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Evaluation of Azido 3-Deoxy-d-manno-oct-2-ulosonic Acid (Kdo) Analogues for Click Chemistry-Mediated Metabolic Labeling of Myxococcus xanthus DZ2 Lipopolysaccharide

Metabolic labeling paired with click chemistry is a powerful approach for selectively imaging the surfaces of diverse bacteria. Herein, we explored the feasibility of labeling the lipopolysaccharide (LPS) of Myxococcus xanthus—a Gram-negative predatory social bacterium known to display complex outer membrane (OM) dynamics—via growth in the presence of distinct azido (-N3) analogues of 3-deoxy-d-manno-oct-2-ulosonic acid (Kdo). Determination of the LPS carbohydrate structure from strain DZ2 revealed the presence of one Kdo sugar in the core oligosaccharide, modified with phosphoethanolamine. The production of 8-azido-8-deoxy-Kdo (8-N3-Kdo) was then greatly improved over previous reports via optimization of the synthesis of its 5-azido-5-deoxy-d-arabinose precursor to yield gram amounts. The novel analogue 7-azido-7-deoxy-Kdo (7-N3-Kdo) was also synthesized, with both analogues capable of undergoing in vitro strain-promoted azide–alkyne cycloaddition (SPAAC) “click” chemistry reactions. Slower and faster growth of M. xanthus was displayed in the presence of 8-N3-Kdo and 7-N3-Kdo (respectively) compared to untreated cells, with differences also seen for single-cell gliding motility and type IV pilus-dependent swarm community expansion. While the surfaces of 8-N3-Kdo-grown cells were fluorescently labeled following treatment with dibenzocyclooctyne-linked fluorophores, the surfaces of 7-N3-Kdo-grown cells could not undergo fluorescent tagging. Activity analysis of the KdsB enzyme required to activate Kdo prior to its integration into nascent LPS molecules revealed that while 8-N3-Kdo is indeed a substrate of the enzyme, 7-N3-Kdo is not. Though a lack of M. xanthus cell aggregation was shown to expedite growth in liquid culture, 7-N3-Kdo-grown cells did not manifest differences in intrinsic clumping relative to untreated cells, suggesting that 7-N3-Kdo may instead be catabolized by the cells. Ultimately, these data provide important insights into the synthesis and cellular processing of valuable metabolic labels and establish a basis for the elucidation of fundamental principles of OM dynamism in live bacterial cells.


■ INTRODUCTION
The bacterial outer membrane (OM) serves as an important physical barrier, protecting the interiors of Gram-negative cells from the local environment. 1 The OM is thus often the first point of contact for extracellular interactions with other cells and/or substrata. Rather than a static structure, the OM is now known to be a highly dynamic interface, from which OM vesicle 2 and tube 3 projections are made, and for which its fluidity serves as a marker to activate stress-response pathways in a cell. 1 The OM is structured as an asymmetric bilayer, with the periplasmic (interior) leaflet largely composed of phospholipids. The outer leaflet is composed primarily of lip-opolysaccharide (LPS), a glycolipid containing three distinct segments. The proximal portion of LPS is Lipid A, composed of two β-(1→4)-linked N-acetyl-2-deoxy-D-glucosamine residues to which attached acyl chains serve to embed LPS in the membrane. Molecules of LPS are also capped by a distal Oantigen (OAg) polymer, which contributes to antigenic serotyping specificity for diverse species. Finally, these two segments of LPS are linked via the core oligosaccharide (OS). 4 A unifying factor between Gram-negative species is the presence of 3-deoxy-D-manno-oct-2-ulosonic acid (Kdo) in the inner portion of the core OS, a sugar present in 1−4 copies depending on the species. 5 Following its synthesis, Kdo must first be processed by the cytidine monophosphate (CMP)-Kdo synthase KdsB; this enzyme catalyzes the activation of Kdo by addition of CMP (from cytidine triphosphate [CTP]) to the anomeric position of Kdo, allowing for downstream incorporation of this activated form of Kdo into nascent LPS molecules. 6−8 Methods to label the bacterial OM have thus been developed to facilitate study of this dynamic cellular compartment via fluorescence microscopy. 9 A widespread method to visualize the OM is via treatment of intact cells with lipophilic fluorescent dyes that intercalate within the lipid bilayer. While quick and easy to use, such staining does not readily allow for distinction of labeling between the periplasmic and outer leaflets of the OM. Thus, the use of OM-targeted fluorescent proteins has also become a popular OM-labeling technology. Therein, fusion of a fluorescent protein such as mCherry or superfolder GFP (sfGFP)�capable of correctly folding in the periplasm�to an N-terminal OM lipoprotein sorting signal results in targeting of the fusion protein for insertion into the periplasmic leaflet of the OM and fluorescent labeling of the cell periphery. A benefit of such labeling is that it is genetically encoded and can thus be inducible or constitutive. However, localization of such constructs is confined to the periplasmic leaflet of the OM.
Bio-orthogonal click chemistry-mediated labeling of cellsurface LPS has emerged as a valuable addition to the OMvisualization toolbox. When grown in the presence of a synthetic Kdo variant elaborating an azido (-N 3 ) group at position 8 (8-N 3 -Kdo), diverse Gram-negative species are able to take up the synthetic sugar, incorporate it within the core OS of nascent LPS molecules (in place of native Kdo), and successfully display this azido-modified LPS on the cell surface. Accessible azide groups in this metabolically labeled LPS can then be covalently reacted to a fluorophore-linked alkyne to undergo cycloaddition, thus fluorescently labeling the outer leaflet of the OM. Initially, Cu(I)-catalyzed azide−alkyne cycloaddition (CuAAC) was used to label cells elaborating 8-N 3 -Kdo-containing LPS, 10 but this precluded further in vivo experimentation due to Cu(I)-induced cytotoxicity. To circumvent this limitation, strain-promoted azide−alkyne cycloaddition (SPAAC) has been subsequently employed to label 8-N 3 -Kdo via click chemistry in the LPS of treated bacteria without killing the cells, thus permitting live-cell imaging of the OM. 11,12 Gram-negative Myxococcus xanthus is a social bacterium with a multicellular life cycle. Within a swarm group, this bacterium exhibits coordinated behaviors of single cells. Conditions of nutrient depletion trigger the formation of aggregates in which cells are connected via OM tubes and chains of OM vesicles. These aggregates then mature into spore-filled fruiting bodies. Under such conditions, swarms also exhibit predatory behavior and can kill various bacteria and fungi to saprophytically utilize the degradation products. Groups of M. xanthus cells move via extension and retraction of type IV pili (T4P) 13 functioning in concert with secreted exopolysaccharide (EPS) and biosurfactant polysaccharide (BPS), 13−15 whereas single cells glide on surfaces via substratum-coupling and directed transport of the Agl−Glt complex at bacterial focal adhesions. 16,17 Compatible M. xanthus cells can also fuse their cell envelopes and engage in OM exchange; 3 this phenomenon results in the transfer of OM contents that can transiently complement deficiencies in various OM proteins 18 and lipids. 19 All of the above complex physiological characteristics are linked by a requirement for a highly dynamic cell surface. To ultimately be able to study the M. xanthus cell surface in a selectively activable manner, we thus sought to evaluate the feasibility of metabolic labeling and bioorthogonal click chemistry tagging of the OM in this bacterium. Herein, we describe the structure of the M. xanthus DZ2 LPS carbohydrate backbone as well as novel syntheses of Kdo variants for click-mediated metabolic labeling with an azido group at position 8 (8-N 3 -Kdo) or position 7 (7-N 3 -Kdo) of the sugar. Cells incubated with 8-N 3 -Kdo vs 7-N 3 -Kdo, respectively, displayed slower vs faster growth (relative to untreated cells), but still displayed single-cell gliding motility, T4P-dependent group motility, and fruiting body formation. While both sugars demonstrated SPAAC reactivity in vitro, only the cell surfaces of 8-N 3 -Kdo-grown live cells could be SPAAC-labeled with DBCO-linked fluorophores. While neither 8-N 3 -Kdo or 7-N 3 -Kdo were found to inhibit KdsB function, unlike the former, the latter was subsequently shown to not be processed by KdsB, precluding incorporation of 7-N 3 -Kdo into nascent LPS molecules. Despite nonaggregating cells being shown to grow faster than those that do aggregate, cells grown in the presence or absence of 7-N 3 -Kdo were found to aggregate at comparable rates, suggesting possible catabolism of 7-N 3 -Kdo by cells. These data provide important insights into the syntheses and metabolic labeling of the bacterial OM with Kdo sugar azido derivatives, including the necessary processing of these sugars by recipient cells prior to their LPS integration. This structure was confirmed via complete analysis of 2D NMR spectra (gCOSY, TOCSY, ROESY, HSQC, and HMBC) of OAg (Table 1). The methyl content of the OAg was calculated to be ∼60% as determined via the integrals of 1 H signals of the methyl group at GalNAc O-6 and N-acetate. The NMR spectra of the OAg fraction contained many additional small signals, including some associated with a rhamnan, as identified via H-6 signals around 1.3/18 ppm (H/ C).
Fractionated sample corresponding to M. xanthus DZ2 core OS was analyzed by negative-mode electrospray ionization (ESI)−mass spectrometry (MS). Observed peaks were found to fit the published structure of the core OS from M. xanthus DK1622 20 ( Figure S1B)  20 With the presence of at least one Kdo sugar, the LPS of M. xanthus DZ2 (Figure 1) should thus be amenable to metabolic labeling with Kdo azido analogues. Synthesis of 8-Azido-8-deoxy-Kdo (1), i.e., 8-N 3 -Kdo. Given its high commercial price (>10 000 USD/g), we set out to optimize the in-house synthesis of 8-azido-8-deoxy-Kdo (1), Figure 1. Schematic of the M. xanthus DZ2 LPS carbohydrate backbone. Lipid A amino groups are acylated (R) with 13-methyl-C 14 -3OH, C 16 -3OH, or 15-methyl-C 16 -3OH fatty acids (in the ratio ∼3:1:5) as previously reported. 20 However, the exact position of these fatty acids is not known. The acylation pattern of Lipid A −OH groups (★) was not determined herein.
Our synthesis of 4-azido-4-deoxy-D-arabinose (4) was inspired by an approach developed by Kosma and coworkers 24 for 4-azido-containing L-arabinose derivatives. As shown in Scheme 2, commercially available L-xylose was first subjected to a Fischer glycosylation using Dowex H + as the acid catalyst in refluxing MeOH. Accordingly, methyl Lxylopyranoside (8) was obtained in 86% yield as an inseparable α/β anomeric mixture. Tin acetal-mediated regioselective tosylation at the C4 position followed by benzoylation of the resulting diol at both C2 and C3 positions led to fully protected compound 9 in 79% yield after two steps. Importantly, attempts to use a nosyl (Ns) instead of a tosyl (Ts) group at the same position were not successful in our hands as only low yields of the target compound were obtained. S N 2 displacement reaction of the C4 tosylate by sodium azide in DMSO followed by Zempleń deacetylation provided D-arabinose derivative 10 in excellent yield (91% over two steps). 1 H NMR analysis indicated that the pyranose ring was mainly found in the 1 C 4 conformation ( 3 J H1,H2 = 5.6 Hz for the major equatorial α-anomer). Compound 10 was then subjected to acetolysis by treatment with Ac 2 O in the presence of H 2 SO 4 leading to a triacetylated derivative, which was subsequently subjected to Zempleń saponification. The resulting 4-azido-4-deoxy-D-arabinose (4) was transformed into its corresponding 7-N 3 -Kdo counterpart (2) using the Cornforth procedure (Scheme 2) in 61% yield following purification via anion exchange column. 22 NMR analysis showed that 7-N 3 -Kdo (2) was found as a mixture of furanose and pyranose isomers with the α-pyranose as the major species.  (DBCO)-amine under previously established 25 optimal SPAAC reaction conditions (30°C for 30 min, then 60°C for 30 min); this resulted in the formation of the expected adduct in each case after a total of 60 min of reaction (Scheme 3). The "click" reactivity of each sugar was next tested under conditions anticipated for in vivo labeling of M. xanthus cells, for which growth temperature is optimal at 32°C. Both 8-N 3 -Kdo and 7-N 3 -Kdo were incubated with DBCO-amine for 1 h at 32°C, with reaction aliquots removed at 5 min intervals. Thin-layer chromatography (TLC) analysis of the various aliquots indicated that the free sugars were already completely consumed after the first 5 min of the reaction ( Figures S2 and  S3). Each of the synthesized Kdo azido variants tested herein is thus capable of undergoing a SPAAC click reaction under idealized as well as in vivo labeling conditions.

8-N 3 -Kdo and 7-N 3 -Kdo Exert Different Growth Effects on M. xanthus Cells.
To incorporate exogenous azido Kdo variants into nascent LPS molecules, broth cultures are typically supplemented with the desired sugar, allowing permissive bacteria to take up the exogenous sugar and incorporate it during their metabolic cycle. 10 Cells of WT M. xanthus DZ2 were thus grown in shaking cultures in the absence or presence of 8-N 3 -Kdo or 7-N 3 -Kdo. After 24 h of shaking incubation, WT cultures supplemented with 8-N 3 -Kdo displayed ∼52% slower growth relative to control cultures in which no sugar had been added. Conversely, WT growth in the presence of 7-N 3 -Kdo was ∼32% faster than in untreated cultures ( Figure 3A). Importantly, measurements for cells from these cultures revealed comparable ranges of cell-body lengths, indicating no gross differences in morphology for cells subjected to any of the three treatments ( Figure 3B). However, compared to untreated WT cells (median length: 5.797 μm), 8-N 3 -Kdo-grown cells (median length: 6.207 μm) displayed a significantly different distribution of lengths, with values shifted toward slightly longer cells, representing a median increase of ∼7.1% in the latter; conversely, the distribution of lengths for 7-N 3 -Kdo-grown cells (median length: 5.642 μm) was determined to be comparable to that of untreated WT cells, with a median decrease of only ∼2.7% ( Figure 3B). Together, these data indicate that 8-N 3 Figure 3A), once spotted on a solid 1.5% "hard" agar matrix, the production of macroscopic swarm-edge flares was intact for both 8-N 3 -Kdoand 7-N 3 -Kdo-grown cells ( Figure 3C); this suggests that individual cells are still capable of exhibiting gliding motility. Phase-contrast microscopy imaging and single-cell motility tracking on agar pads revealed that gliding speeds were equivalent between WT cells grown in the absence and presence of 8-N 3 -Kdo, but significantly slower in 7-N 3 -Kdogrown cells ( Figure 3D). As untreated and 7-N 3 -Kdo-grown cells were of comparable lengths ( Figure 3B), this may point to differences in gliding machinery activation and/or function stemming from potential differences in the metabolic states of these cells.
On a soft 0.5% agar substratum, swarms of 7-N 3 -Kdo-grown WT cells spread as well as native WT swarms, indicating that T4P-dependent motility of cell groups was not compromised for cells having undergone this treatment ( Figure 3C,E). Conversely, T4P-dependent swarm spreading was partially inhibited in 8-N 3 -Kdo-grown WT cells ( Figure 3C,E). If 8-N 3 -Kdo is indeed incorporated into M. xanthus LPS, this may indicate a role for PEtN (attached to native Kdo at position 8) in T4P-dependent swarm biofilm spreading.
Finally, under nutrient-limiting conditions, the ability of cells from either treatment to aggregate and contribute to fruiting body formation was maintained, indicating the continued capacity of these cells to undertake a developmental cycle ( Figure 3C). Taken together, these results indicate that Kdo azido variants are compatible with the complex life cycle of M. xanthus.

Surfaces of M. xanthus Cells Grown in 8-N 3 -Kdo vs 7-N 3 -Kdo Are Differentially Labeled.
To probe the click reactivity of LPS from cells grown in 8-N 3 -Kdo and 7-N 3 -Kdo, cells grown in either sugar were treated with DBCO-linked fluorescent dyes, namely (i) AFDye 488 (DBCO-AF488), (ii) polyethylene glycol (PEG) 4 -5/6-Texas Red (DBCO-TxRed), and (iii) Cy5 (DBCO-Cy5). This was done in an effort to fluorescently label cell-surface LPS elaborating exposed -N 3 groups via a SPAAC click reaction. As evidenced via live-cell fluorescence microscopy, cells grown in the presence of 8-N 3 -Kdo displayed bright fluorescent peripheral marking with all three DBCO-linked fluorophores, indicative of successfully click-labeled LPS in these cells ( Figure 4A). Importantly, these cells were still motile following the various click and wash steps, demonstrating their continued viability ( Figure 4B). Unexpectedly, cells from 7-N 3 -Kdo-supplemented cultures treated with any of the three DBCO-linked dyes did not display any peripheral fluorescence ( Figure 4A), despite the in vitro SPAAC reactivity of 7-N 3 -Kdo ( Figure S3); this suggests that 7-N 3 -Kdo is not taken up by cells and/or cannot be incorporated into LPS.

M. xanthus Encodes Numerous Major Facilitator Superfamily Proteins. Both E. coli and Klebsiella pneumoniae
were previously shown to be permissive for metabolic labeling via 8-N 3 -Kdo incorporation, with both species known to contain the nan sialic acid uptake operon, whereas native Pseudomonas aeruginosa, which does not contain the nan operon, could not be so labeled. Specific expression of sialic acid transporter NanT�a major facilitator superfamily (MFS) protein�was shown to mediate 8-N 3 -Kdo transport across the IM into the cytoplasm in E. coli; moreover, heterologous expression of E. coli NanT in P. aeruginosa resulted in the latter cells being metabolically labeled with 8-N 3 -Kdo. 26 Given the role of MFS transporter NanT in mediating Kdo azidoanalogue uptake, we thus scanned the M. xanthus genome 27 for NanT homologues as well as proteins containing known Pfam domains MFS_1, MFS_1_like, MFS_2, MFS_3, MFS_4, and MFS_5. This analysis revealed 36 putative MFS transporter proteins encoded by the bacterium (Table S1). Previous RNAseq transcriptomic profiling 28,29 revealed these genes to be expressed during the M. xanthus life cycle (Table S1). While identification of the specific transporter gene(s) responsible for Kdo azido-variant uptake is beyond the scope of the current paper, these data are consistent with diverse transport capabilities for M. xanthus, which are reflective of its broad metabolic capacity as a soil bacterium.

8-N 3 -Kdo (But Not 7-N 3 -Kdo) Is a Substrate for CMP-Kdo Synthase KdsB.
Given the requirement for Kdo to be processed into an activated form by KdsB prior to its incorporation into LPS 6−8 �via the addition of CMP (from CTP), releasing pyrophosphate [PP i ] in the process�we thus carried out an enzymatic assay to determine the suitability of 8-N 3 -Kdo and 7-N 3 -Kdo as substrates for KdsB. In the presence of CTP, KdsB was incubated with either Kdo, 8-N 3 -Kdo, or 7-N 3 -Kdo, with ensuant PP i release quantified via chromogenic reaction with eikonogen reagent followed by spectrophotometric determination of absorbance at 595 nm. 30 Kdo and 8-N 3 -Kdo were shown to be substrates for KdsB, with the latter more inefficiently processed; conversely, 7-N 3 -Kdo could not be processed by the enzyme under these conditions ( Figure  5A). We then forced the experimental conditions to an even greater extent to see if 7-N 3 -Kdo was indeed a substrate, increasing its concentration to 5000 μM (up to ∼12× the K M for Kdo) and the reaction time up to 3 h. However, no reaction was observed ( Figure 5A). The lack of KdsB activity toward 7-N 3 -Kdo may indicate that the hydroxyl group at C7 of Kdo plays a crucial role in the interaction of Kdo with KdsB. Alternatively, the presence of a bulkier -N 3 group at C7 of Kdo may not allow 7-N 3 -Kdo to properly situate in the binding pocket of KdsB 31 (as opposed to Kdo with a -N 3 group at C8 which can likely extend further into the open cavity) ( Figure  S4), indicating that the enzyme may be more permissive to Kdo modifications at C8 versus C7.
Using varying concentrations of Kdo, 8-N 3 -Kdo and 7-N 3 -Kdo (0−5000 μM), we determined the steady-state kinetic parameters of purified KdsB in the presence of each sugar (via monitoring of PP i release as described above), with the enzyme displaying Michaelis−Menten parameters in the presence of Kdo and 8-N 3 -Kdo (as per previous reports 30,32 ); no PP i release was detected in the presence of 7-N 3 -Kdo ( Figure 5B). However, native Kdo still proved to be a more efficient acceptor for CMP compared to 8-N 3 -Kdo ( Figure 5B).
Since each Kdo azido variant was not processed by KdsB to the extent of native Kdo, the inhibitory potential for each analogue on KdsB was tested. KdsB was first preincubated with high concentrations of 8-N 3 -Kdo or 7-N 3 -Kdo (5000 μM), after which substrate mix was added containing CTP and native Kdo at concentrations set at ∼0.5 and 1 K M to favor the detection of substrate competitors. After incubation for 30 min, reactions were quenched with ethanol, followed by detection of released PP i . Under these conditions, neither of the Kdo azido analogues had an inhibitory effect on KdsB ( Figure 5C). For 8-N 3 -Kdo in particular, as it is a substrate for KdsB ( Figure 5A,B), an even larger amount of PPi was released into the reaction mixture (compared to native Kdo alone) ( Figure 5C), suggesting that KdsB is capable of interchangeably metabolizing both Kdo and 8-N 3 -Kdo.
The lack of surface click labeling with DBCO-linked fluorophores for 7-N 3 -Kdo-grown cells ( Figure 5A) is consistent with an inability of this particular Kdo azido analogue to be processed by KdsB, thus precluding its incorporation into nascent LPS molecules. This is likely a broadly applicable finding given the extensive sequence conservation between KdsB proteins from diverse proteobacteria, in particular with respect to key active-site residues implicated in Kdo processing 31 ( Figure S5).

7-N 3 -Kdo May Be Catabolized by M. xanthus Cells.
Within the context of 7-N 3 -Kdo not being incorporated into LPS molecules (Figure 4), we thus revisited the faster growth phenotype of M. xanthus DZ2 grown in the presence of this Kdo azido analogue. Conventional wisdom in working with M. xanthus holds that nonaggregating strains grow faster than those that aggregate. To explicitly test this assertion, we compared the growth curves of WT M. xanthus DZ2 in liquid culture with those of the nonaggregating 13 isogenic mutant strains ΔwzaX (deficient in EPS secretion) and ΩpilA (deficient in T4P assembly and EPS secretion). Both ΔwzaX and ΩpilA were found to achieve exponential-phase growth, and reach stationary phase, sooner than WT ( Figure 6A), consistent with a lack of M. xanthus cell aggregation resulting in faster culture growth. We thus compared the autoaggregative profiles 13 of WT cells grown in the absence and presence of 7-N 3 -Kdo to probe for reduced clumping in the latter. However, cells grown with 7-N 3 -Kdo clumped and sedimented in cuvettes at comparable rates to untreated cells ( Figure 6B), indicating that altered aggregation was not a viable explanation for accelerated growth in the presence of 7-N 3 -Kdo. To support this contention, we compared the growth of the nonaggregating 13 ΩpilA mutant strain of M. xanthus in the absence/presence of Kdo azido analogues. Similar to WT cells ( Figure 3A), ΩpilA cells grew ∼63% slower in the presence of 8-N 3 -Kdo ( Figure 6C). Importantly, ΩpilA mutant cells also grew ∼17% faster in the presence of 7-N 3 -Kdo compared to untreated cells (analogous to WT cells) ( Figure  6C), despite their inability to aggregate. 13 Therefore, the higher density of M. xanthus cells in the presence of 7-N 3 -Kdo may be due to catabolism of this particular supplemented Kdo azido variant resulting in accelerated growth.

■ CONCLUSIONS
We reported the first-ever synthesis of a Kdo variant with a substitution at carbon 7, 7-azido-7-deoxy-Kdo (7-N 3 -Kdo), in addition to greatly improving the synthesis of 8-azido-8-deoxy-Kdo (8-N 3 -Kdo). The eight-carbon skeleton of these Kdo analogues was efficiently built via the Cornforth homologation procedure from the corresponding azido-containing D-arabino precursors, opening an efficient synthetic route to the more cost-effective generation of modified Kdo sugars.
Cells of M. xanthus grown with 8-N 3 -Kdo exhibited slower growth. However, given the capacity of these cells to manifest a multicellular life cycle, this would suggest that depletion of PEtN within the context of the bacterium's total cell-surface Kdo pool is not overly deleterious to the viability of the bacterium. However, the depletion of -PEtN groups and/or the non-native presence of -N 3 groups deep in the inner core OS would be expected to modify the overall cell-surface properties to a certain degree, which may help explain various phenotypic differences ( Figure 3). As cases-in-point, cells with different physical surface properties display differences in single-cell gliding speeds, gliding reversal frequencies, dye-binding profiles, tolerance to antibiotics and oxidative-stress agents, and biofilm organization, as well as T4P formation, stability, and positioning. 13,14,33 The effect of 7-N 3 -Kdo is more perplexing. While it clearly cannot be detected on the cell surface (given its inability to be processed by KdsB and thus not activated for integration into an LPS molecule), in its presence, aggregating and nonaggregating strains alike grew faster than untreated cells. Cells grown with 7-N 3 -Kdo also exhibited slower single-cell gliding motility, despite being of equivalent length to untreated cells.
As cell-surface LPS should be identical between WT cells grown with and without 7-N 3 -Kdo, the overall cell-surface properties between such samples should be comparable. Therefore, differences in cell-surface properties are an unlikely explanation for altered gliding speed. This may instead point to an as-yet-unknown metabolic effect that affects the production and/or function of the gliding motility machinery. Considering this and the faster growth in the presence of 7-N 3 -Kdo (a phenotype that is not due to decreased aggregation), this may point to 7-N 3 -Kdo being used as an energy source by the cells, thus favoring cell growth. Such a contention does however require further investigation to more concretely support it.
The X-ray crystal structure of KdsB may provide insights into the ability of the enzyme to process 8-N 3 -Kdo, but not 7-N 3 -Kdo. Although a -N 3 at Kdo carbon 8 would extend upward into the cavity of the binding pocket, a -N 3 at Kdo carbon 7 would likely abut against the wall of the binding pocket, disfavoring the required stereochemistry at the active site ( Figure S4). Functional Kdo analogues (i.e., those that can be processed by KdsB and have CMP added) have been previously shown with modifications at carbon 3, 5, or 8. 34 Given that CMP is added to carbon 2, and considering the unoccupied space in the binding pocket ( Figure S4), carbon 4 may also support a −N 3 modification without impeding addition of CMP.
Ultimately, our data provide important insights into the improved synthesis of valuable metabolic labels and establish a basis for the elucidation of fundamental principles of OM dynamism in bacteria. ■ METHODS General Synthesis Methods. All of the starting materials and reagents were purchased from commercial sources and used without further purification. Air-and water-sensitive reactions were conducted in oven-dried glassware under an inert atmosphere in anhydrous solvents, which were prepared by supplying the solvents over heat-gun activated 4 Å molecular sieves. These solvents were introduced to the reaction through a dried, argon-filled syringe. Reactions were monitored by TLC with silica gel 60 F 254 0.25 mm precoated aluminum foil plates. Compounds were visualized using UV 254 and orcinol (1 mg·mL −1 ) in a 10% aqueous solution of H 2 SO 4 or Hanessian's stain [2 g of Ce(SO 4 ) 4 (NH 4 ) 4 , 5 g of MoO 4 (NH 4 ) 2 , 200 mL of H 2 O, and 20 mL of H 2 SO 4 ] with heating. Normal-phase flash column chromatography was performed on silica gel 60 Å (15−40 μm). Reversed-phase flash column chromatography was performed on C 18 silica gel (fully capped, 25−40 μm). Anion exchange column chromatography was performed on Serdolit CG-400 I (Cl − form). NMR spectra were recorded at 297 K in the mentioned deuterated solvents with 400 or 600 MHz instruments, employing standard software given by the manufacturer. 1 H and 13 C NMR spectra were referenced to TMS (δ H = δ C = 0.00 ppm) and acetone (δ H = 2.22 ppm; δ C = 30.9 ppm) for spectra in D 2 O. Assignments were based on 1 H, 13 C, HSQC, and COSY experiments. All of the 13 C NMR experiments were 1 Hdecoupled. High-resolution mass spectra (HRMS) were recorded on an ESI-Q-TOF mass spectrometer. All NMR analyses for molecule verifications have been provided ( Figures  S6−S33) (5). D-Arabinose (2.0 g, 13.3 mmol, 1.0 equiv) was suspended in MeOH (48 mL) at 0°C. A freshly prepared solution of AcCl (1.1 mL, 15.5 mmol, 1.16 equiv) in MeOH (9.5 mL) at 0°C was added dropwise to the latter suspension. The mixture was warmed up to rt and stirred for 3 h at rt. TLC analysis (DCM/MeOH 8:2) showed full conversion of the starting material. The reaction was then quenched with pyridine (4 mL), and the resulting mixture was co-evaporated with toluene. The crude was then dissolved in dry pyridine (41 mL) under argon at 0°C. A solution of TsCl (3.8 g, 20.0 mmol, 1.5 equiv) in anhydrous pyridine (41 mL) was then added dropwise. The mixture was warmed up to rt and stirred for 16 h. After verification of the complete consumption of the starting material by TLC (100% EtOAc), the reaction was quenched by adding ice water (3 g) and stirring for 30 min. Water (50 mL) was added, and the aqueous phase was extracted with EtOAc (3 × 50 mL). The organic layers were combined, washed with brine (25 mL), and dried over MgSO 4 . The solvents were evaporated under reduced pressure. The resulting crude was then dissolved in dry DCM (80 mL) under argon. Ac 2 O (12.6 mL, 133 mmol, 10 equiv) was added, and the mixture was cooled down to 0°C. Concentrated H 2 SO 4 (710 μL, 13.3 mmol, 1.0 equiv) was added dropwise. The mixture was stirred at 0°C for 20 min. TLC analysis (100% EtOAc) showed total conversion of the starting material. The reaction mixture was poured on a mixture of ice/water (100 mL). DCM was then evaporated under reduced pressure. The resulting aqueous phase was extracted with EtOAc (3 × 100 mL). The organic phases were combined, washed with a saturated aqueous solution of NaHCO 3 (2 × 75 mL), dried over MgSO 4 , and the solvents were evaporated under reduced pressure. The crude was then purified by silica gel column chromatography (100% hexanes to hexanes/EtOAc 5:5) to give compound 5 (4.0 g, 70% over three steps, α/β = 6:1) as a pale yellow oil. Analytical data agreed with those previously reported. 21 Synthesis of 1,2,3-Tri-O-acetyl-5-azido-5-deoxy-D-arabinofuranose (6). A solution of compound 5 (3.2 g, 7.5 mmol, 1.0 equiv) and NaN 3 (1.84 g, 28.4 mmol, 3.8 equiv) in dry DMSO under argon was stirred at 80°C for 16 h. After verification of the consumption of the starting material by TLC (hexanes/acetone 6:4), the reaction mixture was diluted with DCM (100 mL). The organic phase was washed with H 2 O (2 × 100 mL), dried over MgSO 4 , and the solvents were evaporated under reduced pressure. The crude was then purified by silica gel column chromatography (hexanes/EtOAc 8:2 to 4:6) to give compound 6 (1.6 g, 71%, α/β = 6:1) as a pale yellow oil. Analytical data agreed with those previously reported. 21 Ammonium 8-Azido-3,8-dideoxy-D-manno-oct-2-ulopyranosylonate (1). To a solution of compound 6 (455 mg, 1.51 mmol, 1.0 equiv) in MeOH (9.4 mL) was added a 25% w/w solution of NaOMe in MeOH (65.3 μL, 0.3 mmol, 0.2 equiv). The resulting mixture was stirred at rt for 30 min. TLC analysis (DCM/MeOH 9:1) showed full conversion of the starting material. The reaction was then quenched by the addition of Dowex H + resin until pH ≈ 7 was reached. The suspension was then filtered over Celite and the solvents were evaporated under reduced pressure to give 5-azido-5-deoxy-Darabinose (3), which was directly used in the next step without further purification. Azide 3 (265 mg, 1.5 mmol, 1.0 equiv) was dissolved in distilled H 2 O (4.5 mL) containing Na 2 CO 3 (394 mg, 3.7 mmol, 2.5 equiv). Oxaloacetic acid (242 mg, 1.8 mmol, 1.2 equiv) was added in small portions over 5 min. After adding a few drops of a 10 N aqueous NaOH solution until the pH increased to 11.0 (as checked with a pH meter), the reaction mixture was stirred at rt for 2 h. Then, the pH was decreased to 5.0 using AcOH. Catalytic NiCl 2 ·6H 2 O (0.01 equiv) was added and the mixture was stirred at 50°C for 1 h. Then, the pH was increased again to 8.0 with a saturated aqueous solution of NH 4 OH. After cooling the mixture to rt, it was passed through an anion exchange resin column CG-400 (HCO 3 − ). It was first eluted with H 2 O to remove the excess of azide 3 and then eluted with aqueous NH 4 HCO 3 (0.05 N to 0.30 N). The fractions were pooled according to TLC analysis (CHCl 3 /MeOH/H 2 O 10:10:3 as revealed with Hanessian's stain) and freeze-dried to provide ammonium Kdo derivative 1 (163 mg, 39%) as a white amorphous solid. Analytical data agreed with those previously reported. 10,22 Synthesis of Methyl L-Xylopyranoside (8). L-Xylose (1.0 g, 6.7 mmol, 1.0 equiv) was dissolved in anhydrous MeOH (200 mL) under Ar at rt. Dowex H + resin (500 mg) was added, and the mixture was refluxed for 16 h. The reaction mixture was cooled down to rt and then filtered over Celite. The solvents were evaporated under reduced pressure. The residue (anhydrous loaded) was purified through silica gel flash chromatography (DCM/MeOH/acetone 16 (9). A Dean-stark apparatus was fixed to a flask in which Bu 2 SnO (1.4 g, 5.7 mmol, 1.1 equiv) and methyl xylopyranoside 8 (921 mg, 5.6 mmol, 1.0 equiv) were suspended in anhydrous toluene (18 mL). After the mixture was refluxed for 6 h, it was cooled down to rt and the solvents were evaporated under reduced pressure. To a solution of this residue in anhydrous THF (8.4 mL), a solution of TsCl (1.3 g, 6.8 mmol, 1.2 equiv) in anhydrous THF (11 mL) was added dropwise under Ar in the presence of DMAP (1 mg, 5 μmol, 0.001 equiv) and left to stir at rt for 96 h. Following reaction completion according to TLC (100% EtOAc), the solvents were evaporated under reduced pressure. The residue (dryloaded) was purified using silica gel flash chromatography. First, the column was eluted with toluene to remove most of the dibutyltin salts, and then with the eluent system (100% EtOAc) to give methyl 4-O-para-toluenesulfonyl-α-L-xylopyranoside contaminated with tin salts. The latter crude diol was dissolved in anhydrous pyridine (17.5 mL) under Ar. After cooling the mixture to 0°C, BzCl (2.0 mL, 18 mmol, 3.1 equiv) was added dropwise followed by DMAP (7.1 mg, 0.058 mmol, 0.01 equiv). The reaction mixture was warmed to rt and stirred for 16 h. After reaction completion according to TLC (hexanes/EtOAc 5:5), it was diluted with EtOAc (50 mL). The organic phase was washed with a 1 N aqueous HCl solution (2 × 50 mL), followed by a saturated aqueous NaHCO 3 solution (50 mL), and then with brine (50 mL). The organic layers were dried using MgSO 4 and then evaporated under reduced pressure. The resulting residue was purified by silica gel flash chromatography (hexanes/EtOAc 9:1 to 6:4) to give derivative 9 (2.3 g, 79% over 2 steps, α/β = 5:2) as a white amorphous solid: R f 0.4 (hexanes/EtOAc 6:4); 1 (10). To a solution of derivative 9 (17.5 g, 33.3 mmol, 1.0 equiv) in anhydrous DMSO (50 mL), NaN 3 (8.1 g, 125 mmol, 2.5 equiv) was added, and the mixture was stirred under Ar at 90°C overnight. Then, the reaction mixture was diluted with DCM (200 mL) and the organic phase was washed with distilled water (2 × 100 mL) and a saturated aqueous NaHCO 3 solution (100 mL). The organic layer was dried using MgSO 4 , and the solvents were evaporated under reduced pressure. The resulting residue was purified by silica gel flash chromatography (hexanes/EtOAc 7:3 to 1:1) to provide methyl 4-azido-2,3-di-O-benzoyl-4-deoxy-D-arabinopyranoside (S1, 14.0 g, quantitative, α/β = 3.5:1.5) as a white amorphous solid: R f 0. Synthesis of 4-Azido-4-deoxy-D-arabinopyranose (4). To a solution of diol 10 (5.2 g, 27 mmol, 1.0 equiv) dissolved in DCM (275 mL), Ac 2 O (26 mL, 275 mmol, 10.0 equiv) was added and the reaction mixture was cooled down to 0°C. A solution of concentrated H 2 SO 4 (1.5 mL, 27 mmol, 1.0 equiv) was added, and the reaction was warmed to rt and stirred for 20 min. After diluting the mixture with DCM (50 mL), the organic phase was washed with a saturated aqueous NaHCO 3 solution (3 × 100 mL) and brine (100 mL). The combined organic layers were dried over MgSO 4 , and the solvents were evaporated under reduced pressure. The residue was purified by silica gel flash chromatography (hexanes/EtOAc 8:2 to 1:1) to give 1,2,3-tri-O-acetyl-4-azido-4-deoxy-D-arabinopyranoside (S2, 3.8 g, 50%, α/β 1.5:3.5) as a white amorphous solid: R f 0.3 (Hex/EtOAc 6:4); 1 H NMR (600 MHz, CDCl 3 , βanomer) δ 6.31 (d, 3 4 v/v, 125 mL), a NaOMe solution was added (25 wt % in MeOH, 577 μL, 2.52 mmol, 0.2 equiv), and the reaction was stirred overnight at rt. The reaction mixture was quenched with Dowex H + resin until neutrality was reached. The resin was filtered off through Celite and the solvents were evaporated under reduced pressure. The residue was purified by reversed-phase (C 18  Ammonium 7-Azido-3,7-dideoxy-D-manno-oct-2-ulopyranosylonate (2). Azide 4 (350 mg, 2.0 mmol, 1.0 equiv) was dissolved in distilled H 2 O (6.0 mL) containing Na 2 CO 3 (526 mg, 4.9 mmol, 2.5 equiv). Oxaloacetic acid (317 mg, 2.4 mmol, 1.2 equiv) was added in small portions over 5 min. After adding a few drops of a 10 N aqueous NaOH solution until the pH increased to 11.0 (as checked with a pH meter), the reaction mixture was stirred at rt for 2 h. Then, the pH was decreased to 5.0 using AcOH, catalytic NiCl 2 ·6H 2 O (0.01 equiv) was added, and the mixture was stirred at 50°C for 1 h. After cooling the mixture to rt, it was passed through an anion exchange resin column CG-400 (HCO 3 − ). It was first eluted with H 2 O to remove the excess of azide 4 and then eluted with aqueous NH 4 HCO 3 (0.05 to 0.30 N). The fractions were pooled according to TLC analysis (CHCl 3 /MeOH/H 2 O 10:10:3; revealed with Hanessian's stain) and freeze-dried to provide ammonium Kdo derivative 2 (340 mg, 61%) as a white amorphous solid. According to NMR, compound 2 was obtained as a mixture of f urano and pyrano anomers with αpyranose as the major form.  ) at an initial OD 600 of 0.02. Tubes were closed with sliding plastic caps, with the caps sealed to the tubes using Parafilm. Culture tubes were then inclined on a tube rack and incubated with shaking for 24 h. To remove culture rings accumulating on the side of the glass tubes, tubes were briefly mixed via vortex every 6−8 h to dissociate the adhered ring and ensure cells remained in the liquid phase. Upon harvest, 100 μL of culture volume was diluted in 900 μL of TPM to help dissociate clumps, followed by reading of the OD 600 in a disposable polystyrene cuvette to determine the final culture density using a spectrophotometer.

Synthesis of Methyl 4-Azido-4-deoxy-D-arabinopyranoside
Growth Curve Analysis. Wild-type (DZ2), 35 ΔwzaX (TM469), 36 and ΩpilA (TM293) 13 strains were initially grown overnight in 12.5 mL CYE cultures in a shaking incubator (32°C , 220 rpm, 125 mL flask). The next day, each strain was subcultured into a 150 mL CYE starting volume (in a 1 L flask) to a starting OD 600 of 0.05, and grown for 86 h (32°C, 220 rpm). Culture density was measured every 4 h via determination of the OD 600 . Prior to cultures reaching an OD 600 of 1.0, 1 mL of culture volume was removed at each time point and transferred to a disposable spectrophotometer cuvette; cuvettes were then inserted into foam microtube racks, floated in a Bransonic 32 Ultrasonic cleaner bath (Branson), sonicated for 30 s to break up cell aggregates, after which OD 600 was determined. At each time point following culture attainment of an OD 600 of 1.0, 100 μL of culture was transferred to a cuvette, and diluted with 900 μL of TPM buffer (10 mM Tris-HCl, pH 7.6, 8 mM MgSO 4 , and 1 mM KH 2 PO 4 ), prior to sonication and OD 600 determination as described.
Extraction of LPS for Backbone and OAg Structure Determination. LPS was extracted by stirring 2−3 g of lyophilized cell mass in 200 mL of 45% aqueous phenol at 75°C for 20 min. The ensuing mixture was dialyzed for several days in running tap water, after which acetic acid was added to 10% final concentration. The precipitate was then removed by centrifugation at 8000 rpm for 30 min, followed by dialysis against distilled water and lyophilization. This freeze-dried sample was then dissolved in 10 mL of ddH 2 O, after which AcOH was added to a final concentration of 2%. The sample was heated for 2 h at 100°C until clear solution and precipitate formed, then cooled. The precipitate was removed by centrifugation at 10 000 rpm, followed by product separation on a Biogel P6 column (2.5 × 60 cm 2 ) in 1% aqueous AcOH, then lyophilized.
To prepare OAg for structure determination, LPS was hydrolyzed by 2% aqueous AcOH (100°C, 1−2 h, until a precipitate was formed). The precipitate was removed by centrifugation and then separated on a Biogel P6 column (2.5 × 60 cm 2 ) in 1% aqueous AcOH with a refractive index detector.
Anion Exchange Chromatography. Sample up to 50 mg was injected into a HiTrap Q column (Amersham, two columns, by 5 mL each, connected in series) in water at 3 mL· min −1 , washed with water for 5 min, and then eluted with a linear gradient from water to 1 M aqueous NaCl over 1 h with UV detection at 220 nm and spot test on silica TLC plate with development by dipping in 5% H 2 SO 4 in ethanol and heating with heat gun until brown spots become visible. Samples were desalted on Sephadex G-15 column (1.6 × 60 cm 2 ) in 1% aqueous AcOH with a refractive index detector.
NMR Spectroscopy for LPS Structure Determination. For LPS structure determination, NMR experiments were carried out on a Bruker AVANCE III 600 MHz ( 1 H) spectrometer with 5 mm Z-gradient probe with acetone internal reference (2.225 ppm for 1 H and 31.45 ppm for 13 C) using standard pulse sequences cosygpprqf (gCOSY), mlevphpr (TOCSY, mixing time 120 ms), roesyphpr (ROESY, mixing time 500 ms), hsqcedetgp (HSQC), hsqcetgpml (HSQC-TOCSY, 80 ms TOCSY delay), and hmbcgplpndqf (HMBC, 100 ms long-range transfer delay). The resolution was kept <3 Hz/pt in F2 in proton−proton correlations and <5 Hz/pt in F2 of H−C correlations. The spectra were processed and analyzed using the Bruker Topspin 2.1 program. Monosaccharides were identified by COSY, TOCSY, and NOESY cross-peak patterns and 13 C NMR chemical shifts. Amino group location was concluded from high-field signal position of aminated carbons (CH at 45−60 ppm). Connections between monosaccharides were determined from transglycosidic NOE and HMBC correlations.
Mass Spectrometry for LPS Structure Determination. For LPS analysis, ESI-MS was obtained using a Waters SQ Detector 2 instrument. Samples were injected in 50% aqueous MeCN with 0.1% TFA. Cone voltage was set to 60 V for nondestructive MS or to 160 V to observe fragmentation of polymers.
Determination of Neutral and Amino Sugars as Alditol Acetates. Monosaccharides were detected as reduced and acetylated derivatives (alditol acetates). Polysaccharide sample (0.2−1.0 mg) with inositol internal standard was hydrolyzed with 3 M aqueous TFA (120°C, 3 h), dried, reduced with NaBD 4 , reagent-destroyed with 0.5 mL of AcOH, solution-dried under the stream of air, dried twice with the addition of MeOH (1 mL), acetylated with 0.2 mL of Ac 2 O and 0.2 mL of pyridine for 30 min at 100°C, dried, and analyzed by GC-MS on Thermo Trace 1310 instrument with ITQ1100 ion trap detector, capillary column HP-5, 160−260°C by 4°C·min −1 . Phenotypic Analyses. For cultures grown in glass tubes, cells were sedimented via centrifuge (4000 × g, 5 min) and resuspended in TPM buffer to an OD 600 of 2.0, followed by spotting 5 μL of each resuspension to the appropriate matrix and incubation at 32°C of the Parafilm-sealed plate. To analyze gliding motility flare formation, cells were spotted to 15  Phenotype plates were imaged on an Olympus SZX16 stereoscope with ILLT base. To image gliding flares, objective 2× and zoom 8× were used with illumination between the oblique and brightfield lenses. T4P-dependent swarm spreading was captured at objective 0.5×, zoom 0.7×, with dark-field illumination. Fruiting bodies were imaged using objective 0.5×, zoom 2×, with oblique illumination. Measurements of T4Pdependent swarm surface area were obtained using Olympus CellSens software with the three-point circle method.
SPAAC "Click" Chemistry Labeling of Cells Grown with Kdo Azido Variants. For cultures grown in glass tubes with Kdo azido variants, the calculated culture volume was removed and sedimented in a centrifuge (4000 × g, 5 min) so that resuspension in 10 μL would yield an OD 600 of 1.0. Cell pellets were then twice washed with TPM buffer and sedimented (4000 × g, 5 min). Pellets were then resuspended either in 10 μL of TPM (for nonfluorophore controls) or 10 μL of DBCO-fluorophore solution (30 μM stock solution prepared in TPM). Resuspended cells were incubated in the dark with rocking (170 rpm) at 37°C for 30 min to promote SPAAC click chemistry reactions between DBCO and the -N 3 groups on Kdo analogues; the cells were then sedimented via centrifuge and washed twice with TPM as described above. For live-cell fluorescence microscopy of clicked cells, the final pellet was resuspended in 10 μL of CYE broth.
Single-Cell Motility and Fluorescence Analysis. For phase-contrast and fluorescence microscopy on agar pads, cells from (non)clicked samples were sedimented and resuspended in TPM buffer to OD 600 1.0, spotted (3 μL) on a glass coverslip, and then overlaid with a pad of 1.5% agar prepared with (TPM). For motility analysis, the cells were left to adhere for 15 min prior to imaging at 32°C using an Axio Observer 7 fluorescence microscope (Zeiss) with a 40× Plan Apochromat 1.3 oil objective VIS-IR M27 for single-cell motility and objective α Plan-Apochromat 100×/1.46 Oil DIC (UV) M27 for fluorescence analysis, with Axiocam 512 camera, TL LED as light source, and Zen Black software (Zeiss). DBCO-AF488 fluorescence was imaged using the 38 green fluorescent protein reflector, filter excitation wavelength 450−490, and filter emission wavelength 500−550. DBCO-TxRed fluorescence was imaged using the 92 HE/DAPI/GFP/mCherry reflector, filter excitation wavelength 583−600, and filter emission wavelength 617−758. DBCO-Cy5 fluorescence was imaged using the 50 Cy5 reflector, filter excitation wavelength 625− 655, and filter emission wavelength 665−715. Images for single-cell gliding speed analysis were taken at 45 s intervals for 62 frames. Cell gliding speeds were calculated using the MicrobeJ module for Fiji. 37 Images for fluorescence analysis were taken at 30 s intervals. Gliding cell montages were generated using Fiji.
Cell Length Analysis. Images to be analyzed for cell length were acquired as part of the initial frames for the movies acquired for the determination of gliding speed (see above). Prior to analysis, images were processed in FIJI to optimize cell detection via the "Enhance Contrast" function (0.3%), followed by background subtraction (Rolling ball radius: 10.0 pixels, light background). Cells were measured using the FIJI MicrobeJ module. 37 The parameters used for length detection were set to "Basic", with the "MaxEntropy" method applied to threshold the images. The area selected for cell detection was set to 2−15 μm. Automatically detected objects were manually curated to remove any selections that were either background artifacts or merged cells to ensure only individual cells were analyzed. The length of each detected and retained object was then recorded in the results panel under the column "SHAPE.length" in micrometers.
Autoaggregation Testing. Real-time autoaggregation analysis was carried out as previously described. 13 In brief, for WT cells grown with(out) 7-N 3 -Kdo, resuspensions from 10 mL cultures (grown for 24 h) were adjusted to OD 600 0.5 in CYE medium, with 1 mL of this resuspension transferred to a disposable spectrophotometer cuvette. Samples were strongly aspirated/ejected in the cuvette for 10 s via micropipette, followed by immediate OD 600 determination (t = 0). Subsequent OD 600 values were read every 10 min thereafter, up to 90 min. Cuvettes were covered and left undisturbed in a cuvette box atop the lab bench in between time points. All OD 600 values were normalized against the initial OD 600 reading obtained at t = 0 for each sample.
KdsB Enzyme Analysis of Kdo Azido Analogues. Corynebacterium glutamicum R163 harboring the expression plasmid pJKB72 was used to produce KdsB from Escherichia coli K-12 fused to a C-terminal histidine tag. 38 Protein was produced and purified through immobilized metal ion affinity chromatography using the protocol described by Mamat and co-workers. 39 The preparation was dialyzed (MWCO 6−8 kDa membrane) against Tris-HCl (20 mM) pH 7.5, NaCl (100 mM), MgCl 2 (5 mM), and 2-mercaptoethanol (2 mM) at 4°C and concentrated with a centrifugal filter (MWCO 10 kDa). Protein was quantified with the Bradford protein assay and purity (>95%) was evaluated with a 12% SDS-PAGE gel. The final preparation was stored at −80°C. To monitor KdsB enzymatic activity to evaluate Kdo azido analogues as potential substrates or inhibitors, we relied on the eikenogen assay described by Yi and co-workers. 30 This colorimetric assay quantifies pyrophosphate (PP i ) in the reaction mixture. The assay was performed in a 96-well clear, round bottom polystyrene microplate format. The reaction buffer consisted of glycine−NaOH (100 mM) pH 10.0 and MgCl 2 (5 mM). First, 10 μL of KdsB and 10 μL of Kdo/Kdo azido analogues in reaction buffer were added to the corresponding wells and preincubated for 10 min at room temperature. Then, 20 μL of CTP in reaction buffer was added. Each reaction mixture (40 μL) contained 100 nM KdsB and varying concentrations of CTP, Kdo, and Kdo azido analogues. The reaction was incubated at room temperature for the indicated times and an equal volume (40 μL) of ice-cold absolute ethanol was added to quench the reaction. Then, assay reagents were added as reported and the plate was incubated at room temperature for 20 min to allow color to develop. Absorbance was measured at 595 nm. Data were fitted to the Michaelis−Menten model using nonlinear regression.
Genomic Analyses. A dataset of 3,662 reference bacterial genomes was downloaded using the Prokaryotes.txt file on 2021-12-07 from NCBI. The Translated CDS files belonging to phylum Proteobacteria were subjected to offline hmmscan (E-value cutoff of 1 × 10 −5 ) against the Pfam-A v34.0 database (downloaded: 2021-03-24), followed by parsing via hmmscanparser.sh, and arranging in the form of protein architecture using in-house scripts. KdsB proteins were identified using the Pfam domain CTP_transf_3 (PF02348). From the total output, 38 KdsB proteins were selected from well-known organisms belonging to the phylum Proteobacteria. Alignments were performed using MUSCLE 40 and alignment was visualized using CHROMA. 41 The NanT protein from E. coli (P41036.2) was used as a query to search for homologs in the M. xanthus genome using an E-value cutoff of 1 × 10 −5 , which led to the identification of five NanT homologs in M. xanthus DZ2. Translated CDS files were subjected to offline hmmscan (E-value cutoff of 1 × 10 −5 ) against the Pfam-A v34.0 database, followed by parsing via hmmscan-parser.sh, and arranging in the form of protein architecture using in-house scripts. Proteins with Pfam domains MFS_1 (PF07690), MFS_1_like (PF12832), MFS_2 (PF13347), MFS_3 (PF05977), MFS_4 (PF06779), and MFS_5 (PF05631) associated with MFS transporters were identified from this output, leading to the identification of 31 putative MFS proteins.
Putative MFS transporters identified in M. xanthus with associated RNAseq transcriptomic profiling over the course of development (Table S1) (XLSX) 1