Spinning sugars in antigen biosynthesis: characterization of the Coxiella burnetii and Streptomyces griseus TDP-sugar epimerases

The sugars streptose and dihydrohydroxystreptose (DHHS) are unique to the bacteria Streptomyces griseus and Coxiella burnetii, respectively. Streptose forms the central moiety of the antibiotic streptomycin, while DHHS is found in the O-antigen of the zoonotic pathogen C. burnetii. Biosynthesis of these sugars has been proposed to follow a similar path to that of TDP-rhamnose, catalyzed by the enzymes RmlA, RmlB, RmlC, and RmlD, but the exact mechanism is unclear. Streptose and DHHS biosynthesis unusually requires a ring contraction step that could be performed by orthologs of RmlC or RmlD. Genome sequencing of S. griseus and C. burnetii has identified StrM and CBU1838 proteins as RmlC orthologs in these respective species. Here, we demonstrate that both enzymes can perform the RmlC 3’’,5’’ double epimerization activity necessary to support TDP-rhamnose biosynthesis in vivo. This is consistent with the ring contraction step being performed on a double epimerized substrate. We further demonstrate that proton exchange is faster at the 3’’-position than the 5’’-position, in contrast to a previously studied ortholog. We additionally solved the crystal structures of CBU1838 and StrM in complex with TDP and show that they form an active site highly similar to those of the previously characterized enzymes RmlC, EvaD, and ChmJ. These results support the hypothesis that streptose and DHHS are biosynthesized using the TDP pathway and that an RmlD paralog most likely performs ring contraction following double epimerization. This work will support the elucidation of the full pathways for biosynthesis of these unique sugars.

The sugars streptose and dihydrohydroxystreptose (DHHS) are unique to the bacteria Streptomyces griseus and Coxiella burnetii, respectively. Streptose forms the central moiety of the antibiotic streptomycin, while DHHS is found in the O-antigen of the zoonotic pathogen C. burnetii. Biosynthesis of these sugars has been proposed to follow a similar path to that of TDP-rhamnose, catalyzed by the enzymes RmlA, RmlB, RmlC, and RmlD, but the exact mechanism is unclear. Streptose and DHHS biosynthesis unusually requires a ring contraction step that could be performed by orthologs of RmlC or RmlD. Genome sequencing of S. griseus and C. burnetii has identified StrM and CBU1838 proteins as RmlC orthologs in these respective species. Here, we demonstrate that both enzymes can perform the RmlC 3'',5'' double epimerization activity necessary to support TDP-rhamnose biosynthesis in vivo. This is consistent with the ring contraction step being performed on a double epimerized substrate. We further demonstrate that proton exchange is faster at the 3''-position than the 5''-position, in contrast to a previously studied ortholog. We additionally solved the crystal structures of CBU1838 and StrM in complex with TDP and show that they form an active site highly similar to those of the previously characterized enzymes RmlC, EvaD, and ChmJ. These results support the hypothesis that streptose and DHHS are biosynthesized using the TDP pathway and that an RmlD paralog most likely performs ring contraction following double epimerization. This work will support the elucidation of the full pathways for biosynthesis of these unique sugars.
The gammaproteobacterium Coxiella burnetii evolved recently from tick endosymbionts (1) to become an obligate intracellular pathogen of mammals (2). This bacterium infects a wide range of mammals causing reproductive failures (3,4). It is a significant economic pathogen of sheep, goats, and cattle (5), causing spontaneous abortion of pregnancies (6). Humans can be infected by C. burnetii either through aerosols or consumption of meat and dairy products (with 1-10 bacteria sufficient to cause disease) (7,8). C. burnetii in unpasteurized ruminant milk poses a particular risk (9)(10)(11). Humans generally present with a self-limiting febrile illness (12). A small percentage of cases lead to more serious complications such as miscarriage, hepatitis, or endocarditis (2,13,14). C. burnetii infection can be effectively treated with doxycycline (2). However, diagnosis is often challenging as C. burnetii grows only in highly defined media, requires environment-controlled microaerobic incubators (15), and symptoms of infection are generally nonspecific.
From a "One Health" perspective, a vaccine against C. burnetii would be the ideal approach to reduce veterinary and human infection (6). Inactivated whole cell vaccines have proved effective in animals and humans (16,17). These are not licensed for human use in the USA, EU, or UK as they cause severe reactions in seropositive individuals (18). Current efforts to develop novel vaccines are focused on the development of subunit- (19) or epitope-based vaccines (20,21). A very strong subunit vaccine candidate is the C. burnetii polysaccharide O-antigen. A complete O-antigen is required for immune evasion and efficient infection of mammalian cells (22,23). Serial passaging of C. burnetii selects for mutants that cease producing some or all their O-antigen (24,25). The most common mutants either lack a polymer of the unusual saccharide virenose ("intermediate") or the entire O-antigen (26) (Fig. 1A). Strains lacking the O-antigen are avirulent in the guinea-pig model of infection (27). The O-antigen is the dominant epitope of WT C. burnetii (28), suggesting that any vaccine against C. burnetii will require the O-antigen (28). This highlights the importance of a deepened understanding of the biosynthesis of this polysaccharide.
The C. burnetii O-antigen characteristically contains two C-3 methylated/hydroxymethylated sugars, virenose and dihydrohydroxystreptose (DHHS; Fig. 1B) (29,30). Virenose is also found in Bacillus cereus and Streptomyces albaduncus (31,32). Biosynthetic pathways have been proposed in these two organisms, and a similar pathway has been proposed in C. burnetii (33). No equivalent pathway has been proposed for DHHS biosynthesis. The most similar known sugar to DHHS is streptose (Fig. 1B), the central saccharide unit of the aminoglycoside antibiotic streptomycin (34), produced by Streptomyces griseus. Streptose differs from DHHS in having no hydroxyl at C-6 and being oxidized at the C-3 hydroxymethyl group. Streptose is synthesized from TDP-glucose, in a manner analogous to the biosynthesis of rhamnose (Fig. 1C) (34)(35)(36)(37). A reasonable path to DHHS synthesis would involve a C-4 oxidase of TDP-glucose, a 5''-mono or 3'',5''-double epimerase (38), and an enzyme similar to the streptose synthase (37) (Fig. 1, D and E). Although it remains possible that the epimerase might also perform the ring rearrangement, the only precedent for this reaction was performed by an enzyme from a different structural class (39).
Two regions (cbu0825 to cbu0856 and cbu1831 to cbu1838) have been suggested as coding for DHHS synthesizing genes (26). The cbu1831 to cbu1838 region contains two genes that show high sequence identity to genes in the rhamnose biosynthesis pathway. CBU1834 has over 60% sequence identity to characterized RmlA orthologs (33) and so is highly likely to be a glucose-1-phosphate thymidylyltransferase. TDP-glucose has also been proposed to be a precursor of virenose (33) and so is not necessarily characteristic of DHHS biosynthesis. CBU1838 shows 27 to 53% identity with characterized RmlC orthologs (Table S1). With this sequence identity, this protein is highly likely to be an ortholog of RmlC and act as a TDP-D-xylo-4-hexulose 3,5epimerase. This activity is not required for virenose biosynthesis. As C. burnetii does not produce rhamnose, it is unlikely that it would be required for rhamnose biosynthesis. This leaves DHHS biosynthesis as the only reasonable function of CBU1838.
RmlC is part of a family of epimerases that use a deprotonation/reprotonation mechanism (40). RmlC uses a histidine base to deprotonate the substrate and a tyrosine acid to reprotonate from the opposite side of the sugar (41)(42)(43)(44). The same residues are likely involved in epimerizing at both 3'' and 5'' positions: the sugar ring flips between the two reactions to present the second site to be epimerized to the catalytic base and acid (41). These amino acids are highly conserved in the family (Fig. 2). RmlC orthologs have been characterized biochemically and structurally from a range of species (Table 1) (38,42,43,(45)(46)(47)(48)(49). Paralogs that selectively epimerize at either the 3 0 or 5 0 carbon have also been characterized (44,50,51). These studies have highlighted likely residues for binding to the TDP carrier (43,52,53). For streptose and DHHS biosynthesis, only epimerization at the 5'' position is strictly necessary: the C3-C4 bond is broken in the streptose biosynthesis step (35), which will remove the stereocentre at C3. RmlC paralogs for streptose (StrM) and DHHS (CBU1838) biosynthesis have been proposed bioinformatically (33) (e.g., by KEGG (54)); however, such assignments can be inaccurate and so experimental confirmation is desirable (55).
Here, we tested the hypothesis that StrM and CBU1838 are orthologs of RmlC. We expected that they would perform only the epimerization step and not perform the structural rearrangement activity that has yet to be assigned. As both enzymes contain all the catalytic residues proposed for 3'',5''double epimerization, we expected that they might perform both transformations. We demonstrate in vivo and in vitro that these enzymes are competent to perform the double epimerization. Using NMR, we demonstrate that, surprisingly, proton-deuteron exchange at the 3'' position is faster than at the 5'' position for both enzymes. The structures of CBU1838 and StrM show that they form an active site highly similar to previously characterized enzymes, in keeping with the biochemical data. These data support a 3'',5'-double epimerized substrate for the streptose ring rearrangement.  (26). B and C, the unusual monosaccharides virenose and dihydrohydroxystreptose (DHHS) from the C. burnetii O-antigen (B) and the related sugar streptose (C). The differences between DHHS and streptose are highlighted with red circles. D and E, proposed biosynthetic pathways for TDP-linked streptose (D) and DHHS (E). The equivalent Escherichia coli enzymes for each step are shown in blue in between the two panels; proposed enzymes for the Streptomyces griseus (34) and C. burnetii (this study) pathways, where known, are shown in black. Chemical structures drawn using BIOVIA Draw v16.1. GalNAc, N-acetyl-galactosamine; GlcNAc, N-acetylglucosamine; Glc, glucose; KDO, 2-keto-3-deoxy-D-mannoctanoic acid; Man, mannose; Vir, virenose.

CBU1838 complement RmlC in Escherichia coli
We firstly tested whether CBU1838 can complement the E coli RmlC ortholog RfbC. We cultured WT MG1655 E. coli and rfbC mutant bacteria, and rfbC mutants complemented with CBU1838. We extracted soluble small molecules from each culture. WT MG1655 showed negligible levels of TDP-glucose and moderate levels of TDP-rhamnose. The rfbC mutant showed increased TDPglucose levels and reduced TDP-rhamnose levels ( Fig. 3; p < 0.0001 and p < 0.05 respectively, ANOVA test comparing all tested samples). This is expected for a mutant at this essential step of the pathway. When the rfbC mutant was complemented with CBU1838, the WT sugar nucleotide levels were restored (Fig. 3). In contrast, complementation with the RmlC paralog EvaD, which preferentially epimerizes  (50) and 4HMZ (44) (MtRmlC, AoEvaD, and SbChmJ, respectively) and the PDB files from CBU1838 and SgStrM solved in this study. Chain A was used in each case. Secondary structure elements for MtRmlC and SgStrM are noted above and below the sequence alignments, respectively. The strand exchange between chains is highlighted with a blue box. The residues that interact with TDP and citrate and which are likely the catalytic acid and base are indicated with black arrows, sky blue dotted arrows, and pink dashed arrows, respectively. Figure generated using ESPript v3.0222 (72); a red box/white character relates strict identity, a black bold character relates similarity in a group, and filled in yellow characters relates similarity across groups. the 5'' position, was not able to restore WT sugar nucleotide levels. These data gave good confidence that CBU1838 performs both 3'' and 5'' epimerizations of TDP-6-deoxy-Dxylo-4-hexulose, as expected.

StrM and CBU1838 complement RmlC in vitro
We next tested whether StrM and CBU1838 can perform the function of E. coli RmlC in a coupled assay. We used the test enzyme coupled to RmlB and RmlD in a catalytic cascade to convert TDP-glucose to TDP-rhamnose. Both StrM and CBU1838 complemented RmlC in these assays effectively (Fig. 4). Identical concentrations of StrM and RmlC gave a similar effect. A higher concentration of CBU1838 was required to achieve the same effect. This is consistent with the expected function of CBU1838, epimerizing a sugar that retains the 6''-OH group. These data further support the concept that StrM and CBU1838 perform both 3'' and 5'' epimerizations of TDP-6-deoxy-D-xylo-4-hexulose.
CBU1838 shows a reduced rate compared to RmlC and StrM, consistent with its modified substrate We determined the kinetic constants for StrM and RmlC, in comparison to other characterized enzymes. We used purified TDP-6-deoxy-D-xylo-4-hexulose and coupled the test enzymes to RmlD. Data from StrM, RmlC, CBU1838, and EvaD all fitted very well to the Michaelis-Menten equation, with no evidence of cooperativity (Fig. S1). StrM and RmlC showed a similar k cat , with StrM having a slightly lower K M ( Table 2). CBU1838 showed a lower k cat and a clearly higher K M ( Table 2). This is consistent with its expected 6''-hydroxy substrate. As expected, the 5''-specific epimerase EvaD showed a substantially lower k cat . We did not observe any activity with the 3''-specific epimerase ChmJ (Fig. S2). Previous studies have demonstrated that ChmJ has no 5''-epimerization activity (44), and that the coupling enzyme RmlD only reduces double-epimerized products (56,57). The kinetic constants provide further evidence that StrM is likely an ortholog of RmlC. CBU1838 can perform this activity, but its likely native substrate is 6''hydroxylated.     RmlC, StrM, and 1838 epimerize at both the 3'' and 5'' positions To give further confidence in the activity of CBU1838 and StrM, in a novel assay, we monitored their activity directly using NMR and GC-MS without employing a coupling agent. We performed 1 H-NMR experiments in deuterated buffer. As RmlC uses a deprotonation-protonation mechanism with different proton acceptors and donors, we expected to observe a loss of signal from protons bound to the 5'' and 3'' carbon atoms. We observed solvent exchange at both positions for RmlC, StrM, CBU1838, and EvaD, with relative rates in the same order as the rates observed in a coupled assay (Figs. 5A and S3). Surprisingly, the rate of exchange at the 3'' position was significantly faster than the rate at the 5'' position for RmlC, StrM, and CBU1838 (Table 3); for EvaD, the rate of exchange at the two positions was similar. When the reaction was permitted to proceed to completion, RmlC, StrM, CBU1838, and EvaD all showed almost complete exchange at both 3'' and 5'' positions (Fig. 5B). In contrast, the 3'' specific enzyme ChmJ showed almost complete exchange at the 3'' position, with very little exchange observed at the 5'' position. GC-MS analysis of enzyme reactions run to completion confirmed that exchange had occurred at both positions for CBU1838, StrM, and RmlC (Fig. S4). Consistent with the NMR data and previous work (44), ChmJ showed strong exchange at the 3'' position only. These data strongly support a double epimerase activity for both CBU1838 and StrM.

Sugar backbone rearrangement is not observed with either StrM or CBU1838
In neither NMR nor GC-MS data was there any evidence of a rearrangement of the sugar backbone. The proposed biosynthetic pathways of both StrM and CBU1838 require such a rearrangement, catalyzed by either the epimerase or a synthase/reductase for each pathway. Our direct detection of product formation and chemical structure analysis might have allowed us to detect this rearrangement. However, as the fivemembered rings are likely to be substantially disfavored at equilibrium, the possibility remains that rearranged structures were present but below the limit of detection.
The structures of StrM and CBU1838 are consistent with their role as double epimerases We determined the structures of StrM and CBU1838 by X-ray crystallography, in the presence and absence of TDP (Table S2). Both proteins show an overall dimeric architecture consistent with previously solved members of this family (Fig. 6A). Both proteins show a strand exchange in common with orthologs. A phylogenetic tree (Fig. S5) suggests that CBU1838 groups with other 3'',5'' epimerases, while StrM is closer in sequence to the single site epimerases. The sequence identities and structure RMSDs are consistent with this   ( Table S1). TDP binds to both proteins in a similar manner (Fig. 6, B and C). TDP is held in place by a network of amino acid side chains a (F21*, R25*, F28*, E30*, Q48, R61, H64, Y140, K170) from both protomers (starred residues from the domain swapped strands). Additional interactions are formed through bound solvent from N50, Q72, Y134, and E145. These are all strongly conserved across RmlC orthologs (Fig. 2). The CBU1838-TDP structure showed a citrate molecule (carried over from crystallization) bound to the active site. This molecule occupies the same space as the substrate in previously solved structures. The citrate interacts with S52, H64, K74, and Y134 from the protein and TDP (Fig. 6D). These amino acids are well conserved among orthologs, and H64/ Y134 are the catalytic base and acid proposed from previous studies (41)(42)(43)(44). The conformation of these side chains is very similar to that seen in complexes of other enzymes with substrate analogs (Fig. S6). These structures give further confidence that CBU1838 and StrM are indeed orthologs of RmlC.
They have a very similar fold, show clear and specific binding to TDP, and retain the key catalytic residues in the correct orientation.

Discussion
Streptose and DHHS are monosaccharides produced by a single organism each. Both sugars are proposed to be synthesized through modified TDP-rhamnose pathways. S. griseus and C. burnetii contain putative orthologs of this biosynthetic pathway. Both organisms contain a single TDP-6-deoxy-Dxylo-4-hexulose 3'',5''-epimerase ortholog (StrM and CBU1838). We sought to determine whether these enzymes perform both epimerizations and whether either might also perform the sugar ring rearrangement activity. Answering these questions will provide further insight into the biosynthesis of these unusual sugars.
CBU1838 showed robust double epimerase activity both in vitro and in vivo. CBU1838 complemented an rfbC (rmlC) mutation, restoring flux through the pathway and consuming cellular TDP-glucose and TDP-6-deoxy-D-xylo-4-hexulose (Fig. 3). CBU1838 and StrM were competent to replace RmlC in vitro in a biosynthetic pathway (Fig. 4). It was notable that substantially more CBU1838 was required to restore activity than RmlC or StrM. Similarly, CBU1838 showed a lower k cat and higher K M than either RmlC or StrM (Table 2). This is consistent with the proposed role of CBU1838, as the DHHS pathway requires a substrate with a 6''-hydroxyl. CBU1838 contains a methionine residue (M123) in place of a phenylalanine conserved among other RmlC orthologs, which would provide space for the hydroxyl. The k cat and K M values that we obtained for StrM and RmlC are in the same order as those observed for other RmlC paralogs (Table S3). It is notable that our data were collected at 37 C, while the comparative data were collected at 21 to 25 C. It is likely that other enzymes would react faster at higher temperatures but might show a higher K M (58).
Our NMR data show that CBU1838 and StrM have similar activity to E. coli RmlC, with CBU1838 again showing a reduced comparative activity. The rate of proton exchange at the 3'' position was faster than that at the 5'' position for all three enzymes (Fig. 5 and Table 3). The Mycobacterium tuberculosis RmlC showed faster epimerization at the 5'' position (41). Dong et al. suggested that in the likely catalytic conformation, the 5'' proton is more acidic than the 3'' proton (41). Our experiments measured the loss of proton signal by NMR, while the previous study measured the product masses by GC-MS. Furthermore, our study used chemically prepared TDP-6-deoxy-D-xylo-4-hexulose, which has only recently become commercially available. This may explain some differences between our study and previous work. We note that as all the reactions involve proton transfers at all steps (40), there may be a strong kinetic isotope effect (59) on either or both epimerizations. As our methods for differentially measuring the two epimerizations rely on deuterium, this confounding effect cannot be excluded. The structures of CBU1838 and StrM are highly similar to previously solved structures of paralogs from other species. TDP is held in place by residues that are strongly conserved (Figs. 2 and 6) (43,48,52). Although we were not able to determine a structure with a TDP-linked sugar, the side chains lining the sugar binding (and catalytic) cavity adopt conformations that are highly similar to those in previous structures (38, 41, 43-45, 48, 50, 51, 53). The proposed catalytic residues, H64 and Y134, occupy a very similar space to the equivalent residues in substrate analog cocrystal structures (41,53). As their local hydrogen-bonding networks are also conserved, it is likely that they will also be catalytic in these proteins. The presence and conformation of these catalytic residues cannot predict the specificity of the enzymes. While one example of a 5''-specific enzyme showed a mutation that altered the conformation of an important active site side chain (50), 3''specific epimerases have shown a complete complement of active site residues (44,51). Our development of a functional assay in E. coli allowed us to confirm the activity of StrM and CBU1838 in vivo (Fig. 3); although the 5''-specific enzyme EvaD was expressed to the same levels as the active enzymes, it was not competent to complement rmlC. However, this assay did not distinguish between StrM (likely a direct RmlC ortholog) and CBU1838 (which is expected to use the 6''-hydroxylated analog).
The evidence presented in this and previous studies strongly suggests that both CBU1838 and StrM act as TDP-4-keto-(6deoxy)-glucose 3'', 5'' double epimerases. The lower rates observed for CBU1838 are consistent with its natural substrate retaining the 6''-hydroxyl group; an activity to generate this substrate has not yet been identified. This suggests that the streptose and DHHS synthase enzymes (Fig. 1) accept a substrate that is epimerized at both the 3'' and 5'' positions. Although our data suggest that these enzymes cause proton exchange at the 3'' carbon more efficiently than the 5'' carbon, we cannot exclude that the kinetic isotope effect could be confounding. Our results provide powerful evidence that DHHS biosynthesis in Coxiella has convergently evolved to use a similar pathway to streptose biosynthesis in S. griseus.
The key challenge that this presents is to identify the enzymes that provide the two remaining activities to generate DHHS.

Gene cloning and synthesis
The gene fragments for rmlB, rmlC, and rmlD were amplified from E. coli DH5α (NEB) genomic DNA (gDNA) by PCR. The CBU1838 gene fragment was codon-optimized using IDT's tools and ordered as a gBlock from IDT and amplified by PCR. Primer sequences are provided in Table S4. Gene fragments were cloned into pNIC28-Bsa4 (Addgene #26103, gift of Opher Gileadi, SGC Oxford) using ligation-independent cloning, following published methods (60). Briefly, 16.5 μg pNIC28-Bsa4 plasmid was digested with 60 U BsaI (NEB) in 100 μl following the manufacturer's protocol and incubated for 2 h at 50 C. To generate the ligation-independent cloning cohesive ends in both the plasmid and PCR product, a mixture of 1 μl water, 5 μl BsaI-digested plasmid/PCR-pure DNA, 2 μl 5X T4 DNA polymerase buffer (Fermentas), 1 μl 25 mM dGTP (insert) or dCTP (plasmid), 0.5 μl 100 mM DTT, 0.5 μl T4 DNA polymerase (Fermentas) was prepared and incubated at 22 C for 30 min, then 20 min at 75 C. Two microliters of treated DNA product was added to 1 μl of treated plasmid. The mixture was incubated at room temperature for 10 min followed by transformation into 5-alpha competent cells (NEB). The E. coli DH5α rmlD contains an inactivating point mutation. This was corrected by site-directed mutagenesis using the QuikChange Lightening kit (Agilent), following the manufacturer's instructions.
TDP-sugar epimerase genes for S. griseus strM, Amycolatopsis orientalis evaD, and Streptomyces bikiniensis chmJ were codon optimized using an in-house Python script (https:// github.com/njharmer/CodonOptimise) and cloned into pET28b with an additional SUMO tag, pET28b, and pET21a respectively by Twist Bioscience. Plasmids were transformed into the expression strain E. coli BL21 (DE3) (Novagen) using kanamycin (50 μg/ml) for selection. For complementation of E. coli rfbC mutants, TDP-sugar epimerase genes for strM, evaD, and CBU1838 were cloned into the cloning vector pTWIST-A-MC under the control of the lac operon, with a strong ribosome-binding site calculated by the RBS Calculator (De Novo DNA). The Genbank files for all plasmids are available in Supplementary Data.

Preparation of electrocompetent E. coli mutant cell lines and complementation
ΔrfbC E. coli strains (Keio collection; Horizon Discovery #OEC4987-200827372) were grown overnight on LB plates with 50 μg/ml kanamycin. Colonies were inoculated from LB plates into 10 ml fresh LB broth and incubated overnight at 37 C with shaking at 225 rpm. One milliliter of overnight culture was inoculated into 100 ml fresh LB medium with kanamycin and incubated at 37 C with shaking at 225 rpm until A600 was approximately 0.6. Cells were harvested by centrifugation (4200g, 10 min, 4 C). The cells were resuspended in 40 ml prechilled autoclaved Milli-Q water. Cells were harvested by centrifugation as above and resuspended in 40 ml prechilled autoclaved 10% (v/v) glycerol. A minimum of three 10% glycerol washes were conducted before cells were resuspended in 500 μl 10% (v/v) glycerol. Complementation plasmids (40 ng) were added to 50 μl electrocompetent cells, vortexed for 30 s, and incubated on ice for 10 min. The suspension was transferred to a prechilled 0.1 cm electroporation cuvette (Thermo Scientific #5510-11). A single pulse of 1.8 kV was given by a Gene Pulser Xcell Microbial System (Bio-Rad #1652662). Cells were recovered by adding 1000 μl prewarmed LB medium. The complemented cell suspension was incubated at 37 C for 1 h shaking at 225 rpm. Transformed cells were grown on LB agar plates supplemented with 100 μg/ml ampicillin overnight at 37 C.

Detecting expression of complemented E. coli
Complemented E. coli strain colonies were inoculated into 100 ml bottles containing 10 ml LB medium, 100 μg/ml ampicillin, and incubated overnight at 37 C with shaking at 225 rpm. One milliliter of each overnight culture was inoculated into individual 500 ml conical flasks containing 100 ml LB medium supplemented with ampicillin and incubated at 37 C with shaking at 225 rpm. When the A600 reached 0.6, gene expression was induced by adding 200 μM IPTG. Cultures were incubated at 20 C with shaking at 225 rpm overnight. Fifty milliliter aliquots of overnight cultures were harvested by centrifugation (4500g, 30 min, 4 C). Cells were lysed using 1 ml BugBuster Master Mix (Millipore #71456-4), following the manufacturer's instructions. The soluble and insoluble fractions were separated by centrifugation at 16,000g for 30 min at 4 C. Samples were separated on a 4 to 12% ExpressPlus PAGE gel (GenScript #M41212) following the manufacturer's instructions. Samples were transferred to a nitrocellulose membrane (Sartorius Stedim Biotech #11327-41BL) using a Pierce G2 Fast Blotter (Thermo Scientific # 15146375) using the preprogrammed protocol. Antibodies were added using an iBind Western device (ThermoFisher #SLF1000), following the manufacturer's recommendations. The primary and secondary antibodies used were anti-penta His (Qiagen #34660) and goat anti-mouse IgG (LiCOR #926-68070) at a dilution of 1:1000. Blots were imaged using an Odyssey Clx imaging system (LiCOR).

Sample preparation for LC-MS QQQ analysis
Complemented E. coli strain colonies were inoculated into 100 ml bottles containing 10 ml LB medium, 100 μg/ml ampicillin for complemented strains, and incubated overnight at 37 C with shaking at 225 rpm. One milliliter of each overnight culture was inoculated into individual 500 ml conical flasks containing 100 ml LB medium supplemented with kanamycin and ampicillin as necessary and incubated at 37 C with shaking at 225 rpm until A600 reached 0.6. Expression was induced by adding 200 μM IPTG. Cultures were incubated at 20 C with shaking at 225 rpm overnight. Fifty milliliter aliquots of overnight cultures were harvested by centrifugation (4500g, 30 min, 4 C). Cellular contents were extracted into 1 ml 50% (v/v) acetonitrile. Cell suspensions were incubated at 25 C for 30 min and centrifuged at 20,000g for 30 min at 4 C. The supernatant was filtered using Millex Non-Sterile Low Protein Binding Hydrophilic LCR (PTFE) Membrane (0.45 μm) filters (Merck #SLLHR04NL) into 1.5 ml MS vials with silicone/PTFE septa and stored at −20 C for later analysis.
Quantitative LC-MS QQQ analysis: Quantitative analysis was performed using an Agilent 6420B triple quadrupole (QQQ) mass spectrometer (Agilent Technologies, Palo Alto) coupled to a 1200 series Rapid Resolution HPLC system. Five microliter of sample extract was loaded onto an Agilent Poroshell 120 HILIC-Z, 2.7 μm, 2.1 × 150 mm analytical column (Agilent Technologies #673775-924). For detection using negative ion mode, mobile phase A comprised 90% (v/v) LC-MS grade acetonitrile with 10 mM ammonium acetate and 5 μM medronic acid, and mobile phase B was 100% water (LC-MS grade) also with 10 mM ammonium acetate and 5 μM medronic acid. The following gradient was used: 0 min -10% B; 6 min -35% B; 10 to 13 min -40% B; 15 min -10% B; followed by 5 min re-equilibration time at a flow rate of 0.25 ml min −1 with the column held at 25 C for the duration. The QQQ source conditions for electrospray ionization were as follows: 350 C gas temperature with a drying gas flow rate of 11 L min −1 and a nebulizer pressure of 35 psig. The capillary voltage was 4 kV.

MS data analysis
Data analysis was undertaken using Agilent MassHunter Quantitative Analysis software (version B.07.01, SP1). Data were normalized to the internal standard, 13 C TDP-glucose, and the optical density of each culture pre-extraction. TDPglucose, TDP-rhamnose, GDP-mannose, GDP-fucose, and UDP-glucose concentrations were calculated using standard calibration curves. For the other compounds (where a standard was not available), normalized peak areas were compared.

Expression and purification of TDP-sugar epimerases
RmlB, RmlC, RmlD, StrM, EvaD, and ChmJ were expressed in 500 ml of ZYM-5052 auto-induction media supplemented with 100 μg/ml kanamycin following the methods of (61). Each flask was inoculated with 10 ml of an overnight culture and grown at 37 C with agitation at 200 rpm until A600 reached 0.6. Cultures were further incubated at 20 C for 18 h with agitation. Cells were harvested by centrifugation at 4500g for 30 min at 4 C. The pellet was resuspended in binding buffer (20 mM Tris-HCl, 500 mM NaCl, 10 mM imidazole, pH 8.0) and lysed by sonication (SONIC Vibra cell VCX130). The sample was clarified by centrifugation (24,000g for 30 min at 4 C). The soluble fraction was purified using an ÄKTAxpress chromatography system (Cytiva). The sample was purified using a 1 ml HisTrap crude column (Cytiva). After loading sample, the column was washed with binding buffer, and the protein step eluted into binding buffer supplemented with 250 mM. The eluate was purified over a Superdex 200 pg 16/ 600 size-exclusion column (Cytiva) and eluted isocratically into 10 mM Hepes, 500 mM NaCl, pH 7.5 (for coupled assays and crystallization) or 10 mM Na x HPO 4 , 500 mM NaCl, pH 7.5 for NMR studies. The eluted protein was concentrated using a Vivaspin centrifugal concentrator (Generon) and stored at −20 C with 20% (v/v) glycerol for enzymatic assays or stored at −80 C in small aliquots without any glycerol for crystallization or NMR.
All data were collected on a Bruker Neo 600 MHz spectrometer equipped with TCI cryoprobe. Standard Bruker proton experiments were carried out at 298 K (25 C). Periodic 1 H NMR experiments (64 scans) were carried out in 3 mm NMR tubes. 1 H NMR spectra were recorded every 10 min over a 90 min period, set using the Topspin multi_zgvd command. Later intervals were set manually to record at 2 h, 4 h, 8 h, 18 h, and 24 h, as appropriate. TopSpin 4.0.6 was used for data processing using command 'efp' followed by automatic phase correction ('apk') and automatic baseline correction ('abs'). The residual water peak at δ 4.7058 ppm was used as reference for spectra calibration. A set of selected diagnostic signals from no-enzyme control samples was integrated for calibration of the relative abundance of each signal. An integral of H-6 thymidine peak ( 1 H) was used for peak intensity calibration of the rest of integrals of diagnostic signals, and GraphPad Prism v8.1.2 was used to analyze the data obtained.
Deuterium-incorporation analysis GC-MS 0.5 mg TDP-6-deoxy-D-xylo-4-hexulose (Carbosynth) was incubated in deuterated buffer alone or with the addition of either 1 μM of EcRmlB, EcRmlC, SgStrM, CBU1838, or AoE-vaD in 250 μl final volume. After 18 h, enzymes were inactivated and removal by ethanol precipitation followed by centrifugation at 14,000g for 10 min. Sodium borohydride (2 mg) was added to each sample for the reduction of ketomoeity followed by incubation at room temperature for 2 h, with shaking at 120 rpm. The reaction was terminated with few drops of glacial acetic acid, followed by 20 μl 10% (v/v) acetic acid in methanol. Solvents were evaporated under reduced pressure using GeneVac for 1 h at 35 C, and 25 μg myo-inositol was added to the sample as an internal standard. The sugar-nucleotide bond was hydrolyzed by using 250 μl 2 M TFA followed by incubation at 100 C for 1.5 h. After cooling the samples, solvents were evaporated again and 100 μl isopropanol was added before a final evaporation in order to remove the traces of TFA for 1 h.
A second reduction was achieved by addition of 250 μl 1 M ammonia, containing 10 mg/ml sodium borohydride for 1 h and then 250 μl acetic acid-methanol (1:9, v/v) was added. The samples were dried under reduced pressure (GeneVac, 1 h at 40 C) and the addition of acetic acid-methanol followed by drying was repeated once again. This was followed by addition of 250 μl methanol (without acetic acid) twice with evaporating the solvent off each time (20 min each). The resulting sample was acetylated by incubation at 120 C for 20 min in a mixture of 100 μl acetic anhydride and 100 μl pyridine. After this, approximately 200 μl toluene was added, and the mixture was concentrated under reduced pressure. The addition of toluene and evaporation was repeated at room temperature.
The alditol acetates were separated by partition between equal volume of methylene chloride and water. After partition, the methylene chloride layer was left to evaporate slowly at room temperature. Finally, samples were dissolved in acetone and analyzed by GC-MS.

Crystallization
Protein concentrations used for crystallization were 5.85 mg/ml (SgStrM) and 5.2 mg/ml or 7.8 mg/ml (CBU1838). Crystals were grown using the microbatch method using an Oryx8 crystallization robot (Douglas Instruments). Crystals were grown in hydrophobic plates (Douglas Instruments #VB-Silver-1/1) and covered in a 1:1 mix of paraffin oil and silicone oil (Molecular Dimensions). Crystals were grown either as a 1:1 mixture of protein and mother liquor, or as a 3:2:1 mixture of protein, mother liquor, and seed stocks where seeds had been prepared from previous crystals. The successful cocrystallization conditions, substrates soaking conditions, and cryoprotectants used are detailed in Table S5.

X-ray data collection and structure determination
Data were collected at Diamond Light Source (Didcot) at 100 K using Pilatus 6M-F detectors (Table S2). All data were processed using XDS (62). CBU1838 data were subjected to anisotropic ellipsoidal truncation using the STARANISO server (63). The resolution limit for StrM datasets was set at a point where CC ½ was above 0.3 in the highest resolution shell. CBU1838 data were processed to include regions where the mean I/σ was systematically above 1.2. Further data processing and structural studies was carried out using the CCP4 program package (64) and the CCP4i2 interface (65). Molecular replacement was used to provide initial phases for datasets using Morda (66) and MolRep (67). The best models for CBU1838 and StrM were 2IXI (41) (54% identity) and 4HMZ (44) (52% identity), respectively. Density fitting was performed using Coot (68) and refined with REFMAC5 (69). Model validation was carried out with internal modules of CCP4i2 and Coot, employing MolProbity calculations (70).

Data availability
All data underpinning this work are publicly available. Structure coordinates and structure factor files are deposited with the Protein Data Bank (accession numbers: 7PA1, 7P7I, 7P7W, 7P9L, 7P9P, and 7P9Y). Enzymatic and mass spectrometry data are available as Supplementary Files or from Open Research Exeter (doi: http://doi.org/10.24378/exe.3724).