Structural, mechanistic, and physiological insights into phospholipase A-mediated membrane phospholipid degradation in Pseudomonas aeruginosa

  1. Florian Bleffert
  2. Joachim Granzin
  3. Muttalip Caliskan
  4. Stephan N Schott-Verdugo
  5. Meike Siebers
  6. Björn Thiele
  7. Laurence Rahme
  8. Sebastian Felgner
  9. Peter Dörmann
  10. Holger Gohlke  Is a corresponding author
  11. Renu Batra-Safferling  Is a corresponding author
  12. Karl-Erich Jaeger
  13. Filip Kovacic  Is a corresponding author
  1. Institute of Molecular Enzyme Technology, Heinrich Heine University Düsseldorf, Forschungszentrum Jülich GmbH, Germany
  2. Institute of Biological Information Processing - Structural Biochemistry (IBI-7: Structural Biochemistry), Forschungszentrum Jülich GmbH, Germany
  3. Institute for Pharmaceutical and Medicinal Chemistry, Heinrich Heine University Düsseldorf, Germany
  4. Centro de Bioinformática y Simulación Molecular (CBSM), Faculty of Engineering, University of Talca, Chile
  5. John von Neumann Institute for Computing (NIC), Jülich Supercomputing Centre (JSC), and Institute of Bio- and Geosciences (IBG-4: Bioinformatics), Forschungszentrum Jülich GmbH, Germany
  6. Institute of Molecular Physiology, and Biotechnology of Plants (IMBIO), University of Bonn, Germany
  7. Institute for Plant Genetics, Heinrich Heine University Düsseldorf, Germany
  8. Institute of Bio- and Geosciences, Plant Sciences (IBG-2), and Agrosphere (IBG-3), Forschungszentrum Jülich GmbH, Germany
  9. Department of Microbiology, and Immunobiology, Harvard Medical School, United States
  10. Department of Molecular Bacteriology, Helmholtz Centre for Infection Research, Germany
  11. Institute of Bio- and Geosciences (IBG-1: Biotechnology), Forschungszentrum Jülich GmbH, Germany

Abstract

Cells steadily adapt their membrane glycerophospholipid (GPL) composition to changing environmental and developmental conditions. While the regulation of membrane homeostasis via GPL synthesis in bacteria has been studied in detail, the mechanisms underlying the controlled degradation of endogenous GPLs remain unknown. Thus far, the function of intracellular phospholipases A (PLAs) in GPL remodeling (Lands cycle) in bacteria is not clearly established. Here, we identified the first cytoplasmic membrane-bound phospholipase A1 (PlaF) from Pseudomonas aeruginosa, which might be involved in the Lands cycle. PlaF is an important virulence factor, as the P. aeruginosa ΔplaF mutant showed strongly attenuated virulence in Galleria mellonella and macrophages. We present a 2.0-Å-resolution crystal structure of PlaF, the first structure that reveals homodimerization of a single-pass transmembrane (TM) full-length protein. PlaF dimerization, mediated solely through the intermolecular interactions of TM and juxtamembrane regions, inhibits its activity. The dimerization site and the catalytic sites are linked by an intricate ligand-mediated interaction network, which might explain the product (fatty acid) feedback inhibition observed with the purified PlaF protein. We used molecular dynamics simulations and configurational free energy computations to suggest a model of PlaF activation through a coupled monomerization and tilting of the monomer in the membrane, which constrains the active site cavity into contact with the GPL substrates. Thus, these data show the importance of the PlaF-mediated GPL remodeling pathway for virulence and could pave the way for the development of novel therapeutics targeting PlaF.

Editor's evaluation

This study provides new insights into how a bacterial phospholipase called PlaF degrades membrane phospholipids in a controlled fashion to allow bacteria to alter their membrane composition to adapt to changing conditions. Inas much as PlaF is important for virulence, it will be interesting to see if the comprehensive biochemical and structural analysis in the current paper will aid in the development of a class of antibiotics targeting PlaF.

https://doi.org/10.7554/eLife.72824.sa0

Introduction

Biological membranes are steadily changing and adapting to environmental and developmental conditions (Eickhoff and Bassler, 2018; Parsons and Rock, 2013). These changes affect processes indispensable for cell life, such as nutrient uptake (Higgins, 1992), chemical signaling (Venturi and Fuqua, 2013), protein secretion (Krampen et al., 2018), folding (Mackenzie, 2006), interaction with hosts (Baxter et al., 2015), and antibiotic resistance (García-Fernández et al., 2017). An important mechanism to maintain membrane functionality in bacteria is the alteration of lipid composition (Rowlett et al., 2017; Schniederjans et al., 2017; Zhang and Rock, 2008). The adjustment of the fatty acid (FA) composition of glycerophospholipids (GPLs) upon thermal adaptation represents one of the most important mechanisms of membrane lipid homeostasis (Sinensky, 1974; Cossins, 1994). Adaptive changes in membrane GPL composition were observed under numerous other conditions, including environmental stresses (Rowlett et al., 2017), the transition from planktonic to sessile lifestyle (Benamara et al., 2014), and heterologous protein production (Kanonenberg et al., 2019).

De novo synthesis of GPLs is the main pathway used to tune the proportions of different lipid classes in bacteria (Zhang and Rock, 2008; Jeucken et al., 2019). Furthermore, bacteria rapidly alter their membrane GPL composition by chemical modifications (cis-trans isomerization and cyclopropanation) of acyl chains in GPLs to respond to environmental changes (Zhang and Rock, 2008). However, the bacterial pathway for remodeling of GPLs involving a rapid turnover of the acyl chains of GPLs is unknown. Interestingly, such a pathway was discovered in eukaryotes by W. E. Lands more than 60 years ago (Lands, 1958). This Lands cycle involves PLA-catalyzed deacylation of membrane GPLs to mono-acyl GPLs (lysoGPLs) followed by lysophospholipid acyltransferase (LPLAT)-mediated acylation of lysoGPL to yield a new GPL molecule with acyl chain composition different from the starting GPL (Lands, 1958). Despite the importance of this metabolic process in different animal and plant tissues, it took nearly 50 years before the enzymes involved in phospholipid remodeling were discovered (Shindou and Shimizu, 2009). Fourteen different mammalian LPLAT with specificities for different GPL head groups were reported to be involved in the Lands cycle (Hishikawa et al., 2008; Valentine et al., 2022). The recently published structure of human LPLAT provided the first insights into the molecular mechanism by which lysoGPL is acylated to GPL (Zhang et al., 2021). At least 16 mammalian PLAs (cytosolic and calcium-independent families) that may act on the membrane GPLs with different substrate profiles and tissue expression patterns are known (Clark et al., 1990; Song et al., 1999; Underwood et al., 1998; Ohto et al., 2005). Some PLAs have a suggested role in the remodeling of membrane GPLs (Asai et al., 2003), while others are involved in producing lipid mediators and bioenergetics (Murakami et al., 2020). Detailed computational studies revealed that human iPLA2β is allosterically activated by binding to the membrane, which is required to extract a single GPL molecule from the membrane and subsequent hydrolysis (Mouchlis et al., 2015).

Whereas extensive studies have been carried out for secreted bacterial PLAs acting as host-cell effectors (Istivan and Coloe, 2006), only limited information is available for the enzymes from the intracellular PLA family (Flores-Díaz et al., 2016). Previously, we reported that periplasmic TesA from Pseudomonas aeruginosa is a multifunctional enzyme with lysoPLA activity (Kovačić et al., 2013). However, this enzyme has no PLA activity, and therefore it is most likely not related to membrane GPL remodeling (Leščić Ašler et al., 2010). We recently published a novel intracellular PLA from P. aeruginosa whose function for remodeling of GPLs still needs to be experimentally analyzed (Weiler et al., 2022). Comprehensive lipidomic profiling of 113 Escherichia coli strains with deleted or overexpressed lipid metabolism genes did not reveal the identity of an intracellular PLA suitable for the Lands cycle (Jeucken et al., 2019). Here, we describe PlaF from P. aeruginosa (Kovacic et al., 2016; Bleffert et al., 2019) as the first cytoplasmic membrane-bound PLA with a role in virulence and GPL remodeling pathway in bacteria. We determined the crystal structure of PlaF (Kovacic et al., 2016; Bleffert et al., 2019) as a basis to provide mechanistic insights into PLA-mediated membrane phospholipid degradation related to bacterial virulence.

Results

PlaF is an integral cytoplasmic membrane-bound enzyme

We previously purified PlaF from the Triton X-100 solubilized membranes of a P. aeruginosa strain carrying the p-plaF expression plasmid (Kovacic et al., 2016; Bleffert et al., 2019). Here, we show that catalytically active PlaF is an intrinsic integral membrane protein as it was absent in the soluble fraction of the P. aeruginosa p-plaF (Figure 1a) and remained membrane-associated after treatment of PlaF-containing membranes with denaturation agents (Na2CO3 or urea), which destabilize weak interactions between peripheral proteins and the membrane (Figure 1b). To identify if PlaF is associated with the inner or outer membrane, P. aeruginosa p-plaF membranes were fractioned by ultracentrifugation in a sucrose density gradient. Western blot analysis of the cytoplasmic membrane protein SecG (Bleves et al., 1996), and the outer membrane-associated Lipid A (Matsushita et al., 1978) combined with PlaF activity measurement revealed that the majority of PlaF was in the cytoplasmic membrane fractions (#9–13) (Figure 1c). As expected, the Lipid-A-containing fractions (#1–3) showed negligible PlaF activity (Figure 1c), overall demonstrating that PlaF is a cytoplasmic integral membrane protein. Proteolysis experiments in which P. aeruginosa p-plaF cells with a chemically permeabilized outer membrane were treated with trypsin revealed a time-dependent degradation of PlaF (Figure 1d). These results suggest that PlaF is likely anchored to the cytoplasmic membrane via a transmembrane (TM) domain at the N-terminus predicted from sequence analysis (Kovacic et al., 2016), and its catalytic C-terminal domain protrudes into the periplasm.

Subcellular localization of PlaF.

(a) PlaF is a membrane protein of Pseudomonas aeruginosa. The membrane (M) and soluble fractions (SFs) of cell extracts from P. aeruginosa p-plaF, and the empty vector control strain (EV) were separated, analyzed by immunodetection with anti-His6-tag antibodies, and by esterase activity assay. The membrane protein marker P. aeruginosa XcpQ was detected with anti-XcpQ antibodies. (b) PlaF is an integral membrane protein of P. aeruginosa. The crude membranes of P. aeruginosa p-plaF were treated with sodium carbonate, urea, Triton X-100, or MES buffer control followed by ultracentrifugation (S, supernatant; M, membrane proteins). PlaF was detected as in (a). (c) PlaF is a cytoplasmic-membrane protein of P. aeruginosa. The membrane fractions of P. aeruginosa p-plaF and the EV strains were isolated and separated by ultracentrifugation in a sucrose density gradient. The esterase activity was assayed as in (a). P. aeruginosa SecG, and outer membrane lipid A were used as markers for cytoplasmic, and outer membranes, and detected by Western blotting using anti-SecG, and anti-Lipid A antibodies, respectively. Inlet: A model of PlaF cellular localization. All values are mean ± standard deviation (S.D.) of three independent experiments measured in triplicates. (d) The catalytic domain of PlaF is exposed to the periplasm. P. aeruginosa p-plaF cells with permeabilized outer membrane were treated with trypsin for the indicated periods, and PlaF was detected as described in (a).

PlaF is a PLA1 involved in the alteration of membrane GPL composition as determined by global lipidomics

The previously reported carboxylesterase activity of PlaF (Bleffert et al., 2019) was here further analyzed using different PLA substrates. PlaF, purified with n-octyl-β-D-glucoside (OG) as described previously (Kovacic et al., 2016), showed PLA1 but no PLA2 activity toward the artificial substrates specific to each of these two phospholipase families (Figure 2a) and the natural phospholipid diacyl phosphatidylglycerol containing pentanoic and oleic acid at the sn-1 and sn-2 positions, respectively (Figure 2—figure supplement 1). The substrate profile of PlaF against natural di-acyl GPLs commonly occurring in P. aeruginosa membranes (Benamara et al., 2014) was determined with a spectrum of substrates (see legend to Figure 2b). In vitro, purified PlaF preferably hydrolyzed GPLs containing medium-chain FAs (C12 and C14) and showed comparable activities with phosphatidylethanolamine (PE), phosphatidylglycerol (PG), and phosphatidylcholine (PC) (Figure 2b).

Figure 2 with 2 supplements see all
Phospholipolytic activity profiling of PlaF.

(a) PlaF is a phospholipase A1. Enzyme activities of PlaF were measured fluorimetrically using artificial PLA1, and PLA2 substrates containing either ethanolamine (PE) or choline (PC) head groups. The control enzymes were PLA1 of Thermomyces lanuginosus, and PLA2 of Naja mocambique. Results are means±S.D. of three independent measurements performed with at least three samples. (b) PlaF releases FAs from naturally occurring bacterial GPLs. PLA activity of PlaF was measured by quantification of released FAs after incubation of PE, PG, and PC substrates containing FAs with different chain lengths (C12–C18). (c) PlaF changes GPL composition of Pseudomonas aeruginosa cells. Crude lipids extracted from P. aeruginosa wild-type (WT), ΔplaF, and ΔplaF::plaF membranes were quantified by Q-TOF-MS/MS using an internal standard mixture of GPLs. PlaF substrates are elevated in ΔplaF and depleted in ΔplaF::plaF, while modified GPLs show inverse response than GPL substrates. The GPL amount (nmol) was normalized to mg of crude lipids, and optical density (Supplementary file 3). FA composition of GPL is depicted as XX:Y, where XX defines the number of carbon atoms, and Y defines the number of double bonds in FAs bound to GPL. Results are mean ± S.D. of four biological replicates of WT, ΔplaF, and three of the ΔplaF::plaF. T-test of normally distributed values, ** p<0.01, * p<0.05. FA, fatty acid; GPL, glycerophospholipid.

To examine the role of membrane-bound PlaF in the regulation of the membrane GPL composition in vivo, we constructed the P. aeruginosa deletion mutant ΔplaF lacking the entire plaF gene by homologous recombination, and a complemented ΔplaF::plaF strain as a control (Figure 2—figure supplement 2). The activity assay showed ~90% loss of PLA1 activity in the mutant strain, and restoration of activity in ΔplaF::plaF slightly above the wild-type (WT) level (Figure 2—figure supplement 2). These findings indicate that PlaF is a major but not the only intracellular PLA1 in P. aeruginosa.

The quantitative mass spectrometric (Q-TOF-MS/MS) analysis of total GPLs isolated from four biological replicates of P. aeruginosa WT, ΔplaF, and ΔplaF::plaF cells revealed significant differences in membrane GPL composition (Figure 2c, Supplementary files 1-3). Statistical analysis of 323 GPL molecular species identified six significantly (p<0.05) accumulating GPLs, varying in the composition of head groups (PE, PG, PC, and phosphatidylinositol, PI), length, and unsaturation of acyl chains, in P. aeruginosa ΔplaF. Interestingly, these GPLs were present at low concentrations in the cells which may explain why they were not detected in the previous lipidomic analyses of P. aeruginosa GPLs (Benamara et al., 2014; Le Sénéchal et al., 2019). In the complemented strain (ΔplaF::plaF), these GPLs were depleted compared to the ΔplaF, although not to the WT level (Supplementary file 2). These results strongly indicate that PlaF specifically hydrolyses low abundant GPLs in vivo. We furthermore observed that the other seven PE, PG, and PC species, which belong among the most abundant P. aeruginosa GPLs (Benamara et al., 2014; Le Sénéchal et al., 2019), were significantly depleted (Figure 2c) in P. aeruginosa ΔplaF, and their concentrations were significantly elevated in complementation strain (Figure 2c). This may explain why the net GPL contents of the WT and the ΔplaF strain were not significantly (p=0.67) different. Significantly affected GPLs in the ΔplaF strain account for ~11% (mol/mol) of the total GPL content, indicating the profound function of PlaF in membrane GPL remodeling.

Our quantitative lipidomics results, which revealed that several PE, PG, and PC molecular species accumulated or were depleted in ΔplaF, together with in vitro PLA activity data of PlaF with various PE, PG, and PC substrates, indicate that PlaF might be a major PLA involved in the Lands cycle (Figure 2d). Thus, the six low-abundant PE, PG, and PC species that accumulated in ΔplaF might be PlaF substrates. PlaF-mediated hydrolysis of these substrates yields lysoGPL intermediates. Acylation of these lysoGPLs by an unknown acyltransferase will produce modified GPLs typical to P. aeruginosa. The absence of lysoGPL intermediates in ΔplaF will lead to the depletion of modified GPLs (Figure 2d).

PlaF is a potent virulence factor of P. aeruginosa affecting in vivo toxicology

We next addressed the question of whether PlaF contributes to the virulence of P. aeruginosa by using the G. mellonella infection model and the bone marrow-derived macrophages (BMDMs) viability assay. The results revealed a remarkable difference in the survival of G. mellonella larvae infected with P. aeruginosa WT or ΔplaF. While ΔplaF was avirulent during 20 hr of infection, the majority of the larvae (~80 %) did not survive 20 hr after infection with the P. aeruginosa WT (Figure 3a). The viability assays with P. aeruginosa-infected BMDMs showed a significantly (p<0.01) stronger killing effect of P. aeruginosa WT compared to ΔplaF 6 hr after infection (Figure 3b). As expected, the complemented strain (ΔplaF::plaF) restored the loss of virulence of ΔplaF in G. mellonella, and BMDM assays (Figure 3a and b). Comparison of the growth curves of P. aeruginosa ΔplaF, and the WT in nutrient-rich medium (Figure 3—figure supplement 1) showed that PlaF most likely does not reduce virulence by affecting the growth of P. aeruginosa.

Figure 3 with 2 supplements see all
PlaF is a novel virulence factor of Pseudomonas aeruginosa PAO1.

(a) Left: P. aeruginosa ΔplaF strain is less virulent than the respective wild-type (WT) strain in a Galleria mellonella larvae virulence assay. Kaplan-Meier plot of representative data of at least two experiments with 10 larvae per group. PBS treated and untreated larvae served as infection and viability controls, respectively. Right: Statistical analysis of the survival at the 20 hr using three independent experiments with 10 larvae each. (b) P. aeruginosa ΔplaF strain is less cytotoxic to bone marrow-derived macrophages (BMDMs) than the WT strain in cell culture. The BMDM cells (5×105) were infected with 5×105 bacteria in a 24-well plate, and lactate dehydrogenase activity in supernatants was determined as a measure of BMDM death. The ΔplaF phenotype could be complemented with P. aeruginosa ΔplaF::plaF. PBS or Triton-X100 (1% v/v) treated cells served as viability or 100% killing controls, respectively. Results are the representative data of two independent experiments (n=10). One-way ANOVA analysis, *** p<0.001, ** p<0.01, ns, not significant; PBS, phosphate-buffered saline.

A BLAST search revealed PlaF orthologs in more than 90% of all sequenced P. aeruginosa genomes, including 571 clinical isolates (Supplementary file 4). Furthermore, we found PlaF homologs in pathogens from the Pseudomonas genus (P. alcaligenes, P. mendocina, and P. otitidis), and other high-priority pathogens (Acinetobacter baumannii, Klebsiella pneumoniae, and Streptococcus pneumoniae) (Figure 3—figure supplement 2). These results indicate that PlaF is a novel and potent P. aeruginosa virulence factor, which is conserved in important pathogens and, therefore, might be a promising target for developing novel broad-range antibiotics.

Crystal structure of PlaF homodimer in the complex with natural ligands

To gain insights into the PlaF structure, we crystallized the OG-solubilized PlaF protein purified from P. aeruginosa membranes as described previously (Bleffert et al., 2019). The structure was refined at a resolution of up to 2.0 Å (Table 1). The final model in the asymmetric unit consists of two protein molecules (PlaFA and PlaFB), which are related by improper twofold non-crystallographic symmetry (Figure 4a). Active site cavities of both the monomers reveal non-covalently bound ligands—myristic acid (MYR), OG, and isopropyl alcohol (IPA) in PlaFA; and undecyclic acid (11A), OG, and IPA in PlaFB (Figure 4a, Supplementary file 5). These FAs are the natural ligands from the homologous organism P. aeruginosa that were co-purified with PlaF, as confirmed by gas chromatography-mass spectrometric (GC-MS) analysis of organic solvent extracts of purified PlaF (Figure 4—figure supplement 1). Compared to the protein chains, the bound FAs have higher average B-factor values for 11A (89.0 Å2) and MYR (66.6 Å2), indicating different flexibility of the ligands bound to the active sites of the two PlaF molecules.

Table 1
Data collection and refinement statistics on PlaF.
X-ray data
Beamline/detectorID29, ESRF (Grenoble, France)/DECTRIS PILATUS 6M
Wavelength (Å)/monochromatorλ=0.96863/channel-cut silicon monochromator, Si (111)
Resolution range (Å)47.33–2.0 (2.05–2.0)*
Space groupI 41 2 2
Unit cell (a=b), c (Å); α=β=γa=133.87 c=212.36; 90°
Total reflections669,964 (47,385)
Unique reflections65,113 (4527)
Multiplicity10.3 (10.5)
Completeness (%)100.0 (100.0)
Mean I/sigma (I)24.6 (2.5)
Wilson B-factor (Å2)38.3
R-merge (%)5.3 (91.3)
R-meas (%)5.6 (100.6)
Refinement
R-work (%)16.3 (23.15) (2.071–2.0)*
R-free (%)18.57 (27.81)
Number of atoms5187
Macromolecules4831
Ligands123
Water233
Protein residues620
RMS (bonds)0.008
RMS (angles)1.07
Ramachandran favored (%)99
Ramachandran outliers (%)0
Clashscore3.14
Average B-factor (Å2)49.1
Macromolecules (Å2)48.8
Ligands (Å2)79.2
Solvent (Å2)47.9
  1. *

    Values in parentheses are for the highest resolution shell.

Figure 4 with 5 supplements see all
Overall structure of dimeric PlaF with bound endogenous FA ligands.

(a) A unique N-terminal helix comprising a putative transmembrane helix (αTM1, L5–L27, gray) flanked by charged residues (K2, R3) on one side and, on another side, the juxtamembrane helix (αJM1, A28–L37, yellow). αJM1 links the αTM1 with the catalytic domain, which consists of an α/β-hydrolase (blue, α-helices; green, β-strands, and gray, loops), and a lid-like domain (brown). Ligands bound in the active site cleft are shown as ball-and-sticks (oxygen, red; carbon of OG, MYR, and IPA, green, orange, and blue, respectively). Thick gray lines roughly depict the membrane borders. (b) Dimer interface. Interactions involving TM-JM helices are predominantly hydrophobic with four weak H-bonds (indicated by a red asterisk) detected mostly in the αJM1. R83 is the only residue outside of the JM-TM helix involved in interactions. Residues of the PlaFB molecule are indicated in italics. A detailed list of interactions is provided in Supplementary file 6. (c) A model suggesting the orientation of PlaF in the membrane. The water molecules are indicated as green spheres. The transparent surface of PlaF was colored as in (a). PlaF is rotated by 180° along the normal to the membrane compared with Figure 4. (d) Interaction network within the ligand-binding cleft of PlaFA. MYR is linked via H-bond with the catalytic S137, and via hydrophobic interactions with OG. The sugar moiety of OG from PlaFA forms H-bonds with V33 of PlaFA, which is interacting with V33 and G36 of PlaFB. The part of the cleft in the direction of the opening 3 is occupied by several water molecules (W, yellow spheres). The cleft accommodates one IPA molecule bound to the water. Arrows indicate two openings not visible in this orientation. The cleft was calculated using the Pymol software and colored by elements: carbon, gray; oxygen, red; nitrogen, blue. FA, fatty acid.

The N-terminal 38 amino acids form a long, kinked helix that comprises the putative TM (αTM1) and the JM (αJM1) helices, connecting the catalytic domain with the membrane (Figure 5a). The kink angle in the TM-JM helices is the main difference between the two monomers (Figure 4—figure supplement 2) and is likely caused by crystal packing effects (Figure 4—figure supplement 2). Dimerization is mediated primarily via hydrophobic interactions between symmetry-nonrelated residues from the TM-JM domains of two monomers (Figure 4b, Supplementary file 6), consistent with the hydrophobic effects that dominate in the stabilization of dimeric TM domains (MacKenzie et al., 1997). In addition, four weak H-bonds (Figure 4b) between JM residues stabilize the PlaF dimer. The TM-JM helices adopt a coiled-coil-like conformation (Figure 4—figure supplement 2), where the αTM1 crosses its counterpart at V14 to form an elongated X-shaped dimer interface with the buried surface area of 656 Å2 per monomer. The full-length PlaF dimer represents a unique structure, as neither a relevant match to the TM-JM helix (Figure 4—figure supplement 3) nor the membrane-spanning coiled-coil structure of the TM-JM dimer has been reported previously.

Figure 5 with 2 supplements see all
PlaF oligomeric states and their enzymatic activity.

(a) PlaF forms dimers in cell membranes.In vivo cross-linking experiments were performed by incubating Pseudomonas aeruginosa p-plaF or the empty vector control (EV) cells with different concentrations of DMP cross-linker followed by immunodetection of PlaF with anti-PlaF antiserum. (b) In vitro cross-linking of purified PlaF. Purified PlaF was incubated with DMP, BS2G, and BS3 cross-linking reagents or buffer control (ø) for 90 min, and the samples were analyzed by SDS-PAGE. Molecular weights of protein standard in kDa are indicated. (c) PlaF homodimerization, and activity are concentration-dependent. Protein-protein interactions of purified PlaF were monitored by measuring the changes in thermophoresis (ΔFnorm, gray circles) using the MST method. The MST results are mean ± S.D. of two independent experiments with PlaF purified with OG. Esterase activity (black squares) of PlaF was measured in three independent experiments using 4-methylumbelliferyl palmitate substrate. Dissociation (KD) and activation (Kact) constants were calculated using a logistic fit of sigmoidal curves.

The crystal structure of PlaF is indicative of a specific orientation in the membrane

The catalytic domain of PlaF adopts a canonical α/β-hydrolase fold (Ollis et al., 1992; Figure 4a) with three α-helices forming a distinct lid-like domain that covers the active site (Figure 4a). Despite the high homology of the catalytic domain, the lid-like domain varies significantly between PlaF homologs (Figure 4—figure supplement 4), as observed previously for other lipolytic enzymes (Figure 4—figure supplement 4; Chow et al., 2012). Furthermore, the lid-like domain shows a less ordered structure, as suggested by comparatively higher B-factors (Figure 4—figure supplement 5). This is likely a consequence of the lack of stabilizing interactions between the charged residue-rich (23 of the 77 residues) lid-like domain and the hydrophilic head groups of membrane GPLs in the native membrane environment. The TLS (translation-libration-screw-rotation) model revealed higher disorder in the TM-JM domains, presumably also due to the missing interactions with the hydrophobic membrane (Figure 4—figure supplement 5). No ordered water molecules in the vicinity of αTM1 (Figure 4c) and the presence of several charged and polar residues adjacent to αTM1 suggest a model where the TM-JM domain spans through the membrane with the catalytic domain localized on the membrane surface (Figure 4c).

Ligand-mediated interaction network connects dimerization and active sites

The active site of PlaF comprises the typical serine-hydrolase catalytic triad with S137, D258, and H286 interacting through H-bonds (Jaeger et al., 1994; Supplementary file 7). Interestingly, S137 shows two side-chain conformations, where one conformer is within the hydrogen bond distance of the FA ligand (Figure 4d, Supplementary files 5 and 7). Additionally, S137 forms H-bonds with residues I160, D161, and A163 located in the lid-like domain. The active site cleft in PlaF is formed by residues from the helix αJM1, the α/β-hydrolase and the lid-like domains (Figure 4d, Supplementary file 8). In PlaF, the large T-shaped active site cleft formed by residues from the JM helix, the α/β-hydrolase, and the lid-like domains is amphiphilic, making it compatible with binding the bulky GPL substrates. Three openings are observed in the cleft—one, close to the catalytic S137, lined with residues from the loops preceding αE, and αF; second, in the middle pointing toward the putative membrane, lined mostly with polar residues of the loops preceding αB, and αF; and third, at the dimer interface, comprising residues from αJM1, and the loop preceding αF of the lid-like domain. The third opening accommodates a pseudo-ligand OG (Figure 4d), which with its pyranose ring interacts with residue V33 of PlaFA, which in turn participates in dimerization via interactions with V33 and T32 of PlaFB (Figure 4b). The alkyl chains of OG and MYR bound in the active site cleft are stabilized via hydrophobic interactions (Figure 4d). Finally, the H-bond interaction of catalytic S137 with the carboxyl group of MYR completes an intricate ligand-mediated interaction network bridging the catalytic (S137) and dimerization (V33) sites in PlaF (Figure 4d). The crystal structure presented thus suggests a role of dimerization and ligand binding in regulating PlaF function, which was subsequently analyzed biochemically.

The PlaF activity is affected by dimerization

To investigate the oligomeric state of PlaF in vivo, we performed cross-linking experiments in which intact P. aeruginosa p-plaF cells were incubated with the cell-permeable bi-functional cross-linking reagent dimethyl pimelimidate (DMP). Western blot results revealed the presence of monomeric and dimeric PlaF in DMP-treated cells, whereas dimers were absent in untreated cells (Figure 5a and Figure 5—figure supplement 1). Size exclusion chromatography showed that PlaF was extracted from the membranes with detergent and purified by IMAC elutes as a monomer (Figure 5—figure supplement 2). Incubation of this purified PlaF for 90 min with bi-functional cross-linkers of different lengths (DMP; bis(sulfosuccinimidyl) glutarate, BS2G or bis(sulfosuccinimidyl) suberate, BS3) resulted in the formation of a substantial amount of PlaF dimers, suggesting spontaneous dimerization in the solution (Figure 5b). Microscale thermophoresis (MST) measurements were performed in which the fluorescence-labeled PlaF was titrated with an equimolar concentration of non-labeled PlaF to quantify spontaneous dimerization. The results revealed a sigmoidal binding curve from which a dissociation constant KD=637.9±109.4 nM was calculated, indicating weak binding (Figure 5c). Measurements of the esterase activity of PlaF samples used for MST experiments revealed that the specific activity of PlaF strongly decreased with increasing PlaF concentrations (Figure 5c). Enzyme activity measurements were employed to calculate the activation constant Kact=916.9±72.4 nM. The similar dissociation and activation constants support a model in which PlaF activity is regulated through reversible dimerization in vitro.

FAs induce dimerization and inhibit PlaF

To investigate the effect of FA ligands on the activity of PlaF, we used mM concentrations of FAs with different chain lengths (C5–C15) in a competitive inhibition assay. PlaF was strongly inhibited (>80%) with FAs containing 10–14 carbon atoms (Figure 6a), while the shorter and longer FAs showed only moderate to weak inhibition (Figure 6a). To explore the underlining mechanism, we performed kinetic inhibition studies with increasing concentrations of decanoic acid (C10). The results showed that C10 FA lowered maximal hydrolysis rates (vmax) as expected for a competitive inhibitor. Yet, elevated binding constants (Km) in the presence of higher concentrations of C10 FA indicate that PlaF undergoes allosteric changes affecting the binding of FAs (Figure 6b, Supplementary file 9). We examined whether inhibitory FAs affect dimerization by cross-linking of PlaF in the presence of C10, C11, and C12 FAs. The results of SDS-PAGE revealed a significantly higher amount of dimeric PlaF in FA-treated than in untreated samples (Figure 6c). These results suggest a potential regulatory role of FAs on PlaF activity via FA-induced dimerization, which agrees with the previously demonstrated lower activity of the PlaF dimer compared to the monomer (Figure 5).

FAs exert an inhibitory effect on PlaF and trigger dimerization.

(a) Inhibition of PlaF with FAs. Esterase activity of PlaF was measured in the presence of 7.5 mM FA (C5–C15); an untreated PlaF sample was set as 100%. The results are mean ± S.D. of three experiments with three samples each. (b) Kinetic studies with FA C10 show evidence of mixed-inhibition. Double-reciprocal plots of initial reaction velocities measured with the p-NPB substrate and FA C10 inhibitor at concentrations in a range of 0–7.5 mM. (c) The effect of FAs on PlaF dimerization. PlaF samples incubated with FAs (C10–C12), dimethyl sulfoxide (DM, DMSO used to dissolve FAs), and purification buffer (B, dilution control) were cross-linked with dimethyl pimelimidate (DMP). FA, fatty acid.

The tilt of monomeric PlaF in a lipid bilayer permits direct GPL access to the active site

To better understand the molecular mechanism of PlaF activation through monomerization, we performed a set of 10 independent, unbiased 2 μs long MD simulations starting from dimeric or monomeric PlaF embedded in an explicit membrane with a GPL composition similar to the native P. aeruginosa membrane (Figure 7a). The simulations revealed only minor intramolecular structural changes in monomeric and dimeric PlaF compared to the initial structure (RMSDall atom <4.0 Å) (Figure 7—figure supplement 1, Supplementary file 10). Spontaneous monomerization was not observed during the MD simulations (Figure 7—figure supplement 1), in line with the sub-nanomolar dissociation constant and the simulation timescale. However, in 8 and 6 out of 10 simulations started, respectively, from PlaFA or PlaFB, a tilting of the monomer for ~25° toward the membrane was observed (Figure 7b, left and Figure 7—figure supplement 1). This tilting motion cooperatively with rotation of PlaF (Video 1) results in the active site cleft of the catalytic domain being oriented perpendicularly to the membrane surface, such that GPL substrates can have direct access to the active site through the opening at the dimer interface (Figure 7a, right). In dimeric PlaF, this opening is, according to the model suggested from the X-ray structure, at >5 Å above the membrane surface (Figure 7a) so that the diffusion of a GPL from the membrane bilayer to the cleft entrance in this configuration is thermodynamically unfavorable. In all MD simulations started from the tilted PlaF monomer, the protein remains tilted (Figure 7b, right and Figure 7—figure supplement 1), which corroborates the notion that the tilted orientation is preferred over the respective configuration in di-PlaF.

Figure 7 with 2 supplements see all
Molecular dynamics (MD) simulations and PMF computations of PlaF in the lipid bilayer.

(a) Structures used for MD simulations. di-PlaF: Crystal structure oriented in the membrane by the PPM method. PlaFA: Chain A from PlaF dimer oriented as in the dimer. The entrance of the active site cleft is more than 5 Å above the membrane bilayer surface. t-PlaFA: Extracted monomer A oriented using the PPM method. Cocrystallized MYR, 11A, and OG (depicted in pink), although not included in the simulations, are shown in the figures to highlight the orientation of the active site cleft. Arrows between the structures reflect the predicted states of equilibria under physiological conditions in Pseudomonas aeruginosa. Percentages of the different states are obtained from the molecular simulations (see main text and (e)). (b) MD simulations of monomeric PlaF. Time course of the orientation of monomeric PlaF with respect to the membrane starting from the PlaFA configuration as observed in the structure (left). In 80% of the trajectories, the monomer ends in a tilted configuration (marked with *). When starting from t-PlaFA (right), in all cases, the structure remains tilted. This shows a significant tendency of the monomer to tilt (McNemar’s Χ2=6.125, p=0.013). (c) Potential of mean force (PMF) of monomer tilting. The distance between the COM of Cα atoms of residues 33–37 (yellow, and gray spheres) and the COM of the C18 of the oleic acid moieties of all lipids in the membrane (continuous horizontal line in the membrane slab) was used as a reaction coordinate. The shaded area shows the standard error of the mean obtained by dividing the data into four independent parts of 50 ns each. The yellow shaded regions are the integration limits used to calculate Ktilting (Equation 5). The spheres in the PMF relate to monomer configurations shown in the inset. (d) PMF of dimer separation. The distance between the COM of Cα atoms of residues 25–38 of each chain was used as the reaction coordinate. The shaded area shows the standard error of the mean obtained by dividing the data into four independent parts of 50 ns each. Insets show representative structures at intermediate reaction coordinate values. (e) Percentage of PlaF monomer as a function of total PlaF concentration in the membrane according to the equilibria shown in (c) and (d). The monomer percentage was computed according to Equations 7–11 (see Materials and methods and SI for details). The red line shows the experimentally determined PlaF concentration under overexpressing conditions in P. aeruginosa p-plaF, while the blue-dashed region shows an estimated span for the PlaF concentration in P. aeruginosa wild-type (see Materials and methods for details). Calculated percentages are shown in (a).

Video 1
MD simulation of monomeric PlaFA in GPL bilayer.

Blue and red spheres indicate head groups of GPLs in two leaflets of the bilayer.

To further explore the transition of the monomeric PlaFA to its tilted orientation (t-PlaFA), we calculated the free energy profile or potential of mean force (PMF) for the tilting process by using umbrella sampling and post-processing the distributions with the WHAM method (Suzuki, 1975; Grossfield, 2016). As reaction coordinate, the distance (d) of the top of the JM domain (residues 33–37) to the membrane center was chosen. Distances of ~37 and ~17 Å were calculated for non-tilted PlaFA using the crystal structure and t-PlaFA using the structure obtained from the unbiased MD simulations where tilting spontaneously occurred, respectively. The converged and precise (Figure 7—figure supplement 1; SEM<0.4 kcal mol–1) PMF revealed two minima at d=19.6 and 30.6 Å, with t-PlaFA favored over PlaFA by 0.66 kcal mol–1 (Figure 7c). The free energy barrier of ~1.2 kcal mol–1 explains the rapid transition from PlaFA to t-PlaFA observed in the unbiased MD simulations. The equilibrium constant and free energy of PlaF tilting are Ktilting=3.35 and a ΔGtilting=–0.8±0.2 kcal mol–1. These results suggest a model in which PlaF is activated after monomerization by tilting with respect to the membrane surface, which allows substrate access to its catalytic site.

Estimating the ratio of monomeric and dimeric PlaF in the cell

To investigate if dimerization-mediated PlaF inhibition is dependent on PlaF concentration in the GPL bilayer, we calculated the free energy profile of dimerization, similarly as for the tilting process. For this, the distance (r) between Cα atoms of the JM region of the two chains was used as a reaction coordinate. The converged (Figure 7—figure supplement 1) and precise (SEM<1.4 kcal mol–1) PMF revealed that di-PlaF is strongly favored at r=9.5 Å (–11.4 kcal mol–1) over the monomer (Figure 7d), fitting with the distance of 9.9 Å observed in the crystal structure of PlaF. From the PMF, the equilibrium constants (Ka=1.57×107 Å2; KX=2.58×105) and free energy (ΔG=–7.5±0.7 kcal mol–1) of PlaF dimerization were computed (Equations 1–3), taking into consideration that KX and ΔG relate to a state of one PlaF dimer in a membrane of 764 lipids, according to our simulation setup. Experimentally, a concentration of one PlaF dimer per ~3786 lipids in P. aeruginosa plaF-overexpressing cells (Bleffert et al., 2019) was determined. However, the concentration in P. aeruginosa WT is likely 100- to 1000-fold lower, as we could not detect PlaF by Western blot (Figure 7—figure supplement 2). Under such physiological conditions and considering that the equilibria for dimer-to-monomer transition and titling are coupled (Figure 7a), between 74% and 96% of the PlaF molecules are predicted to be in a monomeric, tilted, catalytically active state in P. aeruginosa (Figure 7e). Our quantitative real-time-PCR results revealed that plaF is constitutively expressed in P. aeruginosa WT at a much lower level than sigma factors rpoD and rpoS (Savli et al., 2003; Figure 7—figure supplement 2). This agrees with previous global proteomic and transcriptomic results (Erdmann et al., 2018). As a catalytically active form of PlaF is favored in the WT, PlaF is likely involved in the constant remodeling of membrane GPLs.

Discussion

PlaF catalyzed remodeling of membrane GPLs

Employing lipidomic profiling of P. aeruginosa WT and the plaF gene deletion mutant, we found substantial changes in membrane GPL composition consistent with in vitro PLA1 activity of PlaF and its integral cytoplasmic membrane-localization. The present understanding of bacterial PLAs is limited to extracellular (ExoU, YplA, and SlaA; Istivan and Coloe, 2006; Sawa et al., 2016) and outer membrane (PlaB and OMPLA; Snijder et al., 1999; Schunder et al., 2010) enzymes with a proposed role in host-pathogen interactions, but, so far, bacterial PLA proteins tethered to the cytoplasmic membrane were not described (Jeucken et al., 2019).

Although bacterial enzymes catalyzing de novo GPL synthesis, their physiological functions and biochemical mechanisms are becoming increasingly well understood (Jeucken et al., 2019), information about GPL turnover enzymes remains largely obscure. Several of our findings indicate that PlaF plays a hitherto unexplored role in the membrane remodeling (Figure 8) that becomes especially apparent during virulence adaptation.

A model of PlaF-mediated membrane GPL remodeling.

PlaF is anchored with the TM helix to the inner membrane of Pseudomonas aeruginosa (Figures 1c and 4c), where it forms an inactive dimer (Figure 5c). Monomerization (Figure 5c) and subsequent spontaneous tilting (Figure 7) lead to activation. Binding of dodecanoic acid (C12) to monomeric PlaF triggers dimerization (Figure 6c) and inhibits enzymatic activity (Figure 6a). Tilting constrains the active site cavity of PlaF to the membrane surface such that GPL substrates can enter (GPL1, Figure 2), which are hydrolyzed to FA and lysoGPL1. A yet unknown acyl transferase possibly acylates lyso-GPL1 to yield modified GPL2 (Figure 2). GPL, glycerophospholipid.

  1. Deletion of plaF gene in P. aeruginosa leads to accumulation of several low abundant PE, PG, and PC molecular species (Figure 2c). PE, PG, and PC with different acyl chain lengths (C12–C18) were hydrolyzed by PlaF in vitro (Figure 2b). A low in vitro PLA1 activity of PlaF (µU/mg) is expected for an enzyme that could irreversibly damage the membrane.

  2. The P. aeruginosa ΔplaF strain revealed several depleted GPLs (Figure 2c), which may be explained assuming that lysoGPLs generated by PlaF activity are missing in this strain for further acylation to yield modified GPLs.

  3. FAs with 10–14 carbon atoms inhibit PlaF activity in vitro (Figure 6a). As PlaF can produce such FAs in vivo (Figure 2c), it is reasonable to assume that their cellular function is related to the regulation of PlaF activity by product feedback inhibition. This phenomenon is well known for lipolytic (Ruiz et al., 2004; Markweg-Hanke et al., 1995) and other central metabolic enzymes (Rose, 1971; Van Schaftingen and Hers, 1981; Alam et al., 2017).

  4. PlaF is constitutively expressed (Figure 7—figure supplement 2 and Erdmann et al., 2018) at low levels suggesting that PlaF-catalyzed GPL remodeling may have general importance for P. aeruginosa physiology.

  5. The P. aeruginosa ΔplaF strain shows strongly impaired killing of G. mellonella and human macrophages compared to WT (Figure 3), thus revealing the important function of PlaF-mediated GPL remodeling for P. aeruginosa virulence.

It is well known that the global diversity of GPL acyl chains in eukaryotes derives from de novo synthesis (Kennedy pathway) and remodeling (Lands cycle) pathways, which are differentially regulated (Jacquemyn et al., 2017). In the Lands cycle, GPLs are targeted by PLA and acyltransferases that respectively remove and replace acyl chains in GPLs by a recently described mechanism (Zhang et al., 2021; Mouchlis et al., 2015). We suggest that PlaF is the PLA that alters P. aeruginosa membranes by hydrolysis of the main classes of GPLs, namely PE, PG, and PC. Although the observed changes may be caused by the absence of PlaF in the membrane of P. aeruginosa, it is more likely that PlaF directly hydrolyses GPLs as only low concentrations of PlaF were detected in the cell (Figure 7—figure supplement 2). The exact molecular function of PlaF in GPL-remodeling and the regulation of virulence of P. aeruginosa remains unknown. One possibility is that PlaF tunes the concentration of low-abundance GPL species in the membrane, creating a suitable membrane environment for the stabilization of membrane proteins or protein complexes (Corradi et al., 2019). In addition, PlaF-generated GPLs might have a more sophisticated function for membrane-embedded virulence-related proteins. This was demonstrated for ABHD6, a human membrane-bound PLA, which controls the membrane concentration of lipid transmitter 2-arachidonoylglycerol involved in regulating the endocannabinoid receptor (Marrs et al., 2010). Notably, human ABHD6 and PlaF share ~50% sequence similarity and hydrolize similar substrates (Bleffert et al., 2019).

Although PlaF is an important enzyme involved in GPL metabolism, future research should reveal (i) which acyltransferase is involved in the acylation of lysoGPLs produced by PlaF, (ii) if PlaF has acyltransferase activity as described for cPLA2γ involved in the Lands cycle in humans (Asai et al., 2003), and (iii) if periplasmic lysophospholipase TesA (Kovačić et al., 2013) and the recently discovered intracellular PLA PlaB (Weiler et al., 2022) are involved in the Lands cycle.

Structural insights into dimerization and ligand-mediated regulation of PlaF activity

The high-resolution structure of PlaF with the natural ligands (FAs) bound to its active site represents the first dimeric structure of a full-length, single-pass TM protein (Figure 4). It contributes to our understanding of the role of TM-JM domain-mediated dimerization for the biological activity of single-pass TM proteins, which is undisputed in bacteria and eukaryotes, yet, poorly understood at the atomic level due to the lack of full-length dimeric structures (Bocharov et al., 2017; Fink et al., 2012). The present structure-function relationship of single-pass TM dimers derives from structural data of isolated TM helices without their soluble domains. Therefore, their biological relevance remains questionable (Bocharov et al., 2017).

The crystal structure of PlaF reveals unprecedented details of interactions between the membrane-spanning TM-JM domains and underlines the role of PlaF for degradation of membrane GPLs. The TM and JM domains are not distinct but fold into a long kinked α-helix (Figure 4a). This is different from the structure of a human epidermal growth factor receptor (EGFR), the only structure of an isolated TM-JM domain, in which TM and JM helices are connected by an unstructured loop (Bragin et al., 2016; Endres et al., 2013). The mechanism undergoing PlaF dimerization likely differs from the EGFR family, although it is not excluded that the truncation of soluble domains might destabilize the TM-JM dimer of EGFR, leading to structural changes. We identified intramolecular interactions of 13 residues from the catalytic domain of PlaF with the JM domain, which clearly demonstrates the stabilizing role of the soluble domain on the TM-JM helix. Sole interactions of TM-JM helices result in the formation of a coiled-coil structure (Figure 4b) that stabilizes the PlaF dimer by burying the surface of 656 Å2, which is slightly larger than the interface of the glycophorin TM helix dimer (400 Å2) without the JM region (MacKenzie et al., 1997). The biological relevance of PlaF dimerization is corroborated by crosslinking experiments with P. aeruginosa cells, which revealed the in vivo occurrence of PlaF dimer (Figure 5a). Furthermore, enzyme activity measurements and MST analysis of protein-protein interactions revealed that the activity decreases and dimerization increases as a function of increasing PlaF concentration in vitro (Figure 5c). These findings open the question of regulation of dimerization-mediated PlaF inhibition in vivo and the role of membrane GPLs and their hydrolytic products in this process. Homodimerization mediated via TM-JM interactions was previously shown to be required for activation of single-span TM proteins from receptor tyrosine kinase (Li and Hristova, 2010) and ToxR-like transcriptional regulator (Buchner et al., 2015). However, structural and mechanistic details remained unknown.

A metabolic role of PlaF related to the liberation of FAs and lysoGPLs from membrane GPL substrates addresses the question of regulating PlaF function by substrates or products. A dimer interface with mainly hydrophobic interactions and a few H-bonds detected in the JM region (Figure 4b) seems to be designed to interact with amphipathic GPLs. However, it remains to be elucidated if PlaF-GPL interactions regulate PlaF dimerization and its activity as shown for interactions of SecYEG with cardiolipin and bacteriorhodopsin with sulfated tetraglycosyldiphytanylglycerol (Corradi et al., 2019; Essen et al., 1998).

C10–C14 FAs exert competitive inhibition as in vitro effectors of PlaF (Figure 6a) and enhance dimerization (Figure 6c) in the concentration range (0.5–7.5 mM) similar to the intracellular concentration of FAs in E. coli (~2–4 mM) (Lennen et al., 2013). The dimerization-triggering function of FAs is strengthened by observing a mixed-type inhibition (Figure 6b), which indicates that FAs affect PlaF not only by binding to the active site but also by modulating the oligomerization equilibrium (Gabizon and Friedler, 2014). Interestingly, we identified FA ligands in the PlaF structure bound to the PlaF active site cleft (Figure 4) that were copurified with PlaF from P. aeruginosa (Figure 4—figure supplement 1). Furthermore, we identified an OG molecule, used for purification, in the active site of PlaF. The pseudo-ligand (OG) and natural products (FAs) form an intricate interaction network connecting the catalytic (S137) with the dimerization site (S29, T32, and V33) in the JM domain (Figure 4e). Although the static structure of dimeric PlaF cannot explain how FAs trigger dimerization, we speculate that in vivo, the position of the OG molecule is occupied by FAs, which facilitates the interaction between the two JM-helices, stabilizing the dimer.

Atomistic model of PlaF catalyzed hydrolysis of membrane GPLs

The question remains of how does the PlaF dimer-to-monomer transition activate PlaF in the GPL bilayer? The active sites in the crystal structure of di-PlaF already adopt catalytically active conformations (Figure 4a), suggesting that the activation of PlaF most likely does not involve structural rearrangements of the active site. To unravel a possible effect of the structural dynamics of PlaF in the membrane on enzyme regulation by dimerization, we performed extensive MD simulations and configurational free energy computations on dimeric and monomeric PlaF embedded into a GPL bilayer mimicking the bacterial cytoplasmic membrane. While structural changes within di-PlaF and monomeric PlaF were moderate (Supplementary file 10), monomeric PlaF spontaneously tilted as a whole toward the membrane, constraining the enzyme protein in a configuration with the opening of the active site cleft immersed into the GPL bilayer (Figure 7a and b). A configuration similar to t-PlaF was observed for monomeric Saccharomyces cerevisiae lanosterol 14α-demethylase, a single TM spanning protein acting on a membrane-bound substrate (Monk et al., 2014). In t-PlaF, GPL can access the active site cleft directly from the membrane with the sn-1 acyl chain entering the first (Wittgens et al., 2017). This is unlikely in di-PlaF, in which the opening of the active site cleft is >5 Å above the membrane (Figure 7e). There, a GPL would need to leave the bilayer into the water before entering the active site cleft, which is thermodynamically unfavorable.

Based on the experimental evidence, we propose a hitherto undescribed mechanism by which the transition of PlaF between a dimeric, not-tilted to a monomeric, tilted configuration is intimately linked to the modulation of the PlaF activity. This mechanism, to the best of our knowledge, expands the general understanding of mechanisms of inactivation of integral single-pass TM proteins and differs from suggested allosteric mechanisms implying structural rearrangements (even folding), mostly in the JM domain, upon ligand binding as underlying causes for functional regulation (Bocharov et al., 2017). Rather, for PlaF, monomerization followed by a global reorientation of the single-pass TM protein in the membrane is the central, function-determining element.

Based on computed free energies of association (Figure 7d) and tilting (Figure 7c), and taking into account the concentration range of PlaF in P. aeruginosa, PlaF preferentially exists as t-PlaF in the cytoplasmic membrane (Figure 7e). Increasing the PlaF concentration in the membrane will thus shift the equilibrium toward di-PlaF. This result can explain the observations that PlaF, an enzyme with membrane-disruptive activity, is found in only very low amounts (Figure 7—figure supplement 2) in WT P. aeruginosa cells and that overproduction of PlaF in P. aeruginosa is not harmful to the cells.

Implications for drug development

Based on our observation that P. aeruginosa ΔpalF shows strongly attenuated virulence, we suggest that interfering with PlaF function might be a promising target for developing new antibiotics against P. aeruginosa. This class of antibiotics should be potent assuming that GPL remodeling plays a global role in the virulence adaptation in bacteria through simultaneous regulation of virulence-related processes (Benamara et al., 2014; Le Sénéchal et al., 2019; El Khoury et al., 2017; Blanka et al., 2015). Analogously, eukaryotic PLAs regulating inflammatory pathways through the release of arachidonic acid were recently suggested as potential targets of anti-inflammatory drugs (Mouchlis and Dennis, 2016). Our structural and mechanistic studies provide a basis for targeting PlaF by competitive inhibition and interfering with dimerization (Gabizon and Friedler, 2014; Hopkins and Groom, 2002).

Materials and methods

Cloning, protein expression, and purification

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Molecular biology methods, DNA purification, and analysis by electrophoresis were performed as described previously (Kovacic et al., 2016). For the expression of PlaF, P. aeruginosa PAO1 (WT) cells transformed (Choi et al., 2006) with plasmid pBBR-pa2949 (Kovacic et al., 2016), here abbreviated as p-plaF, were grown overnight at 37°C in lysogeny broth (LB) medium supplemented with tetracycline (100 µg/ml) (Bleffert et al., 2019). The total membrane fraction of P. aeruginosa p-plaF was obtained by ultracentrifugation, membranes were solubilized with Triton X-100, and PlaF was purified using Ni-NTA IMAC and buffers supplemented with 30 mM OG (Bleffert et al., 2019). For biochemical analysis, PlaF was transferred to Tris-HCl (100 mM, pH 8) supplemented with 30 mM OG (Table 2).

Table 2
Material used in this work.
MaterialOrdering details
Galleria mellonella larvaeFauna Topics GmbH, order number: 527
Trypsin, porcine, MS gradeMerck, order number: 650279
Anti-SecG antiserumgift of R. Voulhoux, CNRS AMU LCB, Marseille
Anti-lipid A antibodiesAcris Antibodies, Herford, Germany, order number: BP 2235
Ni-NTA agaroseMacherey–Nagel, Düren, Germany, order number: 745400
n-Octyl-β-D-glucosideMerck, order number: 850511P
para-Nitrophenyl butyrateSigma-Aldrich, order number: N9876
GlycerophospholipidsAvanti Polar Lipids, Alabaster, USA
NEFA-HR(2) kitWako Chemicals, Richmond, USA, order number: 999-34691
[N-((6-(2,4-DNP)amino)hexanoyl)-1-(BODIPYFL C5)-2-hexyl-sn-glycero-3-phosphoethanolamine]Thermo Fisher Scientific Inc, Waltham, MA, order number: A10070
1-O-(6-BODIPY558/568-aminohexyl)-2-BODIPYFL C5-Sn-glycero-3-phosphocholineThermo Fisher Scientific Inc, Waltham, MA, order number: A10072
N-((6-(2,4-dinitrophenyl)amino)hexanoyl)-2-(4,4-difluoro-5,7-dimethyl-4-bora-3a,4a-diaza-s-indacene-3-pentanoyl)–1-hexadecanoyl-sn-glycero-3-phosphoethanolamine triethylammonium saltThermo Fisher Scientific Inc, Waltham, MA, order number: D23739
Thermomyces lanuginosus PLA1Sigma-Aldrich, order number: L3295
Naja mocambique mocambique PLA2Sigma-Aldrich, order number: P7778
Dimethyl pimelimidateMerck, order number: 80490
Bis(sulfosuccinimidyl) glutarateThermo Fisher Scientific, order number: 21610
Bis(sulfosuccinimidyl) suberateThermo Fisher Scientific, order number: 21586
PD-10 columnsMerck, order number: GE17-0851-01
NHS Labeling KitNanoTemper, Munich, Germany, order number: MO-L011
4-Methylumbelliferyl palmitateSigma-Aldrich; order number: M7259
CytoTox 96 non-radioactive cytotoxicity assayPromega, order number: G1780
NucleoSpin RNA Preparation KitMacherey–Nagel, Düren, Germany, order number: 740955
RNase-Free DNase KitQiagen, Hilden, Germany, order number: 79254
Ambion DNA-Free DNase KitThermo Fisher Scientific, Darmstadt, Germany, order number: AM1906
Maxima First Strand cDNA Synthesis KitThermo Fisher Scientific, Darmstadt, Germany, Order number: K1641
SYBR Green/ROX qPCR Master MixThermo Fisher Scientific, Darmstadt, Germany, order number: K0221
N-Methyl-N-(trimethylsilyl) trifluoroacetamideSigma-Aldrich; order number: 69479

SDS-PAGE, zymography, and immunodetection

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The protein analysis by electrophoresis under denaturation conditions (Laemmli, 1970), in-gel esterase activity (zymography), and immunodetection by Western blotting were performed as described previously (Kovacic et al., 2016). The protein concentration was determined by UV spectrometry using a theoretical extinction coefficient of PlaF containing a C-terminal His6-tag of 22,920 M–1 cm–1 (Bleffert et al., 2019).

Enzyme activity assays, inhibition, and enzyme kinetic studies

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Esterase activity assays with p-nitrophenyl FA esters as substrates were performed in 96-well microtiter plates as described previously (Kovacic et al., 2016). Phospholipid substrates purchased from Avanti Polar Lipids (Table 2) were prepared for PLA activity assays (25 µl enzyme+25 µl substrate) performed as described previously (Jaeger and Kovacic, 2014). The amount of FAs released by the PLA activity of PlaF was determined using the NEFA-HR(2) Kit (Wako Chemicals, Richmond, USA) (Bleffert et al., 2019). PLA1 and PLA2 activities of PlaF were measured using fluorescent substrates purchased from Thermo Fisher Scientific Inc (Table 2): PLA1-PE, [N-((6-(2,4-DNP)amino)hexanoyl)-1-(BODIPYFL C5)-2-hexyl-sn-glycero-3-phosphoethanolamine]; PLA2-PC, 1-O-(6-BODIPY558/568-aminohexyl)-2-BODIPYFL C5-Sn-glycero-3-phosphocholine; and PLA2-PE, N-((6-(2,4-dinitrophenyl)amino)hexanoyl)-2-(4,4-difluoro-5,7-dimethyl-4-bora-3a,4a-diaza-s-indacene-3-pentanoyl)-1-hexadecanoyl-sn-glycero-3-phosphoethanolamine triethylammonium as described by da Mata Madeira et al., 2016. Measurements were performed using a plate reader in 96-well plates at 25°C by combining 50 µl of the substrate with 50 µl PlaF (0.7 µg/ml), or control enzymes, the PLA1 of Thermomyces lanuginosus (5 U/ml) and the PLA2 or Naja mocambique mocambique (0.7 U/ml).

Inhibition

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The inhibition of PlaF by FAs was assayed by combining FA dissolved in DMSO (20-fold stock solution) with para-nitrophenyl butyrate (p-NPB)substrate solution followed by the addition of the PlaF sample (8 nmol) and spectrophotometric enzyme activity measurement using p-NPB substrate (Tian and Tsou, 1982). In control experiments, all compounds except FA were combined to assess PlaF activity in the absence of FA. Inhibition constants were calculated by fitting enzyme kinetic parameters obtained by varying FA concentration (0, 0.5, 1.5, 2.5, 5, and 7.5 mM) for different substrate concentrations (0.05, 0.1, 0.2, 0.3, 0.5, and 1 mM) (Kenakin, 2012).

Subcellular localization

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Membranes from P. aeruginosa WT and p-plaF (PlaF overproduction strain) were isolated as described previously (Kovacic et al., 2016). To separate integral from peripheral membrane proteins, total cell membranes were incubated for 30 min at room temperature with: 10 mM Na2CO3 (pH 11), 4 M urea (in 20 mM MES buffer pH 6.5) or 2% (w/v) Triton X-100 (in 20 mM MES buffer pH 6.5). After the incubation, the samples were centrifuged for 30 min at 180,000g to separate membranes from solubilized proteins.

The separation of the inner and outer membrane was performed with a discontinuous sucrose gradient by ultracentrifugation at 180,000g for 72 hr and 4°C (Viarre et al., 2009). The sucrose gradient consisted of 1.5 ml fractions with 35%, 42%, 46%, 50%, 54%, 58%, 62%, and 65% (w/v) sucrose in 100 mM Tris-HCl, pH 7.4. Isolated membranes from P. aeruginosa WT were suspended in buffer containing 35% (w/v) sucrose and loaded on the top of the discontinuous sucrose gradient. Fractions were collected from the bottom (pierced tube), and sucrose concentration was determined with a refractometer (OPTEC, Optimal Technology, Baldock, UK). To determine the orientation of catalytic PlaF domain P. aeruginosa p-plaF cells (10 ml culture with OD580nm 1 grown in LB medium at 37°C) were harvested by centrifugation (4000g, 4°C, 5 min) and suspended in 1 ml Tris-HCl buffer (50 mM, pH 7.5, 10% sucrose (w/v)) followed by shock freezing with liquid nitrogen (Eichler and Wickner, 1998). Cells were thawed to room temperature and centrifuged (4000g, 4°C, 5 min) followed by incubation of the pellet for 1 hr on ice in Tris-HCl buffer (30 mM, pH 8.1, sucrose 20% (w/v) EDTA 10 mM). Trypsin (20 µl, 1 mg/ml) was added to the suspension containing the cells with the permeabilized outer membrane and incubated at room temperature for up to 5 hr. The proteolytic reaction was stopped with onefold SDS-PAGE sample buffer and incubation for 10 min at 99°C. Immunodetection of SecG with anti-SecG antiserum (gift of R. Voulhoux, CNRS AMU LCB, Marseille) and lipid A antibodies (BP 2235, Acris Antibodies, Herford, Germany) was performed as described above for PlaF using the respective antisera at 1/2000 and 1/1000 dilutions.

Cross-linking assays

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In vitro cross-linking using the bifunctional cross-linking reagents DMP was performed as previously described (de Jong et al., 2017) with the following modifications. PlaF (10 µl, 15.5 µM) purified with OG was incubated with 6 µl freshly prepared DMP (150 mM in 100 mM Tris-HCl, pH 8.4), BS2G (5 mM in 100 mM Tris-HCl, pH 8.0) and BS3 (5 mM in 100 mM Tris-HCl, pH 8.0) for 90 min (Table 2). The cross-linking reaction was terminated with a 5 µl stop solution (50 mM Tris-HCl, 1 M glycine, NaCl 150 mM, pH 8.3). For in vivo cross-linking, P. aeruginosa p-plaF and EV strains were grown in LB medium at 37°C to OD580nm 1. Cells were harvested by centrifugation (10 min, 4000g, 4°C), suspended in 1/20 volume of Tris-HCl (pH 8.3, 100 mM, NaCl 150 mM), and treated with the same volume of freshly prepared cell-permeable cross-linking reagent DMP (0, 20, 30, and 50 mM in Tris-HCl buffer 100 mM, pH 8.4) for 2 h. The cross-linking reaction was terminated with the same volume of stop solution (50 mM Tris-HCl, 1 M glycine, 150 mM NaCl, pH 8.3).

Analysis of concentration-dependent dimerization

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Purified PlaF (20 µl, 50–60 µM) was transferred from the purification buffer into the labeling buffer (Na-PO4 20 mM, pH 8.3) supplemented with OG (30 mM) using PD-10 columns (GE Healthcare, Solingen, Germany) according to the manufacturer’s protocol. Labeling was performed by incubating PlaF with 15 µl dye (440 µM stock solution) for 2.5 hr using the NHS Labeling Kit (Table 2). PlaF was then transferred into a purification buffer using PD-10 columns. Non-labeled PlaF was diluted with the same buffer in 16 steps by combining the same volume of the protein and buffer, yielding samples with concentrations from 26.9 µM to 1.6 nM. Samples containing 100 nM labeled PlaF were incubated for 16 hr at room temperature in the dark, and MST experiments were performed using the Monolith NT.115 device (NanoTemper, Munich, Germany) with the following setup: MST power, 60%; excitation power 20%; excitation type, red; 25°C. Constants were calculated according to the four-parameter logistic, nonlinear regression model using Origin Pro 2018 software.

The enzymatic activity of PlaF samples used for MST analysis was assayed by combing 15 µl of enzyme and 15 µl 4-methylumbelliferyl palmitate (4-MUP, 2 mM) dissolved in purification Tris-HCl (100 mM, pH 8) containing 10% (v/v) propan-2-ol (Table 2). Fluorescence was measured for 10 min (5 s steps) using a plate reader in black 96-well microtiter plates at 30°C.

Construction of a P. aeruginosa ∆plaF, and ∆plaF::plaF strains

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The mutagenesis vector pEMG-ΔplaF (Figure 2—figure supplement 2) was generated with upstream and downstream regions of plaF gene amplified by standard PCR using Phusion DNA polymerase, a genomic DNA of P. aeruginosa PAO1 as a template, and primer pairs 5′-ATATATGAATTCTCTGCTCGGCGCGAAACGCAGCGP-3′/5′-ATATATACGCGTGGGTGTCCGAAGGCTTCAGGAAAAAAGGGGC-3′ and 5′-ATATATACGCGTAAACGCGAACCGGCGCCTGGG-3′/5′-CTGGATGAATTCTGGCCTGGACACCGACAAGGAAGTGATCAAGG-3′, respectively. DNA fragments upstream and downstream of the plaF gene were cloned into the pEMG vector by ligation of DNA fragments hydrolyzed with EcoRI restriction endonuclease. P. aeruginosa PAO1 (WT) cells were transformed with the pEMG-ΔplaF and P. aeruginosa ΔplaF mutant strain was generated by homologous recombination (Martínez-García and de Lorenzo, 2011). Generation of pUC18T-mini-Tn7T-Gm-plaF plasmid (Figure 2—figure supplement 2) for recombination of plaF gene containing 128 bp upstream region of plaF with a chromosome of P. aeruginosa ΔplaF. A DNA fragment containing the upstream region and plaF gene was amplified using primer pair 5′-AATAGAGCTCACCGCCGTCCTTAGGTTC-3′/5′-AATAGAGCTCCGTTTTCAGCGACCGGC-3′ from the genomic DNA of P. aeruginosa PAO1. Both primers contained the restriction site SacI for cloning into the pUC18T-mini-Tn7T-Gm (gifts from Herbert Schweizer, Addgene plasmids #63121, #64968, and #64946). P. aeruginosaplaF was transformed with pUC18T-mini-Tn7T-plaF-Gm and helper plasmid pTNS2 encoding the Tn7 site-specific transposase ABCD by tri-parental conjugation and the positive clones were identified by PCR using primer pair 5′-GCACATCGGCGACGTGCTCTC-3′/5′-CACAGCATAACTGGACTGATTTC-3′. The gentamycin-resistance gene was excised from P. aeruginosaplaF::plaF-Gm by Flp-recombinase produced from pFLP3 plasmid (Choi et al., 2005).

G. mellonella virulence model

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G. mellonella larvae (Table 2) were sorted according to size and split into groups of 10 in Petri dishes. P. aeruginosa WT, the ΔplaF, and the ΔplaF::plaF strains were grown overnight and sub-cultured to mid-log phase in LB media at 37°C. The bacteria were washed twice with PBS and adjusted to OD600 0.055, which equals 5×104 bacteria/µl. This suspension was diluted in PBS to the infection dose of 500 bacteria per 10 µl, which were injected into the hindmost left proleg of the insect. Hereby, PBS injections were used as infection control and untreated larvae as viability control. If more than one larvae was dying within the control group, the experiment was repeated. The survival of larvae incubated at 30°C was monitored (Koch, 2014).

Cytotoxicity assay

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BMDMs were isolated from the bones of C57BL/6 mice and cultured in RPMI supplemented with 20% (v/v) conditioned L929 medium to allow for differentiation into macrophages for at least 7 days. BMDMs were seeded at a concentration of 5×105 cells in a 24-well plate. The BMDMs cells (n=10) were infected with 5×105 bacteria (cultivated overnight in LB medium at 37°C), which accounts for MOI 1 (Mittal et al., 2016). PBS treated cells served as viability control. Supernatants were taken at 0, 1, 3, and 6 hr post-infection. LDH levels were determined (n=6) using the CytoTox 96 Non-Radioactive Cytotoxicity Assay according to the manufacturer’s protocol. As 100% killing control, uninfected cells were lysed with 1% (v/v) Triton-X100. Statistical analysis was performed using a one-way ANOVA to determine significant changes of normally distributed values obtained from two independent experiments with 10 samples each.

Growth curves

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The growth of P. aeruginosa WT and ΔplaF cultures in Erlenmeyer flasks (agitation at 160 rpm) was monitored by measuring OD580nm for 24 hr. OD580nm was converted to colony-forming units by multiplying with the factor 8×108 experimentally determined for P. aeruginosa PAO1 strain from our laboratory.

Quantitative real-time-PCR

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RNA was isolated from P. aeruginosa PA01 and ΔplaF grown overnight (37°C, LB medium) with the NucleoSpin RNA Preparation Kit and genomic DNA was quantitatively removed using RNase-Free DNase Kit and Ambion DNA-Free DNase Kit according to the manufacturer’s recommendations (Table 2). One µg of RNA was transcribed into cDNA using the Maxima First Strand cDNA Synthesis Kit (Table 2). For the quantitative real-time-PCR (qRT-PCR), 50 ng of cDNA was mixed with SYBR Green/ROX qPCR Master Mix (Table 2) to a total volume of 20 μl and qRT-PCR was performed as described previously (Savli et al., 2003). Following primers were used for rpoD (3′-CAGCTCGACAAGGCCAAGAA-5′, CCAGCTTGATCGGCATGAAC), rpoS (3′-CTCCCCGGGCAACTCCAAAAG-5′, 3′-CGATCATCCGCTTCCGACCAG-5′) and plaF (3′-CGACCCTGTTGCTGATCCAC-5′, 3′-ACGTCGTAGCTGGCCTGTTG-5′).

Lipidomic analysis of GPLs extracted from cell membranes

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The cells of P. aeruginosa WT, ∆plaF, and ∆plaF::plaF cultures grown overnight in 15 ml LB medium (Supplementary file 3) at 37°C were harvested by centrifugation at 4000g and 4°C for 15 min and suspended in 2 ml ddH2O followed by boiling for 10 min to inactivate phospholipases. Cells were harvested by centrifugation (4000g, 4°C, 15 min) and total lipids were extracted from the cell pellet (Gasulla et al., 2013). Briefly, after boiling the water was removed by centrifugation (4000g, 4°C, 15 min). Lipids were extracted with CHCl3:CH3OH=1:2 (v/v) and the organic phase was collected. The extraction was repeated with CHCl3:CH3OH=2:1 (v/v) and the organic phases were combined. One volume of CHCl3 and 0.75 volumes of an aqueous solution containing 1 M KCl and 0.2 M H3PO4 were added to the combined chloroform/methanol extracts. Samples were vortexed and centrifuged (2000g, 5 min). The organic phase was withdrawn and the solvent of the lipid extract was evaporated under a stream of N2. Total lipids were dissolved in CHCl3:CH3OH=2:1 (v/v). GPLs were quantified by Q-TOF mass spectrometry (Q-TOF 6530; Agilent Technologies, Böblingen, Germany) as described elsewhere (Gasulla et al., 2013). Statistical analysis of the GPL amount was performed using the T-test and the Shapiro-Wilk method to determine significant changes of normally distributed values obtained from four P. aeruginosa WT lipidome and four ∆plaF samples. Ratio of PlaF and GPLs was calculated knowing GPLs extraction yield of 40 µg GPLs per 1 ml P. aeruginosa p-plaF (OD580nm 1) and PlaF purification yield of ~1 µg from 1 ml P. aeruginosa p-plaF culture with OD580nm 1 (Bleffert et al., 2019).

GC-MS analysis of FA

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FAs were extracted from PlaF purified from 13 g P. aeruginosa p-plaF cells with OG using four parts of organic solvent (CHCl3:CH3OH=2:1). Extraction was repeated three times, the chloroform extracts were combined, chloroform was evaporated, and FAs were dissolved in 200 µl chloroform. The chloroform extract was mixed with 10 volumes of acetonitrile and filtered through a 0.2 µm pore size filter. For GC-MS analysis, FA extracts and standards (C10-, C11-, C14-, C15-, C16-, and C18-FA; C16-, C18-, and C20-primary fatty alcohol) were converted into their trimethylsilyl esters and trimethylsilyl ethers, respectively. 900 µl of the sample or standard solution (CHCl3:acetonitrile=1:5) was mixed with 100 µl N-methyl-N-(trimethylsilyl) trifluoroacetamide and heated to 80°C for 1 hr. The GC-MS system consisted of a Trace GC Ultra gas chromatograph, TriPlus autosampler, and an ITQ 900 mass spectrometer (Thermo Fisher Scientific, Waltham, MA). Analytes were separated on a Zebron-5-HT Inferno column (60 m × 0.25 mm i.d., 0.25 µm film thickness, Phenomenex, USA). Helium was used as carrier gas at a constant gas flow of 1.0 ml/min. The oven temperature program was as follows: 80°C; 5°C/min to 340°C, held for 5 min. The injector temperature was held at 290°C, and all injections (1 µl) were made in the split mode (1:10). The mass spectrometer was used in the electron impact (EI, 70 eV) mode and scanned over the range m/z 25–450 with an acquisition rate of 3 microscans. The transfer line and ion source were both kept at 280°C. Data processing was performed by the use of the software XCalibur 2.0.7 (Thermo Fisher Scientific). FAs from the PlaF sample were identified by comparison of their retention times and mass spectra with FA standards.

Reaction of purified PlaF (620 µl, 300 µg/ml) with 1-(9Z-octadecenoyl)-2-pentadecanoyl-glycero-3-phospho-(1′-rac-glycerol) (PG15:0-18:1, 0.5 mM) in 4 ml NEFA buffer was conducted for 24 hr at 37°C followed by extraction of FAs, derivatization, and GC quantification. FAs were transferred to 15 ml Falcon tubes by dissolving in 500 µL CH2Cl2 twice. After evaporation to dryness the remaining fatty acids were derivatized to their methyl esters according to Funada et al. with modifications (Funada and Hirata, 1999). Briefly the residues were dissolved in 1 ml 1 M sulfuric acid in methanol. For esterification the Falcon tubes were placed in an ultrasonic bath for 30 min. The fatty acid methyl esters (FAMEs) were extracted after addition of 3.3 ml water and 1.7 ml hexane by vigorous shaking on a Vortex for 1 min. The upper organic phase was withdrawn and dried over sodium carbonate. An aliquote was directy used for GC-MS analysis. A 1 mM fatty acid mixture in methanol (C10:0, C12:0, C14:0, C16:0, C18:0, C17:0 cyc (9,10), C18:1 cis-Δ9, C18:1 trans-Δ9, C18:1 trans-Δ11, C18:2 cis,cis-Δ9,12, C18:2 trans,trans-Δ9,12 and C18:3 cis,cis,cis-Δ9,12,15) was diluted to 50, 100, 200 and 400 µM and derivatized in the same manner as above. The Agilent GC-MS system consisted of a gas chromatograph 7890A and an autosampler G4513A coupled to a quadrupole mass spectrometer MS G3172A (Agilent, CA, USA). Analytes were separated on a SGE BPX70 column (30 m x 0.32 mm i.d., 0.25 µm film thickness, Thermo Fisher Scientific, USA). Helium was used as carrier gas at a constant gas flow of 1.5 ml/min. The oven temperature program employed for analysis of FAMEs was as follows: 120°C; 20°C/min to 160°C; 3°C/min to 200°C; 20°C to 220°C, held for 8.7 min. The injector temperature was held at 250°C, and all injections (1 µl) were made in the split mode (1:10). The mass spectrometer was used in the electron impact (EI) mode at an ionizing voltage of 70 eV. Analytes were scanned over the range m/z 50 - 400 with a spectrum recording interval of 4 scans/sec. The GC interface temperature was held at 250°C. The MS source and quadrupole temperatures were kept at 280°C and 150°C, respectively. Data processing was performed by use of the software ChemStation E.02.02.1431 (Agilent, CA, USA). Fatty acids from PlaF samples were identified by comparison of their retention times and mass spectra with those of fatty acid standards and published data (Yang et al., 2013; Benamara et al., 2014; Chao et al., 2010). Quantification of FAMEs C16:0 (1), C17:0 cyc(9,10) (4), C18:0 (5) and C18:1 trans-Δ11 (6) (Figure 1) were performed by external calibration with the corresponding reference compounds. C18:1 cis-Δ11 (7) was quantified by use of the calibration curve of oleic acid (C18:1 cis-Δ9) justified by the almost congruent calibration curves of elaidic acid (C18:1 trans-Δ9) and C18:1 trans-Δ11.

Crystallization, data collection, structure determination, and analysis

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PlaF purified with OG was crystallized as described previously (Bleffert et al., 2019). The X-ray diffraction data were recorded at beamline ID29 of the European Synchrotron Radiation Facility (ESRF, Grenoble, France) and processed as described (Bleffert et al., 2019). The structure was determined by molecular replacement using the automated pipeline ‘MrBUMP’ from the CCP4 package (Keegan et al., 2011). In detail, a combination of PHASER (McCoy et al., 2007), REFMAC (Murshudov et al., 1997), BUCCANEER (Cowtan, 2006), and SHELXE (Hübschle et al., 2011) resulted in an interpretable electron density map to expand the placed model by molecular replacement using the model built with HsaD from Mycobacterium tuberculosis (PDB code: 2VF2) (Lack et al., 2010). Phase improvement was achieved by running several cycles of automated model building (ARP/wARP, CCP4) and refinement using the PHENIX (Adams et al., 2011) package. The model was further corrected by manual rebuilding using the program COOT (Emsley and Cowtan, 2004). Detailed statistics on data collection and refinement are provided in Table 1. None of the residues is in disallowed regions according to Ramachandran plots generated with MolProbity (PHENIX) (Adams et al., 2002). The secondary structure was defined according to Kabsch and Sander (Kabsch and Sander, 1983). Interaction surface area was determined by PISA server (Krissinel and Henrick, 2007). Coordinates and structure factors for PlaF have been deposited in the Protein Data Bank under accession code 6I8W.

Identification of structural homologs of PlaF

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PlaF structural homologs were defined as protein structures from a non-redundant subset of PDB structures with less than 90% sequence identity to each other (PDB90 database, 12.10.2020) with a Z-score higher than 2 according to the DALI server (Holm and Rosenström, 2010). Sequence alignment based on structural superimposition of all 357 homologs of PlaFB (all 340 homologs of PlaFA were among PlaFB homologs) was used to identify proteins with homology in TM-JM helix of PlaF (residues 1–38). To evaluate homology, 39 3D structures with partial conservation of TM-JM helix were superimposed with the PlaF structure using Pymol (http://www.pymol.org) (Figure 4—figure supplement 3).

Sequence analysis

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A protein sequence of PlaF was used for a BLAST search of Pseudomonas Genome Databank (https://www.pseudomonas.com/) to identify PlaF orthologs in 4660 sequenced P. aeruginosa genomes. Pseudomonas Genome Databank BLAST search was extended to all pathogenic Pseudomonas species designated as those with assigned risk group 2 according to the German classification of prokaryotes into risk groups. NCBI BLAST (https://blast.ncbi.nlm.nih.gov/Blast.cgi) was used to identify PlaF homologs in other pathogenic bacteria.

Molecular dynamics simulations of dimer and monomers

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The crystal structure of the PlaF dimer was used as the starting point for building the systems for molecular dynamics (MD) simulations. Five missing C-terminal residues on both chains were added by using MODELLER (Sali and Blundell, 1993), and all small-molecule ligands were removed. The dimer was oriented into the membrane using the PPM server (Lomize et al., 2012). From the so-oriented dimer structure, chain B was deleted to obtain a PlaFA monomer in a dimer-oriented configuration; in the same way, chain A was deleted to keep PlaFB. Additionally, the PlaFA and PlaFB monomers were oriented by themselves using the PPM server, yielding tilted configurations (t-PlaFA and t-PlaFB). These five starting configurations, di-PlaF, PlaFA, PlaFB, t-PlaFA, and t-PlaFB, were embedded into a DOPE:DOPG=3:1 membrane with CHARMM-GUI v1.9 (Jo et al., 2009) resembling the native inner membrane of Gram-negative bacteria (Benamara et al., 2014; Murzyn et al., 2005). A distance of at least 15 Å between the protein or membrane and the solvation box boundaries was considered. KCl at a concentration of 0.15 M was included in the solvation box to obtain a neutral system. The GPU particle mesh Ewald implementation from the AMBER16 molecular simulation suite (Le Grand et al., 2013; Darden et al., 1993) with the ff14SB (Maier et al., 2015) and Lipid17 (Dickson et al., 2014; Skjevik et al., 2016; Case, 2017) parameters for the protein and the membrane lipids, respectively, were used; water molecules were added using the TIP3P model (Jorgensen et al., 1983). For each protein configuration, 10 independent MD simulations of 2 µs length were performed. Covalent bonds to hydrogens were constrained with the SHAKE algorithm (Ryckaert et al., 1977) in all simulations, allowing the use of a time step of 2 fs. Details of the thermalization of the simulation systems are given below. All unbiased simulations showed stable protein structures (Figure 7—figure supplement 1) and membrane phases, evidenced by electron density and order parameter calculations (Figure 7—figure supplement 1). The area per lipid through all simulations calculated for the leaflet opposite to the one where PlaF was embedded was 61.3±0.13 Å2 (mean ± SEM), similar to values reported previously (Murzyn et al., 2005).

Thermalization and relaxation of simulated systems

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Initially, systems were energy-minimized by three mixed steepest descent/conjugate gradient calculations with a maximum of 20,000 steps each. First, the initial positions of the protein and membrane were restrained, followed by a calculation with restraints on the protein atoms only, and finally a minimization without restraints. The temperature was maintained by using a Langevin thermostat (Quigley and Probert, 2004), with a friction coefficient of 1 ps–1. The pressure, when required, was maintained using a semi-isotropic Berendsen barostat (Berendsen et al., 1984), coupling the membrane (x-y) plane. The thermalization was started from the minimized structure, which was heated by gradually increasing the temperature from 10 to 100K for 5 ps under NVT conditions, and from 100 to 300K for 115 ps under NPT conditions at 1 bar. The equilibration process was continued for 5 ns under NPT conditions, after which production runs were started using the same conditions.

Structural analysis of MD trajectories

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All analyses were performed by using CPPTRAJ (Roe and Cheatham, 2013). The distance between the centers of mass (COM) of residues 25–38 Cα atoms of the chains in the dimer structure was evaluated (Figure 7—figure supplement 1); this residue range corresponds to the solvent-accessible half of helix TM-JM (Figures 7a and 8). For the monomer structures, the angle with respect to the membrane normal was assessed. For this, the angle between the membrane normal and the vector between the COM of residues 21–25 and residues 35–38 was calculated (Figure 7b).

PMF and free energy calculations of dimer dissociation

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For calculating a configurational free energy profile (PMF) of the process of dimer dissociation, 36 intermediate states were generated by separating one chain of the dimer along the membrane plane by 1 Å steps, resulting in a minimum and maximum distance between the chain COM of 40.8 and 68 Å, respectively. The generated structures represent the separation process of the PlaF dimer. To sample configurations along the chain separation in a membrane environment, each intermediate state was embedded into a membrane of approximately 157×157 Å2 by using PACKMOL-Memgen (Schott-Verdugo and Gohlke, 2019), and independent MD simulations of 300 ns length each, with a total simulation time of 10.8 µs. Umbrella sampling simulations were performed by restraining the initial distance between chains in every window with a harmonic potential, using a force constant of 4 kcal mol–1 Å–2 (Torrie and Valleau, 1977); the distance between the COM of Cα atoms of residues 25–38 of each monomer was used as a reaction coordinate, being restrained in every simulation. Values for the reaction coordinate, representing the intermonomer distance r, were recorded every 2 ps and post-processed with the Weighted Histogram Analysis Method implementation of A. Grossfield (WHAM 2.0.9) (Suzuki, 1975; Grossfield, 2016), removing the first 100 ns as an equilibration of the system. The kernel densities showed a median overlap of 8.2% between contiguous windows (Figure 7—figure supplement 1), well suited for PMF calculations (Chen and Kuyucak, 2011). The error was estimated by separating the last 200 ns of data in four independent parts of 50 ns each and then calculating the standard error of the mean of the independently determined energy profiles.

The association free energy was estimated from the obtained PMF following the membrane two-body derivation from Johnston et al., 2012 and our previous work (Pagani and Gohlke, 2018). The PMF of dimer association is integrated along the reaction coordinate to calculate an association constant (Ka), which is transformed to the mole fraction scale (Kx) taking into account the number of lipids NL per surface area A, and this value is used to calculate the difference in free energy between dimer and monomers (ΔG), according to Equations 1–3:

(1) Ka=||Ω||(2π)20Drew(r)kBTdr
(2) Kx=KaNLA
(3) ΔG=RT ln(Kx)

where r is the value of the reaction coordinate, w(r) is the PMF at value r, D is the maximum distance at which the protein is still considered a dimer, kB is the Boltzmann constant, and T is the temperature at which the simulations were performed. The factor Ω(2π)2 considers the restriction of the configurational space of the monomers upon dimer formation in terms of the sampled angle between the two chains in the dimeric state (Equation 4) and the accessible space for the monomers, (2π)2.

(4) Ω=maxθa-minθa*maxθb-minθb

In Equation 4, the angle θa is defined as the angle formed between the vectors connecting the COM of chain b with the COM of the chain a and with the COM of residues 25–38 of the latter chain; θb is defined analogously starting from the COM of chain a. A value for ||Ω|| of 0.55 computed from Equation 4 indicates the fraction of the accessible space that the PlaF monomers have in the dimeric state compared to when both chains rotate independently [(2π)2].

PMF and free energy calculations of monomer tilting

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The initial conformations used in every window for calculating the PMF of the monomer tilting were obtained from the first microsecond of MD simulations of replica 10 of PlaFA (oriented as in the di-PlaF crystal structure) where spontaneous tilting occurred. The distance d along the z-axis between the COM of Cα atoms of residues 33–37 of the monomer with the membrane center was used to select 22 intermediate tilting configurations. d significantly correlates (R2=0.997, p<0.001) with the angle formed by the second half of helix αJM1 of the monomer (residues 25–38) and the normal vector of the membrane (Figure 7—figure supplement 1). The starting conformations were extracted from the representative trajectory, taking the respective snapshots where d and the angle showed the least absolute deviation to the average value obtained by binning d in windows of 2 Å width and with an evenly distributed separation of 1 Å. The distance d was restrained for every configuration by a harmonic potential with a force constant of 4 kcal mol–1 Å–2, and sampling was performed for 300 ns per window. The data were obtained every 2 ps and analyzed as described above, resulting in 8.6% of median overlap between kernel densities of contiguous windows (Figure 7—figure supplement 1). The error was estimated in the same way as for the dimerization (see above).

For calculating the free energy difference between the obtained basins, the PMF of monomer tilting was integrated using Equations 5 and 6 (Doudou et al., 2009):

(5) Ktilting=B1e w(d)kBTdrB2e w(d)kBTdr
(6) ΔGtilting=RT ln Ktilting

where d is defined as above, w(d) is the value of the PMF at that distance, and B1 and B2 represent the basins for the tilted and split configurations, respectively. The integration limits B1 and B2 included each basin portion below half of the value between the basin minimum and the energy barrier separating the basins, respectively (Figure 7c, yellow shaded regions).

PlaF dimer versus monomer proportion under in vivo conditions

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The dimer to monomer equilibrium of PlaF in the membrane results from the coupling of the following equilibria:

(7) 2M KaD Ka=DM2
(8) MKtiltingMtilted Ktilting=MtiltedM

yielding,

(9) D Ka Ktilting22Mtilted

where D, M, and Mtilted represent the PlaF dimer, ‘split’ monomer, and tilted monomer, respectively, with Ka and Ktilting being the dimer association and monomer tilting equilibrium constants, obtained from the PMF calculations. Based on the association constant computed according to Equation 7, Ka=[D]/[M]2=1.57×107 Å2, with [D] and [M] as area concentrations of dimer and monomer, respectively, the proportion of PlaF dimer versus monomer in a live cell of P. aeruginosa can be computed. Experimentally, 40 µg GPLs per 1 ml P. aeruginosa p-plaF (OD580nm 1) were extracted, and a PlaF purification yield of ca. 1 µg from 1 ml P. aeruginosa p-plaF culture with OD580nm was obtained (Bleffert et al., 2019; Supplementary file 3). Considering the molecular weight of PlaF of 35.5 kDa and assuming 750 Da as the average molecular weight of membrane GPL, this relates to a concentration under overexpressing conditions of ~5.28×10–4 PlaF monomers per lipid. Under non-overexpressing conditions, the concentration of PlaF monomers is estimated to be at least 100- to 1000-fold lower, that is, 5.28×10–6 to 5.28 × 10–7 PlaF monomers per lipid. Considering that the area per lipid in a PE:PG=3:1 membrane at 300K is approximately 61 Å2 per leaflet (or 30.5 Å2 in a bilayer, computed in this work and Murzyn et al., 2005), the total area concentration of PlaF molecules then is

(10) T = 2D+ M=1.73 x 10-8, 1.73 x 10-7PlaFÅ2.

Expressing the association constant in terms of the monomer concentration using Equation 7 yields

(11) Ka=T-M2M2 2KaM2+M-T=0,

and solving the quadratic equation then results in

(12) M=-1+1+8KaT4Ka=1.25×10-8,6.00×10-8PlaFÅ2

and

(13) D=T-M2=2.43×10-9,5.66×10-8PlaF dimerÅ2,

These results show that in live cells, the fraction of PlaF in the monomeric (dimeric) state is between 35% and 72% (65% and 28%), where the PlaF monomer is considered to be in the ‘split’ configuration with respect to the membrane normal.

As the tilting of the PlaF monomer is energetically favorable compared to the ‘split’ configuration and, hence, depletes the concentration of ‘split’ PlaF monomers, the dimeric PlaF concentration will decrease (Figure 7a). To quantitatively consider the effect of the tilting, we express the overall equilibrium constant for the processes shown in Figures 7a and 8, and described in Equations 7–9 as

(14) K=Ka Ktilting-2=DMtilted2,

where

Ktilting=MtiltedM=3.35, equivalent to Gtilting = -0.72kcalmol, computed according to Equation 5.

Following the same procedure as before then yields

Mtilted=1.66×10-8,1.28×10-7PlaFÅ2
D=3.83×10-10,2.28×10-8PlaF dimerÅ2

showing that in live cells, the fraction of PlaF in the tilted monomeric (dimeric) state is between 74% and 96% (26 and 4%). A graphical representation of the percentage of protein as a tilted monomer with respect to the protein concentration in the membrane is shown in Figure 7e.

Data availability

Diffraction data have been deposited in PDB under the accession code 6I8W. All data generated or analysed during this study are included in the manuscript and supporting file. Sequencing data are embedded in Fig. S1b. Source Data file "Table S1 - lipidome" has been provided for Figure 2. It contains the numerical data used to generate the figure 2c. Source data used to calculate the potentials of mean force and their corresponding simulation trajectory files shown in Figure 7 and Figure 7—figure supplement 1 are accessible at the DSpace instance researchdata.hhu.de under DOI:http://doi.org/10.25838/d5p-31.

The following data sets were generated
    1. Granzin J
    2. Batra-Safferling R
    (2019) RCSB Protein Data Bank
    ID 6I8W. Crystal structure of a membrane phospholipase A, a novel bacterial virulence factor.
    1. Schott-Verdugo S
    2. Gohlke H
    3. Batra-Safferling R
    4. Jaeger KE
    5. Kovacic F
    (2022) HHU ResearchData
    Structural and mechanistic insights into bacterial phospholipase A involved in membrane phospholipid degradation and virulence.
    https://doi.org/10.25838/d5p-31

References

  1. Book
    1. Case DA
    (2017)
    AMBER Reference Manual
    University of California.
  2. Book
    1. Cossins AR
    (1994)
    Temperature Adaptation of Biological Membranes. Proceedings of the Meeting Held in Cambridge under the Auspices of the Society for Experimental Biology in Conjunction with Its US/Canadian Counterparts
    Portland Press.
    1. Darden T
    2. York D
    3. Pedersen L
    (1993)
    Particle Mesh Ewald - an N. Log(N) method for Ewald sums in large systems
    J Chem Physics 98:10089–10092.
    1. Emsley P
    2. Cowtan K
    (2004) Coot: model-building tools for molecular graphics
    Acta Crystallographica. Section D, Biological Crystallography 60:2126–2132.
    https://doi.org/10.1107/S0907444904019158
    1. Jaeger KE
    2. Kovacic F
    (2014)
    Determination of lipolytic enzyme activities
    Methods in Molecular Biology 1149:111–134.
    1. Kenakin TP
    (2012)
    Enzymes as Drug Targets
    Pharmacology in Drug Discovery 10:105–124.
    1. Koch G
    (2014)
    Assessing Pseudomonas virulence with nonmammalian host: Galleria mellonella
    Methods in Molecular Biology 1149:681–688.
    1. Lands WE
    (1958)
    Metabolism of glycerolipides; a comparison of lecithin and triglyceride synthesis
    The Journal of Biological Chemistry 231:883–888.
    1. Suzuki Y
    (1975)
    Inborn error of glycolipid metabolism
    Horumon to Rinsho. Clinical Endocrinology 23:885–891.

Decision letter

  1. Arun Radhakrishnan
    Reviewing Editor; University of Texas Southwestern Medical Center, United States
  2. Olga Boudker
    Senior Editor; Weill Cornell Medicine, United States
  3. Lei Zheng
    Reviewer; University of Texas Health Science Center at Houston, McGovern Medical School, United States

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Decision letter after peer review:

Thank you for submitting your article "Evidence for a bacterial Lands cycle phospholipase A involved in membrane phospholipid remodeling" for consideration by eLife. Your article has been reviewed by 2 peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Olga Boudker as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: Lei Zheng (Reviewer #1).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

This study provides new insights into the mechanisms and regulation of phospholipid remodeling in bacteria by a phospholipase PlaF. The reviewers were enthusiastic about the several new findings in this study and have provided several suggestions to address in a revised submission. The major concern, which is not necessarily a negative, is that the manuscript includes a massive amount of data and the connection is at times hard to follow. The strengths of the manuscript are the comprehensive biochemical and structural analysis of how PlaF functions, whereas some of the other data are in comparison, more preliminary in nature. The authors should consider focusing the manuscript (including the title) on these strengths.

Please see specific points to address in the reviews from both reviewers below.

Other Specific Comments from Reviewer #1:

You have included a massive amount of data in the manuscript. The biochemical and structural data are really nice! Other data are interesting, but preliminary, which may dilute the novelty of your work. I would suggest focusing on the molecular mechanism of PlaF, which may help sharpen the paper.

Other Specific Comments from Reviewer #3:

In addition to add experiments as above, it is important to rearrange the manuscript for more clear and smooth flow. It is also advisable to describe more accurately what is known on Lands cycle in mammals.

Reviewer #1 (Recommendations for the authors):

In this manuscript, Kovacic et al., report structure and functional studies of PlaF from Pseudomonas aeruginosa. They found that PlaF functions as a novel phospholipase A in the bacterial inner membrane with its catalytic domain facing towards the periplasm. They determined the X-ray structure of PlaF to 2A resolution. The structure shows a dimeric conformation with free fatty acid (FFA) molecules bound in the substrate-binding pocket. Based on the structure, they performed very nice studies using biochemical assays and MD simulation to propose a regulatory mechanism of PlaF via transmembrane helix-mediated dimerization. In general, this is a very interesting and comprehensive study. The works provide new insights into the GPL homeostasis on the bacterial cell membrane. The conclusion is supported by most of the data, except some issues as follow:

1. The structure looks well-resolved based on the statistics. However, the electron density of FFA in Chain B is very weak, barely seen at 1 σ. Does FFA have different occupancy between Chain A and Chain B? Does this variation contribute to the conformational changes between the two monomers?

2. The hypothesis of GPL remodeling in Figure 2d is too speculative and is not supported by their data in Figure 2c. Is it possible the changes of GPLs are caused by the depletion of PlaF on the membrane?

3. A similar concern regarding Figure 3. The phenotype of the plaF KO strain may not establish the PlaF protein as an antibacterial target. Is the susceptibility caused by the loss of PlaF catalytic activity or simply due to the depletion of plaF? Authors need to be cautious when discussing potential drug binding sites based on the structure in the discussion.

4. It is unclear how FFA binding facilitates PlaF dimerization. It would be important to discuss it based on the structure and biochemical data.

5. It is unclear any ligands have been included in the MD simulation. FFA or OG?

6. The title is vague and a better one may help highlight the major findings of this work.Reviewer #3 (Recommendations for the authors):

The work consists of three parts, 1)phospholipase activity, possibly involved in remodeling pathway of bacterial GPL, 2) structural insight of fatty acid-induced dimeric form with weak catalytic activity, and 3) virulent nature of Pseudomonas against insect and bone marrow-derived macrophages. All are new findings, but the relation of three topics is not wrriten clear enough, and each point needs further detailed analyses as follows.

1. For the phospholipase activity, authors have better determine enzyme activities for individual GPL with different fatty acid species using LC-MS with appropriate internal standards. It has to be shown more clearly whether only sn-1 is cleaved, but not sn-2, and whether PLaF does not have transacylase activity. It is also critical to make several mutants of active site, and measure both carboxyesterase and PLA activities in parallel.

2. It is not clear whether dimer formation is just an artifact by over-expression. How PLaFa and PlaFb are formed? What is biological significance of fatty acid effect at these high concentrations? Is it not detergent effects with certain HLB?

3. The mechanism of virulent effect has to be more clearly determined. Does it relate to phospholipase, or caroboxyesterase, or both? Does PlaF induce membrane lysis of macrophages or insect cells by producting lysoGPL?. More mechanical insights are necessary.

4. Discussion is necessary how the remodeling enzyme plays also virulent roles.

https://doi.org/10.7554/eLife.72824.sa1

Author response

This study provides new insights into the mechanisms and regulation of phospholipid remodeling in bacteria by a phospholipase PlaF. The reviewers were enthusiastic about the several new findings in this study and have provided several suggestions to address in a revised submission. The major concern, which is not necessarily a negative, is that the manuscript includes a massive amount of data and the connection is at times hard to follow. The strengths of the manuscript are the comprehensive biochemical and structural analysis of how PlaF functions, whereas some of the other data are in comparison, more preliminary in nature. The authors should consider focusing the manuscript (including the title) on these strengths.

Please see specific points to address in the reviews from both reviewers below.

Other Specific Comments from Reviewer #1:

You have included a massive amount of data in the manuscript. The biochemical and structural data are really nice! Other data are interesting, but preliminary, which may dilute the novelty of your work. I would suggest focusing on the molecular mechanism of PlaF, which may help sharpen the paper.

Other Specific Comments from Reviewer #3:

In addition to add experiments as above, it is important to rearrange the manuscript for more clear and smooth flow. It is also advisable to describe more accurately what is known on Lands cycle in mammals.

We have thoroughly revised the manuscript as suggested by the reviewers. In particular, the title was modified, the introduction extended, the discussion shortened and focused on the main results. Additional information on the Lands cycle in eukaryotes is now provided in the introduction. Please note that the first structure of a lysophospholipid acyltransferase (LPLAT, Zhang, Q., et al., The structural basis for the phospholipid remodeling by lysophosphatidylcholine acyltransferase 3. Nat Commun, 2021. 12(1): p. 6869.) and classification of LPLATs (Valentine, W.J., et al., Update and nomenclature proposal for mammalian lysophospholipid acyltransferases, which create membrane phospholipid diversity. Journal of Biological Chemistry, 2022. 298(1): p. 101470.) were published during the revision of this manuscript. Furthermore, despite many known mammalian PLAs, the relationship of these enzymes and Lands cycle is still not well understood.

Reviewer #1 (Recommendations for the authors):

In this manuscript, Kovacic et al., report structure and functional studies of PlaF from Pseudomonas aeruginosa. They found that PlaF functions as a novel phospholipase A in the bacterial inner membrane with its catalytic domain facing towards the periplasm. They determined the X-ray structure of PlaF to 2A resolution. The structure shows a dimeric conformation with free fatty acid (FFA) molecules bound in the substrate-binding pocket. Based on the structure, they performed very nice studies using biochemical assays and MD simulation to propose a regulatory mechanism of PlaF via transmembrane helix-mediated dimerization. In general, this is a very interesting and comprehensive study. The works provide new insights into the GPL homeostasis on the bacterial cell membrane. The conclusion is supported by most of the data, except some issues as follow:

1. The structure looks well-resolved based on the statistics. However, the electron density of FFA in Chain B is very weak, barely seen at 1 σ. Does FFA have different occupancy between Chain A and Chain B? Does this variation contribute to the conformational changes between the two monomers?

Thank you for pointing this out. The electron density of FFA in chain B is indeed less well-defined than that of FFA in chain A. However, we believe that the confusion here is due to the calculated electron density maps provided by the PDB, which look very weak around the region of the FFA in molecule B compared to the electron density maps we obtained directly from the Phenix refinement program, which we have used for the interpretation. The main reason for the discrepancies between our maps and those from the PDB is that different programs are used for bulk-solvent correction and overall scaling during refinement. We will gladly provide the MTZ file with the original phases at the request of the reviewer.

No occupancy was refined for the FFA molecules, but the overall B-factors show significant differences in comparison. For the UNDECANOIC ACID of chain B, the average B factor is 89.03 Å2 (average B factor of the surrounding protein chain is 49.29 Å2) in contrast to the MYRISTIC ACID of chain A which has an average B factor of 66.63 Å2 (average B factor of the surrounding protein chain: 47.61 Å2). The average B factor thus shows that the electron density map of FFA(B) is considerably poorly defined, i.e., the FFA has a high flexibility or lower occupancy, as pointed out by the reviewer. We cannot say with certainty or rule out the possibility that these variations are responsible for the differences between the two protein molecules since the overall structure is also influenced in part by crystal packing effects.

We have added this information in the results on page 7 (lanes 10-12).

2. The hypothesis of GPL remodeling in Figure 2d is too speculative and is not supported by their data in Figure 2c. Is it possible the changes of GPLs are caused by the depletion of PlaF on the membrane?

We have now modified figure 2 by removing the speculative part about the putative Lands cycle.

If we keep in mind that PlaF is present in P. aeruginosa cells at very low amounts (below the detection limit of a Western blot, see Figure 7-supplementary figure 2), it is unlikely that such considerably large changes of GPL content are due to the absence of PlaF. However, it cannot be excluded that the absence of PlaF in the cytoplasmic membrane of P. aeruginosa ΔplaF is responsible for the observed GPL changes. We have now commented on that in the discussion (page 12, lanes 28-30).

Our model predicts that PlaF which is anchored to the cytoplasmic membrane modifies this membrane by hydrolysing GPLs. This appears plausible and in agreement with a) in vitro catalytic activity of PlaF against a range of cytoplasmic GPLs (Figure 2), and b) decreased PLA activity of the ΔplaF strain compared to the wild type (Figurre 2-supplementary figure 2).

3. A similar concern regarding Figure 3. The phenotype of the plaF KO strain may not establish the PlaF protein as an antibacterial target. Is the susceptibility caused by the loss of PlaF catalytic activity or simply due to the depletion of plaF? Authors need to be cautious when discussing potential drug binding sites based on the structure in the discussion.

We have now removed a large part of the discussion about ligand binding sites of PlaF as potential drug binding sites.

We are aware that further experimental evidence is needed to establish PlaF as an antibacterial target and these experiments are ongoing. Our preliminary results point to the importance of PlaF as a virulence regulator in P. aeruginosa. We observed that PlaF strongly affects virulence-related phenotypes, namely biofilm production and architecture (Author response image 1) and swimming motility (Author response image 2). Additionally, we observed an extended lag-phase during the growth of P. aeruginosa ΔplaF compared with the wild-type, which may be explained by impaired iron transport in ΔplaF (Author response image 3).

Author response image 1
PlaF is a novel P. aeruginosa biofilm effector.

(A) Biofilm amount of P. aeruginosa ΔplaF and the wild-type strain cultivated in 96-well MTP (LB medium, 37°C, without aeration) was quantified by staining the cells attached to MTP with crystal violet. The results are mean ± standard deviation of five biological replicates, each measured eight times. Statistical analysis was performed using the t-test, * p < 0.05. (B) Biofilm architecture was analysed by CLSM after 24, 72, and 144 h growth at 37ºC in a flow cell with a continuous supply of LB medium. Shown are representative figures of two biological replicates analysed by imaging three different sections (100 x 100 µm) of the cover glass.

Author response image 2
P. aeruginosa ΔplaF mutant strain is impaired in swimming motility.

The growth of the strains on LB agar swimming plates was quantified using ImageJ and expressed as swimming area, *** p < 0.00005, n = 10.

Author response image 3
Iron-acquisition is disbalanced in P. aeruginosaplaF.

A) Strains were grown statically at 37°C in 96-well microtiter plates for biofilm formation or in Erlenmeyer flasks under planktonic conditions for 24 h. Pyoverdine concentration was quantified in cell-free supernatant of P. aeruginosa PA01 and ∆plaF grown in plastic 96-well microtiter plates (biofilm) or in Erlenmeyer flasks (planktonic) at 37°C for 24 h. Results are mean ± S.D. B) P. aeruginosa ΔplaF shows a lower intrecellular iron concentration than the wild type. Iron was quantified [Tamarit, J., et al., Anal Biochem, 2006] in cells isolated from the cultures grown 18 h in LB medium at 37 C. C) The growth curves of P. aeruginosa PA01 and ∆plaF (n = 3) in M9 minimal medium under iron-limiting conditions and in the presence of 100 µM FeSO4 were determined by measuring the optical density at 580 nm. Results are mean ± S.D. The growth of P. aeruginosaplaF and PA01, as well as P. aeruginosaplaF and ∆plaF + FeSO4, differ significantly (p < 0.05).

To understand how PlaF modulates the virulence of P. aeruginosa on a molecular level, we have performed a global proteomic analysis of wild-type and ΔplaF strains. The results revealed a pleiotropic effect of PlaF deletion on many virulence-related pathways, among them attachment, signalling, iron homeostasis, chemotaxis (Figures 4 – 6). We suggest that changes in GPL composition modulate the function of cytoplasmic membrane proteins, which in turn triggers a cellular response. These still unpublished results are part of a manuscript in preparation for submission and can be provided to the reviewers upon request.

We decided not to include proteomics results in the eLife manuscript as the manuscript already contains many data as outlined by the reviewers.

4. It is unclear how FFA binding facilitates PlaF dimerization. It would be important to discuss it based on the structure and biochemical data.

We agree that the here presented dimeric structure of PlaF cannot provide a clear answer on how FFA binding facilitates PlaF dimerisation. FFA inhibition kinetics indicate that additionally to competitive inhibition (blocking of the active site by bound fatty acid), there is a non-competitive inhibition component which might implicate that FFAs trigger the formation of the inactive dimer.

Our observation that the dimer interface and the catalytic site are connected through bound FFA and a pseudo-ligand (octyl glucoside) provides a structural hint about the link of dimerisation, activity and FFA binding.

This was now briefly discussed on page 14, lanes 30-32.

5. It is unclear any ligands have been included in the MD simulation. FFA or OG?

We did not include any fatty acid or detergent in our simulations. They are shown in the figures to visualise the active site of PlaF and a putative path for substrate access. This information was now added in the caption of figure 7 and the M and M part.

6. The title is vague and a better one may help highlight the major findings of this work.

We now changed the title to highlight the structural and mechanistic results.

Reviewer #3 (Recommendations for the authors):

The work consists of three parts, 1)phospholipase activity, possibly involved in remodeling pathway of bacterial GPL, 2) structural insight of fatty acid-induced dimeric form with weak catalytic activity, and 3) virulent nature of Pseudomonas against insect and bone marrow-derived macrophages. All are new findings, but the relation of three topics is not wrriten clear enough, and each point needs further detailed analyses as follows.

1. For the phospholipase activity, authors have better determine enzyme activities for individual GPL with different fatty acid species using LC-MS with appropriate internal standards. It has to be shown more clearly whether only sn-1 is cleaved, but not sn-2, and whether PLaF does not have transacylase activity. It is also critical to make several mutants of active site, and measure both carboxyesterase and PLA activities in parallel.

Thank you for your suggestion. We have now performed an experiment in which 1-(9Z-octadecenoyl)-2-pentadecanoyl-glycero-3-phospho-(1'-rac-glycerol) (PG_15:0_18:1) was hydrolysed by PlaF following quantification of fatty acids by GC-MS. The experiment was challenging because of a large amount of purified PlaF needed.

In two independent experiments, 0.4 µM pentanoic acid was detected after hydrolysis by PlaF and FA extraction. The concentration of oleic acid (18:1) in PlaF-treated samples was not higher than in the blank samples. Oleic acid is bound at sn2 position in the lipid and pentanoic acid at sn1 position; thus, we can conclude that PlaF is a PLA1 as shown previously (see Figure 2a). We have now added these results as Figure 2- supplementary figure 1 and comment on them in the main text (page 5, lanes 7-9).

The colorimetric NEFA assay which we have used to measure PLA activity is a standard assay and was applied to characterize other PLAs [1, 2]. The advantage of the NEFA assay is its sensitivity and, in comparison to a GC-MS-based assay, the NEFA assay does not involve an error-prone fatty acid extraction step.

We have previously shown that the variant PlaF-S137A with the catalytic serine replaced by a non-functional alanine does not possess esterase activity [3]. Furthermore, a cell lysate from P. aeruginosa ΔplaF showed 40% lower esterase activity than that of the wild type strain (Author response image 4). As PlaF is a one-domain protein and has a catalytic triad conserved among many esterases and PLAs, we propose that identical residues are involved in PLA and esterase activities.

Author response image 4
Esterase activity of P. aeruginosa ΔplaF strain.

P. aeruginosa PAO1 and ΔplaF strains were grown overnight in LB-medium at 37°C. Cells were harvested by centrifugation, suspended in Tris‑HCl buffer (100 mM, pH 8) to equal cell count, and enzyme activities were determined with p-nitrophenyl butyrate as the substrate. The results are mean ± standard deviations of three biological replicates, each measured three times.

During the revision of this manuscript, we have published an article describing the putative mechanism of GPL hydrolysis by PlaF [4]. There, we report the results of a mutational analysis of the active site of PlaF, which lead us to suggest how GPL enters the active site pocket of PlaF and how the products are likely released to prepare PlaF for a new catalytic cycle.

At present, we do not have any indications as to an acyltransferase activity of PlaF. We regard the determination of this enzymatic activity and, if detected, of its physiological consequences as a subject of a follow-up study.

2. It is not clear whether dimer formation is just an artifact by over-expression. How PLaFa and PlaFb are formed? What is biological significance of fatty acid effect at these high concentrations? Is it not detergent effects with certain HLB?

We agree with the possibility that overexpression leads to dimer formation. Unfortunately, under laboratory conditions, PlaF is produced by P. aeruginosa wild type at low concentrations (Figure 7-supplementary figure 2); therefore, we have used mild overexpression (using the low copy number plasmid pBBR1mcs-3) to capture PlaF dimers in the cell. Our results suggest equilibrium between dimeric and monomericPlaF in the cell membrane. in vitro results indicate that this equilibrium is shifted towards dimer formation by raising the concentrations of either PlaF or fatty acids. Although we do not entirely understand the structural prerequisites for dimerisation, we know that PlaF forms dimers and dimerisation regulates its activity.

An in vitro effect of fatty acids on PlaF activity and dimerisation fits a product feedback mechanism which would make sense for regulation of PlaF, as it is potentially toxic to P. aeruginosa. Although precise data about the intracellular concentration of FFA in P. aeruginosa are not available, Fang L. and co-authors [5] recently published that the intracellular concentration of FFAs in E. coli is 0.5 – 1 mg/l (2 – 4 mM for palmitic acid). This is in the range of the concentrations used in our experiments.

The structure of di-PlaF in which a) hydrophobic residues of TM helices form intermolecular interactions, b) polar residues of juxtamembrane domains of two PlaF molecules interacts with each other, and c) charged residues are preceding the TM helix agrees with the topology suggested for several TM helix dimers (see figure 1 in reference [6]). Therefore, we rationalise that the PlaF dimer detected in the described crystal structure represents a physiological state.

The effects of detergents or membrane solubilising compounds on the structure of membrane proteins is the subject of a long-standing debate. Octyl glucoside (OG) which is among the most commonly used detergents in structural biology [7], was chosen here for purification and crystallisation because of its stabilising impact on PlaF. Notably, PlaF remained in the active and native form during several weeks of crystallisation in the presence of OG detergent. We did not screen other detergents with different hydrophilic-lipophilic balance values and thus cannot provide a statement about the correlation of HLB and PlaF activity, stability and dimerisation. However, in vivo crosslinking experiments showed that PlaF dimers can be formed in the absence of OG (Figure 5a).

3. The mechanism of virulent effect has to be more clearly determined. Does it relate to phospholipase, or caroboxyesterase, or both? Does PlaF induce membrane lysis of macrophages or insect cells by producting lysoGPL?. More mechanical insights are necessary.

4. Discussion is necessary how the remodeling enzyme plays also virulent roles.

We have shown that PlaF is associated with the cytoplasmic membrane of P. aeruginosa cells; hence, it cannot come into direct contact with the host cells. Therefore, we suggest that PlaF-mediated virulence is not related to the simple lysis of host cells during infections caused by P. aeruginosa.

Although PlaF shows PLA and esterase activities, its cellular localisation and lipidomics results suggest that GPLs are native substrates of PlaF. In our view, PlaF regulates the virulence of P. aeruginosa by modulating the phospholipid composition of the cytoplasmic membrane with which it is associated. Although we do not entirely understand how these changes of GPL content affect the physiology of P. aeruginosa, our global proteomic analysis of wild-type and ΔplaF strains revealed that cells respond to the absence of PlaF by changing the production rates of many virulence-related proteins. This notion is in accordance with the widely accepted assumption that the function and structure of membrane proteins (e.g. two-component sensors, transporters) are regulated through protein-lipid interactions. Our preliminary proteomics results agree with these suggestions (for more details, see the answer to the third question of reviewer 1).

We have now discussed the possible role of PlaF-mediated remodelling of GPL on the regulation of virulence proteins in P. aeruginosa (page 13, lanes 1-3).

References

1. Kuhle, K., et al., Oligomerization inhibits Legionella pneumophila PlaB phospholipase A activity. J Biol Chem, 2014. 289(27): p. 18657-66.

2. Schunder, E., et al., Phospholipase PlaB is a new virulence factor of Legionella pneumophila. Int J Med Microbiol, 2010. 300(5): p. 313-23.

3. Kovacic, F., et al., A membrane-bound esterase PA2949 from Pseudomonas aeruginosa is expressed and purified from Escherichia coli. FEBS Open Bio, 2016. 6(5): p. 484-93.

4. Ahmad, S., et al., Substrate Access Mechanism in a Novel Membrane-Bound Phospholipase A of Pseudomonas aeruginosa Concordant with Specificity and Regioselectivity. J Chem Inf Model, 2021. 61(11): p. 5626-5643.

5. Fang, L., et al., Genome-scale target identification in Escherichia coli for high-titer production of free fatty acids. Nat Commun, 2021. 12(1): p. 4976.

6. Bocharov, E.V., et al., Helix-helix interactions in membrane domains of bitopic proteins: Specificity and role of lipid environment. Biochim Biophys Acta Biomembr, 2017. 1859(4): p. 561-576.

7. Moraes, I., et al., Membrane protein structure determination – the next generation. Biochim Biophys Acta, 2014. 1838(1 Pt A): p. 78-87.

https://doi.org/10.7554/eLife.72824.sa2

Article and author information

Author details

  1. Florian Bleffert

    Institute of Molecular Enzyme Technology, Heinrich Heine University Düsseldorf, Forschungszentrum Jülich GmbH, Jülich, Germany
    Contribution
    Formal analysis, Investigation, Validation, Writing – review and editing
    Competing interests
    No competing interests declared
  2. Joachim Granzin

    Institute of Biological Information Processing - Structural Biochemistry (IBI-7: Structural Biochemistry), Forschungszentrum Jülich GmbH, Jülich, Germany
    Contribution
    Data curation, Formal analysis, Investigation, Validation, Writing – review and editing
    Competing interests
    No competing interests declared
  3. Muttalip Caliskan

    Institute of Molecular Enzyme Technology, Heinrich Heine University Düsseldorf, Forschungszentrum Jülich GmbH, Jülich, Germany
    Contribution
    Formal analysis, Investigation, Writing – review and editing
    Competing interests
    No competing interests declared
  4. Stephan N Schott-Verdugo

    1. Institute for Pharmaceutical and Medicinal Chemistry, Heinrich Heine University Düsseldorf, Duesseldorf, Germany
    2. Centro de Bioinformática y Simulación Molecular (CBSM), Faculty of Engineering, University of Talca, Talca, Chile
    3. John von Neumann Institute for Computing (NIC), Jülich Supercomputing Centre (JSC), and Institute of Bio- and Geosciences (IBG-4: Bioinformatics), Forschungszentrum Jülich GmbH, Jülich, Germany
    Contribution
    Formal analysis, Investigation, Validation, Writing – review and editing
    Competing interests
    No competing interests declared
  5. Meike Siebers

    1. Institute of Molecular Physiology, and Biotechnology of Plants (IMBIO), University of Bonn, Bonn, Germany
    2. Institute for Plant Genetics, Heinrich Heine University Düsseldorf, Düsseldorf, Germany
    Contribution
    Formal analysis, Investigation, Validation, Writing – review and editing
    Competing interests
    No competing interests declared
  6. Björn Thiele

    Institute of Bio- and Geosciences, Plant Sciences (IBG-2), and Agrosphere (IBG-3), Forschungszentrum Jülich GmbH, Jülich, Germany
    Contribution
    Formal analysis, Investigation, Validation, Writing – review and editing
    Competing interests
    No competing interests declared
  7. Laurence Rahme

    Department of Microbiology, and Immunobiology, Harvard Medical School, Boston, United States
    Contribution
    Supervision, Validation, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5374-4332
  8. Sebastian Felgner

    Department of Molecular Bacteriology, Helmholtz Centre for Infection Research, Braunschweig, Germany
    Contribution
    Formal analysis, Investigation, Validation, Visualization, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0030-2490
  9. Peter Dörmann

    Institute of Molecular Physiology, and Biotechnology of Plants (IMBIO), University of Bonn, Bonn, Germany
    Contribution
    Supervision, Validation, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5845-9370
  10. Holger Gohlke

    1. Institute of Biological Information Processing - Structural Biochemistry (IBI-7: Structural Biochemistry), Forschungszentrum Jülich GmbH, Jülich, Germany
    2. Institute for Pharmaceutical and Medicinal Chemistry, Heinrich Heine University Düsseldorf, Duesseldorf, Germany
    3. John von Neumann Institute for Computing (NIC), Jülich Supercomputing Centre (JSC), and Institute of Bio- and Geosciences (IBG-4: Bioinformatics), Forschungszentrum Jülich GmbH, Jülich, Germany
    Contribution
    Conceptualization, Funding acquisition, Supervision, Validation, Writing - original draft, Writing – review and editing
    For correspondence
    gohlke@uni-duesseldorf.de
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-8613-1447
  11. Renu Batra-Safferling

    Institute of Biological Information Processing - Structural Biochemistry (IBI-7: Structural Biochemistry), Forschungszentrum Jülich GmbH, Jülich, Germany
    Contribution
    Conceptualization, Formal analysis, Investigation, Validation, Visualization, Writing - original draft, Writing – review and editing
    For correspondence
    r.batra-safferling@fz-juelich.de
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-8597-4335
  12. Karl-Erich Jaeger

    1. Institute of Molecular Enzyme Technology, Heinrich Heine University Düsseldorf, Forschungszentrum Jülich GmbH, Jülich, Germany
    2. Institute of Bio- and Geosciences (IBG-1: Biotechnology), Forschungszentrum Jülich GmbH, Jülich, Germany
    Contribution
    Funding acquisition, Project administration, Supervision, Writing – review and editing
    Competing interests
    No competing interests declared
  13. Filip Kovacic

    Institute of Molecular Enzyme Technology, Heinrich Heine University Düsseldorf, Forschungszentrum Jülich GmbH, Jülich, Germany
    Contribution
    Conceptualization, Data curation, Formal analysis, Funding acquisition, Investigation, Methodology, Project administration, Resources, Supervision, Validation, Visualization, Writing - original draft, Writing – review and editing
    For correspondence
    f.kovacic@fz-juelich.de
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0313-427X

Funding

Deutsche Forschungsgemeinschaft (267205415)

  • Holger Gohlke
  • Karl-Erich Jaeger
  • Filip Kovacic

European Synchrotron Radiation Facility

  • Renu Batra-Safferling

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

This study was funded by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation), Project CRC 1208 (number 267205415) to FK and KEJ (subproject A02), and HG (subproject A03). The authors are grateful to the beamline scientists at the European Synchrotron Radiation Facility (Grenoble, France) for assisting with the use of beamline ID29. The authors thank R Voulhoux (CNRS AMU LCB, Marseille) for providing anti-SecG antiserum, P Dollinger (HHU Düsseldorf) for help with MST measurements, M Modri (HHU Düsseldorf) for help with PG_15:0_18:1 assay, and M Dick (HHU Düsseldorf) for assistance in setting up biased MD simulations. The authors are grateful for computational support by the ‘Zentrum für Informations und Medientechnologie’ at the Heinrich-Heine-Universität Düsseldorf and the computing time provided by the John von Neumann Institute for Computing (NIC) to HG on the supercomputer JUWELS at Jülich Supercomputing Centre (JSC) (user IDs: HKF7; VSK33; HDD18; plaf).

Senior Editor

  1. Olga Boudker, Weill Cornell Medicine, United States

Reviewing Editor

  1. Arun Radhakrishnan, University of Texas Southwestern Medical Center, United States

Reviewer

  1. Lei Zheng, University of Texas Health Science Center at Houston, McGovern Medical School, United States

Publication history

  1. Preprint posted: June 23, 2021 (view preprint)
  2. Received: August 5, 2021
  3. Accepted: May 10, 2022
  4. Accepted Manuscript published: May 10, 2022 (version 1)
  5. Version of Record published: May 25, 2022 (version 2)

Copyright

© 2022, Bleffert et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Florian Bleffert
  2. Joachim Granzin
  3. Muttalip Caliskan
  4. Stephan N Schott-Verdugo
  5. Meike Siebers
  6. Björn Thiele
  7. Laurence Rahme
  8. Sebastian Felgner
  9. Peter Dörmann
  10. Holger Gohlke
  11. Renu Batra-Safferling
  12. Karl-Erich Jaeger
  13. Filip Kovacic
(2022)
Structural, mechanistic, and physiological insights into phospholipase A-mediated membrane phospholipid degradation in Pseudomonas aeruginosa
eLife 11:e72824.
https://doi.org/10.7554/eLife.72824

Further reading

    1. Biochemistry and Chemical Biology
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    Lang Pan et al.
    Research Article

    Background:

    Few studies have assessed the role of individual plasma cholesterol levels in the association between egg consumption and the risk of cardiovascular diseases. This research aims to simultaneously explore the associations of self-reported egg consumption with plasma metabolic markers and these markers with the risk of cardiovascular disease (CVD).

    Methods:

    Totally 4778 participants (3401 CVD cases subdivided into subtypes and 1377 controls) aged 30–79 were selected based on the China Kadoorie Biobank. Targeted nuclear magnetic resonance was used to quantify 225 metabolites in baseline plasma samples. Linear regression was conducted to assess associations between self-reported egg consumption and metabolic markers, which were further compared with associations between metabolic markers and CVD risk.

    Results:

    Egg consumption was associated with 24 out of 225 markers, including positive associations for apolipoprotein A1, acetate, mean HDL diameter, and lipid profiles of very large and large HDL, and inverse associations for total cholesterol and cholesterol esters in small VLDL. Among these 24 markers, 14 were associated with CVD risk. In general, the associations of egg consumption with metabolic markers and of these markers with CVD risk showed opposite patterns.

    Conclusions:

    In the Chinese population, egg consumption is associated with several metabolic markers, which may partially explain the protective effect of moderate egg consumption on CVD.

    Funding:

    This work was supported by the National Natural Science Foundation of China (81973125, 81941018, 91846303, 91843302). The CKB baseline survey and the first re-survey were supported by a grant from the Kadoorie Charitable Foundation in Hong Kong. The long-term follow-up is supported by grants (2016YFC0900500, 2016YFC0900501, 2016YFC0900504, 2016YFC1303904) from the National Key R&D Program of China, National Natural Science Foundation of China (81390540, 81390541, 81390544), and Chinese Ministry of Science and Technology (2011BAI09B01). The funders had no role in the study design, data collection, data analysis and interpretation, writing of the report, or the decision to submit the article for publication.

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    Doublecortin (DCX) is a microtubule (MT)-associated protein that regulates MT structure and function during neuronal development and mutations in DCX lead to a spectrum of neurological disorders. The structural properties of MT-bound DCX that explain these disorders are incompletely determined. Here, we describe the molecular architecture of the DCX–MT complex through an integrative modeling approach that combines data from X-ray crystallography, cryo-electron microscopy, and a high-fidelity chemical crosslinking method. We demonstrate that DCX interacts with MTs through its N-terminal domain and induces a lattice-dependent self-association involving the C-terminal structured domain and its disordered tail, in a conformation that favors an open, domain-swapped state. The networked state can accommodate multiple different attachment points on the MT lattice, all of which orient the C-terminal tails away from the lattice. As numerous disease mutations cluster in the C-terminus, and regulatory phosphorylations cluster in its tail, our study shows that lattice-driven self-assembly is an important property of DCX.