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Lipids play critical roles in several major chronic diseases of our times, including those that involve inflammatory sequelae such as metabolic syndrome including obesity, insulin sensitivity, and cardiovascular diseases. However, defining the substrate specificity of enzymes of lipid metabolism is a challenging task. For example, phospholipase A2 (PLA2) enzymes constitute a superfamily of degradative, biosynthetic, and signaling enzymes that all act stereospecifically to hydrolyze and release the fatty acids of membrane phospholipids. This review focuses on how membranes interact allosterically with enzymes to regulate cell signaling and metabolic pathways leading to inflammation and other diseases. Our group has developed “substrate lipidomics” to quantify the substrate phospholipid specificity of each PLA2 and coupled this with molecular dynamics simulations to reveal that enzyme specificity is linked to specific hydrophobic binding subsites for membrane phospholipid substrates. We have also defined unexpected headgroup and acyl chain specificity for each of the major human PLA2 enzymes, which explains the observed specificity at a structural level. Finally, we discovered that a unique hydrophobic binding site—and not each enzyme’s catalytic residues or polar headgroup binding site—predominantly determines enzyme specificity. We also discuss how PLA2s release specific fatty acids after allosteric enzyme association with membranes and extraction of the phospholipid substrate, which can be blocked by stereospecific inhibitors. After decades of work, we can now correlate PLA2 specificity and inhibition potency with molecular structure and physiological function.
The ASBMB 2021 Bert and Natalie Vallee Award in Biomedical Sciences honors our work on how phospholipase A2 (PLA2) acts on substrate in membranes and micelles. Lipids play critical roles in the metabolic syndrome including obesity, insulin sensitivity and type 2 diabetes, numerous cardiovascular diseases, and fatty liver disease including nonalcoholic steatohepatitis; these constitute the major chronic diseases of our times, and all of these involve sequelae of inflammation. Over 47,000 distinct molecular species of lipids have been identified by the LIPID MAPS Consortium (www.lipidmaps.org), so defining substrate specificity of enzymes of lipid metabolism is a challenging task.
). We have recently employed “substrate lipidomics” coupled with molecular dynamics (MD) to reveal enzyme specificity linked to highly specific hydrophobic binding sites or “subsites” for the sn-2 fatty acyl chains in membrane phospholipid substrates (
). We discovered unexpected headgroup and acyl chain specificity for each of the major human phospholipase A2 enzymes that explains the observed specificity at a new atomic level. A unique hydrophobic binding site—and not each enzyme’s catalytic residues or polar headgroup binding site—dominates each enzyme’s specificity. Each PLA2 shows unique specificity for its required fatty acid ranging from proinflammatory omega-6 arachidonic acid (AA) or anti-inflammatory fish oil omega-3 eicosapentenoic acid and docosahexenoic acid; others favor membrane remodeling linoleic acid, antibacterial saturated fatty acids, or oxidized fatty acids in low-density lipoproteins. Each PLA2 releases a specific fatty acid after the enzyme associates with membranes causing an allosteric effect and extracts a single phospholipid substrate into its catalytic site. Stereospecific inhibitors (
) have been developed for the specific sites of each enzyme and studied kinetically. After decades of advances in lipid research, we can now correlate PLA2 specificity and inhibition potency with molecular structure and physiological function using a novel lipidomics platform that provides a paradigm for protein–membrane lipid interactions in general.
PLA2s constitute a superfamily of enzymes: some are degradative, some are biosynthetic, and others are signaling enzymes (Fig. 1) (
). All of the PLA2s act stereospecifically to hydrolyze the fatty acid at the middle or sn-2 position of membrane phospholipids. When they produce as products lysophospholipids and free fatty acids, they are degradative enzymes. When the enzyme is coupled to an acyltransferase that specifically puts a polyunsaturated fatty acid back in that sn-2 position to make a new phospholipid that has been remodeled, then they “in effect” become biosynthetic enzymes. When the products are converted by other enzymes to ligands that activate G protein–coupled receptors such as lyso-phosphatidic acid or PGE2, then they are considered signaling enzymes.
The Group Numbering System was originally developed to differentiate the PLA2s that had been described at the time (
) as more unique PLA2s were discovered, so it roughly reflects the order in which the different PLA2s were discovered. Today, there are over 16 Groups and many Subgroups that are comprised of distinct PLA2s (
). For simplicity, the PLA2s are sometimes referred to by general names as belonging to one of six Types using their more generic names (also commonly listed in the order that they were characterized): secreted phospholipase A2 (sPLA2); cytosolic phospholipase A2 (cPLA2); calcium-independent phospholipase A2 (iPLA2); platelet-activating factor (PAF) acetyl hydrolase, also known as lipoprotein-associated PLA2 (Lp-PLA2), lysosomal PLA2, and adipose PLA2, but if one wants to be specific about the particular PLA2, the designation by Type would then be preceeded by the Group Number (
). Numerous laboratories around the world, but particularly in Japan, France, the Netherlands, and the United States, contributed to this early development, and complete references to the initial discovery of each of these Types and authoritative reviews by others on the major Groups and Types are listed elsewhere (
It should be noted that alternate naming systems have been used by many laboratories including first listing the Type followed by a Greek letter to indicate the specific Group/Subgroup (i.e., most commonly cPLA2α instead of Group IVA (GIVA) and iPLA2β instead of Group VIA). Another naming system was developed independently for iPLA2 which describes enzymes based on sequence similarities rather than on catalytic substrate similarities based on a classic well-studied potato (patatin) triacylglycerol lipase resulting in “patatin-like PLA2” (PNPLA) for which Group VIA iPLA2 is known as PNPLA9. Other PNPLA enzymes actually exhibit different activities than PLA2, for example, PNPLA3 is actually a triacylglycerol lipase rather than a PLA2, and there is enormous interest in this enzyme currently because of its association with nonalcoholic fatty liver disease and its advanced form nonalcoholic steatohepatitis (
This review of recent work in our laboratory will focus on the four major human phospholipase A2Types studied, but each of these four are the major studied human form in their Group and have been among the most well-studied and well-characterized examples. They are all highly purified recombinant human enzymes that we study in vitro in our laboratory. The four enzymes and their subcellular localization are depicted in Figure 2 (
). The first is the GIVA cPLA2 which associates with the Golgi and specifically releases AA, a proinflammatory fatty acid that leads to inflammation and has a major role in signaling. Another PLA2 that is localized in the cytosol, but is sometimes associated with mitochondria, is the Group VIA iPLA2; it releases unsaturated fatty acids and is heavily involved in membrane remodeling and implicated in mitochondrial functioning.
As has been noted, there are multiple forms (designated as Groups and Subgroups) of each of the four Types of PLA2 discussed here, but the secreted PLA2Type includes the largest number of well-studied distinct Groups and Subgroups. The first PLA2s to be characterized were secreted enzymes from various snake venoms and porcine pancreas, and the human Group IIA, which was originally cloned from human synovial fluid, is perhaps the most well studied (
). The secreted enzymes are formed in the Golgi and are secreted to the outside of cells where they act to release both saturated and unsaturated fatty acids. It has also been suggested that sometimes they undergo endocytosis and may act intracellularly as well. In this review, we have limited ourselves to discussing the Group V sPLA2 because among the sPLA2s, it has been the focus in our laboratory. Interestingly, it is secreted by macrophage cells (which also express the intracellular GIVA cPLA2 and Group VIA iPLA2), and macrophages additionally secrete the Lp-PLA2. The Group VIIA Lp-PLA2 is the fourth enzyme Type we have studied extensively. It is known as the secreted Lp-PLA2, and it associates with low-density lipoproteins and high-density lipoproteins in the blood stream and specifically releases only oxidized or short-chain fatty acids. This enzyme was independently discovered and named as PAF acetyl hydrolase because of its potent ability to hydrolyze PAF releasing acetate from the sn-2 position (
) that if we are going to study PLA2s, and at that time we were focused on the secreted PLA2 purified from cobra venom (Naja naja naja), that since it was water soluble, it is going to interact with a phospholipid substrate which resides only in membranes or micelles and not freely in solution. Thus, the enzyme must first associate with the lipid aggregate and the first step for a phospholipase to act requires that the water-soluble enzyme associate with the membrane or the micelle. In Figure 3, the lipid–water interface is pictured as a mixed micelle with phospholipids in red and nonionic surfactant Triton-X100 molecules in yellow, though the same principle holds for bilayer membranes or phospholipid vesicles as well as the monolayer surface of lipoprotein particles and lipid droplets.
The four PLA2 enzymes we will discuss herein are all water soluble. The first step depends on the concentration of enzyme and membranes or micelles. Note that the phospholipids in red laterally diffuse around the surface of the micelle or membrane very rapidly until a single phospholipid is “sucked” into the catalytic site. Catalysis occurs, and the products diffuse back around the surface of the micelle or membrane. The second step occurs when the enzyme is associated with the surface, and this step depends kinetically on the surface concentration of the specific phospholipid substrate in the surface of the micelle. This realization led our group to propose “surface dilution kinetics” (
) to analyze water-soluble enzymes acting on substrates localized to the lipid–water interface. This conceptual approach can be applied to other proteins that associate with membranes and especially to other enzymes of lipid metabolism (
) and seems equally applicable to substrates located on the outer surface of bilayer membranes, micelles, lipoproteins, lipid droplets, etc.
Deuterium exchange mass spectrometry applied to protein–membrane and protein–substrate interactions
About a decade ago, our laboratory proposed using the technique of deuterium exchange mass spectrometry (DXMS) to look specifically for the first time at the interaction of proteins with individual phospholipid substrates and their interactions with the membrane (
Quite simply, one takes the PLA2 in aqueous solution, then mixes it with phospholipid vesicles, allows them to associate, dilutes in D2O, and then measures the rate of deuterium exchange with the amide protons in the amino acids in separate peptides in the polypeptide backbone of the enzyme. In essence, where the enzyme interacts with the membrane or where a phospholipid is pulled up into the enzyme active site, the rate of deuterium exchange is changed, both by proximity and by conformational changes that can be quite complicated. Peptides show increases and others show decreases in deuterium exchange depending on whether that portion of the enzyme is more exposed or less exposed to solvent. However, decreases are especially noteworthy when they result from decreased accessibility due to hydrophobic interactions (
) shown in Figure 4 that applies to all four of the enzymes under discussion. Quite simply, the water-soluble enzyme, when mixed with a phospholipid vesicle, causes a conformational change in the enzyme indicated by a square to a circle. Note that this very large membrane is antithetical to the original proposal of allosterism by Monod, Wyman, and Changeux (
), which was pictured as a small allosteric site, for example, with O2 binding to other subunits in tetrameric hemoglobin or perhaps a Ca2+ binding to another site causing a conformation change in a large enzyme. In the case of membranes, it is a very large membrane that causes a conformational change in a relatively much smaller protein as illustrated in Figure 4.
Once associated, the phospholipid molecules move very rapidly by lateral diffusion around the surface of the membrane until a single one is “sucked” into the catalytic site as shown, catalysis occurs, the products diffuse back in the membrane, and the cycle keeps repeating itself. We have suggested that this model is a general model applicable to all four of the PLA2 enzymes discussed herein and perhaps other membrane-associated enzymes as well.
In Figure 5, the catalytic domain of human GIVA cPLA2 was placed in a cube of water and a substrate sn-1 palmitoyl, sn-2 arachidonyl phosphatidylcholine (PAPC) molecule in black was docked in its active site and the cPLA2 was docked to a membrane patch as suggested by the DXMS results. The membrane patch consisted of a full sn-1 palmitoyl, sn-2 oleoyl phosphatidylcholine (POPC) bilayer membrane in purple (although only a slice of the membrane is shown, so the enzyme and substrate can be seen clearly). MD simulations were carried out for 300 ns. See Movie S1.
The first thing one notices is how rapidly the purple phospholipids in the membrane move with lateral diffusion, but basically staying oriented. In contrast, the phospholipid in the catalytic site is much more restricted in motion as is the whole protein as it struggles to find the minimal energy conformation of both the phospholipid ligand and the enzyme. The AA is seen in its characteristic curved form in its final conformation with its four cis double bonds interacting primarily with aromatic amino acid side chains with π-π stacking.
In a separate experiment, the PAPC substrate in its final energy minimized docked conformation was pulled out of the active site along a straight trajectory back into the membrane, and then a slight force was applied in the opposite direction to allow this phospholipid to re-enter the active site. This allows one to visualize how the enzyme, docked to a membrane, might pull a single phospholipid out of the membrane and into the catalytic site. See Figure 6 and Movie S2. Note that the red helix in the protein houses the Ser–Asp dyad that is the catalytic machinery and the Ser is right adjacent to the sn-2 carbonyl position oriented to initiate the catalytic event.
The human Group VIA iPLA2 is a very different enzyme, though it has the same Ser–Asp dyad as its catalytic machinery. In Figure 7, we have docked the catalytic domain of iPLA2 with the same PAPC substrate as employed with cPLA2 in its catalytic site as well as the same POPC membrane patch and then carried out MD simulations for 300 ns. See Movie S3. The active site of iPLA2 is a little bit more of an open site than cPLA2, and in the middle of the simulation, one observes a conformational change whereby the enzyme opens up to accommodate the sn-2 fatty acid chain, in this case AA, in a distinct subsite that binds very specifically to the sn-2 fatty acid. The sn-1 fatty acid, palmitate in this case, is less restricted, and it appears to extend some into the membrane patch. Note that the red helix on the enzyme is buried in the membrane, so one might visualize the membrane as allowing the tail of the sn-1 chain to protrude into the membrane, whereas the sn-2 fatty acid, which is the leaving or cleaved fatty acid, is bound entirely in its sn-2 subsite.
Lipidomics and MD simulations
We recently developed a new “substrate lipidomics” assay for PLA2 which allows us to look at any natural or synthetic phospholipid alone or in mixtures, rather than relying on the traditional specifically radioactively labeled phospholipids (
). Figure 8A shows the results of a cPLA2 assay of an equimolar mixture of five phospholipids, each with the same sn-1 palmitic acid and sn-2 AA, but varying in their polar group including zwitterionic and anionic head groups, in mixed micelles with a nonionic surfactant. The specific activity of the enzyme is the same toward all five substrates within experimental error. This observation was counterintuitive because previously the phospholipases were often identified with the kind of phospholipid that they hydrolyze based on the polar group. However, here it seems that the specificity is not with the polar group, but rather it is dictated by the specific acyl chain in the sn-2 position that is the leaving group.
This is shown in Figure 8B where for cPLA2, the best fatty acid on the sn-2 position is AA by far. However, when we compared the same fatty acids with iPLA2 in Figure 8C, linoleic acid was by far the best substrate and while the enzyme worked somewhat on AA, it was poorer than on linoleic acid.
This can be explained if we look at the results of 1-μs MD simulations and the conformation of the phospholipids in the catalytic site. Shown in Figure 9A is cPLA2 with 1-palmitoyl, 2-arachidonyl phosphatidylethanolamine (PAPE) where the polar group is surrounded by charged and polar amino acids (
). That is why the active site accommodates all the different polar groups so well and apparently almost equally. But the specificity is in the arachidonyl group in the sn-2 position, where it has its four double bonds that are all cis, arranged such that they interact and cause great π-π stacking with the aromatic side chains that are pictured in this catalytic site. In stark contrast as shown in Figure 9B, for iPLA2 with the same PAPE substrate, the AA is found bound with some specificity, but in a “scrunched” or not-ideal “curved” conformation. For both enzymes, the sn-1 palmitic acid is found localized in a distinct subsite.
This can be understood and explained more clearly if the association of these two enzymes with the phospholipid substrate sn-1 palmitoyl, sn-2 linoleoyl phosphatidylcholine (PLPC) containing a sn-2 linoleic acid is compared. Figure 9D shows the same subsite for iPLA2 with linoleic acid in the phospholipid chain, and the two cis double bonds in linoleic acid are aligned very nicely with aromatic tyrosine side chains and perfect π-π stacking, explaining why it binds even better than AA.
However, when the same PLPC phospholipid is docked in the active site of cPLA2 in the same manner that PAPE was docked and the 1-μs simulation is initiated, the phospholipid immediately diffused from the active site and moved into the membrane. See Figure 9C. In other words, the difference in having two less carbons and two less double bonds in linoleic acid (18:2) than AA (20:4) was enough to reduce the affinity of this fatty acid chain dramatically so that it no longer could bind over any reasonable time period within the active site.
Secreted and lipoprotein-associated PLA2s
Our laboratory has used similar approaches to study the human Group V secreted sPLA2, which is a much smaller 13-kDa protein containing seven disulfide bonds and which utilizes a His–Asp for hydrolysis along with a required Ca2+. Its specificity favors linoleic acid in its sn-2 position like iPLA2 (
), and most of the sn-1 fatty acyl chain remains associated with the membrane. However, despite its differences from the two cytosolic enzymes discussed earlier in the study, it appears to follow the same general characteristics in its interaction with substrate phospholipid and membranes (
). Current work in our laboratory is aimed at more fully characterizing the unique specificity of the secreted enzyme for both the sn-1 and sn-2 fatty acyl groups and comparing it in more detail with that of the c- and i-PLA2s.
DXMS has shown that the human Group VIIA lipoprotein-associated PLA2 undergoes a conformational change when it associates with phospholipid vesicles and many additional changes when associated with human high-density lipoproteins (
). This enzyme does not hydrolyze “normal” saturated, unsaturated, or polyunsaturated fatty acids but rather has great specificity for very-short-chain fatty acids in the sn-2 position such as acetate (in PAF) and oxidized fatty acids such as oxovaleryl phosphatidylcholine. However, this 45-kDa extracellular enzyme utilizes a catalytic triad consisting of Ser His Asp in carry out hydrolysis. Current work in our laboratory is aimed at more fully characterizing the unique specificity of this enzyme for the sn-2 fatty acyl groups.
MD simulations reveals specificity of inhibitors for PLA2s based on the sn-2 subsite
Our laboratory has spent considerable effort over the years designing and studying a variety of inhibitors with specificity for each of the PLA2s, often in collaboration with Professor George Kokotos from Athens. Refer to the study by Dennis et al. (
), namely pyrrophenone, which is a widely used pyrrolidine GIVA cPLA2 inhibitor; octylthiotrifluorophosphonate, which is a Group VIA iPLA2 inhibitor; and Ly315920, which is an indole developed as a Group IIA sPLA2 inhibitor, but which also potently inhibits Group V sPLA2.This study showed that the lipidomics assay works extremely well and enlarges enormously the range of phospholipid substrates available to replace the traditional radioactive-based PLA2 assay on PAPC that was used over the years, but for which commercial sources of substrate are no longer available.
Earlier, we extended our DXMS coupled with MD simulations to study PLA2 inhibitors (
). With cPLA2, we examined the then best traditional inhibitor pyrrophenone and a designed substrate analogue oxoamide and concluded that the oxoamide binds in the substrate site with the polar moiety in the polar subsite and the alkyl chain in the sn-2 subsite, but the pyrrophenone bound more distal from the active site (
). However, with iPLA2, we discovered that for an aromatic trifluoroketone, the trifluoroketone moiety bound in the polar subsite, but the aromatic group bound in a distinct region where it was in π-π contacts with the enzyme’s aromatic sidechains, whereas an alkyl trifluoroketone bound in an adjacent region in contact with the enzyme’s aliphatic sidechains (
Computer-aided drug design guided by hydrogen/deuterium exchange mass spectrometry: A powerful combination for the development of potent and selective inhibitors of group VIA calcium-independent phospholipase A2.
). We postulated that these two inhibitors bound in distinctive parts of the hydrophobic acyl chain sites differently if they were analogues of saturated or unsaturated chains. In our recent work on substrate specificity (
) described in Figure 9D, where the optimal binding of an sn-2 linoleic acid (L) in PLPC was described in close contacts with the enzyme’s aromatic side chains, we had also observed that an sn-2 myristic acid (M) in sn-1 palmitoyl, sn-2 myristoyl phosphatidylcholine bound with close contacts in a distinct region of the same sn-2 subsite surrounded entirely by aliphatic sidechains in the enzyme active site. In hindsight, the two fluroketone inhibitors (aromatic and alkyl) discriminated these two parts of the sn-2 subsite of iPLA2 (
) in the same manner as the two best substrates for iPLA2 did.
In conclusion, we have summarized the results of DXMS, MD simulations, and lipidomics analysis reported in several recent papers that have led us to suggest that a water-soluble enzyme associates with membranes, micelles, or other lipid–water interfaces, whereby the membrane causes a conformational change in the enzyme, and aided by rapid lateral diffusion of the phospholipids in the surface of the membrane, a single phospholipid is sucked up into the catalytic site. Furthermore, each enzyme has a very well-defined and unique sn-2 leaving fatty acid subsite. This hydrophobic sn-2 subsite differs dramatically for each PLA2 and affords the unique leaving group specificity to it. This subsite results in dramatically different specificity of each PLA2. In contrast, the polar site, which is rather similar for each enzyme in terms of predominately charged and polar sidechain amino acids, has less specificity.
Furthermore, the sn-1 subsite has to bind saturated fatty acid sidechains mainly consisting of palmitic acid (16:0) and stearic acid (18:0) for which the length of the fatty acid chain is the only difference, though sometimes the sn-1 position is occupied by monounsaturated and polyunsaturated fatty acids. For some of the enzymes, particularly for sPLA2, this is a rather shallow site and most of the fatty acid chain resides in the membrane. This concept is consistent with the overall finding that potent PLA2 inhibitors exhibit Type and Group specificity based on the uniqueness of the hydrophobic site at the sn-2 position, and this is where catalysis occurs for PLA2s. For all of these enzymes, it appears that as soon as catalysis occurs, the products diffuse into the membrane and the cycle repeats itself as indicated in Figure 4. In short, this manuscript has summarized the evidence that membranes allosterically activate enzymes, that lipidomics can identify substrate specificity, and that a specific sn-2 hydrophobic subsite in each enzyme determines its unique specificity.
The authors declare that they have no conflicts of interest with the contents of this article.
I wish to thank all of my students, coworkers, and collaborators who contributed so much to our studies on phospholipase A2 over the past 50 years, but especially a former postdoctoral fellow and project scientist Varnavas Mouchlis, many of whose contributions are highlighted in this manuscript. I also wish to thank my long-term colleagues in my laboratory professor Oswald Quehenberger and Aaron Armando for their leadership in our UCSD LIPID MAPS LIPIDOMICS CORE and advice on the lipidomics and mass spectrometric aspects of the recent work discussed herein. I am also very appreciative of the late professor Virgil Woods and Sheng Li (UC San Diego) who collaborated on our first use of deuterium exchange mass spectrometric approaches, professor Andrew McCammon’s (UC San Diego) collaboration and advice on our molecular dynamics studies over many years, professor George Kokotos’ (University of Athens) collaboration on creating specific phospholipase inhibitors, and valuable conversations with professor George Carman (Rutgers) on lipid enzymes and professor Jean-Pierre Changeux (Collège de France and Institut Pasteur) on the concept of allosterism applied to membranes. I also wish to thank Milda Simonaitis and Jenny Truong for their help with this manuscript.
I knew Bert Vallee as a professor at Harvard Medical School and admired his approach to mechanistic enzymology, especially exemplified by his work on alcohol dehydrogenase. I am grateful to the nominators and the ASBMB leadership for my selection as the recipient of the 2020/2021 Bert and Natalie Vallee Award for basic biomedical research and to the Vallee Foundation for its support of the annual award.
Funding and additional information
Our laboratory’s work on phospholipase A2 was supported by NIH’s National Institute of General Medical Sciences over these past 45 years via RO1-GM20501-44 and its recent renewal as a MIRA R35 GM139641-01 . The content is solely the responsibility of the author and does not necessarily represent the official views of the National Institutes of Health.
MD simulation of GIVA cPLA2with PAPC and POPC membrane. Molecular dynamics simulation of GIVA cPLA2 with PAPC docked in the active site and associated with a POPC bilayer membrane (purple) in accord with the results of deuterium exchange mass spectrometry. Note for visual clarity only a small number of the membrane POPC molecules is shown. Adapted from the study by Mouchlis et al. (
SMD simulation of GIVA cPLA2extracting PAPC into the active site. After the optimal binding mode of PAPC in the active site of GIVA cPLA2 was determined by molecular dynamics simulations as illustrated in Movie S1, the PAPC was pulled out of the active site along a trajectory toward the POPC bilayer membrane where it would have initially been associated in accord with the results of deuterium exchange mass spectrometry. The movie illustrates how the PAPC reenters the active site along that same trajectory when subjected to a small force in the reverse direction. Adapted from the study by Mouchlis et al. (
MD simulation of GVIA iPLA2with PAPC and POPC membrane. Molecular dynamics simulation of GVIA iPLA2 with PAPC docked in the active site and associated with a POPC bilayer membrane in accord with the results of deuterium exchange mass spectrometry. Note for clarity in observing the conformational change that iPLA2 exhibits during the 300-ns simulation, the membrane POPC molecules are not shown. Adapted from the study by Mouchlis et al. (
Computer-aided drug design guided by hydrogen/deuterium exchange mass spectrometry: A powerful combination for the development of potent and selective inhibitors of group VIA calcium-independent phospholipase A2.