|Home | About | Journals | Submit | Contact Us | Français|
Adipose phospholipase A2 (AdPLA or Group XVI PLA2) plays an important role in the onset of obesity by suppressing adipose tissue lipolysis. As a consequence, AdPLA-deficient mice are resistant to obesity induced by a high fat diet or leptin deficiency. It has been proposed that AdPLA mediates its antilipolytic effects by catalyzing the release of arachidonic acid. Based on sequence homology, AdPLA is part of a small family of acyltransferases and phospholipases related to lecithin:retinol acyltransferase (LRAT). To better understand the enzymatic mechanism of AdPLA and LRAT-related proteins, we solved the crystal structure of AdPLA. Our model indicates that AdPLA bears structural similarity to proteins from the NlpC/P60 family of cysteine proteases, having its secondary structure elements configured in a circular permutation of the classic papain fold. Using both structural and biochemical evidence, we demonstrate that the enzymatic activity of AdPLA is mediated by a distinctive Cys-His-His catalytic triad and that the C-terminal transmembrane domain of AdPLA is required for the interfacial catalysis. Analysis of the enzymatic activity of AdPLA toward synthetic and natural substrates indicates that AdPLA displays PLA1 in addition to PLA2 activity. Thus, our results provide insight into the enzymatic mechanism and biochemical properties of AdPLA and LRAT-related proteins and lead us to propose an alternate mechanism for AdPLA in promoting adipose tissue lipolysis that is not contingent on the release of arachidonic acid and that is compatible with its combined PLA1/A2 activity.
Adipose tissue plays an important role in the regulation of energy balance. Although most other tissues can hydrolyze triacylglycerols for their own use, adipose tissue alone can hydrolyze triacylglycerols to export glycerol and fatty acids for use by other tissues. The net rate of triacylglycerol hydrolysis in adipose tissue (or lipolysis) is a determinant of whole-body insulin sensitivity (1) and hence a potentially important therapeutic target in the treatment of obesity. Adipose tissue lipolysis proceeds through the sequential action of the triacylglycerol hydrolases, desnutrin/adipose triglyceride lipase (2–4), hormone-sensitive lipase (5, 6), and monoglyceride lipase (7). The rate of adipose tissue lipolysis is carefully controlled by endocrine mediators, such as insulin and catecholamines, and locally through the action of autocrine/paracrine signaling mediators (8).
In addition to defining the structure and permeability of cells and cellular organelles, phospholipids are a source of locally acting signaling mediators. Hydrolysis of fatty acids linked to the sn-1 or sn-2 positions of phospholipids by phospholipase A1 (PLA1)3 or A2 (PLA2) enzymes, respectively, leads to formation of lipid signaling mediators, such as free fatty acids and lysophospholipids (for a review, see Ref. 9). Adipose phospholipase A2 (AdPLA) classified as Group XVI PLA2 (PLA2G16) (9) is the most abundant PLA2 in adipose tissue (10, 11), and its expression increases during adipose differentiation (12). Expression of AdPLA is also up-regulated by feeding and by insulin in vivo (11). Importantly, AdPLA was shown to control adipose tissue lipolysis via the production of eicosanoid mediators (11). The level of the prostaglandin PGE2 was shown to be markedly reduced in the adipose tissue of AdPLA−/− mice (11). PGE2 activates prostaglandin E receptor 3, which suppresses lipolysis in a cAMP-dependent manner (13). As a result, AdPLA−/− mice have increased rates of lipolysis and fatty acid oxidation and higher energy expenditure than wild type mice (11). AdPLA−/− mice are lean and resistant to diet-induced obesity (11). Moreover, crossing AdPLA−/− mice with leptin-deficient (ob/ob) mice generates AdPLA−/−,ob/ob double knock-out mice that gain considerably less weight than single knock-out ob/ob leptin-deficient mice (11). These data suggest that AdPLA is an important regulator of the rate of adipose tissue lipolysis via production of prostanoid mediators.
The enzymatic activity of AdPLA was initially described to be a calcium-dependent PLA2 utilizing a His-Cys catalytic dyad (10). Later reports described AdPLA to be a calcium-independent phospholipase with combined PLA1, PLA2, and transacylase activities; its PLA1 activity was reported to be higher than its PLA2 activity (14). The sequence and enzymatic activity of AdPLA resemble those of a small family of proteins related to lecithin:retinol acyltransferase (LRAT). All members of the LRAT family have been shown to catalyze PLA1, PLA2, or acyltransferase reactions (15–18). Thus, the LRAT family could be appropriately described as a phospholipase A/acyltransferase family based on a recent proposal by Uyama et al. (19). In this case, AdPLA would be referred to as phospholipase A/acyltransferase-3.
The inhibitory effect of AdPLA expression on adipose tissue lipolysis was proposed to be a result of the production of PGE2 through the release of arachidonic acid from the sn-2 position of phospholipids (11). However, the significant PLA1 activity of AdPLA led to suggestions that AdPLA might have other functions in controlling cellular metabolism and lipid accumulation in addition to production of free arachidonic acid (14, 20). In one such example, overexpression of AdPLA in human embryonic kidney 293 (HEK293) cells (21) leads to peroxisome dysfunction and a decrease in ether-linked triglycerides and plasmalogens (22). Importantly, these effects were not accompanied by a significant release of arachidonic acid in such cells.
AdPLA was originally identified as being specifically expressed in phenotypic revertants of H-ras-transformed cells; thus, it was characterized as a class II tumor suppressor (23) and referred to as H-ras-revertant gene 107 (H-Rev107) or H-Ras-like suppressor 3 (HRASLS3). Expression of AdPLA is down-regulated in several tumor cell lines (24–27); however, overexpression of AdPLA induces growth inhibition of transformed cells (28) and apoptosis of ovarian carcinoma cells (29). The relationship between the enzymatic activity and the tumor suppressive effect of AdPLA is not well understood.
To gain a better understanding of the physiological role of AdPLA, we investigated its structure and enzymatic properties. Based on sequence analysis, members of the LRAT family were predicted to have a circular permutation of the catalytic domain found in members of the superfamily of proteins related to Escherichia coli NlpC and Listeria monocytogenes P60 proteins (NlpC/P60) (30, 31). Several of the NlpC/P60 enzymes were suggested to utilize a conserved Cys-His pair or a Cys-His-His triad in their catalytic mechanism (32, 33). In the case of LRAT and other LRAT family members, the Cys residue was shown to act as a nucleophile and form a covalent thiol-acyl intermediate in the catalytic process (20, 34). A truncated fragment of AdPLA lacking the transmembrane domain was recently characterized by a solution NMR structure (35) and x-ray crystallography (20). Both structures show that AdPLA conforms to the permuted papain domain fold seen in a subset of NlpC/P60 proteins and predict the location of residues potentially involved in the catalytic triad of AdPLA.
Herein, we further describe the enzymatic mechanism of AdPLA by presenting a novel crystal structure of AdPLA that provides additional support for its enzymatic mechanism. We describe the expression and purification of a soluble full-length form of AdPLA that displays robust enzymatic activity, and we examine the PLA1/PLA2 specificity of AdPLA for various natural phospholipid substrates. We provide experimental support for the role of the residues proposed to be involved in the catalytic mechanism, and we probe the structural dynamic difference between full-length and truncated AdPLA based on hydrogen/deuterium exchange experiments coupled to mass spectrometry (DXMS). Finally, we evaluate the possible role of AdPLA in the generation of arachidonic acid in adipose tissue in light of its activity and substrate preference.
We cloned human AdPLA cDNA from a full-length expressed sequence tag clone available from the Open Biosystems Mammalian Gene Collection (MGC) (MGC: 118754, IMAGE: 40000132). A truncation of human AdPLA (T-AdPLA) retaining amino acid residues 1–134 was amplified and cloned through ligation-independent cloning in the vector pTB-MalE (36) digested with SspI. The ligation places T-AdPLA downstream of an N-terminal fusion partner consisting of a hexahistidine-tagged maltose-binding protein (MBP) and a tobacco etch virus (TEV) protease cleavage site. Alternatively, the full-length AdPLA (FL-AdPLA) cDNA was cloned in-frame with MBP and separated by a TEV cleavage site to generate a construct coding for an MBP-FL-AdPLA fusion protein. Mutagenesis of the MBP-FL-AdPLA protein was performed using a QuikChange II XL site-directed mutagenesis kit (Stratagene, La Jolla, CA). The expression constructs were sequenced and transformed in Bl21(DE3) E. coli and grown in Terrific Broth under the selection of 100 μg/ml ampicillin and 25 μg/ml chloramphenicol. The primers used for cloning T-AdPLA and FL-AdPLA and for site-directed mutagenesis reactions of MBP-FL-AdPLA are shown in supplemental Fig. S1. The sequence of the expressed MBP-T-AdPLA and MBP-FL-AdPLA proteins are shown in supplemental Fig. S2.
Protein expression was induced using the Overnight Express Autoinduction System (EMD Millipore-Merck KGaA, Billerica, MA) based on a method described previously (37). The bacteria were harvested by centrifugation at 4,000 × g for 10 min at 4 °C. The pellet was resuspended in 50 mm Tris, pH 8.0, 500 mm NaCl, 5 mm β-mercaptoethanol, 10% glycerol, 25 units/ml Benzonase (EMD Millipore), 50 mm arginine and lysed either by two passages in a French press or through sonication at 50% amplitude for 6 × 20 s. The lysate was spun by centrifugation at 19,000 rpm for 30 min to pellet membranes and insoluble complexes and then filtered through a 0.2-μm filter. Purification of MBP-FL-AdPLA or MBP-T-AdPLA fusion proteins was carried out by metal affinity chromatography. The bacterial supernatant was loaded onto tandem 5-ml His Trap HP columns (GE Healthcare) in conjunction with an ÄKTA Xpress purification system at a flow rate of 1 ml/min. The columns were washed with Buffer A (0.2% Triton X-100, 50 mm Tris, pH 8.0, 50 mm NaCl, 5 mm β-mercaptoethanol, 10% glycerol) over 4 column volumes. Another wash with 5% Buffer B (500 mm imidazole, 50 mm Tris, pH 8.0, 50 mm NaCl, 5 mm β-mercaptoethanol, 10% glycerol) was used to remove nonspecifically bound proteins. Elution was carried out with Buffer B in a linear gradient from 5 to 100%. The resulting fractions were examined by SDS-PAGE, and the protein-containing fractions were pooled and filtered through a 0.2-μm filter. The protein solution was concentrated and desalted using a Spin-X UF centrifugal concentrator (20 ml; 5,000 molecular weight cutoff; Corning Inc. Life Sciences, Tewksbury, MA) to a final concentration of 10 mg/ml in 50 mm Tris, pH 8.0, 50 mm NaCl, 5 mm β-mercaptoethanol. MBP-FL-AdPLA or MBP-T-AdPLA fusion proteins were next purified by anion exchange chromatography using tandem 5-ml HiTrap Q HP columns (GE Healthcare). The column was washed using Buffer A (50 mm Tris, pH 8.0, 50 mm NaCl) over 4 column volumes followed by a wash with 10% Buffer B (50 mm Tris, pH 8.0, 1.0 m NaCl, 5 mm β-mercaptoethanol) to remove nonspecifically bound proteins. Elution was carried out with Buffer B in a linear gradient from 10 to 100% at a flow rate of 1 ml/min, and the protein-containing fractions were pooled and filtered through a 0.2-μm filter. Cleavage with TEV to remove the MBP fusion was performed using 30 mg of MBP-T-AdPLA protein in 30 ml of 150 mm NaCl, 25 mm Tris, pH 8.0, 5 mm β-mercaptoethanol supplemented with 600 μg of hexahistidine-tagged TurboTEV (2 mg/ml stock; 10,000 units/mg; Nacalai USA, San Diego, CA) for 48 h at 20 °C with gentle shaking. The hexahistidine-tagged MBP and TurboTEV proteins were removed by metal affinity chromatography using 2 ml of HisPur nickel-nitrilotriacetic acid resin (Fisher Scientific) in sodium phosphate, pH 7.4, 10 mm imidazole, 5 mm β-mercaptoethanol. The eluted untagged T-AdPLA protein was filtered through a 0.2-μm filter and concentrated to 4 ml using a Spin-X UF centrifugal concentrator (20 ml; 5,000 molecular weight cutoff; Corning Inc. Life Sciences) and purified by using size exclusion chromatography on a HiLoad 16/600 Superdex 75 prep grade column equilibrated with a buffer of 50 mm Tris, pH 8.0, 300 mm NaCl, 5 mm dithiothreitol (DTT) and eluted at 0.8 ml/min over 2.5 column volumes. The T-AdPLA-containing fractions were pooled, filtered through a 0.2-μm filter, and concentrated to 10 mg/ml. The resulting untagged T-AdPLA protein is stable at 4 °C for more than 4 weeks. To verify the identity of the purified protein, the recombinant T-AdPLA protein was examined by SDS-PAGE. The purified protein migrated as a single band as identified by Coomassie Blue staining. The protein band was cut from the gel and treated with iodoacetamide to amidate the Cys residues and digested in-gel with l-1-tosylamido-2-phenylethyl chloromethyl ketone-treated trypsin. Analysis of the tryptic fragments by tandem MS/MS identified the purified protein as AdPLA showing 90% peptide coverage of the T-AdPLA protein (supplemental Fig. S3). The T-AdPLA protein was set up for crystallization screens or for enzyme assays.
In the case of FL-AdPLA, cleavage with TEV of the MBP-FL-AdPLA fusion protein results in precipitation of the FL-AdPLA protein; therefore, MBP-FL-AdPLA was purified sequentially by affinity chromatography, ion exchange chromatography, and size exclusion chromatography as described for MBP-T-AdPLA and then used as an intact MBP-FL-AdPLA fusion protein in enzyme assays. The uncleaved MBP-FL-AdPLA protein is stable at 4 °C for several months. We also expressed and purified a C-terminal truncated fragment of AdPLA representing the residues 1–132 of human AdPLA fused to an N-terminal hexahistidine tag denoted as His6-AdPLA(1–132), which has been described previously (20). The plasmid coding for His6-AdPLA(1–132) was obtained as a kind gift from Marcin Golczak and Kris Palczewski. The plasmid was used to express and purify the His6-AdPLA(1–132) protein following the same procedure as that used for MBP-FL-AdPLA.
A purified sample of T-AdPLA was screened for crystallization in Compact Jr. (Emerald Biosystems, Bainbridge Island, WA) sitting drop vapor diffusion plates at 20 °C using 0.5 μl of protein (10 mg/ml in 50 mm Tris, pH 8.0, 300 mm NaCl, 5 mm DTT) and 0.5 μl of crystallization solution equilibrated against 100 μl of the latter. Small prismatic crystals were obtained in 1–2 days from the Wizard 4 screen (Emerald Biosystems) condition 21 (25% polyethylene glycol (PEG) 1500, 100 mm malonate/imidazole/borate 2:3:3 (MIB) buffer, pH 5.0) and in ~1 week from the Wizard 3 screen condition 4 (Emerald Biosystems; 20% PEG 3350, 200 mm ammonium formate). Samples were transferred to a fresh drop composed of 80% crystallization solution and 20% PEG 200 before flash freezing in liquid nitrogen for data collection. Data were collected at the Advanced Photon Source beamline 17-ID using a Dectris Pilatus 6M pixel array detector.
Intensities were integrated using XDS (38), and the Laue class check and data scaling were performed with Aimless (39). The highest probability Laue class was −3m1 and space group P3121 or P3221. Matthews' coefficient (40) (Vm = 2.8/55.6%) indicated that there was a single AdPLA molecule in the asymmetric unit. Structure solution was conducted by molecular replacement with Phaser (41) via the PHENIX (42) interface. All space groups in the Laue class −3m1 were tested using a single molecule from a previously determined AdPLA NMR structure (Protein Data Bank code 2KYT) as the search model. The top solution was obtained in the space group P3121 and was used from this point forward. Because of the anisotropy of the diffraction data, the model was refined with a resolution cutoff of 2.65 Å. Structure refinement and manual model building were conducted with BUSTER (43) and Coot (44), respectively. Structure validation was conducted with MolProbity (45). Figures were prepared using the CCP4MG package (46). The coordinates and structure factors were deposited in the Protein Data Bank under accession code 4FA0.
Proteins were incubated in D2O buffer consisting of 12 mm Tris, 50 mm NaCl in 99% D2O, pDread 7.5 as described previously (47). Exchange experiments were initiated by mixing 2 μg of T-AdPLA, MBP-T-AdPLA, or MBP-FL-AdPLA in protein buffer or with 6 μl of D2O buffer to a final concentration of 75% D2O. The mixtures were incubated at 23 °C for 10, 30, 100, 300, 1,000, 3,000, or 10,000 s, and then the exchange reaction was quenched by adding 12 μl of ice-cold quench solution (0.96% formic acid, 0.8 m guanidine hydrochloride), resulting in samples with final concentrations of 0.58% formic acid and 0.5 m guanidine hydrochloride, pH 2.5. The samples were then immediately frozen on dry ice and stored at −80 °C.
All steps were performed at 0 °C as described previously (47). The digested peptides were separated by C18 column and detected using a Thermo Orbitrap Elite mass spectrometer. Data processing of DXMS experiments utilized specialized software as described previously (DXMS Explorer, Sierra Analytics Inc.) (47).
Liposomes were created using mixtures of phosphatidylcholine (PC) and phosphatidylglycerol (PG) phospholipids (Avanti Polar Lipids, Alabaster, AL). The PC species examined were 18:0-16:0 PC, 16:0-18:0 PC, 16:0-20:4 PC, 16:0-18:2 PC, 18:1-18:1 PC, 16:0 lyso-PC, and 18:0-18:0 diether PC. 100-μl aliquots from 10 mg/ml stock solutions of each of the PC species examined were mixed with an equal amount of 10 mg/ml 18:1-18:1 PG. The phospholipid mixture was dried down in a borosilicate glass test tube under vacuum to remove the solvent (chloroform). The lipid cake was hydrated in the presence of 50 mm Tris, pH 8, 100 mm NaCl, 1 mm CaCl2 for 1 h at a temperature above the solid-fluid transition temperature (Tc) of the phospholipid mixture followed by sonication of the hydrated lipid for 10–30 min in a bath sonicator to produce a suspension of small unilamellar vesicles representing a concentration of 1 mm for the combined phospholipids in a ratio of 1:1 PC/PG. The assay was conducted by incubating purified MBP-FL-AdPLA, T-AdPLA, MBP-T-AdPLA, bee venom PLA2 (Invitrogen), or Thermomyces lanuginosus lipase with sn-1,3 specificity (Lecitase Ultra-Novozymes A/S, Denmark; obtained from Invitrogen) for 15 min with various liposomes in the presence of 5 mm tris(2-carboxyethyl)phosphine in a total volume of 220 μl. The reaction was stopped by adding 1 ml of Dole's reagent (48) (40 ml of isopropanol, 10 ml of heptane, and 1 ml of 1 m H2SO4) and vortexed for 5 min. The organic phase was extracted with 600 μl of heptane and 400 μl of water. Phase separation was induced by centrifugation at 3,000 rpm for 5 min; the organic phase was collected and dried down under vacuum. The residue was resuspended in 200 μl of a mixture of chloroform and methanol (1:1), 10 μl of which was injected into the HPLC system for analysis. Free fatty acids were analyzed by reverse-phase HPLC coupled to a charged aerosol detector using an HPLC1200 system (Agilent Technologies, Santa Clara, CA) equipped with a Halo C8 column (150 × 4.6 mm; 2.7 μm; MAC-MOD Analytical, Inc., Chadds Ford, PA), mobile phase A composed of methanol/water/acetic acid (750:250:4), mobile phase B composed of acetonitrile/methanol/tetrahydrofuran/acetic acid (500:375:125:4), and a flow rate of 0.8 ml/min with a column temperature of 40 °C. The samples were injected in methanol/chloroform (1:1). The following gradient was used: 100% A to 30% A over 40 min, 30% A to 10% A over 20 min, and holding at 10% A for 5 min followed by re-equilibration in 100% A. The eluted lipids were detected using a Corona charged aerosol detector (ESA Biosciences) operated at 200 pA and 35-p.s.i. nitrogen pressure as described (49). Experimental samples consisting of various enzyme preparations or buffer control were set up in triplicate. Authentic fatty acid standards were used to establish the identity of the eluted peaks.
We also investigated the phospholipase activity of AdPLA using a boron-dipyrromethene (BODIPY)-labeled fluorogenic phospholipid substrate, 1-O-(6-BODIPY 558/568-aminohexyl)-2-BODIPY FL C5-sn-glycero-3-phosphocholine (BODIPY PC-A2; A10072, Invitrogen). Liposomes were prepared by the ethanolic injection method (50). Equal volumes of 10 mm 18:1-18:1 PC, 10 mm 18:1-18:1 PG, and 1 mm BODIPY PC-A2 were combined to prepare a lipid mixture containing 10:10:1 molar ratios of 18:1-18:1 PC/18:1-18:1 PG/BODIPY PC-A2. 10 μl of the lipid mixture was injected while vigorously vortexing in a glass tube containing 1 ml of buffer composed of 50 mm Tris, pH 8, 100 mm NaCl, 1 mm CaCl2. 100 μl of liposome substrate was aliquoted in a 96-well plate and examined using a Synergy H1 multimode microplate reader equipped with dispensers (BioTek, Winooski, VT). The liposome suspension was allowed to equilibrate at 37 °C while the fluorescence signal at 488/515 nm was recorded for 1 min to monitor the background emission. No significant increase in background fluorescence emission was observed. Purified MBP-T-AdPLA, T-AdPLA, MBP-T-AdPLA, His6-AdPLA(1–132), and wild type and mutants of MBP-FL-AdPLA were diluted in 50 mm Tris, pH 8, 100 mm NaCl, 1 mm CaCl2; 100 μl of protein was dispensed in the well containing the liposome suspension; and the fluorescence signal at 488/515 nm was recorded for the next 4 min. To calculate the rate of reaction as a function of the amount of enzyme, we used a final concentration of BODIPY PC-A2 of 1.67 μm, whereas the protein concentration ranged from 7.8 to 125 μg/ml (0.12–1.9 μm). A standard curve of the fluorescence signal at 488/515 nm of 3a,4a-diaza-s-indacene-3-pentanoic acid (BODIPY FL C5; D3834, Invitrogen), the cleavage product of BODIPY PC-A2, was used for evaluating the percentage of substrate conversion. To measure enzyme velocities as a function of substrate concentration, we used a titration of substrate concentrations in molar excess of enzyme concentrations. Initial rates of reaction for various substrate concentrations were derived from steady-state kinetic measurements based on rates measured in the first 20 s of the reaction. The assay was carried out in triplicate, and the data were evaluated using the SigmaPlot and GraphPad software packages.
We tested several strategies to express human AdPLA as a recombinant protein in E. coli for structural studies. It was shown that the C-terminal domain of AdPLA is embedded in the membrane, whereas the N terminus of the AdPLA protein is found in the cytoplasm (10, 51). Analysis of the amino acid sequence of AdPLA reveals the absence of a cleavable signal sequence and the presence of a single hydrophobic domain at its C terminus. In accordance with this model, the C-terminal domain is required for targeting AdPLA to the membrane of the endoplasmic reticulum (10) by acting as a both a targeting signal and a membrane anchor domain (51). To increase the yield of soluble protein, we tested the effect of removing the C-terminal hydrophobic domain and expressed both the full-length and the truncated version of AdPLA fusion proteins with a hexahistidine-tagged MBP protein to increase their solubility (52). The linker between the MBP and AdPLA contains a TEV cleavage site (ENLIF(Q/S)), allowing the MBP fusion partner to be cleaved with TEV protease and removed in the final stages of purification. Cleavage with TEV protease produces a protein that retains the sequence of either FL-AdPLA or T-AdPLA and a stretch of three additional amino acids (SNA-) at the N terminus of the recombinant proteins (supplemental Fig. S2).
Our results indicate that the presence of the C-terminal hydrophobic domain adversely affects the solubility of the bacterially expressed AdPLA protein. Although MBP-FL-AdPLA is a soluble protein, upon cleavage with TEV protease, the FL-AdPLA protein precipitated from solution. Inclusion of chaotropic agents and detergents led to only a modest improvement in the solubility of cleaved FL-AdPLA, and attempts to crystallize the intact MBP-FL-AdPLA protein were not successful. On the other hand, cleavage of purified MBP-T-AdPLA with TEV protease produced a soluble protein. The protein was confirmed as T-AdPLA by MS/MS analysis (supplemental Fig. S3). The purified T-AdPLA protein is monomeric in solution based on its elution profile on a size exclusion chromatography column calibrated with protein standards (supplemental Fig. S3). Crystallization screens of T-AdPLA produced small crystals in several conditions. The crystals were indexed as trigonal/hexagonal with a = 60.05 Å and c = 74.06 Å unit cell parameters. Analysis of the axial reflections indicated the presence of a 3-fold screw axis as systematic absences 00l; l = 3n were observed.
The structure solution of T-AdPLA was carried out by molecular replacement with Phaser (41) using the NMR model (Protein Data Bank code 2KYT) of AdPLA as the search model (35, 53). The terminal ends and residues 39–55 were truncated in the search model as they were shown to be disordered in Protein Data Bank code 2KYT. The top solution was obtained for a single monomer in the asymmetric unit in the space group P3121. Intensities were integrated and scaled using XDS and Scala (54), respectively (Table 1). The Wilson B-factor was unusually high (~65 Å2) for data processed to this resolution (2.3 Å). Because of the anisotropy of the diffraction data, the observed electron density maps were not clear in certain regions of the polypeptide chain, making it difficult to discern certain side chains. Therefore, a second data set was collected at a longer wavelength (λ = 1.7463 Å), and phased anomalous difference electron density maps were calculated in an effort to confirm the position of the Cys sulfur atoms and ensure that the sequence register was correctly assigned. Indeed, anomalous difference map peaks were observed greater than 4σ for the two Cys residues Cys89 and Cys113, respectively (Fig. 1) The structure reported here was deposited in the Protein Data Bank under accession code 4FA0.
Overall, the structure is similar to the solution NMR structure of AdPLA (Protein Data Bank code 2KYT) with a root mean square deviation of 1.43 Å between Cα atoms (Pro6 to Val124) as determined using secondary structure matching (55) via the CCP4 interface (56) (Fig. 2). Residues between Pro39 and Lys57 were disordered and could not be fit to the electron density maps. The largest deviations between the AdPLA crystal structure and the solution NMR structure are observed in regions containing loops 27–33, 71–74, 79–87, and 106–112 as shown in Fig. 3, A and C. The crystal structure we describe here also closely resembles the one recently reported by Golczak et al. (20) of an AdPLA protein shorter by two amino acids at its C terminus (Protein Data Bank code 4DOT) with a root mean square deviation of 0.54 Å between Cα atoms (Pro6 to Val124). The main difference between the crystal structure reported by Golczak et al. (20) and the one reported here occurs in a loop region between Val104 and Ser110 as shown in Fig. 3, B and C. This difference could be attributed to the relatively high conformational flexibility of the Val104–Ser110 loop as observed in the values of the average B-factors for the residues encompassing this domain (Fig. 3D).
The crystal structure of AdPLA reported here (Protein Data Bank code 4FA0) reveals an α+β fold composed of three α-helices and five antiparallel β-sheets in a circular permutation of the topology of papain and members of the NlpC/P60 superfamily of cysteine peptidases (31) (Fig. 4). A Cys residue (Cys113), which is highly conserved in both LRAT family proteins and the NlpC/P60 superfamily of proteins, is found on helix α3. In AdPLA and all LRAT-like proteins studied thus far, the conserved Cys113 residue was shown to be required for their catalytic activity (10, 14, 22) by acting as a nucleophile in forming the covalent acyl enzyme intermediate (20, 34). Our model indicates the presence of a well conserved His residue (His23) on a neighboring β-strand that is 3.6 Å from Cys113. A second His (His35), which resides on an adjacent β-strand, is positioned 3 Å from His23. The three Cys113-His23-His35 residues comprising the catalytic triad of AdPLA are found in a conserved arrangement in the active site of NlpC/P60 proteins, such as E. coli Spr (32).
We assayed the contribution of the C-terminal transmembrane domain to the phospholipase activity of AdPLA. For this, we examined the activity of MBP-FL-AdPLA and MBP-T-AdPLA proteins using a continuous assay based on monitoring the release of a fluorescently labeled acyl group from the PLA2-specific substrate BODIPY PC-A2 (the structure is depicted in Fig. 5, top panel). Release of the BODIPY FL pentanoic acid acyl chain from sn-2 of BODIPY PC-A2 results in an increase in the fluorescence signal (excitation, 488 nm; emission, 515 nm) due to decreased intramolecular quenching by fluorescence resonance energy transfer (FRET) of the BODIPY 558/568 dye attached at the sn-1 position. The assay allowed us to conduct a sensitive kinetic assay of AdPLA and to calculate its Vmax (2.5 ± 0.2 μmol/min/mg) and Km (0.7 ± 0.15 μm) using BODIPY PC-A2 as a substrate. These values are within the range of reported values for AdPLA in previous studies (10, 14). To account for the interfacial mechanism of the PLA2-catalyzed reaction, other kinetic models, such as the “surface dilution model” (57, 58), have been developed, but we were not successful in adapting the assay based on detergent-phospholipid mixed micelles to the analysis of the MBP-FL-AdPLA reaction. In conclusion, our findings establish that MBP-FL-AdPLA exhibits characteristics consistent with a physiological role of AdPLA in the hydrolysis of phospholipids.
Next, we used site-directed mutagenesis to examine the effect of mutating the active site residues Cys113, His23, and His35 on the phospholipase activity of AdPLA. Substituting either Cys113, His23, or His35 with alanine or the more sterically conserved substitutions C113S, H23Q, and H35Q resulted in a significantly reduced AdPLA activity of the resulting proteins (Fig. 6). Both Cys113 and His23 (His21 in mouse AdPLA) had been previously implicated to be important for the activity of AdPLA (10, 14). Here, we also demonstrate that His35 is required for AdPLA activity. We propose that the function of His23 is to form an ion pair with Cys113, acting as a base and subsequently as an acid in the reaction cycle. Meanwhile, His35 maintains His23 in its proper orientation as shown for the NlpC/P60 protein Spr (32). Thus, our studies provide experimental support for the requirement of the Cys113, His23, and His35 residues in the enzymatic activity of AdPLA.
Our results indicate that deletion of the C-terminal transmembrane domain of AdPLA severely affects the activity of MBP-T-AdPLA against liposome substrates. Using a liposome-based assay of AdPLA activity, we were not able to detect any hydrolytic activity associated with MBP-T-AdPLA (Fig. 6) or T-AdPLA (not shown). In comparison, MBP-FL-AdPLA had a robust activity (specific activity of 720 pmol/min/mg) (Fig. 6). These results are in agreement with the requirement of an intact C-terminal domain for the activity of AdPLA demonstrated by other studies (10, 14). Thus, the hydrophobic C-terminal domain is necessary for interfacial catalysis by AdPLA.
In recently published findings, Golczak et al. (20) examined the structure and activity of a 1–132 fragment of AdPLA with nearly the same sequence and structure as the T-AdPLA protein described here (Fig. 3, B and C). Although MBP-T-AdPLA, T-AdPLA, and His6-AdPLA(1–132) do not display significant activity against phospholipids embedded in liposomes (results not shown), it was shown that His6-AdPLA(1–132) protein can become acylated in the presence of phospholipids with very short fatty acid chains, such as diheptanoyl-PC (20). These findings suggest that the C-terminal domain of AdPLA does not include residues that participate in the catalytic reaction; however, this domain is required for the association of AdPLA with membranes.
To shed light on the structural differences between T-AdPLA, MBP-T-AdPLA, and MBP-FL-AdPLA, we used a DXMS approach to probe the solvent accessibility of backbone amides of T-AdPLA, MBP-T-AdPLA, and MBP-FL-AdPLA. We incubated purified T-AdPLA, MBP-T-AdPLA, and MBP-FL-AdPLA with buffer supplemented with D2O. Aliquots of the hydrogen/deuterium exchange reaction were quenched, digested with pepsin, and analyzed by LC-MS/MS. Pepsin digestion of the denatured T-AdPLA, MBP-T-AdPLA, and MBP-FL-AdPLA proteins resulted in a complete coverage of the AdPLA protein based on the identification of 36 and 30 peptides derived from the FL-AdPLA or the T-AdPLA fusion partner, respectively. The peptides with the best signal to noise ratio and least overlapping regions for representation of the hydrogen/deuterium exchange data are shown in supplemental Fig. S4. The kinetics of the rates of deuteration of T-AdPLA, MBP-T-AdPLA, and MBP-FL-AdPLA show the same behavior with the exchange steadily increasing with time, consistent with the premise that the FL-AdPLA and T-AdPLA proteins have similar structures (Fig. 7A). Comparison of the exchange rates of MBP-T-AdPLA and T-AdPLA showed a similar rate of solvent accessibility, arguing that the presence of the MBP fusion partner does not affect the folding of T-AdPLA. The rates of deuteration are also consistent with the crystal structure model as shown in Fig. 7, A and B. Residues 39–59 were found to be disordered in the crystal structure and could not be modeled. Using DXMS, we observed high rates of hydrogen/deuterium exchange for the region 39–59 (greater than 80% in 10 s and reaching a maximum after 1,000 s). These fast rates of exchange reflect the inherent flexibility of the region encompassing residues 39–59. The α3 helix (residues 111–121) containing the Cys113 catalytic residue is packed tightly against the β-sheet β2 and thus displays relatively low rates of exchange in T-AdPLA, MBP-T-AdPLA, and MBP-FL-AdPLA. We noticed very subtle differences between MBP-FL-AdPLA versus MBP-T-AdPLA or T-AdPLA as the rates of deuteration for α3 appear to be slightly lower in the case of MBP-FL-AdPLA than those observed for MBP-T-AdPLA or T-AdPLA, perhaps indicating a more compact structure of this domain.
Two domains of AdPLA encompassing residues Ile16–Tyr27 and Tyr66–Glu98 display much lower rates of solvent accessibility compared with the rest of the protein (Fig. 7, A and B, colored green). Both domains also display low conformational flexibility based on average B-factors (Fig. 3D). These results are consistent with the tight packing around the Ile16–Tyr27 domain, which includes the β-sheet β2 and the His23 catalytic residue. In addition, the Tyr66–Glu98 domain of AdPLA adopts a compact structure that excludes water, consistent with a possible role of this domain in additional interactions with the membrane interface. Overall, the T-AdPLA, MBP-T-AdPLA, and MBP-FL-AdPLA proteins appear to have similar solvent accessibility and to adopt similar conformations.
AdPLA was initially identified as a PLA2 enzyme (10); however, later reports suggested that the PLA1 activity of AdPLA is in fact higher than its PLA2 activity (14). The positional specificity, sn-1 versus sn-2 of AdPLA, is important for evaluating its physiological role. In physiological settings, unsaturated fatty acids, such as arachidonic acid, are found primarily in the sn-2 position of phospholipids. Arachidonic acid release is the key argument in explaining the physiological role of AdPLA in PGE2 production; therefore, determining the positional specificity of AdPLA is essential for explaining its physiological function.
The process of determining the positional specificity of phospholipases is affected by the rapid intramolecular rearrangement of lysophospholipids. Following cleavage by PLA1, the 2-acyl lyso-PC product rearranges to form the more thermodynamically favored 1-acyl lyso-PC (59). Subsequent action of PLA1 can result in a second round of hydrolysis and the production of l-α-glycerylphosphorylcholine through the release of the migrated acyl group. Thus, both sn-1- and sn-2-derived fatty acids can be released by a PLA1 enzyme despite its specificity for the sn-1 position of phospholipids. Because AdPLA was reported to release both sn-1 and sn-2 fatty acids, we sought to determine whether the combined PLA1/PLA2 activity could be explained by PLA1 activity alone.
Based on a fluorescence-based continuous assay using BODIPY PC-A2, our data argue that AdPLA displays intrinsic PLA2 activity by accessing the sn-2 acyl group directly. As seen in Fig. 5, MBP-FL-AdPLA can hydrolyze the BODIPY PC-A2, resulting in the increase of the fluorescence of the BODIPY FL dye. Because the mechanism of detection in this assay depends on the reduced quenching of the BODIPY FL dye by FRET of BODIPY 558/568, this implies that either the sn-1 or sn-2 labeled probes of BODIPY PC-A2 were hydrolyzed and released by MBP-FL-AdPLA. BODIPY PC-A2 contains an alkyl group linked through an ether bond in the sn-1 position. The ether bond of BODIPY PC-A2 confers PLA2 specificity by virtue of the fact that AdPLA, like most other PLA2 enzymes, does not hydrolyze ether bonds as found in 18:0-18:0 diether PC (results not shown). Together, these results demonstrate that the release of the BODIPY FL pentanoic acid acyl chain from sn-2 of BODIPY PC-A2 is the result of direct hydrolysis of the sn-2 ester bond by MBP-FL-AdPLA and that MBP-FL-AdPLA has authentic PLA2 activity.
It is important to determine the specificity of AdPLA for physiological substrates incorporated in liposomes that resemble the composition of cellular membranes. Thus, we used a secondary method to assess the activity of MBP-FL-AdPLA using a direct assay of the hydrolysis of natural phospholipid substrates. To determine the PLA1 or PLA2 specificity of MBP-FL-AdPLA, we used PC substrates with mixed acyl chains. We incubated MBP-FL-AdPLA, MBP-T-AdPLA, commercial PLA2 (bee venom), or PLA1 (T. lanuginosus lipase) preparations with liposomes composed of an equimolar mix of 18:1-18:1 PG and various species of PC. We then extracted and examined the fatty acids produced by HPLC using charged aerosol detection.
Our results indicate that MBP-FL-AdPLA displays potent PLA1 and PLA2 activities by catalyzing the hydrolysis of roughly equivalent amounts of stearate (18:0) or palmitate (16:0) whether linked to the sn-1 or the sn-2 position of 16:0-18:0 PC or 18:0-16:0 PC (Fig. 8, A and B). In the case of phospholipids containing unsaturated acyl chains, MBP-FL-AdPLA hydrolyzed the release of slightly more palmitate from the sn-1 position of 16:0-20:4 PC or 16:0-18:2 PC than arachidonate (20:4) or linoleate (18:2) from the sn-2 position, respectively (Fig. 8, D and E). To confirm the specificity of the substrates, we examined the hydrolysis of PC species with mixed acyl groups using authentic PLA1 or PLA2 enzyme preparations. By incubating liposomes composed of PC species with mixed acyl groups with the PLA1 enzyme from T. lanuginosus, we observed the release of fatty acids linked predominantly to the sn-1 position in 18:0-16:0 PC (Fig. 8C) or 16:0-18:0 PC (not shown). Meanwhile, hydrolysis of PC species with mixed acyl groups by PLA2 resulted in the release of fatty acids linked to the sn-2 position of 18:0-16:0 PC (Fig. 8C) and 16:0-18:0 PC (not shown). Comparison of the rates of hydrolysis indicates that MBP-FL-AdPLA does not appear to show any preference for the hydrolysis of arachidonic acid. Our results also show that MBP-FL-AdPLA can hydrolyze PG to release oleate (18:1) (Fig. 8F), indicating that additional phospholipid headgroups other than aminophospholipids can be recognized by AdPLA. As predicted, based on the fluorescent AdPLA assays, the levels of fatty acids measured in all assays of MBP-T-AdPLA activity are equivalent to the buffer control reaction; and therefore, MBP-T-AdPLA does not exhibit significant hydrolytic activity against substrates embedded in liposomes (Fig. 8). In conclusion, our data presented in Figs. 6 and and88 support a model in which AdPLA has combined PLA1 and PLA2 activities as opposed to exclusively PLA1 or PLA2 activity.
Our findings describe the structure and enzymatic activity of AdPLA, an enzyme that plays an important role in the regulation of adipose tissue function. To gain a better understanding of the molecular mechanism of AdPLA, we solved the crystal structure of a soluble fragment of AdPLA to 2.65-Å resolution (Protein Data Bank code 4FA0). The model presented here closely resembles the previously reported NMR model (Protein Data Bank code 2KYT) (Figs. 2 and and3,3, A and C) and the crystal structure model of AdPLA (Protein Data Bank code 4DOT) (Fig. 3, B and C). The model indicates that AdPLA adopts a globular fold consisting of three α-helices and five antiparallel β-sheets (Fig. 4) organized in a circular permutation of the classic papain fold. AdPLA utilizes a conserved Cys-His-His catalytic triad to orchestrate the nucleophilic attack by Cys on the sn-1 and sn-2 acyl groups of phospholipids. In line with this observation, residues Cys113, His23, and His35 are required for the its activity (Fig. 6). The use of a Cys residue for catalyzing the hydrolysis of phospholipids is unique to the phospholipases and acyltransferases found in the LRAT family. For Cys113 to act as a nucleophile, it would require a pKa of less than 7.0; however, the pKa of free Cys of roughly 8.3–8.8 cannot account for the nucleophilic character of this residue. Therefore, the active site of AdPLA provides an environment that is critical for catalysis by decreasing the pKa of Cys113. In papain, such function is provided by a His residue positioned 3.01 Å from the active thiol to form a thiolate-imidazolium ion pair (60, 61). Based on the similarity of AdPLA to papain, we propose that His23 assists in the deprotonation of Cys113. The distance from the Nϵ of His23 to the Sγ of Cys113 is greater than that generally observed in other proteins utilizing a Cys-His pair; nevertheless, the role of the His23-Cys113 ion pair in the catalytic mechanism of AdPLA is supported by the abolished activity of the H23A and H23Q mutants (Fig. 6). We cannot rule out that additional residues or the dipole of the α3 helix (62) could also interact with Cys113 and contribute to its reduced pKa to generate a strong enough nucleophile.
To explore the enzymatic activity of AdPLA, we generated an MBP-fused, full-length recombinant form of AdPLA, MBP-FL-AdPLA, which exhibits robust PLA1 and PLA2 activities against natural substrates (Fig. 8). We characterized the activity of MBP-FL-AdPLA using a continuous fluorescence-based assay, and we observed enzyme kinetics consistent with a physiologically relevant PLA enzyme (Fig. 5). The use of a fluorescent substrate containing a non-hydrolyzable bond in the sn-1 position provides additional support for the intrinsic PLA2 activity of AdPLA (Fig. 5). In addition to the residues involved in catalysis, we determined that the transmembrane domain of AdPLA is also required for interfacial catalysis. As a result, MBP-T-AdPLA and T-AdPLA, which are recombinant fragments of AdPLA that lack the transmembrane domain, have no detectable PLA1/PLA2 activity against liposome substrates (Figs. 6 and and88).
We probed the solvent accessibility of backbone amides of MBP-FL-AdPLA, T-AdPLA, and MBP-T-AdPLA using a DXMS approach. The Tyr66–Glu98 domain shows low rates of hydrogen/deuterium exchange consistent with the tight globular packing around the β-sheet β2 (Fig. 7, A and B). On the other hand, the residues that surround the active site of AdPLA display relatively high rates of hydrogen/deuterium exchange (Fig. 7B). Thus, we propose that in the process of catalysis water can readily access the active site of AdPLA and allow the acyl enzyme-thioester intermediate to decompose to free fatty acid and lysophospholipid. In agreement with this mechanism, AdPLA displays a robust PLA2 activity rather than an exclusive acyltransferase activity as observed in the case of LRAT. Analysis of MBP-FL-AdPLA, MBP-T-AdPLA, and T-AdPLA by DXMS also indicates that there are no large changes in conformation between the different forms of the AdPLA protein; thus, it is likely that the structure we report for T-AdPLA resembles the catalytically active MBP-FL-AdPLA protein (Fig. 7), and we attribute the lack of interfacial catalysis by MBP-T-AdPLA and T-AdPLA to the absence of a membrane targeting domain.
The association of AdPLA with membranes is an essential part of the interfacial catalysis process. With the exception of several calcium-independent PLA2s, such as Group VIC PLA2 (patatin-like phospholipase domain-containing 6 (PNPLA6) (63) and patatin-like phospholipase domain-containing 7 (PNPLA7) (64, 65), most PLA2 enzymes characterized so far are soluble. Generally, secreted PLA2 and Group IVA cytosolic PLA2 target the phospholipid bilayer in response to signals, such as binding of calcium. Other PLA2s, like group VI A PLA2 (calcium-independent PLA2β) or lipoprotein-associated PLA2, are targeted to the membrane via hydrophobic pockets (66, 67). In contrast to most PLA2 enzymes, AdPLA is an integral membrane protein. Like the related tazarotene-induced gene 3 (68) and LRAT proteins (69), AdPLA is targeted to membranes via a C-terminal signal anchor domain to assume a “tail-anchored” membrane topology (10, 51). Truncated AdPLA lacking the membrane-spanning C-terminal domain is not active against phospholipids embedded in liposomes (reported here and in Refs. 10 and 14) as it does not associate with membranes (10). By virtue of the targeting signal being the last domain to exit the ribosome, tail-anchored proteins must be targeted to membranes in a post-translational fashion, i.e. after protein translation has been completed (70). A substantial fraction of AdPLA is found to be localized in the cytosol (51). Thus, post-translational translocation of AdPLA could allow AdPLA to be targeted to subcellular compartments that have different lipid pools, such as the endoplasmic reticulum or peroxisomes (22), and play a role in its mechanism of action.
The structure of AdPLA, the first member of the LRAT family to be structurally characterized (model presented here and in Refs. 20 and 35), appears to be related to the papain-like NlpC/P60 bacterial cysteine peptidases that are involved in remodeling of the bacterial cell wall (31). The evolutionary relationship between the LRAT family of proteins and the NlpC/P60 protein superfamily is intriguing and could provide clues to the function of these related enzymes. For example, our results indicate that AdPLA shares the conformation of the Cys-His-His catalytic triad first described in the case of the classical NlpC/P60 protein, Spr, from E. coli (32). AdPLA also resembles the structure of YiiF, an NlpC/P60 permuted domain protein from E. coli whose structure indicates the presence of a bound stearate molecule (71). These observations suggest that eukaryotic representatives of the LRAT family of proteins are related in both structure and enzymatic mechanism to cysteine hydrolases from the NlpC/P60 family that catalyze various acylation/deacylation reactions involving proteins and lipids.
One of the strongest arguments in favor of a role for AdPLA in the generation of free arachidonic acid in fat is the dramatic decrease in the phospholipase A activity (18% of wild type levels) and PGE2 levels (12% of wild type levels) in adipose tissues of AdPLA−/− mice compared with wild type mice (11). In addition, treatment of adipocytes isolated from AdPLA−/− mice with PGE2 can restore lipolysis and cAMP to wild type levels (11). These observations are consistent with a role of AdPLA in the formation of PGE2 in fat whereby AdPLA−/− mice have a higher net rate of adipose tissue lipolysis as a result of decreased prostaglandin E receptor 3 signaling (11). However, the robust PLA1 activity of AdPLA brings into question its role in the release of arachidonic acid as this fatty acid is generally bound to the sn-2, not the sn-1, position of phospholipids.
In addition to PLA1 and PLA2 activities, it was reported that AdPLA also displays N-acylation activity (14, 20), suggesting that it might have a potential role in the formation of N-acylphosphatidylethanolamine, a precursor of endocannabinoids, an important class of endocrine regulators (20). Nevertheless, AdPLA displays very low N-acylation activity (results not shown and Ref. 19) in comparison with the PLA1/PLA2 activities described here and elsewhere (10, 14) that suggest that in a cellular environment fatty acid hydrolysis would occur more often than acyl transfer to phosphatidylethanolamine. However, we cannot exclude the possibility that AdPLA contributes to the synthesis of endocannabinoid precursors in fat or that there might still be other unidentified acyl acceptors that become esterified by AdPLA.
Other products of the AdPLA-mediated hydrolysis besides free fatty acids could have important regulatory roles in fat. Our studies show that AdPLA is a potent combined PLA1/PLA2 enzyme, leading to the release of free fatty acids and the formation of 2- or 1-lysophospholipids, respectively. In our studies, AdPLA displays very low lysophospholipase activity and thus does not further hydrolyze lysophospholipids to generate l-α-glycerylphosphorylcholine (results not shown and Refs. 10 and 14). Lysophospholipids can become precursors to important bioactive lipid mediators, such as lysophosphatidic acid (LPA) through the action of lysophospholipase D/autotaxin (72). LPA can in turn be acylated by either comparative gene identification-58 (CGI-58) or adiponutrin to produce phosphatidic acid (73–75). Because AdPLA, autotaxin, CGI-58, and adiponutrin are highly expressed in adipose tissue (10, 76–79), the contribution of AdPLA to the formation of LPA or phosphatidic acid in adipose tissue is particularly pertinent. Phosphatidic acid is a central lipid signaling mediator in fat as seen in both the resistance to obesity of CGI-58 knockdown mice (80) and the lipodystrophy seen in patients suffering from Chanarin-Dorfman syndrome due to CGI-58 deficiency (81–83). LPA in its own right is a potent inducer of white adipose tissue formation and a negative regulator of thermogenic, brown fat (84). Mice deficient in the LPA receptor LPA1R are resistant to diet-induced obesity (85). Similarly, AdPLA−/− are lean, and the white adipose tissue of AdPLA−/− mice displays increased thermogenesis and fatty acid oxidation (11), which is consistent with the possibility that AdPLA−/− mice have decreased levels of LPA. This is an attractive hypothesis because it offers an explanation for a role of AdPLA in adipose tissue that is independent of the formation of PGE2. This hypothesis also fits with the lack of positional or acyl chain specificity of AdPLA. The role of AdPLA in the generation of LPA could be explored by examining the levels of LPA and the activation of cognate LPA receptors in the tissues of AdPLA−/− and wild type mice.
We solved the structure of AdPLA, leading us to the identification of the Cys-His-His residues involved in the catalytic activity of AdPLA. Our results demonstrate that the Cys-His-His catalytic triad and the signal anchor domain of AdPLA are both required for its function. We describe the purification of a recombinant form of full-length AdPLA, which we characterize as a combined PLA1/PLA2 enzyme. The potent PLA1 activity exhibited by AdPLA suggests that other products of AdPLA-mediated hydrolysis other than arachidonic acid could also play a role in its antilipolytic effects.
Recent developments in the understanding of the physiological role, enzymology, and structure of AdPLA pave the way for the development of inhibitors of AdPLA. Such inhibitors are expected to have a positive impact on the treatment of obesity by promoting adipose tissue lipolysis and mimicking the phenotype of the AdPLA−/− mice. Although this goal makes AdPLA an attractive therapeutic target in obesity settings, an important caveat is that unchecked lipolysis could lead to ectopic lipid accumulation if not compensated by increased fatty acid oxidation. As a consequence, both AdPLA−/−,ob/ob double knock-out mice and AdPLA−/− mice on a high fat diet exhibit hepatic lipid accumulation and resistance to insulin (11). However, milder increases in lipolysis as seen in mice overexpressing adipose triglyceride lipase can improve insulin sensitivity (86). It is possible that pharmacological inhibition of AdPLA will not elicit lipolysis and ectopic lipid deposition to the same extent as in AdPLA−/− mice maintained on a high fat diet, particularly if treatment with AdPLA inhibitors is administered in conjunction with a low fat diet or with therapies aimed at increasing fatty acid oxidation. To achieve this goal and improve upon these first insights into the AdPLA enzymatic mechanism, the structure of AdPLA in complex with substrates or inhibitors is greatly needed. Such findings would shed light on the residues involved in the binding and substrate specificity of AdPLA. Trapping the enzyme-substrate complex would allow us to observe the formation of the presumed tetrahedral intermediate and to define the position of the oxyanion hole necessary to polarize the carbonyl group of the substrate. In parallel with structural approaches, more studies are also needed to explore the physiological role and the consequences of the disruption of AdPLA in vivo.
We thank Drs. Rick Dobrowsky, Hei Sook Sul, Robert Hanzlik, Jeff Staudinger, and Ross Stein for valuable discussions. We thank Sara Billings for advice and help in the preparation of the manuscript. We thank Dan Brown and Amanda Wise for help in cloning and initial assay development and Dr. Fei Philip Gao for help in designing the protein purification strategy. We thank Drs. Marcin Golczak and Krzysztof Palczewski for the gift of the pET30b-HRASLS3 bacterial expression construct. Use of the IMCA-CAT beamline 17-ID at the Advanced Photon Source was supported by the companies of the Industrial Macromolecular Crystallography Association through a contract with the Hauptman-Woodward Medical Research Institute. Use of the Advanced Photon Source was also supported by the United States Department of Energy, Office of Science, Office of Basic Energy Sciences under Contract DE-AC02-06CH11357.
*This work was supported, in whole or in part, by National Institutes of Health Grants 5P20RR017708-10 and 3P20RR017708-07S1, subawards of grants from the National Center for Research Resources (to A. R. M.), Grant 8 P20 GM103420-10) from the NIGMS (to A. R. M.), and Grant R01 GM20501 (to E. A. D.).
This article contains supplemental Figs. S1–S4.
The atomic coordinates and structure factors (code 4FA0) have been deposited in the Protein Data Bank, Research Collaboratory for Structural Bioinformatics, Rutgers University, New Brunswick, NJ (http://www.rcsb.org/).
3The abbreviations used are: