## 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.

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).

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).

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.