ABSTRACT
In vast areas of the ocean, microbes must adapt to the availability of scarce nutrients, and a key strategy for reducing the cellular phosphorus (P) quota is to remodel membranes by replacing phospholipids with non-P surrogate lipids. A metallophosphoesterase, PlcP, is essential for lipid remodeling in cosmopolitan marine bacteria of the Roseobacter (e.g., Phaeobacter sp. strain MED193) and SAR11 (e.g., Pelagibacter sp. strain HTCC7211) clades, and transcription of plcP is known to be induced by P limitation. In order to better understand PlcP-mediated lipid remodeling, we sought to characterize PlcP for its metal ion requirement and to determine its selectivity for native bacterial phospholipids. Here, we report the occurrence of a highly conserved binuclear ion center in PlcPs from MED193 and HTCC7211 and show that manganese is the preferred metal for metallophosphoesterase activity. PlcP displayed high activity towards the major bacterial phospholipids, e.g., phosphatidylglycerol but also phosphatidic acid, a key intermediate in phospholipid biosynthesis. In contrast, phosphatidylserine and phosphatidylinositol, both of which are rare lipids in bacteria, are not preferred substrates. These data suggest that PlcP undertakes a generic lipid remodeling role during the cellular response of marine bacteria to P deficiency and that manganese availability may play a key role in regulating the lipid remodeling process.
IMPORTANCE Membrane lipids form the structural basis of all cells. In the marine environment, it is well established that phosphorus availability significantly affects lipid composition in cosmopolitan marine bacteria, whereby non-phosphorus-containing lipids are used to replace phospholipids in response to phosphorus stress. Central to this lipid remodeling pathway is a newly identified phospholipase C-type metallophosphoesterase (PlcP). However, little is known about how PlcP activity is regulated. Here, we determined the role of metal ions in regulating PlcP activity and compared PlcP substrate specificities in PlcP enzymes from two model marine bacteria from the marine Roseobacter clade and the SAR11 clade. Our data provide new insights into the regulation of lipid remodeling in these marine bacteria.
INTRODUCTION
Large expanses of the ocean, particularly surface waters, contain submicromolar concentrations of essential nutrients required for the growth of phytoplankton and heterotrophic bacteria (1, 2), including macronutrients (e.g., P) as well as micronutrients (e.g., iron, manganese). Marine bacteria inhabiting these oligotrophic surface waters have developed sophisticated strategies to meet cellular demands for these essential elements (3). For example, many marine microbes express high-affinity membrane transporters, e.g., the ABC transporter PstSCAB for phosphate or SitABCD for manganese uptake, in order to acquire specific nutrients present at low concentration (4, 5). Other marine microbes, such as the diazotrophic marine cyanobacterium Crocosphaera watsonii, can degrade iron-rich metalloproteins to release and recycle iron under limitation of this element (6).
Another important mechanism for adapting to nutrient deficiency is to reduce the cellular requirement for key elements (3). This strategy is now well established in marine phytoplankton and heterotrophic bacteria, whereby membrane phospholipids are replaced by non-P-containing surrogate lipids in response to P deficiency (7–9). In marine phytoplankton, replacement of phospholipids by the sulfur-containing glycolipid sulfoquinovosyl diacylglycerol (SQDG) significantly reduced the cellular quota for P (9). Our previous work has shown that lipid remodeling is also important in cosmopolitan marine heterotrophic bacteria (8). Members of the marine Roseobacter clade and SAR11 clade can replace phospholipids, primarily phosphatidylglycerol (PG) and phosphatidylethanolamine (PE), using non-P surrogate lipids, such as the betaine lipid diacylglyceryl trimethylhomoserine (DGTS), the glycolipids monoglycosyl diacylglycerol (MGDG) and glucuronic acid diacylglycerol (GADG), and ornithine lipids. Central to this lipid remodeling process in these marine heterotrophic bacteria is a phospholipase C-type phospholipase, designated PlcP, which was first described in the soil bacterium Sinorhizobium meliloti (10). It is believed that during lipid remodeling, phospholipids are degraded by PlcP to diacylglycerol (DAG), which then acts as the precursor for the biosynthesis of surrogate lipids in response to P limitation. Indeed, plcP deletion mutants no longer synthesize surrogate nonphospholipids, supporting the essential role of PlcP in lipid remodeling (8, 10).
Although PlcP-mediated lipid remodeling appears to be widespread among marine bacteria, since the plcP gene has been found in diverse groups of marine heterotrophs, including Alphaproteobacteria, Gammaproteobacteria, Flavobacteria, and Verrucomicrobia (8), little is known about how PlcP activity is regulated. At the transcriptional level, in the marine bacterium Phaeobacter sp. strain MED193 and the terrestrial bacterium Sinorhizobium meliloti, the plcP gene is controlled by the two-component system PhoBR, with a Pho box, to which PhoB binds, found upstream of these plcP genes (8, 10). The PlcP protein is annotated as a member of the metallophosphoesterase superfamily (Pfam family PF00149 [11]). Sequence analyses have shown that PlcP (also known as LpxH2) has moderate sequence similarity to a well-characterized member of PF00149, LpxH, an enzyme catalyzing a key step in lipid A biosynthesis in some bacteria (10, 12). Interestingly, recent structural analyses of LpxH have uncovered a conserved binuclear manganese (Mn2+) center, suggesting that manganese may be important for PlcP activity (13, 14).
In this study, we present a detailed characterization of the PlcP proteins from representatives of the marine Roseobacter and SAR11 clades, Phaeobacter sp. strain MED193 and Pelagibacter sp. strain HTCC7211, respectively, both of which are known to employ PlcP for lipid remodeling in response to P deficiency (7, 8). Specifically, we set out to determine the role of metal ions in regulating PlcP activity and to compare PlcP substrate specificities in PlcP enzymes from these two clades.
RESULTS AND DISCUSSION
PlcP is a member of the metallophosphoesterase family.Functional domain analysis of PlcP proteins from Phaeobacter sp. strain MED193 and Pelagibacter sp. strain HTCC7211 revealed the presence of a highly conserved histidine/aspartate/asparagine cage (Fig. 1A), a signature sequence motif found in the metallophosphoesterase family of proteins (PFam 00149). Subsequent phylogenetic analysis (Fig. 1B), comprising representative sequences from key members of the metallophosphoesterase family, including phosphodiesterases and pyrophosphatases, showed that the most closely related protein to PlcP in this family is LpxH, an enzyme catalyzing the formation of key intermediates in lipopolysaccharide biosynthesis. PlcPs from Phaeobacter sp. strain MED193 and Pelagibacter sp. strain HTCC7211 have 25% and 22% identity, respectively, to LpxH from Pseudomonas aeruginosa. Crystal structures of LpxH from Pseudomonas aeruginosa and Haemophilus influenzae have been solved recently, both of which showed the presence of a binuclear Mn2+ center in the active site (13, 14).
Multiple-sequence alignment and functional domain analyses of PlcP proteins. PLC193, PlcP of Phaeobacter sp. strain MED193; PLC7211, PlcP of Pelagibacter sp. strain HTCC7211; PlcPSm, PlcP of Sinorhizobium meliloti (10). (A) Multiple-sequence alignment of PlcP and closely related LpxH enzymes. The 6 conserved motifs are highlighted in gray. The conserved histidine residue in PlcP (H82) is highlighted in green. (B) Neighbor-joining phylogenetic analysis between members of the metallophosphoesterase family (PFam 00149) including proteins closely related to PlcP: PaLpxH, LpxH from Pseudomonas aeruginosa (14); HiLpx, LpxH from Haemophilus influenzae (13). LpxH displays pyrophosphatase activity and acts on UDP-2,3-diacylglucosamine to produce lipid X, a key precursor for the formation of lipid A in lipopolysaccharide biosynthesis. More-distantly related members of the metallophosphoesterase family include the following: MJ0936, which represents a group of novel phosphodiesterases that do not degrade phosphomonoesters (26); Mre11/SbcD, which are bacterial and archaeal DNA phosphodiesterases involved in DNA repair (27, 28); Dbr1, which is a group of phosphodiester nucleases that act on RNA (29); CpdA, CpdB, and cAMP phosphodiesterases, which are cyclic nucleotide phosphodiesterases; ApaH, which represents a group of enzymes with pyrophosphatase and protein phosphatase activities (30); YfcE, which represents a group of small metallophosphoesterases showing phosphodiesterase activity (31); sphingomyelinase, which is a group of hydrolases responsible for breaking down sphingomyelin to phosphocholine and ceramide (32). Numbers indicate bootstrap values (only values of >50 are shown).
We then employed homology modeling to predict metal-binding sites in PlcP193 (Fig. 2). Modeling predicted a highly conserved Mn2+ coordination center that was superimposable onto that of the LpxH (PDB 5K8K) structure. Using the PDBeFold server, we compared the crystal structure of LpxH to that of our model. This returned a Q score of 0.91, with 1 indicating identical structures, and a root mean square deviation (RMSD; a measure of the average distance between atoms) of 0.25, indicating some small differences between the structures in 222 of the 227 residues compared. With a highly comparable structure, key metal-binding residues H12, D40, N81, H82, H117, D119, H201, and H203 are shown to be orientated similarly in the two proteins. Likewise, potentially key differences are observed (Fig. 1; see also Movie S1 in the supplemental material) in the ligand recognition/binding site between the LpxH crystal structure and the PlcP193 model, including His134/Asp136, Typ126/Ala128, and Arg158/Tyr160, respectively, which could play an important role in substrate recognition. Although our modeling procedure is limited to positioning amino acid residues and not metal ion cofactors, given the highly similar overlap of amino acid residues around the metal coordination center, we hypothesized that Mn2+ metal cofactors would likely be coordinated similarly in PlcP193, and so, we have shown these ions superimposed as such (Fig. 2B).
Homology modeling showing the predicted structure of PlcP193 and the metal-binding pocket. The signature arginine residue in LpxH (Arg81) is replaced by a histidine residue in PlcP (His82).
To validate whether Mn2+ is indeed required for PlcP activity, we overexpressed and purified the PlcP proteins from Phaeobacter sp. strain MED193 and Pelagibacter sp. strain HTCC7211 in Escherichia coli (Fig. 3A). The isolated PlcP193 and PlcP7211 enzymes from recombinant E. coli had no activity. However, when divalent metals were included in the enzyme assay buffer, phosphoesterase activity was immediately restored (Fig. 3B). Of all the metals tested in this experiment at a range of concentrations (see Fig. S1 in the supplemental material), Mn2+ gives the highest activity, followed by Zn2+ and Fe2+. Our data agree well with known metal requirements for other characterized proteins within this family. For example, a binuclear Mn2+-Mn2+ center is common in Mre11 and LpxH group enzymes, which are evolutionarily more closely related to PlcP than other members of the family (Fig. 1B). Zn and Fe, on the other hand, have also been found in several proteins of this family, notably GpdQ glycerophosphodiesterase (15). Moreover, our data support the homology modeling prediction and show that PlcP from these marine bacteria is an Mn2+-dependent phosphoesterase. Interestingly, both PlcP193 and PlcP7211 proteins had phosphodiesterase and phosphomonoesterase activities as assessed using a general substrate containing either a phosphate monoester (PNPP) or a phosphodiester bond (NPPC) (Fig. 3C).
PlcP displays manganese-dependent phosphomonoesterase and phosphodiesterase activities. (A) Overexpression and purification of PlcP from Phaeobacter sp. strain MED193 and Pelagibacter sp. strain HTCC7211. M, protein molecular weight marker. Lane 1, cell-free supernatant induced with isopropyl β-d-1-thiogalactopyranoside (IPTG); lane 2, cell-free supernatant without IPTG induction; lane 3, purified PlcP protein (molecular weight estimated to be ∼27 kDa). (B) PlcP activity assays in the presence of various divalent metal ions (1 mM). Values are means ± standard deviations from three replicated measurements. (C) PlcP activity assays using p-nitrophenylphosphorylcholine (NPPC) or p-nitrophenylphosphate (PNPP). Values are means ± standard deviations from three replicated measurements.
Site-directed mutants and PlcP activity profile.In order to identify key amino acid residues in the PlcP protein required for phosphoesterase activity, and to further validate the predicted homology model (Fig. 2), we constructed site-directed mutants of PlcP193 and compared their activities with those of wild-type PlcP193. The data presented in Fig. 4 confirm the key role of the histidine/aspartate/arginine motifs in PlcP activity. Site-directed mutations of H12, D40/D43, H117/H119, and H201/H203 (highlighted in boxes 1, 2, 4, and 5, respectively, in Fig. 1A) to alanine almost completely abolished PlcP activity (Fig. 4E), supporting our assumption from the homology modeling that these residues are integral to the metal-binding site. Similarly, a histidine-to-alanine (H82A; box 3 in Fig. 1A) mutation completely abolished PlcP activity. Interestingly, the activity of H82N (53% of wild-type PlcP193) and H82R (82% of wild-type PlcP193) mutants is largely retained but statistically different from that of the wild-type PlcP (P < 0.01 and P < 0.05, respectively, by Student's t test), suggesting that the presence of a protonated amine group in this position is important in maintaining PlcP activity. The GNXD motif highlighted in box 3 (Fig. 1A) includes an interesting sequence variation in several enzymes of this family, including LpxH (arginine, R), YfcE (cysteine, C), and MJ0936 (asparagine, N). Structural determination of the LpxH-substrate complex revealed that the arginine residue is required for binding of the phosphate group of lipid X (14). In YfcE, the residue in this position is thought to be critical for the enzyme to switch between a phosphomonoesterase and a cyclic nucleotide phosphodiesterase (16). Steady-state kinetic measurements of the H82N and H82R mutants of PlcP193 showed higher affinity (Km = 154.8 ± 4.8 μM and 182.5 ± 5.3 μM, respectively) toward the artificial substrate p-nitrophenylphosphorylcholine (NPPC) compared to the wild-type (Km = 234.2 ± 6.3 μM) PlcP193 (Table 1). This enhanced substrate affinity in these mutants may be attributable to the reduced size of the side chain, from an imidazole ring of histidine to an aliphatic side chain of asparagine and arginine (Fig. 4B to D).
Homology modeling prediction of the metal coordination center in the PlcP193 enzyme (A) and the site-directed mutants of His82 to Ala82 (B), Arg82 (C), and Asn82 (D). Specific activities of site-directed mutants of PlcP193 are measured using p-nitrophenylphosphorylcholine (NPPC) as the substrate (E). Values are means ± standard deviations from three replicated measurements.
Kinetic parameters of wild-type PlcP7211, wild-type PlcP193, and mutant PlcP193 enzymes following the hydrolysis of p-nitrophenylphosphorylcholine (NPPC)a
Substrate profiles of PlcP on phospholipids.Although the use of artificial substrates provides an overview of PlcP phosphodiesterase and phosphomonoesterase activity and how these activities are regulated by metal ions, the native substrates of PlcP are believed to be native bacterial phospholipids in the membrane (7, 8). Phospholipids are degraded by PlcP to generate diacylglycerol (DAG) for the biosynthesis of non-P surrogate lipids in response to P limitation, and a plcP deletion mutant can no long synthesize such surrogate lipids (8). In Phaeobacter sp. strain MED193 and Pelagibacter sp. strain HTCC7211, the native phospholipids are PG and PE. We therefore hypothesized that PG and PE are the preferred native substrates for PlcP. To test this hypothesis, we used a range of phospholipids in addition to PG and PE. These included phosphatidylcholine (PC), a relative uncommon phospholipid in bacteria and absent in the aforementioned marine bacteria, phosphatidic acid (PA), an intermediate in the biosynthesis of phospholipids, and phosphatidylserine (PS) and phosphatidylinositol (PI), both of which are associated primarily with eukaryotes (17, 18). To rule out any potential effect of fatty acid chain length on PlcP activity across different phospholipid species, all lipid substrates contained sn-1 C16:0/sn-2 C16:0 palmitic acid. The data presented in Fig. 5 show that PG is the preferred substrate whereas the enzyme is least active toward PI and PS. Interestingly, PlcP from these two marine bacteria showed comparable, if not higher, activity towards PA (containing a phosphate monoester bond), an essential intermediate in phospholipid biosynthesis. Our data therefore suggest that PlcP in these marine bacteria have a relatively broad substrate specificity and are able to divert primary bacterial lipids (or the precursor, PA) through the conversion to DAG for the synthesis of non-P lipids in response to P limitation.
Specific activity of the degradation of phospholipids by PlcP193 and PlcP7211. Activity was measured by quantifying the formation of the common product diacylglycerol (DAG) in these reactions. PC, phosphatidylcholine; PE, phosphatidylethanolamine; PG, phosphatidylglycerol; PA, phosphatidic acid; PI, phosphatidylinositol; PS, phosphatidylserine. Values are means ± standard deviations from three replicated measurements.
To conclude, our data show that the activity of purified PlcP from two model marine heterotrophic bacteria is dependent on metal ions, particularly manganese, opening up the possibility that manganese availability may play a role in regulating the lipid remodeling process in natural marine systems (Fig. 6). Dissolved manganese concentrations are in the low nanomolar range (0.1 to 4 nM) in marine surface waters, and it is already known that manganese is important for maintaining optimum photosystem II (PSII) activity in marine phytoplankton (19, 20). Thus, the wide occurrence of ABC-type manganese transporters in SAR11 and marine Roseobacter clades might suggest potential competition with marine phytoplankton for manganese (4). Nevertheless, our observation of PlcP dependence on manganese reiterates more generally the importance of trace metals in regulating the activity of enzymes playing key roles in nutrient transformations in marine systems, e.g., zinc in alkaline phosphatase (21) and iron in nitrogenase (6). Moreover, the fact that PlcP appears to be able to degrade several native bacterial lipids that are common in marine bacteria, e.g., PG and PE, suggests that PlcP undertakes a generic lipid remodeling role among marine heterotrophic bacteria adapted to low-P environments.
Schematic overview of the PlcP-mediated lipid remodeling pathway and its regulation in representative marine bacteria. The major lipids in Phaeobacter sp. strain MED193 and Pelagibacter sp. strain HTCC7211 under P-replete conditions are two phospholipids (highlighted in gray), phosphatidylethanolamine (PE) and phosphatidylglycerol (PG). Under P stress, the two-component signal transduction system PhoBR is activated and the phosphorylated PhoB activates not only the expression of the high-affinity ABC transporter for phosphate (PstSABC) but also the transcription of the plcP gene. A conserved phoB binding site in the plcP promoter has previously been identified in these bacteria (8). The purified PlcP protein requires manganese for activity (Fig. 3). Manganese is likely transported into the cell through either the SitABCD or the MntX transporter system, both of which are present in these marine heterotrophic bacteria (4). Active PlcP can convert PE, PG, or its biosynthesis precursor phosphatidic acid (PA) to diacylglycerol (DAG), which serves as the building block for the biosynthesis of alternative P-free surrogate lipids (highlighted in blue), including diacylglyceryl trimethylhomoserine (DGTS) and the glycolipids monoglycosyl diacylglycerol (MGDG) and glucuronic acid diacylglycerol (GADG) (7, 8).
MATERIALS AND METHODS
Cloning, expression, and purification of PlcP.The wild-type plcP from Pelagibacter sp. strain HTCC7211 and the wild-type plcP and site-directed plcP mutants (encoding mutations H12A, D40A, D43A, N81A, H82A, H82N, H82R, H117A, D119A, H201A, H203A, and D220A) from Phaeobacter sp. strain MED193 were codon optimized and chemically synthesized by Genscript. The genes were then inserted into the pET28a expression vector using NdeI and BamHI restriction sites and transformed into E. coli BL21(DE3)-CodonPlus-RIL. To induce the expression of PlcP, 2% (vol/vol) overnight E. coli culture grown on lysogeny broth (LB) was inoculated into 600 ml fresh LB broth. Kanamycin was added to LB medium to a final concentration of 50 mg liter−1. The cultures were then incubated at 37°C with shaking (200 rpm). When the optical density at 600 nm (OD600) reached ∼0.5, IPTG (isopropyl-β-d-thiogalactopyranoside) was added to a final concentration of 0.2 mM; cells were harvested after 4 h at 37°C for PlcP193 and its mutants or after 8 h at 30°C for PlcP7211. Cells were harvested by centrifugation and resuspended in buffer containing 50 mM Tris-HCl–200 mM NaCl, pH 8.0. Cells were disrupted by sonication. Cell debris was removed by centrifugation at 20,000 × g for 20 min, and the supernatant was loaded onto a nickel column (GE Healthcare). After washing with buffer containing 20 mM Tris-HCl (pH 8.0), 200 mM NaCl, and 100 mM imidazole, proteins were eluted with elution buffer (20 mM Tris-HCl [pH 8.0], 200 mM NaCl, 300 mM imidazole). Purified protein was analyzed by SDS-PAGE, and protein concentrations were determined using the Bradford assay.
Bioinformatics and homology modeling.The homologous sequence and conserved domain were identified using the BLASTp software provided by the National Center for Biotechnology Information (http://blast.ncbi.nlm.nih.gov/Blast.cgi). Multiple alignment analyses of the deduced amino acid sequences were performed by ClustalW2 (http://www.ebi.ac.uk/Tools/clustalw2/index.html). Phylogenetic analysis used the neighbor-joining analysis method using Molecular Evolutionary Genetic Analysis 7.1 software (MEGA, version 7.1) (22). The three-dimensional model structures of PlcP193 and the H82A, H82N, and H82R mutants were generated by submitting the respective amino acid sequences to the Phyre 2 protein modeling and structure prediction server (23) and selecting the overall best scoring model in terms of coverage and confidence. All protein structure models were visualized in Chimera (24).
PlcP activity assay using artificial substrates.PlcP activity was measured in 96-well microplates using 1 mM p-nitrophenylphosphorylcholine (NPPC) or p-nitrophenylphosphate (PNPP) as a substrate, containing 1 μM purified enzyme in 50 mM Tris-HCl (pH 9.5), 60% (wt/vol) sorbitol, 1 mM MnCl2. Enzyme activity was measured at 65°C for 30 min, and absorbance was monitored at 405 nm for the formation of p-nitrophenol. One unit of phospholipase activity was defined as the amount of enzyme releasing 1 μmol p-nitrophenol per min under the standard conditions. Km and Vmax values were calculated using Hanes-Wolff plots at various concentrations of substrates (0.02 to 1.0 mM) in three replicates.
PlcP activity assay using phospholipids.A range of phospholipids were used to test PlcP specificity, including 1,2-dipalmitoyl-sn-glycero-3-phospho-rac-(1-glycerol) sodium salt (PG), 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (PC), 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine (PE), 1,2-dipalmitoyl-sn-glycero-3-phosphate sodium salt (PA), 1,2-dipalmitoyl-sn-glycero-3-phospho-l-serine sodium salt (PS), and 1,2-dipalmitoyl-sn-glycero-3-phospho-1′-myo-inositol ammonium salt (PI) (25). The standard reaction mixture (300 μl) contained 2.5 μM purified enzyme in 50 mM Tris-HCl (pH 9.5), 1 mM MnCl2, and phospholipids (0.1 to 0.8 mM) and was incubated for 30 min at 65°C. Phospholipids and the common degradation product, diacylglycerol (DAG), were extracted according to the Folch method using methanol-chloroform-water at a ratio of 1:2:0.6 (vol/vol/vol). The lipid extract was dried under nitrogen gas at room temperature. The dried lipids were resuspended in acetonitrile and ammonium acetate (10 mM, pH 9.2) at a ratio of 95:5 (vol/vol) and analyzed by liquid chromatography-mass spectrometry (LC-MS).
Phospholipid characterization and quantification by LC-MS.Phospholipids and PlcP-hydrolyzed lipid products were analyzed by LC-MS using a Dionex UltiMate 3000 LC system (Thermo Scientific, Waltham, MA) coupled to a Bruker amazon SL electrospray-ion (ESI) trap mass spectrometer (Billerica, MA). A BEH Amide XP column (2.5-μm inner diameter; 3 mm by 150 mm) was obtained from Waters (Milford, MA) and used for the chromatographic separation using a mobile phase consisting of acetonitrile (solvent A) and 10 mM ammonium acetate, pH 9.2 (solvent B). The column was equilibrated for 10 min with 95:5 (vol/vol) A/B prior to sample injection. The separation of phospholipids was conducted using a stepwise gradient starting from 95% (vol/vol) A and 5% (vol/vol) B to 70% (vol/vol) A and 30% (vol/vol) B after 15 min with a constant flow rate of 150 μl min−1. Instrument settings for the positive-ion ESI/MS and MS/MS analysis of phospholipids were as follows: capillary voltage, 4,500 V; end plate offset, 500 V; 8 liters min−1 drying gas at 250°C; nebulizing gas pressure, 15 lb/in2. Data analysis was performed using the Compass data analysis and QuantAnalysis 4.2 software (Bruker, Billerica, MA). PlcP activity toward phospholipids was measured by quantifying 1,2-dipalmitoyl-sn-glycerol (DAG) formation. A standard calibration curve for DAG was generated by correlating peak area to DAG quantity. A concentrated stock solution of DAG was prepared by dissolving it in chloroform at a concentration of 0.5 mg ml−1. The concentrated stock was further diluted in chloroform to generate standards at 0.005, 0.01, 0.02, 0.04, 0.06, 0.08, and 0.1 mg ml−1.
ACKNOWLEDGMENTS
This project has received funding from the European Research Council (ERC) under the European Union's Horizon 2020 research and innovation program (grant agreement no. 726116). We also thank the Program of Study Abroad for Young Scholars sponsored by Zhengzhou University of Light Industry, China, to W.T., the National Natural Science Foundation of China (NSFC, numbers 31728001, 31630012, U1706207), and a Royal Society International Exchanges grant (IEC\NSFC\170213).
FOOTNOTES
- Received 11 May 2018.
- Accepted 17 May 2018.
- Accepted manuscript posted online 25 May 2018.
Supplemental material for this article may be found at https://doi.org/10.1128/AEM.01109-18.
- Copyright © 2018 American Society for Microbiology.
