Structural snapshots of Pseudomonas aeruginosa LptB_2FG and LptB_2FGC reveal insights into lipopolysaccharide recognition and transport

Domain: physics
Published: 2025-12-17
Editorial outcome: accept

Abstract
Gram-negative bacteria are intrinsically resistant to many antibiotics because of densely packed lipopolysaccharides (LPS) in the outer leaflet of their outer membrane (OM), which acts as a highly effective barrier towards the spontaneous permeation of toxic molecules, including antibiotics. LPS are extracted from the inner membrane by the ABC transporter LptB2FGC and translocated across the periplasm via a protein bridge to the OM. While structural studies have elucidated aspects of Lpt function in enterobacteria, little is known about how this system operates in divergent species such as Pseudomonas aeruginosa, a major human pathogen. Here, we report five cryo-electron microscopy structures of P. aeruginosa LptB2FG and LptB2FGC, revealing a rigid body movement in the periplasmic β-jellyroll domains necessary for LPS to shuttle through the periplasmic space. Notably, these structures exhibit a significantly smaller LPS binding cavity compared to previously determined models, suggesting the ligand-unbound states of the transporter. Mass spectrometry and molecular dynamics simulations indicate that the phosphate groups of LPS are the key determinants for binding and that the transporter can also accommodate cardiolipin. Together, these findings reveal previously unappreciated structural diversity in the Lpt system and provide mechanistic insight into how pathogenic Gram-negative bacteria tailor LPS recognition and transport. This understanding offers new avenues for the development of novel inhibitors targeting membrane biogenesis.

Introduction
Infections caused by Gram-negative bacteria are difficult to treat in part, owing to the low permeability of the outer membrane (OM) to antibiotics. Consequently, few antibiotics are active against Gram-negative bacteria, and multidrug-resistant strains are rising. The inner membrane (IM) of Gram-negative bacteria is mainly composed of phospholipids, while the OM is an asymmetric bilayer presenting lipopolysaccharides (LPS) on the outer leaflet and phospholipids in the inner leaflet. The amphiphilic nature of LPS guarantees bacterial protection towards both polar molecules and toxic hydrophobic compounds, such as antibiotics, and is essential for stabilising the overall OM structure.

LPS is a glycolipid consisting of lipid A, core oligosaccharide and O-antigen polysaccharide. Mature LPS is extracted from the outer leaflet of the IM and transported through the periplasm to be inserted into the outer leaflet of the OM by seven essential proteins LptA-G. The IM sub-complex comprises the ATP-binding cassette (ABC) transporter LptB2FGC, which extracts LPS from the IM and delivers it to the periplasmic components of the Lpt system. The periplasmic protein LptA (named LptH in Pseudomonas aeruginosa) forms a continuous bridge with LptC and LptD β-jellyroll domains spanning across the periplasm. Finally, the heterodimer LptDE inserts LPS into the outer leaflet of the OM. In some Gram-negative bacteria, additional lipoproteins contribute to LPS transport efficiency and system stabilisation. For example, LptM has been proposed to act as an accessory factor that stabilises LptD assembly or modulates transport under stress conditions. More recently, the lipocalin YedD was identified as a component of the translocon and is required for optimal LPS transport.

LptB2FGC is a type VI ABC transporter consisting of 5 subunits. The LptB cytoplasmic subunits form the nucleotide-binding domains (NBDs) that capture and hydrolyse ATP. LptF and LptG possess transmembrane helices that create a cavity and a translocation path for LPS, as well as a periplasmic β-jellyroll domain. Differently from other ABC transporters, this sub-complex comprises the single-pass membrane protein LptC with its transmembrane helix situated between the two transmembrane domains (TMDs) of LptF and LptG and plays an important role in regulating the ATPase activity to allow higher LPS extraction efficiency of the protein complex. LptC also contains a β-jellyroll domain, which is in contact with LptF and LptA in the periplasm to form a periplasmic bridge. Structural data for LptB2FGC have largely been derived from enterobacteria such as Escherichia coli, Klebsiella pneumoniae, Enterobacter cloacae, and Shigella flexneri, with only two more divergent homologues characterised in Vibrio cholerae and Acinetobacter baylyi. In these complexes, the periplasmic β-jellyroll domains have been resolved only in E. cloacae and V. cholerae, where LptC interacts directly with LptF to form a simple heterodimer, while LptG appears to act primarily as a structural scaffold.

Genetic studies have shown that P. aeruginosa LptC (Pa-LptC) cannot functionally substitute E. coli LptC, likely due to a lack of assembly with the Ec-LptB2FG complex; Pa-LptC shares only ~20% sequence identity with its E. coli counterpart, highlighting the divergence between these two Lpt systems. Therefore, which β-jellyroll of LptF and LptG partners with Pa-LptC and serves as the LPS exit pathway, has been unclear. The first crystal structure of Pa-LptB2FG suggested that either LptF or LptG could both serve as the lateral LPS exit route. This prompted the proposal of an alternating lateral access model, later discounted by cryo-EM structures that confirmed LptF as the primary LPS exit route.

Previous structural investigations using X-ray crystallography and cryo-EM for LptB2FG and LptB2FGC have successfully unveiled the proposed mechanism for the structural basis of LPS extraction by LptB2FGC. Nevertheless, several unresolved questions persist regarding the LptB2FGC transporter. The true structure of its apo state remains unclear, since all previously published cryo-EM structures contain LPS, while X-ray structures present similarly sized cavities, although LPS-free. This raise concerns as to whether these X-ray structures truly represent apo forms. In addition, the range of configurations adopted by the β-jellyroll domains throughout the LPS transport cycle is not fully understood. The mechanisms that enable LPS to enter the transmembrane binding cavity, particularly when obstructed by the transmembrane helix of LptC, also remain elusive. Moreover, it is uncertain how selective the transporter is for LPS or whether it might recognise other membrane lipids. Finally, it is not clear how LptF and LptG might contribute differently to transport in divergent Gram-negative bacteria.

In the present study, we have employed an integrated approach to address these unresolved issues. Guided by native mass spectrometry (MS), we optimised and purified an LPS-free LptB2FG(C) transporter of P. aeruginosa. Then, using cryo-EM, we uncovered three major structural populations of LptB2FG and two major populations of LptB2FGC featuring a rigid body movement in the β-jellyroll domains. Further comparison with existing structures indicated that our structures represent the ligand unbound states of the transporter. We further used native MS to probe the structure-activity relationship of LPS and studied how membrane lipids such as cardiolipin influence LPS binding. These results, supported by molecular dynamics simulations, reveal critical residues involved in lipid recognition. Our data provide new mechanistic insights into LPS transport and expand the structural landscape of LptB2FGC complexes from non-enterobacterial species. Our findings identify functionally important distinctions in the P. aeruginosa LptB2FGC complex, provide a structural explanation for the previously observed genetic incompatibility, and highlight opportunities for species-specific antibiotic development.

Results
Purification of LPS-free LptB2FG(C)
We initially expressed P. aeruginosa LptB2FGC in the Escherichia coli strain BL21(DE3) and purified this protein complex following a classic two-step purification protocol consisting of immobilised metal affinity chromatography (IMAC) and size-exclusion chromatography (SEC), as indicated from previous studies. The protein complex was initially purified in a buffer containing 0.03% (w/v) n-dodecyl-β-D-maltopyranoside (DDM), following a delipidation protocol to remove bound lipids, especially LPS. We then performed a detergent screen to establish the ideal conditions for native MS analysis. Among the tested detergents, the recently reported [G1], dendritic oligoglycerol detergent ([G1]-OGD) showed an ability to retain non-covalent interactions and allowed for facile detergent removal from proteomicelles, thus yielding well-resolved spectra. Native mass spectra in [G1]-OGD revealed charge states corresponding to both LptB2FG and LptB2FGC complexes, which may be due to either an LptB2FG/LptB2FGC equilibrium or lower expression levels of LptC compared to other subunits. Moreover, we observed adducts ranging from 1.5 to 4.7 kDa, which are consistent with co-purified phospholipids (PLs) and LPS. Overall, the presence of these adducts indicates the potential absence of a truly apo form of the LptB2FG and LptB2FGC complexes from previous studies, which could explain why the LPS binding cavities in the X-ray and cryo-EM structures are of the same size.

In the quest for an in-depth understanding of the mechanism underpinning LptB2FGC-mediated LPS extraction from the outer leaflet of the IM, we aimed to obtain LPS-free LptB2FG and LptB2FGC complex structures. Hence, we overexpressed the protein complex in ClearColi BL21(DE3) cells. ClearColi (genotype: ΔgutQ, ΔkdsD, ΔlpxL, ΔlpxM, ΔpagP, ΔlpxP, ΔeptA, msbA148) cells are able to produce only lipid IVA, in place of the characteristic hexa-acylated enterobacterial lipid A, as the only LPS-related molecule in the outer membrane. We purified the protein, expressed in ClearColi, in DDM following the same procedure as described above. We confirmed with SDS-PAGE that LptC was successfully co-expressed with the other complex subunits. Next, we carried out native MS experiments in [G1]-OGD, which showed the presence of charge states corresponding to both LptB2FG and LptB2FGC. Notably, no lipid-bound species were detected, indicating that we were able to successfully remove lipid IVA or any other membrane lipid potentially associated with the transporter during expression in ClearColi cells. As a result, we purified the ligand-unbound forms of the two protein complexes. With the improved resolution of the mass spectra, we were able to determine the relative abundance of the two protein complexes and estimate a 1:1 ratio between LptB2FG and LptB2FGC.

Purified LptB2FG(C) has ATPase activity
To further confirm that the purified LptB2FG(C) is active, we performed an ATPase assay in both purification and MS buffer, and the values obtained are in line with previously reported ones, verifying that the complexes we obtained are in their active forms.

In addition, previous reports indicated that tagging LptB at the C-terminus compromises its function in E. coli but retains ATPase activity in vitro. To assess this possibility for P. aeruginosa, we performed a functional assay using an lptB conditional mutant (PAO1 rhaSR-PrhaBAD-lptB ΔlptB) transformed with plasmids that express LptB of P. aeruginosa PAO1 or the LptB variant used in this study with or without the C-terminal 6×His tag. Rhamnose and IPTG were used to induce the chromosomal copy of lptB or the lptB allele cloned into the plasmid pME6032, respectively. Functional assays demonstrated that the His-tagged variants successfully complemented the lptB conditional mutant, even though the colonies were slightly smaller than those of the control strains expressing non-tagged LptB. This indicates that the tagged LptB variants are at least partially functional in vivo in P. aeruginosa. Together, these findings confirm that our purified LptB2FG(C) complexes are enzymatically competent in vitro and that the tagged LptB variants support growth in complementation assays, and we therefore used these preparations for the structural and biochemical analyses reported here.

Substrate-free structures of LptB2FG in lipid nanodiscs
Having succeeded with the purification of functional apo states of LptB2FG/LptB2FGC, we reconstituted the protein complexes into lipid nanodiscs and collected single-particle images using cryo-EM. We then processed this cryo-EM data using the Build and Retrieve (BaR) methodology, following the steps described previously. Extensive classification of the single-particle images indicated there are several distinct populations of LptB2FG and LptB2FGC with various conformations coexisting in the cryo-EM sample. Several iterative rounds of classifications allowed us to sort the images based on five distinct conformations, which correspond to three different LptB2FG and two different LptB2FGC states.

Three distinct conformations of LptB2FG, designated as LptB2FG-I, LptB2FG-II, and LptB2FG-III, were captured in our cryo-EM data and solved to resolutions of 3.34 Å, 3.26 Å, and 3.31 Å, respectively. Overall, all the cryo-EM structures of LptB2FG adopt the fold of the previously solved LptB2FG structure, in which two copies of LptB participate in forming the NBDs, whereas LptG and LptF are engaged in creating the TMDs made up of 12 transmembrane helices and the periplasmic β-jellyroll domains. The main difference between the three cryo-EM structures of LptB2FG is in the conformation of the β-jellyroll domain, which appears to perform a rigid body rotation motion with respect to the outer leaflet of the IM, transitioning from one conformational state to the other. The TMD and NBD domains of these three structures are essentially the same. The transmembrane helices create an outward-open cavity with two interfaces: Interface 1, formed by the interaction between TM1LptG and TM5LptF, and Interface 2, established by the connection between TM1LptF and TM5LptG.

In contrast to all the reported structures of LptB2FG determined by cryo-EM, our cryo-EM structures did not reveal any density consistent with the presence of LPS within the outward-facing cavity of the TMDs. This cavity has a volume of 5,473 Å3, approximately half the volume observed in the X-ray structure, suggesting that the previously solved structure may represent an LPS-bound conformation. Intriguingly, our structures of LptB2FG exhibit an open V-shaped cleft at Interface 1, oriented toward the outer leaflet of the inner membrane, while Interface 2 remains entirely closed. This structural feature is expected to facilitate lipids entering the TMD domain of LptB2FGC via Interface 1. These observations are in line with biochemical and crosslinking investigations that establish Interface 1 as the point of LPS entry. The distinctive structural attributes characterising our cryo-EM LptB2FG structures strongly suggest that we have captured the resting state, or substrate-free conformation, of this transporter.

The open and closed states of interfaces 1 and 2 likely involve rotations of two rigid-body groups within the TMD of LptF. The movement of the first rigid-body group (TM1-3) would be linked to the opening and closing of interface 2, while the second rigid-body group (TM4-5) would regulate access to interface 1. In our substrate-free LptB2FG structures, both rigid groups 1 and 2 rotate counterclockwise toward LptG (viewed from the top of the transmembrane domain), causing group 2 to move away from LptG, thereby opening interface 1. Simultaneously, group 1 moves toward LptG, closing Interface 2 and reducing the volume of the transmembrane domain binding cavity.

We identified four lipid molecules in LptB2FG-II and LptB2FG-III, likely originating from the nanodisc preparation used for sample reconstitution. These lipids were modelled as phosphatidylethanolamine (PE) moieties, arranged on the outer surface of the LptB2FG complex. Among these lipids, PE1 was located outside of interface 1, at the level of the outer leaflet of the cytoplasmic membrane. The other three lipids were positioned in the inner leaflet, with PE2 located in the vicinity of interface 1, and PE2 and PE3 at interface 2. In the LptB2FG-I structure, we observed PE1, PE3, and PE4.

Structures of resting conformation of LptB2FGC in lipid nanodiscs
We also obtained two cryo-EM 3D reconstitutions of LptB2FGC at resolutions of 3.26 Å and 3.61 Å, respectively. These two structures were assigned as LptB2FGC-I and LptB2FGC-II. We then refined the cryo-EM structure of these two complexes. Structure alignment of these two structures gives rise to a root mean square deviation (RMSD) of 1.2 Å over 1324 Cα atoms and indicates the main difference between these two cryo-EM structures is in the β-jellyroll structures. These regions appear to perform a rigid body rotation motion toward the outer leaflet of IM, similarly to what we observed for LptB2FG. The TMD and NBD domains of LptB2FGC are similar to those of the LptB2FG structures and represent the resting state. In our LptB2FGC structures, the periplasmic β-jellyroll of LptC was clearly resolved and found to form a trimeric interface: LptC and LptF form the primary LPS transfer path, while a loop from LptG (residues 209–215) binds onto the N-terminal β-jellyroll of LptC (residues 60–69). In contrast, in the LptB2FGC structure from Vibrio cholerae (6MJP), LptC interacts predominantly with LptF, and LptG makes only minimal contact with LptC, resulting in an interface that resembles a heterodimer rather than a heterotrimer.

Intriguingly, in contrast to previously documented LptB2FGC structures, the transmembrane helix of LptC remained unobservable in these maps, indicating its flexibility and a lack of engagement with LptF and LptG at Interface 1. This, in turn, results in an unobstructed entry point for LPS into the TMD cavity. Similar to LptB2FG, we identified four lipids in the transmembrane region of the form I conformation, which likely originate from nanodisc reconstitution. However, for LptB2FGC-II, the lower resolution and particle counts in this conformation prevented confident lipid identification. A comparison of our cryo-EM structure of LptB2FGC-I with the previously reported V. cholerae LptB2FGC (6MJP) reveals a significant conformational difference (RMSD 5.57 Å across 845 Cα atoms), which may reflect distinct functional states.

LPS binding to LptB2FG
To test whether the purified P. aeruginosa LptB2FG(C) complex could still bind LPS after delipidation, we tested the binding in vitro with exogenously added LPS. We performed native MS using both P. aeruginosa LPS (Pa-LPS) and E. coli LPS (Ec-LPS) at defined concentrations to evaluate binding preferences and specificity. Since LPS is a heterogeneous mixture and to avoid the complications in interpreting our native MS analysis, we have purified P. aeruginosa LptB2FG (without LptC) using our established methodology from above.

Native MS analysis revealed robust binding of both LPS types to LptB2FG, with distinct adduct peaks corresponding to lipid-bound complexes. A clear preference was observed for Pa-LPS over Ec-LPS, as shown by higher signal intensity and multiple binding events at equivalent LPS concentrations. This affinity difference suggests that the transporter has adapted to recognise native P. aeruginosa LPS more tightly, likely due to subtle differences in the lipid A or core oligosaccharide structures between species.

Nevertheless, the transporter remains capable of recognising and interacting with Ec-LPS, as also evidenced by the challenges encountered in optimising the purification of the substrate-bound LptB2FGC complex. Overall, these findings are consistent with prior reports showing that LptB2FG(C) complexes purified from E. coli often contain endogenous Ec-LPS irrespective of the species of origin, underscoring the transporter’s baseline promiscuity. Our data refine this understanding by demonstrating that, while LptB₂FG can bind LPS from multiple sources, it exhibits measurable specificity that likely reflects physiological optimisation.

Key determinants of LPS involved in binding to the LptB2FG(C) complex
To dissect the specific features of LPS required for recognition by the LptB₂FG(C) complex, we performed native MS experiments using defined LPS analogues: Re-LPS, lipid A, and 4′-monophosphoryl lipid A (MPLA). These compounds differ progressively in their structural complexity: Re-LPS contains lipid A and two Kdo (3-deoxy-D-manno-oct-2-ulosonic acid) sugars; lipid A lacks the Kdo moieties; and MPLA is missing both Kdo and the phosphate group at position 1 of the glucosamine backbone. Only Re-LPS has been used previously to investigate Lpt system functionality. Because of the lack of commercial availability of these variants from P. aeruginosa, we used E. coli versions for our study.

We incubated the LptB2FG/LptB2FGC solution with increasing concentrations of Re-LPS (0.1 μM to 5 μM) and recorded mass spectra under our optimised conditions. We could detect the presence of 2.2 kDa adduct in both LptB2FG and LptB2FGC charge states, consistent with Re-LPS (MW = 2239 Da) binding. The native mass spectra revealed a higher degree of total Re-LPS binding to LptB2FGC than LptB2FG. To quantify the binding affinity, we determined the apparent dissociation constant (KD) for individual lipid-binding events to LptB2FG and LptB2FGC using a sequential lipid-binding model. In line with total Re-LPS binding quantification, LptB2FGC displayed sub-micromolar binding affinity for Re-LPS (KD(1) = 0.65 ± 0.02 μM). By comparison, the apparent KD of LptB2FG:Re-LPS was more than twofold higher (KD(1) = 1.45 ± 0.05 μM), suggesting that LptC contributes to high-affinity substrate binding. The increase in affinity may result from either a decreased off-rate of LPS from the central cavity or increased binding to periplasmic domains of the transporter. Because our data are equilibrium occupancies, we cannot distinguish between changes in association and dissociation rates here; accordingly, we present the reduced-off-rate explanation as a working hypothesis aligned with prior structural models of LptC-mediated gate opening. While we do not infer periplasmic-domain engagement from our assays, we cannot completely exclude additional interactions under our conditions because the exact TM-helix position is unknown.

We then investigated the structure-binding relationships for protein-substrate interaction to elucidate the essential moieties for LPS binding. We incubated the LptB2FG/LptB2FGC solution with lipid A and 4’-monophosphoryl lipid A (MPLA) at increasing concentrations. Using native MS, we observed both lipids bound to LptB2FGC at both concentrations employed. Quantification of the relative abundance of lipid-bound LptB2FGC indicated a significant decrease of binding affinity for lipid A and MPLA, with the latter being the weakest interacting species. In the case of LptB2FG, we again observed lower overall lipid binding with a similar trend regarding the relative affinities of the different species. These observations suggest that the Kdo units in the inner core of LPS are important moieties for the interaction between LptB2FGC and its substrates. Similarly, the phosphate group on C1 likely engages in critical interactions with residues from LptB2FGC TMDs. The decreased binding of lipid A and MPLA to LptB2FG compared to LptB2FGC also suggests that LptC is actively involved in recognising portions with the Kdo sugars and the C1-bound phosphate group.

Consistent with these binding trends, structural analyses and MD simulations indicate that the phosphate group on the glucosamine unit is positioned in the upper region of the LPS binding pocket and interacts with conserved positively charged residues, such as Lys33 in LptF. Comparisons with existing LptB2FG(C)–LPS structures show species-dependent variation in this interaction, highlighting a flexible but essential recognition mechanism for LPS transport. Together, these data underscore the critical role of both the Kdo sugars and the C1 phosphate group in stabilising LPS binding within the transporter.

Lipid binding to the LptB2FG(C) complex
Recent studies indicated that cardiolipin (CDL) aids in MsbA-mediated LPS transport, so we sought to probe if there is a similar relationship between LptB2FG/LptB2FGC and CDL. In particular, we chose 14:0 CDL to be consistent with the 14 carbon lipid tails of Re-LPS, and incubated this with the LptB2FG/LptB2FGC complex, we could observe ~50% of LptB2FGC bound to CDL at 2.5 μM concentration. Apparent KD calculation indicated a similar trend when comparing LptB2FG and LptB2FGC interaction with CDL to that of LPS. Indeed, CDL binds more tightly to LptB2FGC (KD(1) = 3.07 ± 0.09 μM) than LptB2FG (KD(1) = 4.80 ± 0.14 μM). This finding suggests a role for LptC in interacting with this phospholipid. We set out to test whether CDL influences LPS binding to LptB2FGC. To this end, we added increasing concentrations of CDL (1.0 to 5 μM) to a solution containing LptB2FGC:Re-LPS complex. Following CDL addition, native MS indicated that the total amount of Re-LPS bound to the protein decreased substantially in a concentration-dependent manner. These results suggest that the binding cavity of LptB2FG is capable of recognising CDL.

From a physiological standpoint, these findings suggest that LptB2FGC may be sensitive to local membrane lipid composition, particularly under conditions where cardiolipin is enriched, such as during stress responses, antibiotic treatment, or the stationary phase. This could reflect a regulatory mechanism by which cardiolipin modulates LPS extraction efficiency or prioritises substrate selection under specific conditions.

Our MD simulations point to shared contact sites with Re-LPS, including interactions with charged residues in TM1 and TM5 helices. Whether CDL acts purely as a competitive ligand or as a functional co-factor influencing transport dynamics is an open question, meriting further investigation.

MD simulations highlight key coordinating residues for lipid binding in the central cavity
To explore the molecular basis for lipid recognition within the LptB2FG binding cavity, we performed molecular dynamics (MD) simulations with either Re-LPS, Lipid A, MPLA or CDL bound to the central cavity. We then calculated the protein-lipid contacts made within 4 Å, over the course of three 500 ns simulations. Despite the asymmetry between LptF and LptG subunits, common domains were observed to interact with the bound lipids, with the majority of contacts formed with TM1, TM2 and TM5 helices. For LptF, the bound lipids interacted with I25, I26, G29, R30 and K33 (TM1), L62, I63 and L66 (TM2), D154 and S246 in the periplasmic domain, and L305, I308, L309, M312, L315, A316 and I319 (TM5). For LptG contacts were made with I23, L26, A27, F30, I33, D34, N37 and D38 (TM1), R59, M63, M66 and I70 (TM2), R132 (TM3), K145, R146, E240 and R265 (periplasmic domain) and V308, F312, R315 and D319 (TM5). These residues likely contribute to a combination of hydrophobic and electrostatic interactions, with the arginine and lysine residues engaging with the negatively charged phosphate head groups and the hydrophobic residues interacting with the acyl chains of LPS and CDL. Notably, residues such as I25, K33, and L62 of LptF, and L26, D34, R132, and R317 of LptG correspond to key contact points identified in the S. flexneri, E. coli, A. baylyi and K. pneumoniae structures. Among the 38 cavity-contact residues highlighted by MD (17 LptF; 21 LptG), prior functional data exist for a subset of positions in enterobacteria (although not directly tested in P. aeruginosa), most notably hydrophobic/charged residues lining the cavity whose mutation compromises Lpt function or growth (e.g., LptF I25/R30/L62; LptG L26/R59/I70/R132/V308/F312). These studies collectively support the idea that the central cavity residues we observe contacting acyl chains in MD are functionally important, although numbering and exact identities vary by species/construct.

Although the contact profiles for Re-LPS, Lipid A, MPLA, and CDL overlapped substantially, Re-LPS formed more persistent and extensive interactions, suggesting greater shape and charge complementarity within the binding cavity. In summary, these simulations provide mechanistic insight that complements our experimental data, identifying specific amino acids likely responsible for lipid selectivity.

Methods
LptB2FGC expression and membranes isolation
The plasmids used for overexpression of P. aeruginosa LptB2FG and LptB2FGC were a kind gift from Yihua Huang (Chinese Academy of Sciences, Beijing, China). The plasmids were constructed as previously described (Luo et al., 2017) and consisted of pQLink-LptBFG or pQLink-LptBFGC, whereby each gene (lptB, lptC, lptF and lptG) had been amplified from P. aeruginosa PAO1-LAC genomic DNA (47085D-5 ATCC) using PCR and inserted in the pQlink plasmid by ligation-independent pathway. To allow affinity purification, a 6xHis-tag was attached to the C-terminus of LptB. All plasmids were amplified by transforming them into E. coli Stellar Competent Cells (Takara), and the DNA sequences were verified by Sanger sequencing.

Initial expression and purification were performed adapting previously described protocol. The amplified plasmid was transformed in E. coli BL21(DE3) (Lucigen). Colonies were inoculated into 100 ml LB media and grown overnight at 37°C supplemented with 100 μg/ml ampicillin. One litre of LB in 2 litre shaker flasks was inoculated with 10 ml of overnight culture and grown at 37°C to a density of 0.6-0.7 at 600 nm at 37°C, and the expression of the recombinant protein complex was induced by 0.5 mM isopropyl-β-D-thiogalactopyranoside (IPTG) at 20°C for 16 h. Cells were collected by centrifugation at 5000xg for 10 min at 4°C. Cell pellets were resuspended in buffer containing 150 mM NaCl, 20 mM Tris-HCl (pH 8.0) and stored at −80°C.

Resuspended cells were thawed and supplemented with EDTA-free protease inhibitor cocktail (Roche) and 10 μg/ml DNase I. The cell suspension was passed several times through an M-110 PS microfluidiser (Microfluidics) at 15,000 psi. Insoluble material was pelleted by centrifugation at 20,000xg for 20 min at 4°C. The supernatant was and then ultracentrifuged at 200,000xg for 1 h to collect membrane fractions. Membranes were resuspended in ice-cold buffer containing 250 mM NaCl, 20 mM Tris-HCl (pH 8.0) and 20% glycerol. Resuspended membranes were used either directly or flash frozen in liquid nitrogen and stored at −80°C.

To fully remove LPS adducts, we then expressed LptB2FG and LptB2FGC in ClearColi BL21(DE3) (Lucigen), an E. coli strain holding different gene knockouts that stop the biosynthetic pathway of LPS to a precursor, lipid IVA. The amplified pQLink-LptBFGC plasmid was transformed in ClearColi BL21(DE3) through electroporation, and the plates were incubated for 32-40 h at 37°C. Colonies were inoculated into 100 ml LB media and grown overnight at 37°C supplemented with 100 μg/ml ampicillin. One litre of LB in 2 litre shaker flasks was inoculated with 40 ml of overnight culture and grown at 37°C to a density of 0.6-0.7 at 600 nm at 37°C, and the expression of the recombinant protein complex was induced by 0.5 mM IPTG at 20°C for 16 h. The steps for cell lysis and membrane isolation are the same as the ones described above.

LptB2FG/LptB2FGC purification
The protein was solubilised from the membrane fraction with 250 mM NaCl, 20 mM Tris-HCl (pH 8.0), 20% glycerol, 2% (w/v) DDM, 2% (w/v) OGNG (Anatrace) for 16 h at 4°C. Insoluble material was removed by centrifugation at 20,000xg for 20 min at 4°C. The supernatant was filtered before loading onto a 5 ml HisTrap-HP column (GE Healthcare, Piscataway, NJ) equilibrated in 150 mM NaCl, 20 mM Tris-HCl (pH 8.0), 20 mM imidazole, 1% (w/v) DDM, and 1% (w/v) OGNG. After the clarified supernatant was loaded, the column was initially washed with 500 ml (100 CVs) of 150 mM NaCl, 20 mM Tris-HCl (pH 8.0), 20 mM imidazole, 1% (w/v) DDM, and 1% (w/v) OGNG, and washed again with 50 ml of 150 mM NaCl, 20 mM Tris-HCl (pH 8.0), 80 mM imidazole and 0.1% DDM. The bound protein was eluted with 150 mM NaCl, 20 mM Tris-HCl (pH 8.0), 300 mM imidazole and 0.03% DDM.

The protein was concentrated to 2.5 ml an Amicon Ultra-15 concentrator unit (Millipore) with a molecular cut-off of 100 kDa and buffer exchanged to 150 mM NaCl, 20 mM Tris-HCl (pH 8.0), 10% glycerol and 0.03% DDM using a PD-10 desalting column. The sample was further concentrated and loaded onto the Superdex 200 size exclusion chromatography (SEC) column in 150 mM NaCl, 20 mM Tris-HCl (pH 8.0), 10% glycerol and 0.03% DDM. Protein concentration was measured using a DS-11 FX Spectrophotometer (DeNovix).

Lipids, nucleotides, and peptide preparation
Unless stated otherwise, all lipids [Kdo2-Lipid A (Re-LPS), 1,2-dimyristoyl-sn-glycero-3-phospho-(1’-rac-glycerol) (14:0 PG, DMPG), 1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine (14:0 PE, DMPE) 1’,3’-bis[1,2-dimyristoyl-sn-glycero-3-phospho]-glycerol (14:0 cardiolipin, CDL) used in this study were obtained from Avanti Polar Lipids powders and stock solutions were prepared following a previously established method. Full-length E. coli LPS and P. aeruginosa LPS are obtained from Sigma (catalogue numbers L3012 and L8643, respectively).

Native MS experiments
Purified LptB2FG(C) was buffer exchanged into MS Buffer (200 mM ammonium acetate + 0.04% (w/v) [G1]-OGD) using a centrifugal buffer exchange device (MicroBio-Spin 6, Bio-Rad) as previously described. The freshly buffer-exchanged protein was kept on ice, with protein concentration measured as before. The protein samples were diluted as desired in 200 mM ammonium acetate buffer with detergent as necessary and loaded into a gold-coated capillary Clark borosilicate capillary (Harvard Apparatus) prepared in the laboratory. The experiments were performed using a Q Exactive UHMR Hybrid Quadrupole-Orbitrap mass spectrometer (Thermo Fisher Scientific). Typically, 2 μl of buffer-exchanged protein solution was electrosprayed in negative polarity from in-house prepared gold-coated borosilicate capillaries. The instrument parameters used for data collection were: capillary voltage 1.1 kV, S-lens RF 100%, quadrupole selection from 1000 to 20,000 m/z range, HCD collision energy 200–300 V, source fragmentation 0 V, in-source trapping 200–300 V in negative mode. The ion transfer optics was set as follows: injection flatapole −5 V, inter-flatapole lens −4 V, bent flatapole −2 V, transfer multipole 0 V. The resolution of the instrument was 8750 at m/z = 200 (transient time of 64 ms), argon pressure in the HCD cell was maintained at approximately at 8 × 10–10 mbar and source temperature was kept at 200°C. The noise level was set at 3 rather than the default value of 4.64. Calibration of the instruments was performed using 10 mg/ml solution of caesium iodide in water. Where required, baseline subtraction was performed to achieve a better-quality mass spectrum. Where required, baseline subtraction was performed to achieve a better-quality mass spectrum. Data were analysed using the Xcalibur 3.0 (Thermo Scientific), NaViA, and UniDec (www.unidec.chem.ox.ac.uk) software packages.

Lipid and nucleotide binding experiments were performed by diluting them in 200 mM ammonium acetate supplemented with 0.04% [G1]-OGD. Native MS experiments were performed at a total LptB2FG(C) protein concentration of 3 μM.

Peak intensities were extracted using UniDec, and the ratios of the intensity of the ligand bound peak versus the total intensity of all observed species were calculated. The apparent dissociation constant (KD) values for individual lipid-binding events to LptB2FG and LptB2FGC were calculated through a sequential lipid-binding model using a previously described Python script. When necessary, a Welch’s T-test was used to evaluate statistically significant differences and data were plotted using GraphPad Prism 8.0. Error bars in the plots indicate standard deviation, and all experiments were performed in triplicate (n=3).

ATPase activity assay
The ATPase activity of LptB2FG, LptB2FG(C), and LptB2FG(C) in native MS buffer were monitored using the ATPase Assay Kit from Abcam (ab270551) following the manufacturer’s protocol. To monitor the ATPase activity, all assay components were allowed to warm up to room temperature prior to use, including the protein. The assay buffer consisted of SEC purification buffer or native MS buffer supplemented with 1 mM ATP and 5 mM Mg2+. Solutions were added to each well of a 96-well plate (Greiner 96F-Bottom) in the following order: (i) assay buffer (100 µL), (ii) protein (100 µL of a 0.5 μM solution in the relevant buffer), (iii) wait 30 min, (iv) PiColorLockTM (50 µL), (v) wait 2 min, (vi) stabiliser (20 µL). Half an hour after adding the stabiliser, the absorbance values at 610 nm (A610) were determined using a PerkinElmer EnSight multimode plate reader. To determine the molar amount of phosphate generated by each protein preparation, a standard curve was determined according to the instructions given in the protocol provided with the kit. ATPase activities of all samples were determined using the mean value of the samples according to the linear regression of standards. The data were plotted using GraphPad Prism 8.0. Error bars in the plots indicate standard deviation, and all experiments were performed six times (n=6).

Nanodisc preparation
To assemble LptB2FG(C) into nanodiscs, a mixture containing 10 μM LptB2FG(C), 30 μM membrane scaffold protein (MSP; 1D1), and 300 μM E. coli total extract lipid was incubated for 15 min at room temperature. After, 0.8 mg/ml prewashed Bio-Beads (Bio-Rad) was added. The resultant mixture was incubated for 1 h on ice, followed by overnight incubation at 4°C. The protein-nanodisc solution was filtered through 0.22 μm nitrocellulose filter tubes to remove the Bio-Beads. To separate free nanodiscs from LptB2FG(C)-loaded nanodiscs, the filtered protein-nanodisc solution was purified using a Superdex 200 column (GE Healthcare) equilibrated with 20 mM Tris-HCl (pH 7.5) and 100 mM NaCl. Fractions corresponding to the size of the LptB2FG(C)-nanodisc complex were collected for cryo-EM.

Electron microscopy sample preparation
The LptB2FG(C)-nanodisc sample was concentrated to 1 mg/ml. A 2.5 μl sample was applied to glow-discharged holey carbon grids (Quantifoil Cu R1.2/1.3, 300 mesh), blotted for 5 s, and then plunge-frozen in liquid ethane using a Vitrobot (Thermo Fisher). The grids were transferred into cartridges. For high-resolution data collection, the sample grids were loaded into a Titan Krios cryo-electron microscope operated at 300 kV equipped with a Gatan BioQuantum imaging filter (GIF) and a K3 summit direct electron detector (Gatan). The micrographs were recorded using Latitude software (Gatan) with counting mode at nominal ×81,000 magnification corresponding to a calibrated pixel size of 1.08 Å (super-resolution, 0.54 Å/pixel) and a defocus range of −1 to −2.5 μm. To remove inelastically scattered electrons, the slit width was set to 20 eV. Each micrograph was exposed for 2 s with a total specimen dose of 50 e-/Å2, and 40 frames were captured per specimen area.

Cryo-EM data processing
The LptB2FG(C) dataset was processed using a BaR protocol. Super-resolution image stacks were aligned and binned using patch motion correction (cryoSPARC) with a binning factor of 2 to give a final pixel size of 1.08 Å/pixel. Contrast transfer functions (CTFs) were estimated using the patch CTF in cryoSPARC. After manual inspection to discard poor images and to estimate particle size, the blob picker in cryoSPARC was used to select particles from subsets of micrographs. These particles were classified in several rounds of 2D classification, and clear templates were selected for template picking in cryoSPARC. The template picker was used to select initial particle sets. Several iterative rounds of 2D classification were used to clean these particle sets with different circular masks to account for different particle sizes. Featureless classes were removed from each step to obtain cleaned heterogeneous particle stacks for further processing. The 2D class averages containing clear 2D features were selected and used to generate a Topaz training model. The trained model was then used to pick particles using Topaz extract in cryoSPARC.

From heterogeneous particle sets, particles were classified, and final maps were solved with the use of BaR. In brief, ab initio methods were used to build initial 3D maps from the selected classified particles, and particles were then retrieved based on the maps. To build the initial maps, particles were separated using 2D classification paired with 3D ab initio and heterogeneous classifications. These initial classes were used to retrieve particles from the topaz-picked particle stack. 3D heterogeneous refinement using the ab initio classes, determined from the build phase of BaR, was applied to the cleaned heterogeneous particle sets. The new particle subsets were then cleaned using multiple rounds of 2D and 3D ab initio classifications. Non-uniform refinement using cryoSPARC was used to refine 3D reconstructions. A soft masks that covers the LptB2FG or LptB2FGC were used for local refinement. The resulting maps corresponding to the Lpt transporters exhibited high quality, allowing for the construction of models.

Model building and refinement
The models for LptB2FG-I, LptB2FG-II, LptB2FG-III, LptB2FGC-I, and LptB2FGC-II were constructed based on cryo-EM maps with resolutions of 3.34 Å, 3.26 Å, 3.31 Å, 3.26 Å, and 3.61 Å, respectively. Initial structures of LptB2FG, LptB2FGC were generated using Alphafold2 (using uniport accession numbers Q9HVV6, Q9HXH5, Q9HXH4 and Q9HVV8) and fitted into the density maps as starting models using Chimera. Subsequent model rebuilding was carried out using Coot, and structural refinements were performed using phenix.real_space_refine from the PHENIX suite. The final atomic model was evaluated using MolProbity. Detailed statistics regarding data collection, 3D reconstruction, and model refinement can be found in Supplementary Table S1.

Molecular Simulation setup
All simulations were run using GROMACS 2021. The LptBFG complexes were positioned in a membrane using the Martini 3 coarse-grain (CG) force field and solvated with water and 0.15 M NaCl to neutralise the system. The membranes were constructed using insane with a 7:2:1 ratio of PE:PG:CDL lipids, using the MemProtMD pipeline. An elastic network of 1000 kJ mol−1 nm−2 was applied between all backbone beads between 0.5 and 1 nm. Electrostatics were described using the reaction field method, with a cut-off of 1.1 nm using the potential shift modifier and the van der Waals interactions were shifted between 0.9-1.1 nm. The systems were first energy minimised by the steepest descent algorithm to 1000 kJ mol−1 nm−1 and then simulated for a total of 1 µs. The temperature and pressure were kept constant throughout the simulation at 310 K and 1 bar, respectively, with protein, lipids and water/ions coupled individually to a temperature bath by the V-rescale method and a semi-isotropic Parrinello-Rahman barostat. The final snapshots from the CG simulations were then converted back to an atomistic description using CG2AT2. LPS and CDL coordinates were positioned into the central cavity of LptBFG based on the coordinates from PDB ID 6S8H, using PyMOL. Converted all-atom simulations of the LptBFG complexes were performed without position restraints for a total of 500 ns, and run in triplicate. In all cases, a 2 fs timestep was used, in an NPT ensemble with V-rescale temperature coupling at 310 K and a semi-isotropic Parrinello-Rahman barostat at 1 bar, with protein, water/ions and, if included, lipids coupled individually. Electrostatics were described using PME, with a cut-off of 1.2 nm and the van der Waals interactions were shifted between 1-1.2 nm. The tip3p water model was used, the water bond angles and distances were constrained by SETTLE. H-bonds were constrained using the LINCS algorithm. Analysis was performed using MDAnalysis and visualised in PyMOL (Schrödinger, LLC. 2015).

Generation of plasmids and complementation assays
The DNA fragments for cloning were amplified by PCR using the genomic DNA of P. aeruginosa PAO1 or the plasmid pQLinkN-yh-LptB-KL-6×His-FGC as the template. All constructs generated in this work were verified by DNA sequencing.

The plasmid constructs to express LptB protein variants were generated by directionally cloning the coding sequence of each gene of interest or its recombinant variant into the shuttle vector pME6032, downstream of the IPTG-inducible Ptac promoter. The pME6032 derivatives were introduced into P. aeruginosa by transformation using chemically competent cells.

Complementation assays were performed by pre-culturing the strains of interest in Mueller-Hinton broth supplemented with 0.01% rhamnose until the late-exponential phase. Cells were collected by centrifugation and resuspended in saline at an OD600 = 1, and 5 µL aliquots of serial ten-fold dilutions in saline were plated onto Mueller-Hinton agar plates supplemented or not with 0.0.1% rhamnose or 0.5 mM IPTG. Images were taken after 16 h of incubation at 37°C.

Sequence alignment
Sequence alignments were performed using the NPS@ClustalW webservice and rendered using the ESPript 3.0 webservice.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "The authors have provided a satisfactory rebuttal to my prior criticisms. I'm grateful to see that my comments proved to be useful for the authors in providing a much more interesting revised manuscript. I should mention that the statement in lines 287-288 is redundant considering that the same point was made in lines 282-283. Finally, regarding my query in Reviewer 1 point 4, the authors reply states \"In theory, the phosphate-to-phosphate distance (~10 Å for cardiolipin versus ~6 Å for lipid IVA) can be distinguished in cryo-EM when local resolution is better than 3 Å and the lipid is well ordered.\" However, I ask the authors to consider that their numbers can't be correct as the phosphate groups in cardiolipin are closer together, while those of lipid A are farther apart. The reference I was citing is extended data Figure 9 in Clairfeuille T, et al. Structure of the essential inner membrane lipopolysaccharide-PbgA complex. Nature. (2020) 584: 479-483. This paper resolved some previous errors that had been made by structural biologists who failed to adequately distinguish between cardiolipin and lipid A. Since the present authors are making the same distinctions, they should be clear about what it is that they are comparing. The relevant link is:\nhttps://www.nature.com/articles/s41586-020-2597-x/figures/13", "reviewer_2": "The authors provided a very thorough response, and I have no further significant edits to suggest. I appreciate the detailed description of what manuscript edits were made in response to each point raised.", "reviewer_3": "The authors have improved the manuscript. I appreciate the work the authors put into revising the manuscript, especially by adding complementation studies, including data using native Pseudomonas LPS, and clarifying some important information in the text. Although the manuscript has been improved, there are still significant issues to address to justify several of the authors’ conclusions.\n\nMain issues:\n1) The revised manuscript now shows that the tagged LptB protein complements in Pseudomonas. Specifically, the authors tested if relevant LptB proteins encoded in plasmids support growth of an lptB conditional strain in the absence of inducer. While this is a significant improvement of the study, the function should be more thoroughly tested for the following reasons. Previous studies have shown that although E. coli LptB-His has ATPase activity (in vitro) and complements the deletion of the lptB gene, it is partially functional in cells. It is functional enough to maintain growth but leads to significant defects in LPS transport, as demonstrated by the fact that cells with LptB-His are permeable (sensitive) to antibiotics and lptB-His can have either positive or negative effects in combination with specific lpt mutations. Whether the same is true or not in Pseudomonas remains unknown, but Fig. S3 shows that even though the depletion strain expressing LptB-KL-6×His (used in this study) can grow without inducer, it grows significantly smaller colonies than a strain producing a His-less LptB-KL. They also show that a depletion strain producing LptB-6×His grows better than LptB-KL-6×His but colonies are still smaller than colonies producing LptB. These data suggest that LptB-KL-6×His protein used in this study is partially functional, likely with significant defects in LPS transport based on the growth data. To describe it as functional is not appropriate. The effect of LptB-KL-6×His on the Pseudomonas Lpt transporter function remains uncharacterized. Whether structures would be different with a fully functional LptB is unknown, but it is a concern since the ATPase controls movement of the ABC transporter.\n2) The authors now report that Pseudomonas LPS binds better to the Pseudomonas Lpt complex than the E. coli LPS (Fig S12). Unfortunately, they did not provide quantitative data. I could not find information about where the Pseudomonas LPS came from or how they purified it (unless I missed it, please add), but if the lack of quantitative data results from the fact that the Pseudomonas LPS is a mixture, I do not understand why they did not then purify it from any of the LPS mutants reported in the literature that would yield a more homogenous preparation. The lack of quantification is especially relevant to the issue about the biological significance of cardiolipin binding, which remains unclear mainly because there is no evidence that the authors’ findings are relevant in Pseudomonas cells. Would cardiolipin even have a significant effect if native LPS was used in the in vitro MS experiments?\n3) Lines 295-302: The authors report that LptB2FGC binds ReLPS with ~2-fold higher affinity than LptB2FG. They state that these results suggest “that LptC contributes to high-affinity substrate binding. The increase in affinity may result from either a decreased off-rate of LPS from the central cavity or increased binding to periplasmic domains of the transporter. Since previous structural studies have linked the presence of LptC’s TM helix with lower LPS occupancy in the central cavity28-31, this explanation with greater engagement of periplasmic domains, is more consistent with our data.” I am not sure what “greater engagement of periplasmic domains” means. I presume it refers to “increased binding to periplasmic domains of the transporter”. There is no data supporting that, in their experimental system, LPS is bound to the periplasmic domains of LptF, LptG, or even LptC. Was ATP added to cause LPS extraction and movement to the periplasmic domains as the authors imply? Can the authors rule out that the higher affinity of LptB2FGC for ReLPS is not an artifact of using E. coli LPS instead of the native Pseudomonas LPS? LPS from Pseudomonas has 5 fatty acyl chains, while E. coli LPS has 6. Since the LptC helix makes the cavity larger, it might increase binding to the E. coli LPS. Their conclusion would have to be supported with data using Pseudomonas LPS.\n4) Based on MD simulations, the authors highlight 17 residues in LptF and 21 in LptG as relevant in lipid binding. They do not test the relevance of any of these residues but state: “Some of these residues, including R30, L62 (LptF) and R132 (LptG), are functionally validated: previous studies have shown that their mutation impairs bacterial viability”. How many is “some”? 3 out of 38 residues? Were any of them tested in Pseudomonas? The authors could use the complementation system shown in Fig. S3 to test the importance of residues in Pseudomonas.\n\nMinor issues:\n- Lines 71-72: “an important role in regulating the ATPase activity to allow higher LPS extraction efficiency of the protein complex” is missing the beginning of the sentence.\n- Figure 2d: Is this structure from LptB2FG-I? It should be indicated. Where in the complex is the cross-section shown in Fig. 2d? Is it at the predicted membrane-periplasmic interface? If the cross-section is of LptB2FG-I, please add markings indicating the plane of the cross-section shown in Fig. 2d. Similar issue with Fig. S7a and b. Please mark point of cross-section in the structure of the complex on the left. Also modify the figure legend in Fig. 7 to indicate structures on the right represent a cross-section.\n- Lines 236-237: “resulting in an interface that resembles a heterodimer rather than a heterotrimer.” ??\n- Lines 191 and 258: “closed-up” should be “close-up\".\n- Lines 244-248: “A comparison of our cryo-EM structure of LptB2FGC-I with the previously reported V. cholerae LptB2FGC (6MJP), reveals a significant conformational change, with a RMSD of 5.57 Å across 845 Cα atoms, indicating a significant conformational difference between these two structures. These differences suggest species-specific adaptations and further underscore the structural plasticity of the LptB2FGC complex.” I do not understand why the authors suggest that the conformational change is the result of differences between species as opposed to stemming from the fact that the Vibrio structure has the TM helix of LptC in interface I, while the cryo-EM structure of LptB2FGC-I does not.\n-Lines 269-271: “This affinity difference suggests that the transporter has adapted to recognise native P. aeruginosa LPS more tightly, likely due to subtle differences in the lipid A or core oligosaccharide structures between species.” A supplemental figure showing the two LPS structures would be very helpful to readers and should be included.\n-Lines 353-354: Citations are needed for: “particularly under conditions where cardiolipin is enriched, such as during stress responses, antibiotic treatment, or stationary phase”.\n-Figure 6: Please add “F” of “G” subscript/superscript to TM labels on structures on right to indicate whether they belong to LptF or LptG, respectively.\n-Discussion: I am confused by this section: “It is interesting to note that, although LPS-bound, the recent A. baylyi LptB2FG structure (8FRM) adopts a closed conformation at Interface 2, which is similar to our substrate-free P. aeruginosa structure. The structural analysis suggests that A. baylyi LptB2FG adopts a conformation in which an LPS molecule has already entered the central cavity through interface 1, but the transmembrane helix of LptC (TMc) has not yet inserted. We therefore propose this configuration as between our apo “resting” conformation and the TMc-capped form like observed in V. cholerae LptB2FGC.” Are the authors implying that LPS enters the cavity first and then LptC helix associates with the transporter? Such a model is very different from the current model and would imply that the LptC helix associates with the cavity after LPS enters it and then would have to leave again for LPS extraction. What data supports this model?\nLines 446-447: “provides the molecular basis for LPS recognition and transport” should be “provides molecular basis for LPS recognition and transport”."}, "author_response": "We thank the reviewer for the positive evaluation. We agree that there is a redundancy and have now modified the text accordingly (lines 279-286, main text):\n“To dissect the specific features of LPS required for recognition by the LptB₂FG(C) complex, we performed native MS experiments using defined LPS analogues: Re-LPS, lipid A, and 4′-monophosphoryl lipid A (MPLA) (Figure 4a). These compounds differ progressively in their structural complexity: Re-LPS contains lipid A and two Kdo (3-deoxy-D-manno-oct-2-ulosonic acid) sugars; lipid A lacks the Kdo moieties; and MPLA is missing both Kdo and the phosphate group at position 1 of the glucosamine backbone. Only Re-LPS has been used previously to investigate Lpt system functionality 17,47,48. Because of the lack of commercial availability of these variants from P. aeruginosa, we used E. coli versions for our study.”\nFinally, regarding my query in Reviewer 1 point 4, the authors reply states \"In theory, the phosphate-to-phosphate distance (~10 Å for cardiolipin versus ~6 Å for lipid IVA) can be distinguished in cryo-EM when local resolution is better than 3 Å and the lipid is well ordered.\" However, I ask the authors to consider that their numbers can't be correct as the phosphate groups in cardiolipin are closer together, while those of lipid A are farther apart. The reference I was citing is Extended Data Figure 9 from Clairfeuille et al. (2020). We acknowledge the reviewer’s point that the phosphate-to-phosphate distance in cardiolipin is shorter than in lipid A or lipid IVA, and we appreciate this clarification.\nThat said, we emphasise that our study focuses on ligand-free cryo-EM structures of LptB FGC, and we do not observe density consistent with either cardiolipin or lipid A in the central cavity. The lipid-like densities we do detect are confined to the periphery of the transmembrane domains and are consistent in shape and size with phosphatidylethanolamine (PE), which is expected based on our nanodisc reconstitution protocol. As such, our manuscript does not attempt to discriminate between lipid A and cardiolipin based on density interpretation, and the concern raised does not impact our structural conclusions.\nWe thank the reviewer for the positive assessment and thoughtful feedback throughout the revision process. We’re pleased that our responses and manuscript edits address the points raised.\nWe thank the reviewer for raising this point. We agree that the small-colony phenotype indicates that C-terminally His-tagged LptB variants are partially functional in vivo and should not be described as fully functional. We have therefore tempered the wording throughout to explicitly state “partially functional.”\nIn the revised manuscript we now state (lines 160-166, main text):\n“Functional assays demonstrated that the His-tagged variants successfully complemented the lptB conditional mutant, even though the colonies were slightly smaller than those of the control strains expressing non-tagged LptB. This indicates that the tagged LptB variants are at least partially functional in vivo in P. aeruginosa (Figure S3). Together, these findings confirm that our purified LptB₂FG(C) complexes are enzymatically competent in vitro and that the tagged LptB variants support growth in complementation assays and we therefore used these preparations for the structural and biochemical analyses reported here.”\nRegarding the functionality of the Pseudomonas Lpt transporter and the influence of LptB-KL-6×His on its structure, we fully agree that we cannot exclude subtle structural differences relative to an untagged protein, nor that differences might emerge under transport-active, ATP-turnover conditions. Importantly, we are not inferring in-cell activity from our structures: all cryo-EM data were collected in nucleotide-free conditions and capture apo conformations, so ATP-driven transport states are not interpreted here; characterising potential tag-dependent differences under turnover will be an important goal for future studies.\nWe thank the reviewer for this constructive comment. As correctly noted, preparations of Pseudomonas LPS are inherently heterogeneous. Although several groups have isolated LPS from rough or mutant P. aeruginosa strains, these invariably consist of polydisperse mixtures of glycoforms differing in phosphorylation, acetylation, acylation state, and core composition (e.g., Knirel et al., Eur J Biochem. 2001, 268(17):4708-19; Kropinski et al., Can. J. Microbiol. 1979, 25(3):390-8; Choudhury et al., Carbohydr. Res. 2005, 340(18):2761-72; Mei et al., Microbiol. Spectrum 2023, 11(6):e0177323). To our knowledge, a truly monodisperse P. aeruginosa LPS preparation suitable for quantitative MS has not been achieved, and chemical synthesis of defined lipid A analogues is technically challenging and prohibitively costly.\nIn line with this, we were unable to obtain a homogeneous preparation and therefore proceeded with a commercially available P. aeruginosa LPS (Sigma Aldrich, catalogue no. L8643), which we now specify in the Methods (supplementary information, lines 85-86). Importantly, the degree of heterogeneity in this preparation is not substantially different from that reported for mutant-derived LPS.\nAs requested, we now include a quantitative comparison of P. aeruginosa LPS and E. coli LPS binding (Figure S12, see below and in supplementary information, lines 379-388.). We quantified the different LPS-bound species by integrating the area under the curve for each peak — a widely accepted approach, especially when dealing with spectra exhibiting broad features. Nonetheless, explicitly note that these values should be regarded as semi-quantitative because of LPS heterogeneity.\nUpdated Figure S12. Pa- LptB2FG binds its native LPS with higher affinity than E. coli LPS. Native mass spectra of Pa-LptB FG in presence of P. aeruginosa LPS (top) and E. coli LPS (bottom). 0.5 uL of LPS at 5 mg/ml was added to 10 uL of purified LptB FG at 3 µM concentration. Since LPS is an heterogenous mixture, a number of adduct peaks are observed in both cases. The bar chart on right shows the relative quantification, suggesting that LPS binding is much higher in the case of Pa-LPS compared to Ec-LPS. We explicitly caution that these values are semi-quantitative because of LPS heterogeneity.\nRegarding cardiolipin, we agree that its physiological significance must be interpreted in the context of native P. aeruginosa LPS. Native LPS likely competes more effectively than E. coli Re-LPS. The limitation in obtaining a homogeneous preparation currently prevents testing this directly. The broader peak distributions in the presence of P. aeruginosa LPS also make additional binding experiments technically challenging and difficult to quantify. Nevertheless, our native MS experiments clearly show that cardiolipin binds directly to the Pa-LptB₂FGC complex and displaces Re-LPS in a concentration-dependent manner (Fig. 5b, S15–S16). These data provide the first direct evidence that cardiolipin can engage the Pa-Lpt transporter and modulate LPS binding. Moreover, under conditions of cellular stress, stationary phase and biofilm formation, cardiolipin levels are known to become significantly enriched in the inner membrane (see ref. 53-58 in the main text) and it might effectively compete for transient occupancy of the transporter's binding site, even against a higher-affinity substrate like Pa-LPS. That noted, independent genetic/physiological data support a functional role for cardiolipin in LPS transport: cardiolipin promotes transport across the inner membrane and can mitigate the transport deficit of under-acylated lipid A (e.g., lpxM contexts), providing in vivo plausibility for cardiolipin–Lpt interactions we observe biochemically (ref 51).\nWe have revised the manuscript to explicitly note that our competition experiments were performed with a model LPS substrate (Re-LPS) and thus represent a proof-of-concept rather than a definitive demonstration of in vivo function. We further acknowledge that the precise contribution of cardiolipin in the presence of native Pa-LPS remains an open question for future studies, particularly under physiological conditions (e.g., stress, stationary phase, biofilm formation) where cardiolipin levels are known to be enriched. We now add (lines 439-446, main text):\n“Our lipid-binding studies further revealed that, beyond LPS, the transporter can accommodate cardiolipin (CDL), a lipid previously implicated in MsbA-mediated LPS transport51. The ability of CDL to bind and displace LPS (with a model LPS) in a concentration-dependent manner, combined with MD simulations showing overlapping contact residues, raises the possibility that non-LPS lipids such as CDL might transiently interact with or regulate the transporter under specific cellular conditions (e.g., stress, membrane remodeling)53-58. This proof-of-concept observation adds a layer of physiological relevance to the transporter's substrate selectivity and suggests that LptB₂FGC may function within a lipid-rich regulatory environment, which remains to be tested in future studies.”\nWe thank the reviewer for pointing out this ambiguity. We agree that our original wording was unclear. Our equilibrium measurements show higher fractional LPS occupancy for LptB FGC than LptB FG under nucleotide-free conditions. This is consistent with a kinetic retention effect in which LptC lowers the effective dissociation rate (k_off) and/or increases the effective association rate (k_on) by gating lipid entry. Because our data report equilibrium occupancies, we cannot distinguish between changes in k_on and k_off here; accordingly, we present the reduced-off-rate explanation as a working hypothesis aligned with prior structural models of LptC-mediated gate opening. While we do not infer periplasmic-domain engagement from our assays, we cannot completely exclude additional interactions under our conditions because the exact TM-helix position is unknown. We have revised the relevant sentence in the manuscript accordingly to make this explicit in lines 296-303 of main text:\n“The increase in affinity may result from either a decreased off-rate of LPS from the central cavity or increased binding to periplasmic domains of the transporter. Because our data are equilibrium occupancies, we cannot distinguish between changes in association and dissociation rates here; accordingly, we present the reduced-off-rate explanation as a working hypothesis aligned with prior structural models of LptC-mediated gate opening28-31. While we do not infer periplasmic-domain engagement from our assays, we cannot completely exclude additional interactions under our conditions because the exact TM-helix position is unknown.”\nThe reviewer also asked whether the ~2 fold effect could be an artefact of using E. coli Re-LPS (hexa-acyl lipid A) rather than native Pa-LPS (often penta-acylated under lab conditions, though hexa-/hepta-acyl forms also occur in clinical isolates). To probe acylation-state generality, we re-analysed pre-delipidation native-MS datasets from ClearColi-expressed complexes, where lipid IVA (tetra-acyl) is the sole LPS-related species. In those data, LptB₂FG was predominantly apo (~60%), whereas LptB₂FGC showed markedly increased lipid IVA occupancy (~80% with 2× adducts), indicating that the LptC-dependent enhancement of lipid occupancy extends to a tetra-acyl scaffold. While indirect, this argues against a hexa-acyl-specific artefact.\nWe clarified in the Results (lines 376-382) that “among the 38 cavity-contact residues highlighted by MD (17 LptF; 21 LptG), prior functional data exist for a subset of positions in enterobacteria (although not directly tested in P. aeruginosa), most notably hydrophobic/charged residues lining the cavity whose mutation compromises Lpt function or growth (e.g., LptF I25/R30/L62; LptG L26/R59/I70/R132/V308/F312). These studies collectively support the idea that the central cavity residues we observe contacting acyl chains in MD are functionally important, although numbering and exact identities vary by species/construct.”\nOur study reports apo-state cryo-EM structures with no LPS density and does not include an LPS-bound structure or a mutational analysis; a systematic functional validation is therefore beyond the scope of this work. The complementation assay in Fig. S3 was designed to validate LptB variants for purification and in vivo functionality. Establishing an analogous complementation system to the essential membrane subunits LptF and LptG would require substantial new strain engineering and several months of work. This is outside the scope of the present study and is reserved for future work.\nThis is now fixed.\nThank you for pointing that out. It is LptB2FG-I, and we have put this in the figure legends. We have added mark points in these figures. Main text, line 198 and supplementary information, lines 347-351.\nWe thank the reviewer for highlighting this unclear statement. We intended to describe how, in the V. cholerae structure, the LptG periplasmic β-jellyroll exhibits minimal interaction with LptC, in contrast to the more prominent LptC–LptF interface. We acknowledge that the original wording was confusing and have removed the sentence for clarity.\nThis is now fixed.\nWe thank the reviewer for this insightful comment. To clarify this point, we have revised the manuscript text to read (lines 245-247, main text):\n“A comparison of our cryo-EM structure of LptB₂FGC-I with the previously reported V. cholerae LptB₂FGC (6MJP) reveals a significant conformational difference (RMSD 5.57 Å across 845 Cα atoms), which may reflect distinct functional states.”\nThank you for this suggestion. We have now added a supplemental figure (Figure S13) showing two LPS structures. Lines 389-392, supplementary information.\nReferences have now been added.\nDone.\nWe thank the reviewer for this comment. The current working model suggests that the LptC transmembrane helix (TMc) associates with the cavity prior to LPS entry and later dissociate to allow LPS extraction. However, our apo structure of P. aeruginosa LptB₂FGC displays an open interface I without TMc insertion. This observation led us to consider an alternative model in which LPS might enter the cavity before TMc insertion. The recent A. baylyi LptB₂FG–LPS structure (PDB: 8FRM), although LPS-bound, shows a closed interface II. While these findings raise the possibility of a different sequence of events, we cannot rule out that TMc may still transiently insert into the interface to facilitate LPS capture and release. Additionally, we acknowledge that the order and coordination of TMc insertion and substrate binding may be species dependent. To avoid overinterpretation, we have revised the text to more cautiously state (in main text, lines 429-432):\n“It is interesting to note that, although LPS-bound, the recent A. baylyi LptB₂FG structure (PDB: 8FRM) adopts a closed conformation at interface II, similar to our substrate-free P. aeruginosa structure. Structural analysis suggests that this A. baylyi conformation captures a state in which LPS has entered the cavity through interface I.”\nThis is now fixed."}]

Decision letter