Abstract
Gram-negative bacteria surround their cytoplasmic membrane with a peptidoglycan (PG) cell wall and an outer membrane (OM) with an outer leaflet composed of lipopolysaccharide (LPS)1. This complex envelope presents a formidable barrier to drug entry and is a major determinant of the intrinsic antibiotic resistance of these organisms2. The biogenesis pathways that build the surface are also targets of many of our most effective antibacterial therapies3. Understanding the molecular mechanisms underlying the assembly of the Gram-negative envelope therefore promises to aid the development of new treatments effective against the growing problem of drug-resistant infections. Although the individual pathways for PG and OM synthesis and assembly are well characterized, almost nothing is known about how the biogenesis of these essential surface layers is coordinated. Here we report the discovery of a regulatory interaction between the committed enzymes for the PG and LPS synthesis pathways in the Gram-negative pathogen Pseudomonas aeruginosa. We show that the PG synthesis enzyme MurA interacts directly and specifically with the LPS synthesis enzyme LpxC. Moreover, MurA was shown to stimulate LpxC activity in cells and in a purified system. Our results support a model in which the assembly of the PG and OM layers in many proteobacterial species is coordinated by linking the activities of the committed enzymes in their respective synthesis pathways.
Similar content being viewed by others
Main
The biosynthetic pathways for phospholipids, PG and LPS in Gram-negative bacteria rely on shared precursor pools (Fig. 1a). Therefore, flux through each pathway must be balanced to prevent overconsumption of essential precursors by a single pathway4,5,6. LPS biosynthesis requires both UDP-N-acetylglucosamine (UDP-GlcNAc) and acyl-ACP molecules that are also used for PG and phospholipid biosynthesis, respectively6,7. Additionally, overproduction of LPS results in the toxic accumulation of LPS intermediates in the inner membrane8. Flux through the LPS pathway must therefore be tightly regulated.
a, Schematic representation of the biosynthetic pathways responsible for PG and LPS biosynthesis, showing the committed enzymes MurA and LpxC, and other relevant enzymes (LpxA and MurB). T-bars indicate inhibition by the antibiotics fosfomycin and CHIR-090. The green arrow indicates activation of LpxC by MurA. b, Size-exclusion chromatography in which 7.5 µM of purified F–LpxC and H–MurA were resolved either individually or as a mixture in the presence or absence of 37.5 µM CHIR-090 as indicated. The shifted F–LpxC + H–MurA fractions were subsequently collected, resubjected to size-exclusion chromatography, and the resulting fractions were resolved by SDS–PAGE followed by Coomassie staining. Dotted lines indicate the peak mobilities for F–LpxC (gold), H–MurA (cyan) or the shifted F–LpxC + H–MurA fractions (black) in the presence of CHIR-090. The mobilities of H–MurA and F–LpxC in SDS–PAGE assays are indicated by a cyan or gold arrowhead, respectively. Data are representative of two replicates. For gel source data, see Supplementary Fig. 1. c, Catalytic activity (total ion counts (TIC)) of purified PaLpxC (100 nM) alone (no addition (No addn)) or in the presence of MurA variants (100 nM). Dots indicate the values obtained for three individual replicates, bars indicate the mean, and error bars represent their standard deviation. *P = 0.0109 (unpaired, two-tailed t-test).
Enterobacteria such as Escherichia coli control LPS synthesis through regulated proteolysis of the committed enzyme, EcLpxC, by FtsH4,9. Previous studies of LpxC in P. aeruginosa (PaLpxC), however, showed that it was not proteolysed10. Accordingly, N-terminally His-tagged PaLpxC (H–PaLpxC) accumulated normally in an ftsH-deletion mutant (Extended Data Fig. 1a). Thus, LPS biogenesis in P. aeruginosa seems to be regulated through a mechanism distinct from that in enterobacteria.
PaLpxC is activated by PaMurA
The overproduction of H–EcLpxC but not H–PaLpxC inhibited growth of P. aeruginosa and increased cellular levels of LPS (Extended Data Figs. 1b and 2a), suggesting that PaLpxC is regulated in P. aeruginosa cells through a mechanism ineffective against EcLpxC. To identify possible regulatory factors, H–PaLpxC or H–EcLpxC was produced in P. aeruginosa and interaction partners were identified following affinity purification. PaMurA and PA4701 were the only proteins enriched with H–PaLpxC but not H–EcLpxC (Supplementary Table 1). PA4701 is a non-essential protein (Extended Data Fig. 3a) of unknown function, whereas PaMurA is the essential committed enzyme for PG synthesis11,12 (Fig. 1a and Extended Data Fig. 3b,c). We focused on the PaMurA–PaLpxC interaction and validated it using in vivo pulldown assays with H–PaLpxC and N-terminally Flag-tagged PaMurA (F–PaMurA; Extended Data Fig. 4a). Notably, the co-purification was enhanced when cells were treated with the LpxC inhibitor CHIR-090, suggesting that the drug-bound LpxC enzyme may have a greater affinity for MurA (Extended Data Fig. 4a). Purified His-tagged PaMurA (H–PaMurA; Extended Data Fig. 5a) was also specifically pulled down by Flag-tagged PaLpxC (F–PaLpxC) using anti-Flag resin, indicating that the interaction was direct (Extended Data Fig. 4b). Purified F–PaLpxC and H–PaMurA were subjected to size-exclusion chromatography (SEC) individually or as 1:1 mixtures with or without CHIR-090. In the mixed sample, a large proportion of protein eluted at a volume corresponding to the heterodimer of F–PaLpxC and H–PaMurA (Fig. 1b). Consistent with the pulldown assays, re-application of the heterodimer fractions to the SEC column showed that the complex remained most stable in the presence of CHIR-090 (Fig. 1b). Notably, H–PaMurA stimulated the enzymatic activity of F–PaLpxC (Fig. 1c). This stimulation was specific to PaMurA as the addition of H–EcMurA had no effect (Fig. 1c). We conclude that PaMurA is a direct and specific activator of PaLpxC in vitro.
We reasoned that if PaMurA is an essential activator of PaLpxC in P. aeruginosa cells, then PaMurA depletion should result in a phenotype that resembles the simultaneous inactivation of both essential enzymes. Cells treated with the MurA inhibitor fosfomycin have a PG synthesis inhibition phenotype involving membrane bleb formation and lysis11 (Extended Data Fig. 3d–f). However, cells depleted of PaMurA instead adopted an enlarged, ovoid shape (Extended Data Fig. 3e,f). This phenotype resembled that of cells treated with both CHIR-090 and fosfomycin (Extended Data Fig. 3d–f), suggesting that PaMurA depletion impairs the synthesis of both PG and LPS. Accordingly, PaMurA depletion reduced LPS levels, whereas depletion of the next enzyme in the pathway, PaMurB, did not (Extended Data Figs. 2b and 3b,c) and instead caused a terminal phenotype resembling that following fosfomycin treatment (Extended Data Fig. 3d–f). We conclude that PaMurA is required for the biosynthesis of normal cellular levels of LPS.
PaMurA has two essential functions
Overproduction of wild-type (WT) PaMurA did not increase LPS levels as expected for an activator of PaLpxC (Extended Data Fig. 2c). We suspected this result might be due to the enzymatic activity of PaMurA competing with the LPS pathway for UDP-GlcNAc (Fig. 1a) thereby preventing runaway LPS synthesis. Alternatively, a particular MurA conformer13 may be the activator and it may not be produced in sufficient levels to stimulate LPS synthesis on PaMurA overexpression. These scenarios predict that the overproduction of catalytically inactive or conformationally trapped PaMurA variants should cause lethal levels of LPS production. We therefore searched for toxic PaMurA variants. P. aeruginosa was transformed with a plasmid encoding mutagenized PamurA under the control of an isopropyl-β-d-thiogalactoside (IPTG)-regulated promoter. The resulting library was pooled and grown in liquid medium with inducer. Plasmids that caused lysis were purified from the culture supernatant, and the PamurA genes from those causing an IPTG-dependent growth defect on retransformation were sequenced.
Twenty-three toxic PaMurA variants (designated PaMurA*) were identified (Fig. 2a and Extended Data Fig. 6a). Their growth inhibitory activity was alleviated by a normally lethal concentration of CHIR-090, suggesting that they hyperactivate PaLpxC (Extended Data Fig. 6a). The amino acid changes in the PaMurA* variants mapped around the active site of the enzyme14 (Extended Data Fig. 6b) and included the catalytic cysteine residue (C117) that forms a covalent intermediate with the phosphoenolpyruvate substrate15. A subset of PaMurA* variants were purified (Extended Data Fig. 5a) and found to have markedly reduced enzymatic activity while retaining their ability to activate purified PaLpxC (Extended Data Fig. 6c,d).
a, Viability assay in which serial dilutions of PAO1 harbouring an empty plasmid or the indicated PamurA variant under IPTG-inducible control were plated on LB agar with or without IPTG supplementation as indicated. Data are representative of three biological replicates. b, Ratio of LpxC product to substrate detected by liquid chromatography–mass spectrometry in the aqueous fraction of methanol–chloroform-extracted P. aeruginosa whole-cell lysates. Dots indicate the values obtained for three biological replicates, bars indicate the mean, and error bars represent their standard deviation. c, Model structure of the PaLpxC–PaMurA complex predicted by AlphaFold19. PaLpxC is represented in gold and PaMurA is represented in cyan. PaMurA residues G58 and E406 are highlighted in red and residue C117 is highlighted in navy. PaLpxC active-site residues34 are depicted in orange.
The equivalent of the C117S substitution in PaMurA has been well characterized in other orthologues in which it has been shown to trap the enzyme in a closed conformation bound to its product16,17. We therefore chose to study the potential in vivo activation of PaLpxC by PaMurA(C117S) further. The LpxC substrate (UDP-3-O-(R-3-hydroxydecanoyl)-N-acetylglucosamine) and product (UDP-3-O-(R-3-hydroxydecanoyl)-glucosamine) were extracted and quantified from cells overproducing PaMurA(WT) or PaMurA(C117S). The PaLpxC product/substrate ratio was unchanged in cells overproducing PaMurA(WT) relative to an empty vector control (Fig. 2b and Extended Data Fig. 7). However, overproduction of PaMurA(C117S) led to a 100-fold increase in the PaLpxC product/substrate ratio and increased LPS levels, indicating that this variant is a potent activator of PaLpxC in vivo (Fig. 2b and Extended Data Figs. 2c and 7). We therefore conclude that PaMurA activates PaLpxC in cells but that only catalytically inactive PaMurA variants promote toxic levels of LPS synthesis when they are overproduced owing to their inability to compete with the LPS synthesis pathway for UDP-GlcNAc.
To further characterize the interaction between PaLpxC and PaMurA, we searched for non-toxic derivatives of PaMurA(C117S), reasoning that some would be unable to bind PaLpxC. Survivors of F–PaMurA(C117S) production from a mutagenized plasmid were selected followed by an immunoblot-based screen for isolates producing stable, full-length F–PaMurA(C117S). This procedure identified PaMurA(C117S/G58D) and PaMurA(C117S/E406K). We confirmed that both variants were not toxic when overproduced to levels similar to PaMurA(C117S) (Fig. 2a and Extended Data Fig. 8a). We next purified H–PaMurA(G58D) and H–PaMurA(E406K) variants with intact active sites (Extended Data Fig. 5a). Both retained MurA activity (Extended Data Fig. 8b), and consistent with the genetic results, they failed to activate F–PaLpxC in vitro (Fig. 1c). Notably, however, whereas H–PaMurA(G58D) was unable to bind to F–PaLpxC, H–PaMurA(E406K) exhibited binding activity similar to that of H–PaMurA(WT) (Extended Data Fig. 8c). Thus, the G58D change impairs the ability of PaMurA to bind PaLpxC, whereas the E406K substitution seems to disrupt the activation mechanism following complex formation. The residues G58 and E406 both lie on the same surface of the PaMurA structure, suggesting that this face comprises the PaLpxC-binding interface (Fig. 2c).
The results thus far suggest that PaMurA may have two essential functions: PG synthesis and activation of PaLpxC. This model predicts that a catalytically active variant of PaMurA that cannot activate PaLpxC should fail to complement a PamurA deletion but remain capable of complementing a catalytically dead PaMurA variant that retains its PaLpxC activation function. To test this prediction, PamurAWT was placed under Plac control and the native allele was either deleted or converted to PamurAC117S. We then introduced a plasmid with or without the PamurAG58D gene controlled by an arabinose-inducible promoter (Para). As expected, PaMurA(WT) depletion resulted in a severe growth defect in either the ΔmurA or PamurAC117S backgrounds (Extended Data Fig. 8d). Notably, however, the production of PaMurA(G58D) was able to restore growth on PaMurA(WT) depletion in the context of the PamurAC117S allele, but not the murA deletion (Extended Data Fig. 8d). We therefore conclude that the PaLpxC activation function of PaMurA is essential and separable from its catalytic activity.
LpxC–MurA interaction is conserved
To investigate the conservation of the LpxC–MurA regulatory interaction, we used evolutionary covariation analysis18. However, because both enzymes are conserved throughout Gram-negative bacteria but only a subset are likely to interact, we could not reliably detect residues that covary between LpxC and MurA without first knowing which regions of the two proteins interact; non-interacting pairs among the genomes analysed generated too much ‘noise’ to detect a clear interaction signature. We therefore modelled the structure of the LpxC–MurA complex with AlphaFold19, which predicted a high-confidence structure for PaLpxC–PaMurA but not between the corresponding E. coli proteins (Fig. 2c and Extended Data Fig. 9a,b). The G58 and E406 residues of PaMurA implicated in PaLpxC binding and/or activation lie at the modelled interaction interface (Fig. 2c), supporting the accuracy of the structure prediction. Using this model as a guide, evolutionary covariation analysis of residues predicted to be at the interaction interface was carried out on LpxC–MurA pairs from 8,302 proteobacterial genomes. Each LpxC–MurA pair was then assigned an ‘interaction score’ as an indication of how strongly the two proteins were predicted to interact (Fig. 3a).
a, MurA–LpxC interaction scores calculated from a direct coupling analysis-based approach (Methods) plotted on a maximum-likelihood phylogenetic tree based on concatenated MurA and LpxC sequences generated with FastTree. Experimentally tested interactions are highlighted. b, Purified F–LpxC (2.5 µM) was mixed in a 1:1 ratio with purified H–MurA in the presence or absence of CHIR-090 (5.7 µM) as indicated. The mixtures were pulled down with anti-Flag resin and the input and eluate were subjected to SDS–PAGE and Coomassie staining. Mobilities of H–MurA and F–LpxC are indicated by a cyan or gold arrowhead, respectively. Data are representative of at least two replicates. For gel source data, see Supplementary Fig. 1.
The P. aeruginosa LpxC–MurA pair had a high interaction score as expected, as did those from other Pseudomonadales and a subset of other gammaproteobacteria including the Legionalles, Xanthomonadales and Oceanospirillales orders (Fig. 3a). In addition, high interaction scores were also observed in a subset of Rhodospirillales, an order of alphaproteobacteria that is highly divergent from P. aeruginosa (Fig. 3a). On the basis of the distribution of interaction scores, we estimate that 48% of gammaproteobacteria and 35% of alphaproteobacteria encode LpxC–MurA pairs that are capable of interacting (Extended Data Fig. 9e). To investigate the accuracy of the covariation analysis, three additional LpxC–MurA pairs with high interaction scores (Magnetospirillum kuznetsovii, Xanthomonadales spp. and Legionella pneumophila) and two LpxC–MurA pairs with low interaction scores (E. coli and Acinetobacter baumannii) were purified (Extended Data Fig. 5a) and their ability to interact was tested in vitro. Only those pairs with high interaction scores were observed to interact (Fig. 3b). Furthermore, L. pneumophila LpxC (LpnLpxC), but not EcLpxC, exhibited increased activity in vitro in the presence of its cognate MurA (Extended Data Fig. 9f,g). Thus, we conclude that LpxC and MurA from a variety of proteobacteria interact, suggesting that the regulatory link between the PG and LPS biogenesis pathways is conserved in a variety of Gram-negative bacteria.
Model for balanced LPS and PG synthesis
The LPS biosynthetic pathway shares critical precursors with both the phospholipid and PG biosynthetic pathways (Fig. 1a). In E. coli and other enterobacteria, the need for balanced consumption of acyl-ACP molecules by the LPS and phospholipid biosynthetic pathways has been well documented4,20,21,22,23. By contrast, little is known about how UDP-GlcNAc utilization is coordinated by the LPS and PG biosynthetic pathways to allow uniform cell envelope expansion. We propose that the PaLpxC-activating function of PaMurA serves to limit LPS biosynthesis such that it cannot outrun PG biosynthesis. In support of this model, overexpression of PaMurA(WT), which is capable of competing with the LPS synthesis pathway for UDP-GlcNAc, had no impact on cellular viability (Fig. 2a). Overexpression of the catalytically inactive variant PaMurA(C117S), however, resulted in the imbalanced activation of PaLpxC in vivo, resulting in cell death (Fig. 2a). By limiting the catalytically active population of PaLpxC to PaMurA levels, runaway LPS biosynthesis is not possible. On the other hand, MurA has been shown to be feedback inhibited by the downstream PG precursor UDP-MurNAc24. Under conditions in which the flux of PG precursor synthesis is too high, UDP-MurNAc will accumulate such that it binds to MurA and locks it into a closed conformation similar to that of PaMurA(C117S)13,16. Thus, when PG biosynthesis outpaces LPS biosynthesis, MurA will be subject to feedback inhibition, decreasing its catalytic activity without affecting its ability to activate PaLpxC to stimulate LPS biosynthesis and rebalance the pathways.
In enterobacteria, LpxC is proteolysed by FtsH, which is in turn regulated by at least two accessory proteins: LapB and YejM5,8,25,26. Although FtsH is broadly conserved, LapB and YejM are observed only in a subset of proteobacterial genomes. By contrast, LpxC and MurA are nearly universally conserved in proteobacteria. Our computational analysis suggests that a substantial group of gammaproteobacteria and alphaproteobacteria control LpxC through activation by MurA (Fig. 3a). Why bacteria use different strategies to regulate flux through the LPS biosynthetic pathway is unclear. We suggest, however, that it may reflect differences in their environmental niche. For example, P. aeruginosa preferentially utilizes tricarboxylic acids over carbohydrates as a carbon source27. Thus, activated sugars such as UDP-GlcNAc may be more limiting for P. aeruginosa than for sugar-loving E. coli such that it focuses its regulation of LpxC on the appropriate partitioning of UDP-GlcNAc. It would not be surprising if future studies uncover other as-yet-unknown strategies of LpxC regulation in other organisms.
Conclusions
In conclusion, our work has identified a previously unknown and unexpected regulatory interaction between essential enzymes involved in the production of two different layers of the Gram-negative cell envelope. In addition to uncovering a potential mechanism for coordinating the biogenesis of the cell wall and OM, these findings also have implications for antibiotic development. MurA is the target of the antibiotic fosfomycin and LpxC has been the subject of several drug discovery campaigns that have yielded inhibitors with potent antibacterial activity11,28,29,30,31,32,33. The connection between these enzymes identified here suggests that it may be possible to develop dual-targeting drugs that alter both MurA and LpxC activity to simultaneously disrupt PG and OM assembly to kill P. aeruginosa and/or sensitize it to other antibiotics made ineffective by the barrier function of its envelope. These findings also raise the possibility that there are many more regulatory interactions between enzymes involved in the biogenesis of different cell envelope components waiting to be discovered.
Methods
Media, bacterial strains and plasmids
As indicated, cells were grown in LB (1% tryptone, 0.5% yeast extract, 0.5% NaCl) or minimal M9 medium35 supplemented with 0.2% casamino acids and 0.2% glucose. The following concentrations of antibiotics were used: chloramphenicol, 25 μg ml−1; kanamycin, 25 μg ml−1; gentamycin, 30 μg ml−1 (P. aeruginosa) or 15 μg ml−1 (E. coli); carbenicillin, 200 μg ml−1 (P. aeruginosa) or 50 μg ml−1 (E. coli). Plasmids used in this study are listed in Supplementary Table 2. The bacterial strains used in this study are listed in Supplementary Table 3. All P. aeruginosa strains used in the reported experiments are derivatives of PAO1. Primers used in this study are listed in Supplementary Table 4. Unless stated otherwise, polymerase chain reaction (PCR) was carried out using Q5 polymerase (NEB M0492L) for cloning purposes and GoTaq Green DNA polymerase (Promega M7123) for diagnostic purposes, both according to the manufacturer’s instructions. Plasmid DNA and PCR fragments were purified using the PurePlasmid miniprep kit (CW Biosciences CW0500M) or the DNA Clean-up kit (CW Biosciences CW2301M), respectively. Details on strain and plasmid construction can be found in the Supplementary Information. No statistical methods were used to predetermine sample sizes used for experiments, but sample sizes are in line with field standards.
Isolation of toxic murA alleles
Alleles of murA that are toxic when overexpressed were isolated as described previously36 with slight modifications. First, the murA gene was mutagenized by PCR with Taq polymerase (NEB M0267L) from purified pKH37 plasmid using primer pair 15/76 in five separate reactions. A 500 ng quantity of each of the five resulting murA* pools was then separately used as a megaprimer to amplify 50 ng of pKH37 using Q5 polymerase (NEB M0492L) in 50-µl reactions with the following thermocycler settings: 1) 95 °C for 3 min; 2) 95 °C for 50 s; 3) 60 °C for 50 s; 4) 72 °C for 10 min; 5) repeat steps 2–4 for a total of 25 cycles. Twenty units of DpnI (NEB R0176L) was subsequently added to each reaction and digestion of unamplified DNA was allowed to proceed at 37 °C for 1.5 h. Each reaction was drop dialysed using 0.025-µm mixed cellulose ester membranes (Millipore VSWP02500) floated on Milli-Q water for 20 min. A 12 µl volume of each dialysed product was then separately transformed into 100 µl of NEB-5-alpha electrocompetent E. coli (NEB C2989K), and recovered in SOC outgrowth medium (NEB B9020S), and transformants were selected by plating the outgrowth onto LB agar supplemented with gentamycin. Approximately 1 million colonies from each of the five libraries were separately pooled and the pKH37 derivatives were purified using the PurePlasmid miniprep kit (CW Biosciences CW0500M).
PAO1 was grown in 25 ml LB overnight at 37 °C, centrifuged at 12,000g for 5 min, and resuspended in an equal volume of 300 mM sucrose. The centrifugation and resuspension steps were repeated for a total of four washes. After a final centrifugation at 12,000g for 5 min, the cell pellet was resuspended in 1/20 of the original volume. A 1.2 µg quantity of pKH37-derived libraries was separately transformed into 150 µl of electrocompetent PAO1. Transformation reactions were recovered in LB at 37 °C for 1 h and transformants were selected by plating the outgrowth onto LB agar supplemented with 30 µg ml−1 gentamycin. Approximately 1–2 million colonies from each library were separately pooled.
To identify pKH37 variants that harbour a toxic allele of murA, each PAO1 × pKH37* library was diluted to an optical density at 600 nm (OD600nm) of 0.01 in 3 ml LB supplemented with 30 µg ml−1 gentamycin and grown at 37 °C for 2 h 45 min. IPTG was added to each culture to a final concentration of 1 mM and the incubation at 37 °C was continued for an additional 2 h. Each culture was then subjected to centrifugation at 21,130g for 5 min and released DNA from each library was separately purified from 2.5 ml of the supernatant using the DNA clean-up kit (CW Biosciences CW2301M). The purified DNA was subsequently re-transformed into PAO1 electrocompetent cells prepared as described above and transformants were selected by plating the outgrowth on LB agar supplemented with gentamycin. The resulting colonies were then patched onto LB agar plates supplemented with gentamycin with and without 1 mM IPTG and grown overnight at 37 °C. The murA allele encoded by IPTG-sensitive isolates was determined by Sanger sequencing using primer pair 15/76.
Isolation of MurA(C117S) loss-of-toxicity variants
pKH100 was mutagenized by passaging it through E. coli strain XL1-red. pKH100 was transformed into XL1-red and five separate cultures were inoculated, each using four unique transformants. After propagation overnight, plasmids were purified using the PurePlasmid miniprep kit (CW Biosciences CW0500M) to create five pKH100* libraries. PAO1 was grown in 25 ml LB overnight at 37 °C, centrifuged at 12,000g for 5 min, and resuspended in an equal volume of 300 mM sucrose. The centrifugation and resuspension steps were repeated for a total of four washes. After a final centrifugation at 12,000g for 5 min, the cell pellet was resuspended in 1/20 of the original volume. A 1.2 µg quantity of each pKH100* was separately transformed into 150 µl of electrocompetent PAO1. Transformation reactions were recovered in LB at 37 °C for 1 h and transformants were selected by plating the outgrowth onto LB agar supplemented with gentamycin. Approximately 1–10 million colonies from each library were separately pooled and stored in LB + 10% dimethylsulfoxide (DMSO) at −80 °C.
PAO1 × pKH100* libraries were plated on LB agar supplemented with 30 µg ml−1 gentamycin and 1 mM IPTG, which allowed growth of only approximately 0.3–1% of all colony-forming units. A total of 17 or 18 IPTG-resistant colonies from each pool were streak-purified and the expression level and molecular weight of F–MurA(C117S)* from each isolate were determined by western blot analysis after growth in LB and induction for 1 h in mid-log phase. Only those isolates with expression levels and molecular weights consistent with F–MurA(C117S) were subjected to Sanger sequencing with primers 15 and 76. In three of the five libraries analysed, F–murAWT was recovered. F–murAC117S/G58D and F–murAC117S/E406K were each recovered in one of the five libraries.
Protein purification
Details on protein purification can be found in the Supplementary Information.
Viability assays
Overnight cultures grown at 30 °C or 37 °C were centrifuged at 12,000g for 2 min and resuspended to an OD600nm of 1 in LB. Resuspensions were serially diluted in LB to 10−6 and 5 µl of each dilution was spotted onto LB agar supplemented with the appropriate inducers and antibiotics. Plates were incubated at 37 °C overnight and imaged using a Nikon D3400 camera equipped with a Nikon AF-S micro NIKKOR 40-mm lens.
Western blot analysis
Overnight cultures were diluted to an OD600nm of 0.01 in 3 ml LB, grown at 37 °C for 4 h, and 2 ml was subjected to centrifugation at 12,000g for 3 min. When necessary, IPTG was added to 1 mM or arabinose was added to 0.1% 1 h before collecting the cultures. The cell pellet was resuspended to an OD600nm of 20 in 1× SDS sample buffer (50 mM Tris pH 6.8, 10% glycerol, 2% SDS, 0.2% bromophenol blue, 1% β-mercaptoethanol) and boiled at 100 °C for 10 min. The lysate was subsequently sonicated using a Qsonica Q800R3 sonicator with 30 cycles of 1 s on, 1 s off at 25% amplitude. Protein concentration was quantified using the Non-Interfering protein assay kit (G-biosciences 786-005). A 10 µg quantity of protein was loaded onto a 10% polyacrylamide–SDS gel and subjected to electrophoresis at 150 V for 70 min. Protein was transferred to PVDF membranes using the mixed molecular weight protocol on a Bio-Rad Trans-Blot Turbo transfer system and membranes were blocked in TBST (20 mM Tris pH 8.0, 200 mM NaCl, 0.05% Tween-20) with 5% milk at room temperature for 1 h. Membranes were then probed with anti-His (GenScript A00186-100), anti-Flag (Sigma F7425) or anti-RpoA (BioLegend 663104) diluted 1:3,000, 1:3,000 or 1:100,000, respectively, in TBST + 5% milk at room temperature for 1 h. Membranes were washed three times for 5 min each in TBST and probed with peroxidase-conjugated goat-anti-mouse (Rockland 610-1302) or rabbit TrueBlot: anti-rabbit IgG HRP (Rockland 18-8816-33) diluted 1:3,000 in TBST + 5% milk at room temperature for 1 h. Membranes were washed three times for 5 min each in TBST, developed with SuperSignal West Pico PLUS Chemiluminescent Substrate (Thermo 34580), and imaged using an Azure Biosystems C600 imager.
In vivo co-affinity purification
To identify in vivo interacting partners of PaLpxC in an unbiased manner, two co-affinity purification schemes were used in replicate samples. Specifically, in condition 1, 150 mM NaCl was used in lysis and wash buffers, but in condition 2, 300 mM NaCl was used in lysis and wash buffers. Overnight P. aeruginosa cultures were diluted to OD600nm = 0.01 in 500 ml LB and grown at 37 °C to early logarithmic phase. In the case of PA1009, the culture was induced with 0.5% arabinose 1 h before collection. The entire culture was subjected to centrifugation at 13,000g for 10 min and the pellets were resuspended in 4 ml of lysis buffer (50 mM Tris pH 7.5, 2% glycerol, 5 mM imidazole, 1% Triton X-100, and 150 mM or 300 mM NaCl) and cells were lysed by sonication with a Q125 Qsonica sonicator. Cell debris was pelleted by centrifugation at 21,130g for 15 min at 4 °C and the clarified supernatant was mixed with Ni-NTA resin (Qiagen 30230) rotating end-over-end at 4 °C for 1.5 h. Resin was washed four times with 6 ml of wash buffer (50 mM Tris pH 7.5, 2% glycerol, 25 mM imidazole, 1% Triton X-100, and 150 mM or 300 mM NaCl). Proteins were eluted from the resin in elution buffer (50 mM Tris pH 7.5, 2% glycerol, 250 mM imidazole, 0.1% Triton X-100 and 150 mM NaCl). Eluates were mixed 1:1 with 2× SDS sample buffer (100 mM Tris pH 6.8, 20% glycerol, 4% SDS, 0.4% bromophenol blue), and 40 µl was loaded onto 4–20% polyacrylamide Mini-PROTEAN TGX gels (Bio-Rad 4568094) and subjected to electrophoresis at 80 V for 25 min. The gel was subsequently stained with Coomassie R-250 and entire lanes were excised for mass spectrometry analysis.
Excised lanes were cut into approximately 1-mm3 pieces. Gel pieces were then subjected to a modified in-gel trypsin digestion procedure37. Gel pieces were washed and dehydrated with acetonitrile for 10 min, followed by removal of acetonitrile. Pieces were then completely dried in a speed-vac. Gel pieces were rehydrated with 50 mM ammonium bicarbonate solution containing 12.5 ng µl−1 modified sequencing-grade trypsin (Promega) at 4 °C. After 45 min, the excess trypsin solution was removed and replaced with 50 mM ammonium bicarbonate solution to just cover the gel pieces. Samples were then placed in a 37 °C room overnight. Peptides were later extracted by removing the ammonium bicarbonate solution, followed by one wash with a solution containing 50% acetonitrile and 1% formic acid. The extracts were then dried in a speed-vac (about 1 h). The samples were then stored at 4 °C until analysis.
On the day of analysis, the samples were reconstituted in 5–10 µl of high-performance liquid chromatography (HPLC) solvent A (2.5% acetonitrile, 0.1% formic acid). A nanoscale reversed-phase HPLC capillary column was created by packing 2.6-µm C18 spherical silica beads into a fused silica capillary (100 µm inner diameter × about 30 cm length) with a flame-drawn tip38. After equilibrating the column, each sample was loaded using a Famos auto sampler (LC Packings) onto the column. A gradient was formed, and peptides were eluted with increasing concentrations of solvent B (97.5% acetonitrile, 0.1% formic acid).
As peptides eluted, they were subjected to electrospray ionization and then entered into an LTQ Orbitrap Velos Pro ion-trap mass spectrometer (Thermo Fisher Scientific). Peptides were detected, isolated and fragmented to produce a tandem mass spectrum of specific fragment ions for each peptide. Peptide sequences (and hence protein identity) were determined by matching protein databases with the acquired fragmentation pattern by the software program Sequest v28 (rev. 13; Thermo Fisher Scientific)39. All databases include a reversed version of all the sequences, and the data were filtered to a peptide false-discovery rate of 1–2%. Only proteins that exhibited the following criteria were reported: at least five unique peptides detected in both PA1018 samples; and at least threefold enrichment in both PA1018 samples compared to the corresponding PAO1 and PA1009 samples.
In vivo targeted pulldowns
Overnight cultures of P. aeruginosa strains encoding combinations of H–PaLpxC, F–PaMurA(WT) and F–PaMurA(C117S) were diluted to OD600nm = 0.01 in 50 ml LB and grown at 37 °C for 2.5 h. IPTG was added to each culture to a final concentration of 1 mM and cultures were grown for an additional 30 min at 37 °C before the addition of DMSO to 0.1% and CHIR-090 to 0.5 µg ml−1 where indicated. Cultures were incubated at 37 °C for an additional hour and collected by centrifugation at 12,000g for 10 min. Cells were washed in 1 ml LB and pelleted by centrifugation at 12,000g for 2 min. Cells were resuspended in 1 ml lysis buffer (25 mM Tris pH 8.0, 150 mM NaCl, 2% glycerol, 0.1% Triton X-100, 5 mM imidazole, 0.1% DMSO with or without 0.5 µg ml−1 CHIR-090 as indicated) and lysed by sonication on ice at 40% amplitude with four cycles of 15 s on, 15 s off. The extracts were clarified by two consecutive centrifugation steps at 21,130g for 5 min at 4 °C and protein concentration was determined with the Non-Interfering protein assay kit (G-biosciences 786-005). Supernatants were diluted to 3.5 mg ml−1 protein in 500 µl lysis buffer and mixed with 25 µl packed Ni-NTA resin (Qiagen 30230). Mixtures were incubated at 4 °C for 3 h rotating end-over-end. Beads were washed three times with 500 µl wash buffer (25 mM Tris pH 8.0, 150 mM NaCl, 2% glycerol, 0.1% Triton X-100, 25 mM imidazole, 0.1% DMSO with or without 0.5 µg ml−1 CHIR-090 as indicated). Protein was eluted in 40 µl elution buffer (25 mM Tris pH 8.0, 150 mM NaCl, 2% glycerol, 0.1% Triton X-100, 500 mM imidazole).
A 10 µg quantity of protein per input sample and 10 µl of eluate samples were resolved by SDS–PAGE in 10% polyacrylamide gels. Protein was transferred to PVDF membranes using the mixed molecular weight protocol on a Bio-Rad Trans-Blot Turbo transfer system and membranes were blocked in TBST + 5% milk at room temperature for 1 h. Membranes were then probed with anti-His (GenScript A00186-100) or anti-Flag (Sigma-Aldrich F7425) diluted 1:3,000 in TBST + 5% milk at room temperature for 1 h. Membranes were washed three times for 5 min each in TBST and probed with peroxidase-conjugated goat-anti-mouse (Rockland 610-1302) or rabbit TrueBlot: anti-rabbit IgG HRP (Rockland 18-8816-33) diluted 1:3,000 in TBST + 5% milk at room temperature for 1 h. Membranes were washed three times for 5 min each in TBST, developed with SuperSignal West Pico PLUS Chemiluminescent Substrate (Thermo 34580), and imaged using an Azure Biosystems C600 imager.
In vitro co-affinity purification
Purified F–PaLpxC and H–MurA variants were each diluted to 2.5 µM in co-purification buffer (25 mM Tris pH 8.0, 150 mM NaCl, 10% glycerol, 1 mM dithiothreitol (DTT), 0.5% DMSO) to a final volume of 100 µl. When indicated, CHIR-090 was added to 5.7 µM. Mixtures were incubated on ice for 30 min after which 85 µl of each mixture was added to an equal volume of Anti-Flag M2 Magnetic Beads (Sigma-Aldrich M8823) and incubated at 4 °C for 1 h rotating end-over-end. Beads were collected with a magnetic rack and washed twice with 300 µl co-purification buffer and once with 200 µl co-purification buffer. When indicated, 5.7 µM CHIR-090 was maintained in the wash buffers. Proteins were eluted with 85 µl elution buffer (25 mM Tris pH 8.0, 150 mM NaCl, 10% glycerol, 1 mM DTT, 100 µg µl−1 Flag peptide (Millipore Sigma F3290)) by incubation at room temperature for 30 min rotating end-over-end. Input samples (5 µl) and eluate samples (15 µl) were resolved by SDS–PAGE in 4–20% polyacrylamide Mini-PROTEAN TGX gels (Bio-Rad 4561095) and protein was detected with Coomassie staining.
SEC
F–PaLpxC and H–PaMurA(WT) either individually or in combination were diluted to 7.5 µM each in 500 µl of SEC buffer 1 (10% glycerol, 0.33% DMSO, 1 mM DTT, 25 mM Tris pH 8.0 and 150 mM NaCl). When indicated, CHIR-090 was added to 37.5 µM. The mixtures were subsequently subjected to SEC using an ActaPure system equipped with a Superdex 200 10/300 GL column equilibrated with SEC buffer 2 (10% glycerol, 1 mM DTT, 25 mM Tris pH 8.0 and 150 mM NaCl) and eluate fractions were collected every 0.35 ml. For F–PaLpxC and H–PaMurA(WT) mixtures, three fractions corresponding to the shifted peak (14.1–15.2 ml with no CHIR-090, 13.8–14.8 ml for with CHIR-090) were pooled and concentrated by centrifugation at 4 °C using centrifugal filters with a molecular weight cutoff of 10-kDa (Amicon UFC801024). The concentrated protein was resubjected to SEC on a Superdex 200 10/300 GL column as described above. A 15 µl volume of the relevant fractions was resolved on a 4–20% polyacrylamide Mini-PROTEAN TGX gels (Bio-Rad 4568094) and protein was detected with Coomassie staining.
Growth curves
Cultures grown overnight at 30 °C were pelleted at 12,000g for 2 min and resuspended in fresh LB. Cultures were diluted in 96-well microtitre plates to an OD600nm of 0.01 in 200 µl LB. Cultures were grown in a Tecan Infinite M-Plex plate reader at 37 °C and OD600nm was monitored every 4 min with shaking at a 1-mm orbital for 200 s after each time point.
LPS silver stain
Overnight cultures were diluted to an OD600nm of 0.01 in 25 ml LB cultures and grown for 4 h at 37 °C. For MurA- and MurB-depletion strains grown in the presence of inducer, 1 mM IPTG was present during the entire outgrowth. For MurA and LpxC overexpression strains, 1 mM IPTG or 0.1% arabinose was added after 3 h of outgrowth and incubation was continued for one additional hour. A 20 ml volume of each culture was centrifuged at 12,000g for 5 min and the pellet was washed in 1 ml of LB before centrifugation at 12,000g for 2 min. Pellets were resuspended to an OD600nm of 20 in LDS sample buffer (Invitrogen NP0008) + 4% β-mercaptoethanol and boiled at 100 °C for 10 min. Samples were sonicated with a Q125 Qsonica sonicator at 25% amplitude for 1 s on, 1 s off for 30 cycles and protein concentration was determined using the Non-Interfering protein assay kit (G-biosciences 786-005). A 5 µg quantity of protein was loaded onto a 15% polyacrylamide–SDS gel and subjected to electrophoresis at 150 V for 70 min before western blot analysis of RpoA as described above. For the LPS gel, 50 µl of each sample was mixed with 1.25 µl of proteinase K (NEB P8107S) and incubated at 55 °C for 1 h followed by 95 °C for 10 min. The equivalent volume accounting for 10 µg of protein was resolved on a 4–12% Criterion XT Bis-Tris gel (Bio-Rad 3450124) at 100 V for 1 h 45 min and LPS was detected by silver stain as described previously40. Briefly, the gel was fixed by incubation in 200 ml of 40% ethanol, 5% acetic acid overnight. Periodic acid was added to 0.7% and allowed to incubate for 5 min. The gel was subsequently washed with three changes of 200 ml ultrapure water over the course of 2 h. The gel was impregnated with 150 ml of freshly made staining solution (0.018 N NaOH, 0.4% NH4OH and 0.667% silver nitrate) for 10 min and subsequently washed with three changes of 200 ml ultrapure water over the course of 2 h. The gel was developed with 200 ml freshly prepared developer solution (0.26 mM citric acid pH 3.0, 0.014% formaldehyde) and development stopped in 100 ml of 1% acetic acid. The gel was imaged in a Bio-Rad ChemiDoc MP Imaging System.
Microscopy
Overnight cultures were diluted to an OD600nm of 0.01 in 3 ml LB with 1 mM IPTG as appropriate. Cultures were grown at 37 °C for 2.5 h and, when appropriate, antibiotics were added to the following concentration: 0.5 µg ml−1 CHIR-090, 64 µg ml−1 fosfomycin or 0.5 µg ml−1 CHIR-090 + 16 µg ml−1 fosfomycin. Cultures were grown at 37 °C for an additional 1.5 h and 1 ml was pelleted at 12,000g for 2 min. Cell pellets were resuspended to an OD600nm of 10 in LB, spotted onto an LB + 1.5% agar pad, and imaged on a Nikon Eclipse 50i microscope equipped with a 100× objective and a Photometrics CoolSNAP HQ2 camera. Images were uniformly edited using the Fiji image analysis platform version 2.3.0/1.53q41. Cell width was quantified using the MicrobeJ plugin (version 5.13I) of ImageJ using the following settings: area: 250–max, length: 3–100, width: 5–25, width variation: 0–0.25, width symmetry: 0–1, circularity, 0.01–1. Normality of each population was tested and determined to be lacking by both the D’Agostino and Pearson test and the Anderson–Darling test carried out by Prism (GraphPad Software, LLC). Statistical significance was determined by Kruskal–Wallis test carried out by Prism (GraphPad Software, LLC). Microscopy source images are available at https://doi.org/10.5281/zenodo.7455522 (ref. 42).
Determination of LpxC product and substrate levels in vivo
Overnight cultures were diluted to an OD600nm of 0.01 in 200 ml LB, grown at 37 °C for 3 h, and IPTG was added to a final concentration of 1 mM. After incubation at 37 °C for one additional hour, cells were collected by centrifugation at 12,000g for 5 min. After resuspension in LB, cells were re-pelleted by centrifugation at 12,000g for 2 min. Pellets were then resuspended in 400 µl HPLC-grade H2O. Subsequently, 250 µl chloroform and 500 µl methanol were added to the mixture. Samples were vortexed vigorously, placed on ice for 10 min, and then spun down at 4 °C at 21,000g. A 500 µl volume of the aqueous layer was collected and dried down at 30 °C overnight in a rotary evaporator. Dried material was resuspended in 100 µl ultrapure H2O and the PaLpxC product and substrate were detected by liquid chromatography–tandem mass spectrometry (LC–MS/MS) as described below.
In vitro LpxC activity assay
Purified F–PaLpxC and H–MurA variants were diluted to 200 nM in binding buffer (25 mM Tris pH 8.0, 150 mM NaCl, 6% glycerol) to a final volume of 15 µl and mixtures were allowed to equilibrate to 30 °C for 10 min. The reaction was started by the addition of 15 µl of substrate mixture (25 mM Tris pH 8.0, 150 mM NaCl, 4% DMSO, 200 µM UDP-3-O-(R-3-hydroxydecanoyl)-GlcNAc (Carbosynth MU75071)) and allowed to proceed for 5 or 10 min at 30 °C at which point the reaction was stopped by the addition of 45 µl 2% acetic acid. Stopped reactions were frozen on dry ice, lyophilized, and the lyophilized material was resuspended in 60 µl H2O. PaLpxC product and substrate were detected by LC–MS as described below. The linear range of the reaction was determined as described above with purified F–PaLpxC alone and reactions were stopped after 1, 2, 5, 10, 15 or 20 min (Extended Data Fig. 5b). H–PaMurA(WT) and H–PaMurA(C117S) alone exhibited background levels of LpxC activity, confirming that the observed stimulation of LpxC activity was not due to contamination within the protein preparation or an alternative enzymatic activity of PaMurA (Extended Data Fig. 6e).
Activity assays with the L. pneumophila and E. coli LpxC–MurA pairs were carried out as described above except that concentration of H–MurA variants in the final reaction mixture was 200 nM. L. pneumophila reactions were stopped after 10 min and E. coli reactions were stopped after 1 min.
LC–MS
To quantify the activity of LpxC in vitro, the PaLpxC product and substrate were detected by LC–MS on an Agilent Technologies 1200 series HPLC system in line with an Agilent 6520 Q-TOF mass spectrometer with electrospray ionization–mass spectrometry operating in negative mode. Samples were separated on a Waters Symmetry C18 column (5 µm; 3.9 mm × 150 mm) with a matching column guard using the following method: flow rate = 0.5 ml min−1, 95% solvent A (H2O, 10 mM ammonium formate) and 5% solvent B (acetonitrile) for 5 min followed by a linear gradient of solvent B from 5–80% over 20 min. Agilent MassHunter Workstation Qualitative Analysis software version B.06.00 was used for analysing the MS data. The PaLpxC substrate (UDP-3-O-(R-3-hydroxydecanoyl)-N-acetylglucosamine) was quantified by monitoring the abundance of 776.21 m/z and resolved as a single peak, which was integrated to infer substrate concentration. The PaLpxC product (UDP-3-O-(R-3-hydroxydecanoyl)-glucosamine) was quantified by monitoring the abundance of 734.1944 m/z and often resolved as multiple peaks as reported previously43, all of which were integrated to infer product concentration.
PaLpxC substrate and product from the aqueous fraction of methanol–chloroform-extracted whole-cell lysates were analysed by LC–MS/MS using the same settings as the LC–MS analysis described above with the following adaptations. Parent ions with m/z of 776.1986 (corresponding to the PaLpxC substrate) and m/z of 734.1872 (corresponding to the PaLpxC product) were targeted for MS/MS with a collision energy of 40. Fragment ions between 50 and 850 m/z were analysed. The relative abundance of the PaLpxC substrate and product were quantified by integrating the peaks observed for 776.1986 m/z and 734.1872 m/z, respectively. Raw LC–MS/MS data are available at https://doi.org/10.5281/zenodo.7455522 (ref. 42).
In vitro MurA activity assay
Purified MurA variants were diluted to 200 nM (for PaMurA(WT), PaMurA(G58D) and PaMurA(E406K)) or 2 µM (for PaMurA* variants) in MurA reaction buffer (25 mM Tris pH 7.75, 6% glycerol) to a final volume of 25 µl and allowed to equilibrate to room temperature for 30 min. The reaction was started by the addition of 25 µl of MurA substrate mixture (25 mM Tris pH 7.75, 2 mM UDP-N-acetylglucosamine (Sigma-Aldrich U4375), and 1 mM phosphoenolpyruvate (Sigma-Aldrich 10108294001)) and the reaction was allowed to proceed for 30 min at 37 °C. The reaction was stopped by the addition of 800 µl Lanzetta reagent (0.033% malachite green, 1% ammonium molybdate, 1 N HCl, 0.2% Triton X-100)44 and was incubated at room temperature for 1.25 min before the addition of 100 µl 34% sodium citrate. After an additional 30-min incubation at room temperature, the absorbance at 600 nm (A660nm) of each sample was determined. A standard curve made from NaH2PO4 diluted in ultrapure water was used to quantify the amount of free Pi released by the reaction.
MurA–LpxC complex structure prediction
The structure of the LpxC–MurA complex was predicted using the default parameters of AlphaFold19. The P. aeruginosa PAO1 LpxC and MurA sequences or the E. coli MG1655 LpxC and MurA sequences were separated by a linker containing 15 repeats of Gly-Gly-Gly-Ser, which was not displayed for clarity. The structure was visualized using ChimeraX version 1.1.1 (ref. 45).
Predicting complex interactions
We sought to use evolutionary covariation to predict the organisms in which the LpxC–MurA interaction does or does not exist. We used EVComplex run with EVcouplings v0.0.5 (ref. 18) informed by the AlphaFold model to define an interaction score based on the inferred pairwise interaction terms \({J}_{{ij}}\) of the complex and validate our proposed complex structure.
We first generated multiple sequence alignments of LpxC and MurA using jackhammer v3.1b2 (ref. 46) with five iterations on the Uniref100 dataset47 released in February 2021. Both P. aeruginosa and E. coli MurA (Uniprot: Q9HVW7 and P0A749) and LpxC (Uniprot: P47205 and P0A725) were used as initial sequences. In each case, alignments were constructed with a range of sequence score thresholds from 0.1 to 0.8 bits per residue, which did not meaningfully change which sequences were included for either species or protein. The same sequences were collected when starting from either species. For all analyses, we began with alignments of 54,940 MurA and 23,634 LpxC sequences generated from a relative bitscore of 0.8, and then concatenated MurA and LpxC sequences for each species, resolving ambiguities in pairing by selecting the sequence in each species closest to the original P. aeruginosa homologue. The final concatenated alignment contained 11,834 sequences. From this concatenated multiple sequence alignment, evolutionary couplings were determined using pseudo-likelihood maximization48.
We more closely analysed the top five ranked intermolecular couplings (Jij) of the direct coupling analysis model that correspond to pairs of amino acid positions in structural contact in the AlphaFold2 model, defined as having Cα atoms within 9 Å. Each pair of sequences can be scored on the coupling between MurA position i and LpxC position j on the basis of the appropriate \({J}_{{ij}}\) term from the model. Summing these five scores for each species, we observed a bimodal distribution (Extended Data Fig. 9c). We selected only pairs of sequences whose sum over the five scores exceeded 0.1, totalling 1,689 sequence pairs (Extended Data Fig. 9c), and trained a new EVcomplex model on just these sequences from the concatenated alignment. In this model, five of the top six intermolecular couplings corresponded to predicted contacts in the AlphaFold2 structure (Extended Data Fig. 9d). We defined a final ‘interaction score’ based on the top 20 coupling terms from this refined model and used it to score all 11,834 pairs of sequences in the concatenated alignment again. The sequence analysis files are available at https://doi.org/10.5281/zenodo.7455522 (ref. 42). The top 20 terms were chosen arbitrarily, but this exact choice does not meaningfully affect downstream analyses, and all conclusions about relative interaction scores remain true for choices of top couplings between five and five hundred.
To estimate the percentage of interacting sequences in each clade, we fitted a two-component Gaussian mixture model using sklearn v1.0.2 (ref. 49) to the interaction scores for all sequence pairs and calculated the posterior probability of each sequence being drawn from the upper distribution. We mapped the taxonomic identifier for each sequence pair to its class in the National Center for Biotechnology Information taxonomy database using ETE3 v1.3.2 (ref. 50), identifying 2,459 alphaproteobacteria and 2,781 gammaproteobacteria in the alignment. We then estimated the expected fraction of interacting sequences in each clade as the average of the interaction probabilities of each sequence in that clade.
To visualize the phylogenetic structure of these sequences, we built a maximum-likelihood phylogeny of MurA and LpxC from the concatenated multiple sequence alignment with FastTree version 2.1.11 (refs. 51). Interaction scores were mapped onto the tree in Python v3.8.8 and visualized with the interactive tree of life (ITOL) v5 (ref. 52). Custom code used in this study can be found at https://github.com/samberry19/evcomplex-interaction-scoring or https://doi.org/10.5281/zenodo.7471436 (ref. 53).
Reporting summary
Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.
Data availability
LC–MS/MS source data, microscopy images and sequence analysis files used to derive LpxC–MurA interaction scores are available at https://doi.org/10.5281/zenodo.7455522. Uniprot accession codes for genes used to generate LpxC–MurA interaction scores are included in the Methods and source data. All bacterial strains and plasmids developed in this study are available upon request. Source data are provided with this paper.
Code availability
All code generated in this study is available at https://github.com/samberry19/evcomplex-interaction-scoring or https://doi.org/10.5281/zenodo.7471436.
References
Silhavy, T. J., Kahne, D. & Walker, S. The bacterial cell envelope. Cold Spring Harb. Perspect. Biol. 2, a000414 (2010).
Zgurskaya, H. I., Lopez, C. A. & Gnanakaran, S. Permeability barrier of Gram-negative cell envelopes and approaches to bypass it. ACS Infect. Dis. 1, 512–522 (2015).
Bush, K. Antimicrobial agents targeting bacterial cell walls and cell membranes. Rev. Sci. Tech. 31, 43056 (2012).
Ogura, T. et al. Balanced biosynthesis of major membrane components through regulated degradation of the committed enzyme of lipid A biosynthesis by the AAA protease FtsH (HflB) in Escherichia coli. Molec. Microbiol. 31, 833–844 (1999).
Klein, G., Kobylak, N., Lindner, B., Stupak, A. & Raina, S. Assembly of lipopolysaccharide in Escherichia coli requires the essential LapB heat shock protein. J. Biol. Chem. 289, 14829–14853 (2014).
Raetz, C. R. Biochemistry of endotoxins. Annu. Rev. Biochem. 59, 129–170 (1990).
Anderson, M. S., Bulawa, C. E. & Raetz, C. R. The biosynthesis of gram-negative endotoxin. Formation of lipid A precursors from UDP-GlcNAc in extracts of Escherichia coli. J. Biol. Chem. 260, 15536–15541 (1985).
Guest, R. L., Same Guerra, D., Wissler, M., Grimm, J. & Silhavy, T. J. YejM modulates activity of the YciM/FtsH protease complex to prevent lethal accumulation of lipopolysaccharide. mBio 11, e00598-20 (2020).
Anderson, M. S. et al. UDP-N-acetylglucosamine acetyltransferase of Escherichia coli: the first step in endotoxin biosynthesis is thermodynamically unfavorable. J. Biol. Chem. 268, 19858–19865 (1993).
Langklotz, S., Schakermann, M. & Narberhaus, F. Control of lipopolysaccharide biosynthesis by FtsH-mediated proteolysis of LpxC is conserved in enterobacteria but not in all gram-negative bacteria. J. Bacteriol. 193, 1090–1097 (2011).
Kahan, F. M., Kahan, J. S., Cassidy, P. J. & Kropp, H. The mechanism of action of fosfomycin (phosphonomycin). Ann. N. Y. Acad. Sci. 235, 364–386 (1974).
Strominger, J. L. Enzymic transfer of pyruvate to uridine diphosphoacetylglucosamine. Biochem. Biophys. Acta 30, 645–646 (1958).
Zhu, J. Y. et al. Functional consequence of covalent reaction of phosphoenolpyruvate with UDP-N-acetylglucosamine 1-carboxyvinyltransferase (MurA). J. Biol. Chem. 287, 12657–12667 (2012).
Eschenberg, S. & Schonbrunn, E. Comparative X-ray analysis of the un-liganded fosfomycin-target MurA. Proteins 40, 290–298 (2000).
Wanke, C. & Amrhein, N. Evidence that the reaction of the uDP-N-acetylglucosamine 1-carboxyvinyltransferase proceeds through the O-phosphothioketal of pyruvic acid bound to Cys115 of the enzyme. Eur. J. Biochem. 218, 861–870 (1993).
Eschenburg, S., Priestman, M. & Schönbrunn, E. Evidence that the fosfomycin target Cys115 in UDP-N-acetylglucosamine enolpyruvyl transferase (MurA) is essential for product release. J. Biol. Chem. 280, 3757–3763 (2005).
Schönbrunn, E., Eschenberg, S., Krekel, F., Luger, K. & Amrhein, N. Role of the loop containing residue 115 in the induced-fit mechanism of the bacterial cell wall enzyme MurA. Biochemistry 39, 2164–2173 (2000).
Hopf, T. A. et al. Sequence co-evolution gives 3D contacts and structures of protein complexes. Elife 3, e03430 (2014).
Jumper, J. et al. Highly accurate protein structure prediction with AlphaFold. Nature 596, 583–589 (2021).
Mohan, S., Kelly, T. M., Eveland, S. S., Raetz, C. R. & Anderson, M. S. An Escherichia coli gene (FabZ) encoding (3R)-hydroxymyristoyl acyl carrier protein dehydrase. Relation to fabA and suppression of mutations in lipid A biosynthesis. J. Biol. Chem. 269, 32896–32903 (1994).
Thomanek, N. et al. Intricate crosstalk between lipopolysaccharide, phospholipid and fatty acid metabolism in Escherichia coli modulates proteolysis of LpxC. Front. Microbiol. 9, 3285 (2018).
Mostafavi, M. et al. Interplay of Klebsiella pneumoniae fabZ and lpxC mutations leads to LpxC inhibitor-dependent growth resulting from loss of membrane homeostasis. mSphere 3, e00508-18 (2018).
Zeng, D. et al. Mutants resistant to LpxC inhibitors by rebalancing cellular homeostasis. J. Biol. Chem. 288, 5475–5486 (2013).
Mizyed, S. et al. UDP-N-acetylmuramic acid (UDP-MurNAc) is a potent inhibitor of MurA (enolpyruvyl-UDP-GlcNAc synthase). Biochemistry 44, 4011–4017 (2005).
Fivenson, E. M. & Bernhardt, T. G. An essential membrane protein modulates the proteolysis of LpxC to control lipopolysaccharide synthesis in Escherichia coli. mBio 11, e00939-20 (2020).
Clairfeuille, T. et al. Structure of the essential inner membrane lipopolysaccharide–PbgA complex. Nature 584, 479–483 (2020).
Collier, D. N., Hager, P. W. & Phibbs, P. V. Jr Catabolite repression control in the Pseudomonads. Res. Microbiol. 147, 551–561 (1996).
Kline, T. Potent, novel in vitro inhibitors of the Pseudomonas aeruginosa deacetylase LpxC. J. Med. Chem. 45, 3112–3129 (2002).
McClerren, A. L. et al. A slow, tight-binding inhibitor of the zinc-dependent deacetylase LpxC of Lipid A biosynthesis with antibiotic activity comparable to ciprofloxacin. Biochemistry 44, 16574–16583 (2005).
Langsdorf, E. F. et al. Screening for antibacterial inhibitors of the UDP-3-O-(R-3-hydroxymyristoyl)-N-acetylglucosamine deacetylase (LpxC) using a high-throughput mass spectrometry assay. J. Biomol. Screen. 15, 52–61 (2010).
Erwin, A. L. Antibacterial drug discovery targeting the lipopolysaccharide biosynthetic enzyme LpxC. Cold Spring Harb. Perspect. Med. 6, a025304 (2016).
Krause, K. M. et al. Potent LpxC inhibitors with in vitro activity against multidrug-resistant Pseudomonas aeruginosa. Antimicrob. Agents Chemother. 63, e00977-19 (2019).
Ushiyama, F. et al. Lead optimization of 2-hydroxymethyl imidazoles as non-hydroxamate LpxC inhibitors: discovery of TP0586532. Bioorg. Med. Chem. 30, 115964 (2021).
Hernick, M. & Fierke, C. A. Catalytic mechanism and molecular recognition of E. coli UDP-3-O-(R-3-hydroxymyristoyl)-N-acetylglucosamine deacetylase probed by mutagenesis. Biochemistry 45, 15240–15248 (2006).
Miller, J. Experiments in Molecular Genetics (Cold Spring Harbor Laboratory, 1972).
Yang, D. C., Tan, K., Joachimiak, A. & Bernhardt, T. G. A conformational switch controls cell wall-remodelling enzymes required for bacterial cell division. Mol. Microbiol. 85, 768–781 (2012).
Shevchenko, A., Wilm, M., Vorm, O. & Mann, M. Mass spectrometric sequencing of proteins from silver-stained polyacrylamide gels. Anal. Chem. 68, 850–858 (1996).
Peng, J. & Gygi, S. P. Proteomics: the move to mixtures. J. Mass Spectrom. 36, 1083–1091 (2001).
Eng, J. K., McCormac, A. L. & Yates, J. R. III An approach to correlate tandem mass spectral data of peptides with amino acid sequences in a protein database. J. Am. Soc. Mass Spectrom. 5, 976–989 (1994).
Tsai, C.-M. & Frasch, C. E. A sensitive silver stain for detecting lipopolysaccharide in polyacrylamide gels. Anal. Biochem. 119, 115–119 (1982).
Schindelin, J. et al. Fiji: an open-source platform for biological-image analysis. Nat. Methods 9, 676–682 (2012).
Hummels, K. R. et al. Data for “Coordination of bacterial cell wall and outer membrane biosynthesis”. Zenodo https://doi.org/10.5281/zenodo.7455522 (2022).
Cohen, F. et al. Optimization of LpxC inhibitors for antibacterial activity and cardiovascular safety. ChemMedChem 14, 1560–1572 (2019).
Lanzetta, P. A., Alvarez, L. J., Reinach, P. S. & Candia, O. A. An improved assay for nanomole amounts of inorganic phosphate. Anal. Biochem. 100, 95–97 (1979).
Pettersen, E. F. et al. UCSF ChimeraX: structure visualization for researchers, educators, and developers. Protein Sci. 30, 70–82 (2021).
Johnson, L. S., Eddy, S. R. & Portugaly, E. Hidden Markov model speed heuristic and iterative HMM search procedure. BMC Bioinform. 11, 431 (2010).
Suzek, B. E. et al. UniRef clusters: a comprehensive and scalable alternative for improving sequence similarity searches. Bioinformatics 31, 926–932 (2015).
Ekeberg, M., Lövkvist, C., Lan, Y., Weigt, M. & Aurell, E. Improved contact prediction in proteins: using pseudolikelihoods to inver Potts models. Phys. Rev. E 87, 012707 (2013).
Pedregosa, F. et al. Scikit-learn: machine learning in Python. J. Mach. Learn. Res. 12, 2825–2830 (2011).
Huerta-Cepas, J., Serra, F. & Bork, P. ETE 3: reconstruction, analysis, and visualization of phylogenomic data. Mol. Biol. Evol. 33, 1635–1638 (2016).
Price, M. N., Dehal, P. S. & Arkin, A. P. FastTree: computing large minimum evolution trees with profiles instead of a distance matrix. Mol. Biol. Evol. 26, 1641–1650 (2009).
Letunic, I. & Bork, P. Interactive Tree Of Life (iTOL) v5: an online tool for phylogenetic tree display and annotation. Nucleic Acids Res. 49, W293–W296 (2021).
Berry, S. P. samberry19/evcomplex-interaction-scoring: v1.0 (v1.0). Zenodo https://doi.org/10.5281/zenodo.7471436 (2022).
Acknowledgements
We thank all of the members of the Bernhardt, Rudner and Mekalanos labs for their thoughtful discussions and advice throughout this project, especially L. Marmont for technical insight regarding protein purification. We also thank S. Lory for strains and reagents as well as advice on the genetic manipulation of P. aeruginosa, B. Silva and M. Welsh for assistance with LC–MS/MS analysis, and the Taplin Mass Spectrometry Core Facility at Harvard Medical School for the identification of co-affinity-purified proteins. LC–MS data were acquired on an Agilent 6520 Q-TOF spectrometer supported by the Taplin Funds for Discovery Program. Protein structures were visualized with UCSF ChimeraX, developed by the Resource for Biocomputing, Visualization, and Informatics at the University of California, San Francisco, with support from National Institutes of Health R01-GM129325 and the Office of Cyber Infrastructure and Computational Biology, National Institute of Allergy and Infectious Diseases. This work was supported by the National Institutes of Health (F32AI164630 to K.R.H., AI148752 to S.W., AI083365 to T.G.B., and U19 AI158028 to T.G.B and S.W.), the Chan Zuckerberg Initiative (CZI2018-191853 to D.S.M.) and Investigator funds from the Howard Hughes Medical Institute (T.G.B).
Author information
Authors and Affiliations
Contributions
K.R.H. and T.G.B. designed the study and wrote the manuscript. S.P.B., J.K.M. and D.S.M. devised and carried out the covariation analysis. Z.L. established the methanol–chloroform metabolite extraction protocol. A.T. and S.W. developed the LC–MS protocol. K.R.H. carried out all other experiments and data analysis. All authors read and approved the manuscript.
Corresponding author
Ethics declarations
Competing interests
The authors declare no competing interests.
Peer review
Peer review information
Nature thanks Russell Bishop and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Peer reviewer reports are available.
Additional information
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Extended data figures and tables
Extended Data Fig. 1 Regulation of LpxC in P. aeruginosa differs from that observed in E. coli.
(a) Anti-His immunoblot detecting H-PaLpxC expressed from the native chromosomal locus in wild-type cells or an ftsH deletion mutant. A corresponding blot for RpoA was used as a loading control. Data are representative of 3 biological replicates. (b) Spot titer assay in which serial dilutions of PAO1 harboring an empty plasmid or one encoding His-PalpxC or His-EclpxC under arabinose-inducible control were plated on LB agar supplemented with arabinose and/or the LpxC inhibitor CHIR-090 as indicated. Plates were incubated at 37 °C for 20 h before being imaged. Data are representative of 3 biological replicates. (c) Anti-His immunoblot analysis of His-PaLpxC or His-EcLpxC protein levels in exponentially growing PAO1. Immunoblot for RpoA serves as a loading control. Data are representative of 3 biological replicates. For gel source data, see Supplementary Fig. 1.
Extended Data Fig. 2 LPS levels are altered upon mis-regulation of LpxC.
Silver stain of LPS harvested from exponentially growing cultures and western blot of RpoA from the same samples as a loading control. (a) PAO1 harboring an empty plasmid or one encoding PalpxC or EclpxC under arabinose-inducible control were induced with 0.1% arabinose 1 h prior to harvesting samples. (b) PAO1, PA1118 [∆murA Plac-murA], or PA1135 [∆murB Plac-murB] were grown in the presence or absence of 1 mM IPTG as indicated before samples were processed. (C) PAO1 harboring an empty plasmid or one encoding PamurA(WT) or PamurA(C117S) under IPTG-inducible control were induced with 1 mM IPTG 1 h prior to harvesting samples. Data are representative of 3 biological replicates. For gel source data, see Supplementary Fig. 1.
Extended Data Fig. 3 MurA is essential and its depletion phenocopies simultaneous inhibition of PG and LPS biosynthesis.
(a–c) Growth curves of P. aeruginosa strains in LB with or without IPTG as indicated. Dots represent the average of 3 biological replicates and dashed lines indicate the standard deviation. The following strains were used: (a) PAO1 [WT] and PA1080 [∆PA4701], (b,c) PAO1 [WT], PA1118 [∆murA Plac-murA], and PA1135 [∆murB Plac-murB]. (d) Phase contrast images of P. aeruginosa cells after 1 hr treatment with the indicated antibiotic(s). Scale bar indicates 2 µm. Data are representative of at least 2 biological replicates (e) Phase contrast images of the indicated P. aeruginosa murA or murB depletion strains grown for 4 h in the presence or absence of inducer as indicated. MurB depletion was analyzed as a control to compare the phenotype of inactivating another early step in PG synthesis with that of MurA. Scale bar indicates 2 µm. Data are representative of at least 3 biological replicates (f) Quantification of cell width after 1 hr treatment with the indicated antibiotic(s) or after depletion of MurA or MurB. Each dot represents an individual cell and the median of the population is indicated by a black line. n indicates the number of cells analyzed. For each condition, the cells quantified were derived from a single population and data are representative of biological duplicates.
Extended Data Fig. 4 PaMurA interacts with PaLpxC.
(a) H-PaLpxC in vivo pulldowns. The expression status of H-PaLpxC and F-PaMurA before (input) and after (elution) co-affinity purification using Ni-NTA resin is indicated above the immunoblots. The variant of F-PaMurA produced is indicated by WT for F-PaMurA(WT) or * for F-PaMurA(C117S). When indicated, 0.5 µg/mL CHIR-090 was added to cultures 1 hr prior to harvesting and was maintained in all lysis and wash buffers as detailed in the methods section. The following strains were used to generate the lysates: PA239 (lane 1), PA1013 (lane 2), PA1068 (lane 3), PA1071 (lanes 4 and 6), and PA1121 (lanes 5 and 7). Data are representative of at 3 biological replicates. (b) Purified F-PaLpxC (2.5 µM) was mixed in a 1:1 ratio with purified H-MurA variants in the presence or absence of CHIR-090 (5.7 µM) as indicated. The mixtures were pulled down with anti-FLAG resin and the input and elution subjected to SDS-PAGE and Coomassie staining. Mobilities of H-MurA and F-LpxC are indicated by a cyan or gold carrot, respectively. Data are representative of 3 replicates. For gel source data, see Supplementary Fig. 1.
Extended Data Fig. 5 Purified proteins used in this study and linearity of LpxC activity assay.
(a) Purified proteins used in this study were resolved by SDS-PAGE and protein was visualized with Coomassie staining. Data are representative of at least two 2 replicates. For gel source data, see Supplementary Fig. 1. (b) Time course in which turnover of UDP-3-O-(R-3-hydroxydecanoyl)-N-acetylglucosamine (PaLpxC substrate) to UDP-3-O-(R-3-hydroxydecanoyl)-glucosamine (PaLpxC product) by FLAG-PaLpxC over the course of 20 min was monitored by LC-MS. The R2 value of the linear regression is presented.
Extended Data Fig. 6 Mutations in the PaMurA active site are toxic but can be suppressed by inhibition of PaLpxC.
(a) Spot titer assay in which serial dilutions of PAO1 harboring an empty vector, one encoding PaMurA(WT) or the indicated PaMurA variant were plated on LB agar supplemented with IPTG and/or CHIR-090 as indicated. Plates were incubated at 37 °C for 20 h before imaging. † indicates the presence of a silent mutation in the construct. See Table S2 for details. Data are representative of 3 biological replicates. (b) Crystal structure of E. cloacae MurA (PDB 1EJC)14 in which residues corresponding to the identified PaMurA dominant negative alleles are depicted in gold spheres, or in the case of Cys117, a red sphere. Note that the substitutions all cluster around the active site. (c) MurA activity assay in which purified PaMurA variants (100 nM) were mixed with UDP-GlcNAc (1 mM) and PEP (0.5 mM) and the release of Pi was measured by Lanzetta assay. Dots indicate the values obtained for three individual replicates, bars indicate the mean, and error bars represent their standard deviation. (d) Catalytic activity of purified PaLpxC (100 nM) alone or in the presence of MurA variants (100 nM) assayed by conversion of UDP-3-O-(R-3-hydroxydecanoyl)-N-acetylglucosamine (PaLpxC substrate) to UDP-3-O-(R-3-hydroxydecanoyl)-glucosamine (PaLpxC product). Dots indicate the values obtained for three individual replicates, bars indicate the mean, and error bars represent their standard deviation. (e) LpxC enzymatic activity detected from preparation of MurA variants (100 nM) alone assayed as in panel d. Dots indicate the values obtained for three individual replicates, bars indicate the mean, and error bars represent their standard deviation.
Extended Data Fig. 7 LC-MS/MS analysis of PaLpxC substrate and product.
(a) Chemical structure of the PaLpxC substrate. The acetyl moiety removed by LpxC is highlighted in gold. Boxed regions indicate putative fragment ions highlighted in panels c, d, and h-j. (b) Extracted ion chromatogram (EIC) of UDP-3-O-(R-3-hydroxydecanoyl)-N-acetylglucosamine (PaLpxC substrate, 776.1986 m/z, black line) and UDP-3-O-(R-3-hydroxydecanoyl)-glucosamine (PaLpxC product, 734.1872 m/z, gold line) derived from in vitro reaction mixtures containing the PaLpxC substrate along with purified PaLpxC and PaMurA resolved using LC-MS operating in negative mode. The dashed lines indicate the peaks assigned to the PaLpxC substrate (S) and PaLpxC product (P). (c—d) MS/MS spectrum associated with the panel b peaks S and P, respectively. The parent ion is indicated by an asterisk and fragment ions corresponding to those highlighted in panel a are labeled. (e-g) EICs PaLpxC product and PaLpxC substrate detected by LC-MS in the aqueous fraction of methanol-chloroform extracted whole cell lysates. The dashed lines indicate the peak assigned to the PaLpxC substrate (S) and PaLpxC product peaks (P1 and P2). Note that the PaLpxC product has been previously reported to resolve as two peaks43, both of which were integrated to infer the relative product abundance. Data are representative of biological triplicates. (h-j) MS/MS spectrum associated with the panel g peaks S, P1, and P2, respectively. The parent ion is indicated by an asterisk and fragment ions corresponding to those highlighted in panel a are labeled.
Extended Data Fig. 8 MurA(G58D) and MurA(E406K) impact binding and activation of PaLpxC.
(a) Anti-FLAG immunoblot detecting F-PaMurA variants after 1 h of induction with 1 mM IPTG. A corresponding blot for RpoA was used as a loading control. Data are representative of 3 biological replicates. (b) MurA activity assay in which purified PaMurA variants (100 nM) were mixed with UDP-GlcNAc (1 mM) and PEP (0.5 mM) and the release of Pi was measured by Lanzetta assay. Dots indicate the values obtained for three individual replicates, bars indicate the mean, and error bars represent their standard deviation. The dashed line indicates the average catalytic activity of PaMurA(C117S) observed in Fig S7C. (c) in vitro pulldowns in which purified F-PaLpxC and H-MurA variants were mixed in a 1:1 ratio in the presence or absence of CHIR-090 and processed as in Extended Data Fig. 4b. Data are representative of at least two replicates. (d) Spot titer assay in which serial dilutions of a PAO1 strain harboring a PamurA deletion or PamurA(C117S) allele at the native locus complemented by a chromosomally-integrated, IPTG-inducible copy of PamurA(WT) were plated on LB agar with the indicated supplements. As indicated, the strains also contained an empty plasmid or one encoding PamurA(G58D) under arabinose-inducible control. Plates were incubated at 37oC for 20 h before being photographed. Data are representative of 3 biological replicates. For gel source data, see Supplementary Fig. 1.
Extended Data Fig. 9 PaLpxC and PaMurA are predicted to interact.
(a,b) AlphaFold2 predicted aligned error matrices of the PaLpxC/PaMurA complex and EcLpxC/EcMurA complex structures, respectively. (c) Distribution of initial interaction scores among all LpxC and MurA pairs analyzed. Red bars indicate the sequences used to train the final EVcomplex model and the score corresponding to the PaLpxC/PaMurA pair is indicated by a dashed line. (d) Model structure of the PaLpxC/PaMurA complex predicted by AlphaFold219. PaLpxC is represented in gold and PaMurA is represented in cyan. The top six intermolecular couplings between LpxC/MurA residues from the final EVcomplex model are highlighted in red with red lines connecting the coupling pairs. (e) Distribution of final interaction scores among all LpxC and MurA pairs analyzed. The data fit a two-component Gaussian mixture model (black line) indicating the presence of a population with low interaction scores (blue line) and high interaction scores (orange line). (f) Catalytic activity of purified LpnLpxC (100 nM) alone or in the presence of LpnMurA (200 nM) assayed by conversion of UDP-3-O-(R-3-hydroxydecanoyl)-N-acetylglucosamine to UDP-3-O-(R-3-hydroxydecanoyl)-glucosamine. Dots indicate the values obtained for three individual replicates, bars indicate the mean, and error bars represent their standard deviation. (g) Catalytic activity of purified EcLpxC (100 nM) alone or in the presence of EcMurA (200 nM) assayed by conversion of UDP-3-O-(R-3-hydroxydecanoyl)-N-acetylglucosamine to UDP-3-O-(R-3-hydroxydecanoyl)-glucosamine. Dots indicate the values obtained for three individual replicates, bars indicate the mean, and error bars represent their standard deviation.
Supplementary information
Supplementary Information
This document contains Supplementary Methods, Tables 1–5, and Supplementary References.
Supplementary Fig. 1
Original source images for all electrophoresis gels reported in this study.
Supplementary Data
Source data for Supplementary Table 1.
Rights and permissions
Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/.
About this article
Cite this article
Hummels, K.R., Berry, S.P., Li, Z. et al. Coordination of bacterial cell wall and outer membrane biosynthesis. Nature 615, 300–304 (2023). https://doi.org/10.1038/s41586-023-05750-0
Received:
Accepted:
Published:
Issue Date:
DOI: https://doi.org/10.1038/s41586-023-05750-0
Comments
By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.
