHPF, CHSY1-
CHPF2, CHSY3-CHPF and CHSY3-CHPF2 (Fig. 1a). To further investigate the formation of
potential homo- and heterodimeric CS polymerase complexes, structural models for all possible
combinations were predicted using AlphaFold 2<sup>30,31</sup> (Fig. 1b, Supplementary Fig. 1). Consistent with
the co-expression experiments, in silico analysis suggested the formation of the heterodimeric
complexes CHSY1-CHPF, CHSY1-CHPF2, CHSY3-CHPF and CHSY3-CHPF2 with high
confidence, as indicated by low Predicted Aligned Error (PAE) values (Fig. 1b). This high confidence
in the heterodimeric complex structure was further supported by interface predicted template modeling
(ipTM) values > 0.9 and predicted local distance difference test (pLDDT) values around 89
(Supplementary Table 1). Detailed analysis of the structural models revealed tightly packed
complexes without major clashes, as well as the presence of a long coiled-coil helix that contributes to
the strong interaction between proteins. Predictions for CHSY1-CHSY3 and CHPF-CHPF complexes
resulted in models with moderate confidence, whereas models for CHSY1-CHSY1, CHSY3-CHSY3,
and CHPF2-CHPF2 had lowest confidence based on low ipTM values (< 0.4) and high distances in
PAE plots (Fig. 1b, Supplementary Fig. 1, Supplementary Table 1).

Based on our co-expression and *in silico* experiments, we propose the existence of four heterodimeric
complexes out of the six possible combinations. Interestingly, we always found either CHSY1 or
CHSY3 to interact with either CHPF or CHPF2, whereas complexes between CHSY1 and CHSY3 or
CHPF and CHPF2 were not observed.
<figure><img src="image_1.png" /><figcaption>**Fig. 1:** CS synthases can form heterodimeric complexes. (a) Transient single- and co-expression of secreted CS synthase-encoding constructs in human embryonic kidney Freestyle 293-F cells. Proteins carrying an N-terminal secretion signal, SUMO (Small Ubiquitin-like Modifier) tag and 6x His tag were detected by western blotting using an anti-His-HRP antibody. Expected molecular weights are 103 kDa for secreted CHSY1 (aa68-aa802), 100 kDa for CHPF (aa81-aa775), 102 kDa for CHSY3 (aa157-aa882) and 101 kDa for CHPF2 (aa57-aa772). (b) Structural models for the six potential heterodimeric CS polymerase complexes were predicted using AlphaFold 2<sup>30,31</sup>. Models are shown in cartoon representation with CHSY1 in green, CHPF in grey, CHSY3 in orange, and CHPF2 in light blue. The N-terminal anchoring helices and flexible stem regions, that were omitted during model prediction, were drawn by hand. The corresponding predicted align error matrices are shown below.</figcaption></figure>
Purified CS polymerase complexes show CS chain elongation activity

To further investigate the importance of heterodimer formation between CS synthases and to study
their differential activities, we purified the four different complexes. We routinely obtained 300 µg to
1 mg of protein complexes from 600 mL Freestyle 293-F suspension culture at satisfying purity, as
confirmed by SDS-PAGE analysis (Fig. 2a). To determine the stability of complexes, thermal
unfolding was analyzed using nano-differential scanning fluorimetry (nanoDSF). Melting
temperatures varied between 43°C to 48°C, with highest stability observed for CHSY3-CHPF and
lowest for the CHPF2-containing complexes CHSY1-CHPF2 and CHSY3-CHPF2 (Fig. 2b,
Supplementary Fig. 2a). Next, we performed mass photometry experiments to analyze the oligomeric
state of purified complexes in solution. The predominant peaks were found between 166 and 178 kDa,
suggesting that the majority of complexes were present in a dimeric form (Fig. 2c, Supplementary
Fig. 2b). In order to test if purified CS complexes were functional, we developed a novel Fluorophore-
Assisted Carbohydrate Gel Electrophoresis (FACE) assay to efficiently follow CS chain
polymerization in vitro. Expanding our previous chemo-enzymatic synthesis approach¹⁴, we generated
custom-tailored fluorescent glycopeptides, which served as acceptor substrates for the CS polymerase
complex. The selected peptide sequence EEASGEAS was derived from the chondroitin sulfate
proteoglycan Colony Stimulating Factor 1 (CSF1)³², and was chemically synthesized harboring a N-
terminal TAMRA fluorophore. Using recombinantly expressed and purified enzymes carrying out
glycosaminoglycan biosynthesis, a pentasaccharide linker (GalNAc-GlcA-Gal-Gal-Xyl) was
covalently added onto the CSF1 peptide (Supplementary Fig. 3a-c). The resulting Penta-CSF1
peptide was characterized by mass spectrometry (MS)(Supplementary Fig. 4). All four purified CS
polymerase complexes were able to elongate the CS chain, using Penta-CSF1 as acceptor substrate and
UDP-GlcA and UDP-GalNAc as donor substrates, with the CHSY3-CHPF2 complex generating
slightly shorter CS chains (Fig. 2d, e).
<figure><img src="image_2.png" /></figure>

<figure><img src="image_3.png" /></figure>

<figure><img src="image_4.png" /></figure>

<figure><img src="image_5.png" /></figure>

<figure><img src="image_6.png" /><figcaption>**Fig. 2: Purified CS polymerase complexes are well-folded and catalytic active**</figcaption></figure>

(a) Coomassie-stained SDS-PAGE analysis of purified CS polymerase complexes. The expected molecular weight of 3C protease-cleaved proteins are 88 kDa for CHSY1 (aa68-aa802), 86 kDa for CHSY3 (aa157-aa882), 79 kDa for CHPF (aa81-aa775), and 81 kDa CHPF2 (aa57-aa772). (b) The melting temperature (T<sub>m</sub>) of CS polymerase complexes was determined using nano-differential scanning fluorimetry (nanoDSF). Experiments were performed as technical duplicates (**see Supplementary Fig. 2**). (c) Mass photometry analysis of purified CS polymerase complexes. The calculated molecular masses are indicated in the graph. Experiments were performed as technical duplicates (**see Supplementary Fig. 2**). (d) Schematic illustration of *in vitro* glycosylation assay used to monitor chain elongation by the CS polymerase complexes. The fluorescently labelled Penta-CSF1 peptide, derived from the CSPG colony-stimulating factor 1, served as the acceptor substrate, while UDP-GlcA and UDP-GalNAc were used as donor substrates. Monosaccharide symbols follow the SNFG (Symbol Nomenclature for Glycans) system³³. (e) Catalytic activity of purified CS polymerase complexes was studied using the *in vitro* glycosylation assay described in (d). Reaction products were separated by polyacrylamide gel electrophoresis, and fluorescent peptides were visualized using a fluorescence imager.
Structural insight into CS chain elongation

After demonstrating that purified CS polymerase complexes were well-folded and functional, we sought to determine the 3D structure of a CS polymerase complex using single-particle cryo-electron microscopy (cryo-EM). The CHSY3-CHPF complex was selected for structural and functional studies due to highest thermal stability and *in vitro* chain polymerization activity in our setup. An octasaccharide, (GalNAc-GlcA)₄, representing an acceptor substrate for the GalNAc-T reaction and the product of the GlcA-T reaction, was added to the sample, along with the donor substrate analog UDP prior to EM grid preparation. Cryo-EM analysis provided a 3D reconstruction at approximately 3.0 Å resolution, revealing the overall architecture of the CS polymerase complex with its four GT domains (Fig. 3a, b, Supplementary Fig. 5 and 6, Supplementary Table 2). The experimental structure closely aligns with the AlphaFold 2-predicted model, showing a root-mean square deviation (RMSD) of 1.080 Å across 522 pruned atom pairs (Cα atoms). The C-terminal regions of both proteins were not visible in the cryo-EM map, probably due to a high conformational flexibility (Supplementary Fig. 7). The analysis revealed that CHSY3 and CHPF form a tightly packed complex, with their respective N-terminal and C-terminal GT domains interacting strongly, resulting in a total interaction surface of 6,711 Å², as determined using PDBePSIA³⁴. Both proteins also harbour a middle domain comprising a beta-sheet fold and a long alpha helix. The alpha helix forms a coiled-coil bundle, contributing to the strong interaction between the two proteins. Furthermore, electron density was observed for a UDP molecule and an Mn²⁺ ion in the active site of the N-terminal GT domain of CHSY3, as well as for an Mn²⁺ ion in the C-terminal GT domain of CHSY3. Additionally, electron densities corresponding to three N-glycans were identified on the asparagine residues N279 and N710 in CHSY3 and N138 in CHPF.

<figure><img src="image_7.png" /><figcaption>Fig. 3: Structure of the human CS
polymerase complex CHSY3-
CHPF.</figcaption></figure>

<figure><img src="image_8.png" /><figcaption>b</figcaption></figure>

(a) Cryo-EM map of the heterodimeric CHSY3-CHPF complex. Density covering CHSY3 is shown in orange and CHPF in grey. A UDP molecule is highlighted in blue, and N-glycans are depicted in dark orange. Both proteins contain an N-terminal GT domain with putative D-glucuronic acid transferase (GlcA-T) activity and a C-terminal GT domain with putative N-acetyl-D-galactosamine transferase (GalNAc-T) activity. The complex is shown from three rotated views. (b) Structural model of the complex,

shown in cartoon representation with the same color scheme as in (a). UDP and N-glycans are shown in stick representation. Close-up view shows EM density around UDP and Mn²⁺ ion at a contour level of 0.125.
To gain insights into the catalytic mechanism of CS chain polymerization, we analyzed the different
GT domains in more detail. The N-terminal GT domain of CHSY3 (residues L173-P395) belongs to
the GT31 family, which, according to the Carbohydrate-Active Enzymes (CAZy) database, assembles
diverse inverting β-1,3-glycosyltransferases³⁵. In its active site, we observed clear EM density for a
UDP molecule and a Mn²⁺ ion (Fig. 4a), revealing residues critical for catalyzing CS chain elongation.
These include a DxD motif (D261 and D263), which is commonly found in many GT enzymes³⁶,³⁷.
Additional residues contributing to donor substrate binding in CHSY3 include M178, R187, D231,
Y233, K238, D262 and H394. Importantly, these residues are also conserved in CHSY1, while most of
them are absent in CHPF and CHPF2 (Fig. 4a). This observation suggests that CHPF and CHPF2 are
unlikely to bind the UDP donor substrate and, consequently, are not expected to catalyze glycan
transfer.

The C-terminal domain of CHSY3 (residues K620-S882) belongs to the GT7 family in the CAZy
database, which groups inverting 4-β-N-acetylgalactosaminyltransferases with a GT-A fold³⁵. To
identify residues important for donor substrate binding, we performed a search for structurally related
proteins using the protein structure comparison server DALI³⁸. The chondroitin polymerase from
*Escherichia coli* strain K4 (PDB-ID: 2Z87)³⁹ was identified as a close structural homolog. By
superimposing its crystal structure, which had been determined in complex with a UDP-GalNAc donor
substrate and a Mn²⁺ ion in its active site, we identified key residues in the C-terminal GT domain of
CHSY3, including D718 and D720, which form the DxD motif, as well as P626, R630, R697, E806,
D807, H831 and H834 (Fig. 4b, Supplementary Fig. 8). As with the N-terminal GT domain, all the
residues important for donor substrate binding and activation were conserved in CHSY3 but not in
CHPF and CHPF2 (Fig. 4b).

*In vitro* characterization of CS chain polymerization

To investigate the catalytic roles of the two putative active sites in CHSY3, we constructed structure-
based point mutants targeting these regions. The constructs CHSY3<sup>D261N/D263N</sup> and CHSY3<sup>H394A</sup> carry
mutations in the N-terminal GT domain, while CHSY3<sup>D718N/D720N</sup> and CHSY3<sup>H831A</sup> have amino acid
substitutions in the C-terminal GT domain. Since CHPF does not harbour a DxD motif, it was kept in
its wild-type form for analyzing CS polymerization activity of the CHSY3-CHPF complex. All four
CHSY3<sup>mutant</sup>-CHPF complexes were successfully purified (**Supplementary Fig. 9a**), and their stability
and integrity were confirmed through nanoDSF and mass photometry analysis (**Supplementary Fig.
9b-d**). The ability of these CHSY3<sup>mutant</sup>-CHPF complexes to perform chain elongation was assessed
using our newly developed *in vitro* glycosylation assay.
<figure><img src="image_9.png" /><figcaption>Fig. 4: Identification of catalytic residues in the N- and C-terminal GT domains of CS synthases.</figcaption></figure>

(a) Close-up view into the putative catalytic sites of the N-terminal GT domains of CHSY3, CHSY1, CHPF and CHPF2. The structures are shown in cartoon representation and colored in orange, green, grey and blue, respectively. The UDP ligand in CHSY3 is shown as blue sticks and the Mn²⁺ ion as purple sphere. Residues in CHSY3 important for UDP binding are highlighted in stick representation. The putative active sites in CHSY1, CHPF and CHPF2 are shown from the same perspective as CHSY3. Structures for CHSY3 and CHPF were determined by cryo-EM, whereas models for CHSY1 and CHPF2 were predicted using AlphaFold²³⁰,³¹. (b) Zoom-in into the putative catalytic site of the C-terminal GT domains of CHSY1, CHSY3, CHPF and CHPF2. In order to identify residues important for donor substrate recognition, the active site of a close structural homolog, the chondroitin polymerase from *Escherichia coli* strain K4, is shown alongside (PDB-ID: 2Z87)³⁹. Its crystal structure, shown in red cartoon representation, was solved in complex with UDP-GalNAc, shown in light blue
sticks. Important residues are highlighted in stick representation. Structures for CHSY3 and CHPF were determined by cryo-EM, whereas models for CHSY1 and CHPF2 were predicted using AlphaFold 2. (c) Activity of wild-type and mutant CHSY3-containing CHSY3-CHPF complexes was analyzed by performing *in vitro* glycosylation reactions, followed by FACE analysis. The pentasaccharide peptide Penta-CSF1 was used as acceptor substrate and UDP-GlcA and UDP-GalNAc as donor substrates. Monosaccharide symbols follow the SNFG (Symbol Nomenclature for Glycans) system³³. (d) *In vitro* glycosylation assays of wild-type and mutant CHSY3-containing CHSY3-CHPF complexes using the hexasaccharide peptide Hexa-CSF1 as acceptor substrate and UDP-GlcA and UDP-GalNAc as donor substrates. (e) *In vitro* glycosylation assays to test the ability of different metal ion species to support GalNAc and GlcA transferase reactions.

For this assay, the fluorescent Penta-CSF1 peptide, which carries a pentasaccharide linker (GalNAc-GlcA-Gal-Gal-Xyl), was used to monitor extension with a GlcA unit. Similarly, the Hexa-CSF1 peptide, carrying a hexasaccharide linker (GlcA-GalNAc-GlcA-Gal-Gal-Xyl), was used to follow extension with a GalNAc moiety (Supplementary Figs. 3 and 10).

Upon incubation of the acceptor substrates with the donor substrates UDP-GalNAc and UDP-GlcA in the presence of either wild-type or mutant CHSY3-containing CHSY3-CHPF complexes, the reaction products were separated by gel electrophoresis and visualized by fluorescence imaging. The wild-type CHSY3-CHPF complex efficiently elongated the Penta-CSF1 and Hexa-CSF1 substrates into high molecular weight chains (Fig. 4c,d). In contrast, the CHSY3<sup>D261N/D263N</sup>-CHPF and CHSY3<sup>H394A</sup>-CHPF complexes, which carry mutations in the N-terminal GT domain, were unable to catalyze GlcA transfer (Fig. 4c). Interestingly, incubation of the C-terminal mutants, CHSY3<sup>D718N/D720N</sup>-CHPF and CHSY3<sup>H394A</sup>-CHPF, with both UDP-GlcA and UDP-GalNAc donor substrates resulted into a distinct band shift, indicating the addition of a single GlcA unit to the Penta-CSF1 substrate. This increased migration speed of Hexa-CSF1 compared to Penta-CSF1 reflects the higher negative charge of the elongated product (Fig. 4c). The C-terminal mutants failed to catalyze further chain polymerization, suggesting an inability to transfer GalNAc residues (Fig. 4c). These findings were further confirmed using Hexa-CSF1 as the acceptor substrate. The N-terminal GT mutants CHSY3<sup>D261N/D263N</sup>-CHPF and CHSY3<sup>H394A</sup>-CHPF catalyzed the transfer of a GalNAc residue, producing the Hepta-CSF1 peptide (Fig. 4d). However, no additional chain elongation was observed for CHSY3<sup>D261N/D263N</sup>-CHPF, validating its impairment in GlcA transfer. In contrast, faint additional bands detected for the CHSY3<sup>H394A</sup>-CHPF complex suggest residual, albeit low, GlcA-T activity. The C-terminal mutants CHSY3<sup>D718N/D720N</sup>-CHPF and CHSY3<sup>H394A</sup>-CHPF failed to elongate the Hexa-CSF1 peptide (Fig. 4d). In summary, the *in vitro* chain elongation assays revealed that the N-terminal GT domain exhibits GlcA-T activity, while the C-terminal GT domain exhibits GalNAc-T activity. Importantly, although CHPF was maintained in its wild-type form, the mutant CHSY3-CHPF complexes were unable to catalyze chain elongation, suggesting that CHPF itself cannot perform either GlcA or GalNAc transfer.
Analysis of the CHSY3<sup>mutant</sup>-CHPF complexes in the presence and absence of various metal ions revealed that both GlcA-T and GalNAc-T are metal ion dependent. The highest catalytic activity was observed with Mn<sup>2+</sup>, while an intermediate activity was observed with Mg<sup>2+</sup> and Ca<sup>2+</sup> (Fig. 4e).

To confirm that point mutations in the GlcA-T or GalNAc-T catalytic sites did not affect overall functionality of the CHSY3-CHPF complex, a rescue experiment was designed. The CHSY3<sup>D261N/D263N</sup>-CHPF and CHSY3<sup>D718N/D720N</sup>-CHPF mutants were mixed in equimolar amounts and incubated with Penta-CSF1 peptide, UDP-GalNAc, and UDP-GlcA donor substrates. The reactions were carried out, and the resulting chain polymerization was comparable to that observed for wild-type CHSY3-CHPF complexes (Supplementary Fig. 11). This suggests that the mutations in CHSY3 did not alter the overall stability of the complex. Furthermore, the fact that mixing the GlcA-T and GalNAc-T deficient mutant complexes restored chain polymerization to wild-type levels supports the hypothesis that CS chain elongation follows a disruptive rather than a polymerization mechanism, where the polysaccharide chain detaches from the complex before entering the next catalytic cycle.

## Mutations in CHSY1 and CHSY3 impair cellular CS production

Next, we established an *in cellulo* complementation assay to study the effect of structure-based point mutations in CHSY1 and CHSY3 on cellular CS biosynthesis (Fig. 5a). A human embryonic kidney (HEK293-6E) cell line lacking both CHSY1 and CHSY3 was chosen as test system<sup>40</sup>. We co-transfected these CHSY1/CHSY3 knockout (KO) cells with plasmids encoding either full-length CHSY1 and CHPF or full-length CHSY3 and CHPF. The cell surface CS content was quantified by flow cytometry using a primary anti-CS antibody and an Alexa Fluor 488-labelled secondary antibody. To detect transfection efficiency, cells were permeabilized and labelled with a primary anti-FLAG antibody (all constructions contained an N-terminal FLAG-tag) and a cyanine3 secondary antibody. Approximately 40%-50% of the cells were successfully transfected (Supplementary Fig. 12).

While CS biosynthesis was almost abolished in the CHSY1/CHSY3 KO cell line, CS levels were restored to around 80% and 40% of wild-type cells 36 hours post-transfection with CHSY1 wild-type and CHSY3 wild-type encoding plasmids, respectively (Fig. 5b, c). In parallel, we transfected plasmids carrying mutations in the GlcA-T domain (CHSY1<sup>D171N/D173N</sup>, CHSY1<sup>H304A</sup>, CHSY3<sup>D261N/D263N</sup>, CHSY3<sup>H394A</sup>) or the GalNAc-T domain (CHSY1<sup>D631N/D633N</sup>, CHSY1<sup>H744A</sup>, CHSY3<sup>D718N/D720N</sup>, CHSY3<sup>H831A</sup>), while keeping CHPF in its wild-type form. All mutant complexes failed to restore CS levels (Fig. 5b, c). These results suggest that both GlcA-T and GalNAc-T activities are essential for promoting cellular CS biosynthesis. They also demonstrate that both CHSY1-CHPF and CHSY3-CHPF CS polymerase complexes can perform CS chain elongation *in cellulo*.
<figure><img src="image_10.png" /></figure>

Results were compared to wild-type HEK 293-E cells. (c) Complementation assay as described in (b), but with CHSY3-CHPF (orange bars). Experiments were performed in triplicates (n=3). Error bars indicate standard deviation. A one-sided paired student's t-test was used to determine *p*-values (*<0.05 and **<0.01). Source data is provided.

<caption>**Fig. 5: Studying the effects of mutations in the CS polymerase complex on cellular CS biosynthesis.**</caption>

(a) Schematic illustration of the *in cellulo* complementation assay. Knock-out (KO) of the CHSY1 and CHSY3 genes via CRISPR/Cas eliminates the production of CS chains. KO cells are then co-transfected with either CHSY1 and CHPF or CHSY3 and CHPF encoding plasmids and cell surface CS levels quantified. (b) Complementation of CHSY1/CHSY3 KO cells with CHPF wild-type and CHSY1 wild-type or mutant constructs. Cell surface CS levels in non-transfected cells (blue bars) and co-transfected cells (green bars) were quantified by flow cytometry using a primary anti-CS antibody and an Alexa Fluor 488-labelled secondary antibody.

# Discussion

## *Heterodimeric CS polymerase complexes*

Chain polymerization is a key step in CS biosynthesis, and four genes - CHSY1, CHSY3, CHPF and CHPF2 - have been shown to be involved in CS chain elongation. However, their individual contributions and relative importance remained unclear. In this study, we demonstrate that human CS synthases form heterodimeric complexes. Through co-expression experiments in human embryonic kidney cells, we observed the formation of four out of the six possible heterodimeric CS polymerase complex combinations: CHSY1-CHPF, CHSY1-CHPF2, CHSY3-CHPF and CHSY3-CHPF2 (Fig. 1a). Interestingly, we always found one of the chondroitin sulfate synthases (CHSY1 or CHSY3) to interact with one of the chondroitin sulfate polymerization factors (CHPF or CHPF2), while neither the chondroitin sulfate synthases nor the chondroitin sulfate polymerization factors appeared to form complexes among themselves. Of note, CHSY1 and CHSY3 share a high sequence identity (68.14%), as do CHPF and CHPF2 (59.15%), whereas CHSY1 exhibits only 23.73% sequence identity with CHPF and 24.61% with CHPF2 (Supplementary Fig. 13a). We did not detect the expression of single
CS synthases or the formation of homodimeric complexes (Fig. 1a). In silico analysis of potential CS
polymerase complexes using AlphaFold 2 further supported the formation of the same four complexes
(Fig. 1b, Supplementary Fig. 1). Additionally, cryo-EM studies of the CHSY3-CHPF complex
revealed that the two proteins form a tightly-packed complex with large interaction surfaces.
Importantly, no monomers were observed during cryo-EM data processing. Mass photometry analysis
of the purified CS polymerase complexes further confirmed that these are rather stable and do not
dissociate (Fig. 2c). Together, our results suggest that heterodimeric CS polymerase complexes
represent the predominant and functional form responsible for CS chain polymerization in human
cells. A similar observation has been made for the human heparan sulfate polymerase EXT1-EXT2<sup>27,28</sup>.
Golgi-localized glycosyltransferases frequently form homo- or hetero-oligomeric complexes, which
enhances their catalytic activity, stability and regulates their cellular localization<sup>41,42</sup>.

Previous studies hypothesized that CS synthases form heterodimeric complexes<sup>25,29</sup>. However, based on pull-down experiments with secreted proteins in the cell medium, Izumikawa and co-workers proposed that any of these proteins could interact to form such complexes<sup>25</sup>. In contrast, our co-expression studies and *in silico* analysis indicate that CHSY1-CHPF, CHSY1-CHPF2, CHSY3-CHPF, and CHSY3-CHPF2 are the four primary CS polymerase complexes, showing the highest co-expression and strongest likelihood in AlphaFold2 predictions. We don't observe any evidence for the existence of the heterodimeric complexes CHSY1-CHSY3 or CHPF-CHPF2. This conclusion is further supported by *in vitro* and *in cellulo* assays, which indicate that only CHSY1 and CHSY3 possess catalytic activity, whereas CHPF and CHPF2 have a structural role that are required for enzymatic activity of CHSY1 and CHSY3 (Fig. 4).

Mechanistic insights into CS chain polymerization

In order to learn more about CS chain polymerization at the molecular level, we determined the
structure of the CHSY3-CHPF complex using cryo-EM. Both CHSY3 and CHPF consist of two GT
domains, bridged by a middle domain, and together form a stable heterodimeric complex. The
AlphaFold 2-predicted model fitted well into the experimental EM map. Given the high structural
similarity between the four CS polymerase complex models, the AlphaFold 2 predictions are likely to
be correct and can thus serve as a valuable tool for future structure-based studies.

UDP, the reaction by-product of the GlcA and GalNAc transferase reaction, was added during cryo-EM sample preparation. We identified one UDP molecule bound in the CHSY3 GlcA-T active site. This finding revealed key residues involved in the GlcA-T reaction. In addition, we inferred important amino acid residues for the GalNAc-T reaction through structural comparisons with the *E. coli* K4 chondroitin polymerase (Fig. 4a, b). The GlcA-T domain of CHSY1 and CHSY3 belongs to the CAZy GT31 family, while their GalNAc-T domain is part of the GT7 family. Both, GT7 and GT31 families
generate a β-glycosydic linkage using an inverting mechanism. Based on structural comparison with previously characterized homologs, the residues D252 and D720 in CHSY1, and D342 and D807 in CHSY3, were identified as catalytic base residues that start the reaction cycle⁴³,⁴⁴. A significant C-terminal portion, 48 residues in CHSY3 and 45 residues in CHPF are not visible in the cryo-EM structure, likely due to a high flexibility (Supplementary Fig. 7). The Alphafold 2 model suggests that these regions form α-helices, which are located in close proximity to the active site. The C-termini might thus contribute to acceptor substrate binding, as observed for the GT7 homolog B4GalT7⁴⁴. On the other hand, these helices, when stabilized near the active site, could also block substrate access, thus preventing the glycosyltransferase reaction. Future structural studies on the CS polymerase complex, bound to acceptor substrates of varying length, are needed to characterize acceptor substrate binding in molecular detail. These studies could also reveal an involvement of the flexible C-terminal helices in the transfer reaction.

<figure><img src="image_11.png" /><figcaption>**Fig. 6 Molecular mechanism of CS chain polymerization.**</figcaption></figure>

Illustration of the proposed mechanism for chain elongation by the CS polymerase complex CHSY3-CHPF. In a first reaction step, a GlcA residue is added onto the proteoglycan carrying a pentasaccharide-linker by the N-terminal GT domain of CHSY3. The generated hexasaccharide product is released and binds to the C-terminal GT domain of CHSY3 exhibiting GalNAc transferase activity. Upon GalNAc addition, the proteoglycan dissociates and a new reaction cycle starts. Repetitive addition of GlcA and GalNAc residues leads to the generation of the long chondroitin sulfate backbone.

Based on the distance between the active sites, we propose a disruptive rather than a polymerization-based reaction mechanism (Fig. 6). The same observation was made for the bacterial chondroitin polymerase, whose two active sites are located on opposite sides, around 60 Å apart³⁹. However, in contrast to human CS biosynthesis, this bacterial chondroitin polymerase functions as a monomer. This raises the question: Why have vertebrates evolved a much more sophisticated CS polymerization machinery that relies on four distinct CS synthases? One possible explanation could be that this redundancy provides a protective mechanism against deleterious mutations. Additionally, it seems plausible that early in evolution, all four CS synthases originally possessed catalytic activity. In such a
scenario, the formation of heterodimeric complexes would increase the local concentration of
enzymes, facilitating chain elongation by reducing the substrate's travel path, enhancing the efficiency
of CS biosynthesis.

However, our *in vitro* and *in cellulo* functional analysis indicate that only CHSY1 and CHSY3 are
catalytically active. This conclusion is further supported by the architecture of the catalytic sites. Key
residues in the N-terminal domain (D261, D263 and H394 in CHSY3), which are critical for GlcA
transfer, and in the C-terminal domain (D718, D720 and H831 in CHSY3), which are essential for
GalNAc transfer, are highly conserved in CHSY1 and CHSY3 but are absent in CHPF and CHPF2
(Supplementary Fig. 13b)⁴⁵. We speculate that only human CHSY1 and CHSY3 have retained their
catalytic activities, while CHPF and CHPF2 gradually lost their enzymatic function due to
accumulated mutations over time. Nevertheless, CHPF and CHPF2 remain essential for complex
formation and activity, likely due to their role in stabilizing the overall architecture of the CS
polymerase complexes. Of note, the amino acid substitution D633N in CHSY1, previously identified
in a patient with temtamy preaxial brachydactyly syndrome⁴⁶, was found to be critical for CS
polymerization in our *in cellulo* studies. This residue, part of the DxD motif in the GalNAc-T domain,
is essential for CS biosynthesis. This comparison underlines the relevance of the established *in cellulo*
assay for studying the effects of specific mutations. On the other hand, to this date, only a few patient
mutations have been identified in CHSY1/CHSY3⁴⁶,⁴⁷ compared to the heparan sulfate polymerase
EXT1/EXT2²⁷. This is unlikely due to a lower importance of CS compared to HS, but rather to the
redundancy within the system, as CHSY1 can compensate for loss of CHSY3, and vice versa.

Our *in vitro* chain elongation studies suggest that all four purified CS polymerase complexes -
CHSY1-CHPF, CHSY1-CHPF2, CHSY3-CHPF and CHSY3-CHPF2 - are capable of carrying out CS
chain polymerization (Fig. 2a). One possible reason for the existence of multiple CS polymerase
complexes could be tissue-specific expression of the CS genes²⁵,⁴⁸. Another reason could be that
different CS polymerase complexes produce CS chains of varying lengths, as previously suggested by
Ogawa and co-workers⁴⁹. At present, we cannot conclude whether the slight differences in CS
polymerization activity observed among the complexes have biological relevance, as these variations
could also be caused by differences in the thermal stability of the complexes or varying purification
conditions (Fig. 2b,e). To further investigate potential differences in CS polymerase complex activity,
additional *in vitro* kinetic studies using purified complexes, along with novel methodologies for
precise chain-length determination, will be necessary. The presence of four different CS polymerase
complexes in the cell could also allow for differential regulation, thereby fine-tuning chain length and
sulfation motifs. Furthermore, these complexes may interact differently with downstream biosynthetic
enzymes. For example, chondroitin 4-O-sulfotransferase-1, which introduces a sulfate group to
position 4 of GalNAc residues, has been shown to regulate CS chain length⁵⁰. The function of the
middle domain remains elusive, but it may play a role in promoting protein-protein interactions with other cellular components, such as CS biosynthesis enzymes or the CSPG substrate. These questions warrant further investigations.

# Methods

## Structure prediction using AlphaFold 2

To study the formation of homo- and heterodimeric CS polymerase complexes, we performed model predictions for all combinations of the four human CS synthases using AlphaFold 2³⁰. The protein sequences were retrieved from UniProt²⁶ and trimmed to remove the membrane spanning helix. The resulting constructs - CHSY1 (Q86X52, aa68-aa802), CHPF (Q8IZ52, aa81-aa775), CHSY3 (Q70JA7, aa157-aa882) and CHPF2 (Q9P2E5, aa57-aa772) - were used as input sequences in ColabFold, which is based on AlphaFold 2 using the options num_models = 5 and num_cycle = 3⁵¹⁻⁵³,³¹. The best ranked model was used for subsequent analysis in ChimeraX⁵⁴.

## Co-expression study

To test soluble and secreted expression of human homo- and heterodimeric CS polymerase complexes, the coding regions of N-terminal truncated CHSY1 (aa68-aa802), CHPF (aa81-aa775), CHSY3 (aa157-aa882) and CHPF2 (aa57-aa772) were inserted into the pHR-CMV-TetO2 vector⁵⁵, utilizing the restriction sites AgeI and KpnI. The resulting constructs contained an N-terminal chicken RPTP sigma signal peptide, followed by SUMO fusion protein, 6x His tag and a 3C protease cleavage site. CHSY1 and CHSY3 feature an Avi tag (biotin acceptor peptide) at their C-terminus, whereas CHPF and CHPF2 contained another 3C site followed by a twin streptavidin tag. 20 mL of human Freestyle 293-F cells (Thermo Fisher, R79007) were transfected or co-transfected using polyethylenimine (PEI25K, Polysciences, 23966-100) at a PEI to DNA mass ratio of 2:1 (single-transfection) or 2:0.5:0.5 (co-transfections). The cells were then incubated in a shaking incubator at 37°C with 8% CO₂ for 96 h. Supernatants were collected by centrifugation at 500 xg for 5 min at room temperature. Transient expression was analyzed by western blot analysis using an anti-His HRP antibody (Sigma, A7058) at a dilution of 1:2000. Chemiluminescent signal was detected upon addition of Chemiluminescent HRP substrate (Sigma, A7058) on a ChemiDoc™ MP imaging system (Bio-Rad).

## Purification of CS polymerase complexes

600 mL of Freestyle HEK293-F cells (2 x 10⁶ cells/mL) were co-transfected with the constructs and conditions described above. The cell supernatant was recovered by two consecutive centrifugation
steps at 500 xg for 5 min and 6000 xg for 15 min, respectively. The supernatant was filtered through a
0.45 µM sterile filter (Merck), and buffer was added to a final concentration of 50 mM Tris pH 7.8,
150 mM NaCl, 10 mM imidazole, and 10 mM MgCl₂. All subsequent steps were performed at 4°C.
The supernatants were loaded onto a 5 mL HisTrap™ HP column (Cytiva). The column was washed
with 5 column volumes of buffer A (50 mM Tris pH 7.8, 150 mM NaCl) containing 50 mM imidazole.
The CS polymerase complexes were eluted with buffer A containing 250 mM imidazole. CS
polymerase complexes were dialyzed using dialysis membrane (Spectra/Por® MWCO 3.5 kD) in
buffer B (50 mM Tris pH 7.8, 500 mM NaCl) for 16 h at 4 °C in the presence of 500 µg 3C protease. A
second HisTrap™ HP purification step was carried out and protein complexes eluted at an imidazole
concentration of 100 mM. In this step, a 1 mL HisTrap™ HP column was coupled with two 5 mL
HiTrap™ desalting column for buffer exchange. CHSY1-CHPF and CHSY3-CHPF were stored in 20
mM MES at pH 6.5, 150 mM NaCl, while CHSY1-CHPF2, CHSY3-CHPF2 were stored in 50 mM
Tris at pH 7.8, 500mM NaCl. Protein samples were concentrated to 0.3 – 1 mg/mL and flash frozen in
liquid nitrogen before storage at -80°C for subsequent functional and structural analysis. Point
mutations in CHSY3 constructs were introduced using site-directed mutagenesis PCR
(Supplementary Table 3). Mutant CS polymerase complexes were purified using the same protocol
as for the wild-type complexes.

Nano-differential scanning fluorimetry

Thermal stability of purified CS polymerase wild-type and CHSY3 mutant-containing CHSY3-CHPF
complexes were measured using nano-differential scanning fluorimetry (nanoDSF). Protein complexes
were diluted with 20 mM MES pH6.5, 150 mM NaCl buffer to a final concentration of 0.2 mg/mL and
loaded into glass capillaries (Nanotemper). Melting curves were recorded on a Prometheus NT48
device (Nanotemper) using an excitation power of 100% and a temperature gradient from 15 °C to 95
°C using a temperature slope of 1.5 °C per minute. Melting points of duplicate measurements were
analyzed using the PR ThermControl v.2.3.1 software.

Mass photometry analysis

Mass photometry analysis were performed using a Refeyn OneMP instrument. CS polymerase wild-
type and CHSY3 mutant-containing CHSY3-CHPF complexes were diluted with a buffer containing
20 mM MES pH6.5, 150 mM NaCl to a final concentration of around 25 nM. Microscope coverslips
(24 × 50 mm, 170 µm) were thoroughly cleaned with H₂O and isopropanol before being air-dried. A
reusable gasket (Grace Bio-Labs) was placed onto the clean coverslip and 19 µL of the buffer used for
protein dilution were applied. The focal position was identified automatically using Refeyn
AcquireMP 2024.1.1 software and then 1 µL of protein standard or diluted CS polymerase complex was added. Mass calibration was performed using a 1:20 diluted NativeMark™ unstained protein standard solution (Thermo Fisher Scientific). 60 s movies were recorded, corresponding to 6,000 frames, using a regular field-of-view (18 µm²) acquisition setting. The corresponding masses for detected particles was calculated using Refeyn DiscoverMP 2024.1.0 software. Mass kernel density was estimated with an 8 kDa bandwidth.

Chemo-enzymatic synthesis of penta- and hexasaccharide peptides

A peptide derived from colony stimulating factor 1 (CSF1, EEASGEAS) was chemically synthesised
with a TAMRA fluorescent label at the N-terminal position (SB-peptide). The pentasaccharide
(GalNAc-GlcA-Gal-Gal-Xyl) addition was realised in a two-step process. The first step was to prepare
the tetrasaccharide linker as previously described¹⁴. Briefly, 300 µM CSF1 was incubated with 1 µM
XylT1, 8.4 µM β4GalT7, 1mM UDP-Xyl and 1 mM UDP-Gal in a buffer containing 25 mM MES pH
6.5, 125 mM NaCl, 2.5 mM MgCl₂, 2.5 mM MnCl₂ with a final reaction volume of 3 mL. After 16 h
incubation at room temperature, 2 µM β3GalT6, 2 µM GlcAT-1, 1 mM UDP-GlcA and 1 mM UDP-
Gal were added. After 24 h, the sample was loaded onto two Superdex Peptide 10/300 GL columns
(Cytiva), connected in serie and pre-equilibrated in 25 mM MES pH 6.5, 250 mM NaCl. Elution
fractions were analyzed by 25% acrylamide gel electrophoresis and using a fluorescence imaging
system. The tetrasaccharide-CSF1 peptide-containing fractions were pooled and dialysed against 4 L
of dH₂O, lyophilized and resuspended in 1 mL of dH₂O. In a second step, the fifth sugar, GalNAc, was
enzymatically added using purified CHSY3-CHPF complex. A 2 mL reaction mixture containing 300
µM tetrasaccharide-CSF1 peptide, 1 mM UDP-GalNAc, 1 µM CHSY3-CHPF was prepared in 50 mM
MES pH 6.5, 50 mM NaCl, 5 mM MnCl₂ and 2.5 mM MgCl₂. After 16 h incubation in the dark at
37°C, the sample was purified and analyzed as described above. The concentration of Penta-CSF1 was
determined using a titration curve with non-glycosylated peptides, loaded onto a 25% acrylamide gel,
and by correlating the band intensities quantified using Image Lab v.6.1 (Bio-Rad). This Penta-CSF1
peptide served as substrate for the synthesis of the Hexa-CSF1 peptide. A 200 µL reaction mixture
was prepared containing 100 µM Penta-CSF1, 1 mM UDP-GlcA, 1 µM CHSY3-CHPF, 50 mM MES
pH 6.5, 50 mM NaCl, 5 mM MnCl₂ and 2.5 mM MgCl₂. After 16 h incubation in the dark at 37°C, the
reaction products were purified and analyzed as described above. All glyco-peptides were stored at –
20 °C.
### *In vitro* glycosylation assays

Fluorophore-Assisted Carbohydrate Gel Electrophoresis (FACE) assay using chemo-enzymatically synthesized fluorescent Penta-CSF1 and Hexa-CSF1 peptide substrates was used to follow *in vitro* CS chain elongation by purified CS polymerase complexes. CS chain elongation reaction mixture contained 1 µM Penta-CSF1 acceptor substrate, 1 mM UDP-GalNAc and 1 mM UDP-GlcA donor substrates, 125 nM wild-type or mutant CS polymerase complexes and reaction buffer (50 mM MES pH6.5, 50 mM NaCl, 5 mM MgCl₂, 5 mM MnCl₂). The final reaction volume was 20 µL. Reactions for wild-type complexes were incubated for 24 h at 30 °C, and mutant complexes for 4 h at 37 °C, before being stopped by heating the samples to 85 °C for 10 min. 5 µl reaction mix were diluted with 15 µl loading buffer containing 20 % (v/v) glycerol and trace amounts of phenol red, from which 5 µl were loaded onto a 25% polyacrylamide gel. Bands were detected using a ChemiDoc™ MP imaging system (Bio-Rad) and the Alexa 546 nm filter. The bands were quantified using Image lab 6.0.1. To determine the importance of metal ions for GlcA and GalNAc transfer, similar reactions were prepared, including either 10 mM EDTA, 10 mM MgCl₂, 10 mM MnCl₂ or 10 mM CaCl₂. The reactions were incubated for 20 min. For the rescue experiment, CS polymerase complex concentrations varied between 62.5 nM and 125 nM. The reactions were incubated for 2 h.

### Mass spectrometry analysis

We utilized a matrix-assisted laser desorption ionisation (MALDI)-time-of-flight (TOF)/TOF mass spectrometer (Autoflex MAX, Bruker Daltonics, Bremen, Germany). A Bruker proprietary (Nd:YAG) 355 nm smartbeamTM II solid-state laser (maximum frequency: 2 kHz) was used for ionisation. Each mass spectrum was generated by averaging 10,000 laser shots, randomly acquired from various target spots. Glycosylated peptides were analyzed in reflector positive ion mode, detecting the 800-3,000 m/z range. Reaction mixtures containing peptides were diluted in 5 % formic acid (FA) solution to obtain 5 µM samples. The procedure for depositing samples on the MALDI plate (Bruker AnchorChip™) has been described previously⁵⁶. Briefly, a thin layer composed of a saturated solution of α-cyano-4-hydroxycinnamic acid matrix in acetonitrile was prepared. On the TL, 0.8 µL of peptide sample was loaded. Then 0.8 µL of a mix of α-cyano-4-hydroxycinnamic acid and 2,5-dihydroxybenzoic acid matrices were put on each spot. The matrices were left to dry at room temperature. 0.6 µL of a standard calibrator (peptide standard II, Bruker Daltonics) were added next to the samples, as well as 0.6 µL of the matrix solution. Intact peptides spectra were processed using FlexAnalysis™ (version 3.4). The sophisticated detection algorithm of the Numerical Annotation Procedure allowed the identification of peaks with a signal-to-noise ratio threshold of 2 and a maximum number of 500 detected peaks.
### Cryo-EM sample preparation and data collection

Freshly purified CHSY3-CHPF complex, diluted in 20mM MES pH 6.5, 100mM NaCl and 2.5mM $MnCl_2$, was mixed with 10 mM UDP and 500 µM octasaccharide ($GalNAc-GlcA)_4^{57}$ and incubated for 1 h on ice. The final protein concentration was 0.6 mg/mL. 4 µL sample was applied onto a glow-discharged carbon-coated copper Quantifoil® R 1.2/1.3 300 Mesh grid, excess liquid was removed by blotting for 5 s and grid was plunge-frozen in liquid ethane cooled by liquid nitrogen using a Vitrobot Mark IV (FEI) plunge freezer, with the chamber temperature set to 4 °C and 100% humidity. Grids were screened using a 200 kV Glacios (FEI) electron microscope. Data collection was carried out at the European Synchrotron Radiation Facility (ESRF) on a 300 kV Titan Krios G4 (Thermo Fisher Scientific) transmission electron microscope (beamline CM02), equipped with a cold field emission gun, a Selectris X energy filter, and a Falcon 4i electron detector. 12,169 movies were collected with a total electron dose of 50 $e^-/Å^2$ at a magnification of 165,000 x and raw pixel size of 0.73 Å.

### Cryo-EM image processing, structure building and refinement

Cryo-EM data analysis was performed using the cryoSPARC v3.3.1 software. Movies were imported and patch motion correction and patch CTF estimation was carried out. Upon manual inspection, 7,598 micrographs were selected. An AlphaFold 2 predicted model of the CHSY3-CHPF complex was converted into a map using EMAN's pdb2mrc command$^{58}$, low-pass filtered to 8 Å and used for several rounds of template-based particle picking, resulting in a total of 4,534,470 selected particles. Particles were extracted with a box size of 196 pixels and 2x binned. 2D classification was performed to remove junk particles and particles were further sorted using several rounds of heterogeneous refinement. After removing duplicate particles, a final set of 166,015 particles was obtained, which was used for *ab initio* reconstruction and non-uniform refinement, resulting in a 3D reconstruction with a global resolution of 3.0 Å based on a FSC cut-off of 0.143. Local resolution-filtered full map was generated using a B-factor of -40. In addition, local refinements were performed for the N-terminal and C-terminal part of the complex, resulting in maps with an approximate resolution of 2.9 Å and 3.0 Å, respectively. These maps were also local resolution filtered using B-factors of -40. A composite map was generated by adding the N-terminal and C-terminal maps together using the vop maximum command in UCSF ChimeraX v.1.6.1$^{54}$. The AlphaFold 2 predicted model of the CHSY3-CHPF complex was docked into the EM map using UCSF ChimeraX v.1.6.1$^{54}$. The model was trimmed to remove the N-terminal and C-terminal amino acid residues as well as loops for which no EM density was observed. The model was then manually refined using WinCoot$^{59}$. A UDP ligand, two $Mn^{2+}$ ions and three N-acetylglucosamines were placed into corresponding EM density. Several rounds
of real-space-refinement in PHENIX⁶⁰ and manual building in WinCoot were performed. The final model was validated using MolProbity⁶¹. Figures were prepared using UCSF ChimeraX v.1.6.1⁶².

### *In cellulo complementation assay*

A complementation assay was designed to test the ability of wild-type and mutant CHSY1 and CHSY3 proteins to carry out CS chain polymerization inside a cell. Synthetic genes encoding full-length CHSY1, CHSY3, CHPF and CHPF2 were inserted into the pTWIST-CMV-Beta-Globin-WPRE-Neo vector (Twist Bioscience). The constructs also encoded a N-terminal 3x FLAG tag, 8x His tag and a 3C cleavage site. Point mutations were introduced through site-directed mutagenesis PCR (see Supplementary Table 3). HEK293-6E wild-type and CHSY1/CHSY3 double knockout cells⁴⁰ were initially cultured as adherent cells in DMEM Gluta MAX™ medium, containing 20% fetal bovine serum, 1% penicillin and streptomycin at 37 °C and 8% CO₂. The adherent cells were then detached and transferred into suspension culture using FreeStyle™ medium and placed into a shaking incubator set to 37 °C and 8% CO₂. Double knockout cells (2 x 10⁶ cells/mL) were co-transfected with wild-type and mutant-containing CHSY1-CHPF and CHSY3-CHPF constructs using PEI (PEI:DNA₁:DNA₂ = 2:0.5:0.5). After 36 hours, cells were harvested by centrifugation. To detect cell surface CS levels, 1 x 10⁶ cells were incubated with an anti-CS antibody (Sigma, C8035 clone 56, dilution 1:250) for 1 h on ice. Cells were washed twice with a PBS buffer containing 1% (w/v) bovine serum albumin (BSA) and then incubated with an Alexa Fluor 488-labelled anti-mouse secondary antibody (Jackson Immuno Research, 115-545-075) at a dilution of 1:300 for 1 h on ice. After three subsequent wash steps with the same buffer, the cells were fixed using 4% paraformaldehyde (PFA) for 15 min, again followed by three washing steps. Finally, cells were re-suspended in 200 µL PBS buffer containing 1% (w/v) BSA and analyzed. To detect the transfection efficiency, 2 x 10⁶ cells were fixed with 4% PFA for 10 min at room temperature, washed with PBS, and permeabilized with 0.1% Triton X-100 for 10 min at room temperature. After washing with PBS buffer containing 1% (w/v) BSA, the cells were labelled with primary anti-FLAG antibody (Sigma, F1804) at a dilution of 1:300 for 1 h on ice, followed by two wash steps. Next, a cyanine3-conjugated anti-mouse antibody was added at a dilution of 1:300 (Jackson ImmunoResearch, 115-165-068) and cells incubated for 1 h on ice. Cells were finally washed, fixed with 4% PFA for 15 min on ice, and re-suspended in 300 µL wash buffer. Flow cytometry experiments were performed on a Miltenyi biotech flow cytometer and analyzed using the Macs Quant v.2.13.0 software (Miltenyi). The transfection efficiency was quantified by dividing the number of cells successfully transfected by the total cell count. The applied gating strategy is exemplified in Supplementary Fig. 12a. To quantify CS abundance, the median background signal (only anti-Alexa Fluor 488 antibody) was subtracted from the median signal
obtained for staining with both anti-CS antibody and anti-Alexa Fluor 488 antibody. Median values
were calculated for full cell population after omitting a small population of dead cells. Resulting
values were normalized to HEK293-E WT cells (100 %).

Data availability

The atomic coordinates for the CHSY3-CHPF complex have been deposited in the Protein Data Bank (PDB ID: 9Q8Z). The Cryo-EM maps have been deposited in the Electron Microscopy Data Bank under the following IDs: EMD-52913 (composite map), EMD-53018 (consensus map), EMD-53011 (focused refinement of the N-terminal part), and EMD-53012 (focused refinement of the C-terminal part). Mass spectrometry raw data is available through the Figshare repository (https://doi.org/10.6084/m9 figshare.28615220).

Acknowledgments

We thank Hugues Lortat-Jacob for regular feedback on project ideas and his feedback on the
manuscript. We thank Caroline Mas for assistance and access to mass photometry analysis, Michel
Thepaut for access to the nano-differential scanning fluorimetry instrument, Jean-Philippe Kleman for
help to acquire and analyze flow cytometry data, and Manal Hal Majed for performing site-directed
mutagenesis PCR. We thank Shibom Basu and EMBL Grenoble for assistance and access to
AlphaFold 2 predictions. We thank Eleftherios Zarkadas for support during cryoEM specimen
optimization and for cryo-EM data acquisition, and Guy Schoehn for establishing and managing the
IBS-ISBG cryo-electron microscopy platform and for providing access, training and support. We
thank Chrystel Lopin for providing the chemically synthesized octasaccharide compound and Rebecca
Miller for sharing CHSY1/CHSY3 KO cell lines.

This work used the EM facility, the cell imaging platform, the mass spectrometry facility and the
biophysics characterization platform of the Grenoble Instruct-ERIC Center (ISBG ; UAR 3518 CNRS-
CEA-UGA-EMBL) within the Grenoble Partnership for Structural Biology (PSB), supported by
FRISBI (ANR-10-INBS-0005-02) and GRAL, financed within the University Grenoble Alpes
graduate school (Ecoles Universitaires de Recherche) CBH-EUR-GS (ANR-17-EURE-0003). The IBS
Electron Microscope facility is supported by the Auvergne Rhône-Alpes Region, the Fonds Feder, the
Fondation pour la Recherche Médicale and GIS-IbiSA. The cryo-electron microscopy data were
acquired at the CM02 CRG beamline operated by IBS at the European Synchrotron Radiation Facility
in Grenoble. The purchase of this microscope was funded by the EquipEx+ France Cryo-EM project
(ANR-21-ESRE-0046). This work was funded by the Impulscience® Program of the Bettencourt
Schueller Foundation, project name [GlycoPol], by the ANR GlycoLink, grant number [ANR-23-CE44-0011], a Labex-GRAL PhD fellowship, and the "Investissements d'avenir" program Glyco@Alps, grant number [ANR-15-IDEX-02].

## Author contribution

R.W. designed the project. P.D. performed AlphaFold 2 predictions. P.D. and M.W generated expression constructs. P.D and M.B. performed co-expression tests and purified CS polymerase wild-type and mutant complexes. P.D performed nanoDSF experiments. P.D. and S.D.V performed mass photometry experiments. M.B. produced penta- and hexasaccharide peptides. P.D. and M.B performed *in vitro* glycosylation assays. E.B.-E performed mass spectrometry analysis. M.M. and P.D. prepared and screened EM grids. R.W. and P.D. processed cryo-EM data. R.L.C. built the structure and carried out model refinement. P.D., R.L.C. and R.W. analyzed the structure. M.F. and P.D. performed *in cellulo* complementation assay. P.D. and R.W. wrote the manuscript and all authors contributed to its revision.

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