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. 2025 Jun 9;16(24):6059–6065. doi: 10.1021/acs.jpclett.5c00860

Conformational Dynamics and Activation of Membrane-Associated Human Group IVA Cytosolic Phospholipase A2 (cPLA2)

Mac Kevin E Braza †, Edward A Dennis †,‡,*, Rommie E Amaro §,*
PMCID: PMC12183713  PMID: 40489676

Abstract

Cytosolic phospholipase A2 (cPLA2) associates with membranes, where it hydrolyzes phospholipids containing arachidonic acid to initiate an inflammatory cascade. All-atom molecular dynamics simulations were employed to understand the activation process when cPLA2 associates with the endoplasmic reticulum (ER) membrane of macrophages, where it acts. We found that membrane association causes the lid region of cPLA2 to undergo a closed-to-open state transition that is accompanied by the sideways movement of loop 495–540, allowing the exposure of a cluster of lysine residues (K488, K541, K543, and K544), which are known to bind allosteric activator PIP2 from the membrane. The active site of the open form of cPLA2, containing catalytic dyad residues S228 and D549, exhibited a 3-fold larger cavity than the closed form of cPLA2 in aqueous solution. These findings provide mechanistic insight into how cPLA2–ER membrane association promotes major transitions between conformational states critical to allosteric activation and enzymatic phospholipid hydrolysis.

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The interaction between cytosolic proteins and bilayer membranes of intracellular organelles is central to biological function and signaling. Integrating cellular, biochemical, and structural information is of the utmost importance in elucidating protein–membrane interactions at the atomistic level. , In the current study, we investigated how human cytosolic phospholipase A2 (cPLA2), a critical enzyme in the inflammatory cascade, associates with intracellular endoplasmic reticulum (ER) membranes, where it undergoes a critical state transition. The presence of known allosteric regulators such as membrane association, Ca2+, and phosphatidylinositol 4,5-bisphosphate (PIP2) was also elucidated.

Phospholipase A2 (PLA2) constitutes a superfamily of enzymes that hydrolyze the fatty acyl group occupying the sn-2 position of membrane phospholipids. Group IVA (GIVA) cPLA2 exhibits specificity for phospholipids containing arachidonic acid (AA) at the sn-2 position. This specificity can be explained at the molecular level, where several aromatic amino acid side chains form a precise binding site cavity for AA. AA consists of 20 carbons and four cis double bonds that exhibit π–π stacking with the aromatic amino acid side chains in the binding site. At the cellular level, cPLA2 is a water-soluble protein that associates with the surface of the phospholipid bilayer. Once cPLA2 pulls a single phospholipid substrate into its active site, it hydrolyzes and releases it, whereby downstream enzymes bind the AA and initiate the inflammatory cascade. To date, there is only one crystal structure of group IVA cPLA2. It contains both a C2 and a catalytic domain, but a mechanistic understanding of the synergism between these two domains is lacking. Previous studies using both classical and steered molecular dynamics (MD) simulations revealed that the lid region of the membrane-associated cPLA2 was highly flexible before the substrate binding of 1-palmitoyl-2-arachidonoyl-sn-glycero-3-phosphocholine (PAPC or 16:0/20:4 PC).

The Monod–Wyman–Changeux (MWC) model describes allosteric regulation, in which distant ligand binding drives cooperative conformational changes. In the classical MWC allosteric protein nicotinic acetylcholine receptor (nAChR), Ca2+ modulates acetylcholine binding at distinct sites. , Similarly, cPLA2 is allosterically regulated by Ca2+ (∼30 Å from the active site), with PIP2 and membranes also acting as allosteric regulators, as shown by the Dennis group. ,

Hydrogen/deuterium exchange-mass spectrometry (HDX-MS) data for proteins provide qualitative analysis by mapping the average exchange rates of peptides under varying conditions. − Although HDX-MS can be used as an indirect probe of protein allostery and dynamics, it can be limiting. MD simulations help to augment and extend the available HDX-MS, structural, dynamical, and biological data to provide an atomically detailed mechanism of protein dynamics. Furthermore, MD simulations can elucidate allosteric processes under different model types and conformational landscape shifting. ,

In the present study, we carried out all-atom molecular dynamics simulations of aqueous and membrane-associated cPLA2 to complement findings from in vivo, lipidomics, and HDX-MS data. ,− Furthermore, we analyzed the conformational dynamics induced by the three allosteric regulators, namely, membrane association, Ca2+, and PIP2. The detailed computational methods are described in the Supporting Information. In brief, we analyzed three replicates of 500 ns explicit solvent classical MD trajectories for each system, gathering a combined 7.5 μs data set (Table ). In summary, the five systems we simulated were (1) aqueous cPLA2 with bound Ca2+, (2) membrane-associated cPLA2, (3) membrane-associated cPLA2 with Ca2+, (4) membrane-associated cPLA2 with PIP2, and (5) membrane-associated cPLA2 with Ca2+ and PIP2. Furthermore, we describe various conformational dynamics of the C2 and catalytic domains, lid region opening, and loop 495–540.

1. cPLA2–Membrane Systems Employed in the Present Study .

      no. of lipids
    
system Ca2+ 1% PIP2 upper leaflet lower leaflet total no. of atoms simulation length
1 + – – – 424 425 3 replicates × 500 ns
2 – – 400 400 495 230 3 replicates × 500 ns
3 + – 400 400 495 366 3 replicates × 500 ns
4 – + 400 400 495 388 3 replicates × 500 ns
5 + + 400 400 495 110 3 replicates × 500 ns
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a

All systems were simulated using the CHARMM36 force field and built using the CHARMM-GUI Membrane Builder. Membrane compositions were derived from Andreyev et al. Na+ and Cl– ions at 150 mM were added. cPLA2 histidine protonation states were set based on the PROPKA output. The orientation of cPLA2 with respect to the membrane was consistent with the OPM and HDX-MS data.

We built the RAW264.7 macrophage ER membrane environment with its associated cPLA2 (Figure ). The orientation of cPLA2 with the membrane is consistent with the predicted placement from the Orientations of Proteins in Membranes (OPM) database. Furthermore, this is consistent with all previous studies, especially the available cPLA2 HDX-MS data. ,,,− The ER has been reported to be composed of glycerophospholipids, sphingolipids, and sterols at varying concentrations. We used the subcellular lipidome data reported for the ER of RAW264.7 macrophages. Furthermore, it has been reported that PI is the most abundant 20:4-containing glycerophospholipid in RAW264.7 macrophages. , In the most recent study of Murawska et al., they verified that phospholipid 1-stearoyl-2-arachidonoylphosphatidylinositol (SAPI or 18:0/20:4 PI) is the most specific substrate for cPLA2 in the RAW macrophages. The ER membrane contains 10% SAPI. The CHARMM-GUI membrane builder was used to embed cPLA2 in the membrane and build the extensive ER membrane. Although we recognized that the upper and lower leaflets of the ER membrane are probably asymmetric, for our purposes, symmetric membranes were employed in these studies. The exact composition of the lipids employed in the models is reported (Tables S1 and S2).

1.

1

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Human cytosolic phospholipase A2 (cPLA2) and RAW264.7 macrophage ER membrane interactions. (A) Cellular environment of cPLA2. (B) Phospholipid active site catalytic dyad residues (S228 and D549) and allosteric site residues K488, K541, K543, and K544. The lid region is also highlighted (magenta). (C) The membrane lipid bilayer model was assumed in this study, with the exact lipid composition listed in Tables S1 and S2. (D) cPLA2–membrane association and dissociation processes.

We simulated cPLA2 with and without bound Ca2+ in the C2 domain. The C2 domain is composed of an eight-stranded antiparallel β-sandwich. The changes in the RMSF of the several secondary structures at the C2 domain (Figure A,B) were measured. The RMSF quantifies the average deviation of the cPLA2 Cα atom from its mean position during the simulations. Our simulations agree with data for Ca2+ binding at the C2 domain, which rigidifies, stabilizes, and activates cPLA2. The association of the C2 domain with the phospholipid membrane requires Ca2+.

2.

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Effect of Ca2+ and PIP2 on the conformational dynamics of the C2 domain, lid region, and loop 495–540 of cPLA2. (A) Superimposed C2 domain of the system without and with Ca2+. (B) RMSF of β sheets 4–7 (right). The RMSF for the other systems is also shown in Figure S6. The RMSF for the Cα atom was measured from the n = 3 replicates of 500 ns all-atom MD simulations. (C) Superimposed lid region residues 412–457. (D) Superimposed loop region residues 495–540. The top clustered conformation of aqueous cPLA2 (green), cPLA2 with Ca2+ without PIP2 (purple), and cPLA2 with Ca2+ and PIP2 (cyan). Closed, semi-open, and fully open states for the lid region and loop 495–540 are shown.

Ca2+ binding neutralizes the negative electrostatic potential at the loop and β-sheet residues. The hydrophobic interactions between the membrane and the C2 domain allow the association of the now neutral C2 environment due to the lower free energy cost of inserting charged amino acids into the membrane. Furthermore, an increased Ca2+ concentration signals the translocation of cPLA2 from the cytosol to the plasma membrane, Golgi, and ER. We further investigated how the catalytic domain also recognizes the membrane surface.

Our other goal was to understand how the well-known allosteric regulator phosphatidylinositol 4,5-bisphosphate (PIP2) affects the activation process. PIP2 is predominantly concentrated in the plasma membrane, which is a critical regulator of cell signaling. However, a small yet measurable fraction (∼1%) exists in the endoplasmic reticulum (ER), not as a signaling hub but as an intermediate in lipid metabolism. The ER primarily functions as a lipid-processing center, supplying PI for PIP2 biosynthesis at the plasma membrane, thereby maintaining the dynamic equilibrium that is essential for cellular signaling fidelity. In our MD simulations, PIP2 continuously diffuses laterally in the membrane (Figures S1 and S2).

Loop 495–540 exhibits reduced RMSF values when PIP2 is present in the upper leaflet, as compared to the system that does not contain PIP2 (Figure S4B). We note, however, that PIP2 binding is not observed in our 500 ns long simulations. Moreover, this implies that PIP2 remained resistant to extraction into the catalytic site of cPLA2, likely due to its highly negative polarity and the intrinsic time scale of lipid binding exceeding the 500 ns simulation time scale. It is not a good substrate for cPLA2. This suggests that while the portion of the PIP2 in the membrane may associate with the PIP2 allosteric sites, this was not observed but would require extended simulation times to overcome the slower, collective lipid reorganization processes that are essential for binding.

The lid region samples multiple conformations that can interconvert among the closed, semi-open, and open conformations. HDX-MS was used to probe how the different parts of cPLA2, including the lid region, interact with the membrane. , Through a series of HDX-MS experiments, the lid region was found to be a flexible regulated structure with a particular orientation to the membrane. After further analysis of the structural and conformational dynamics of different cPLA2 systems, we found that the lid region and loop 495–540 showed large fluctuations throughout the simulations (Figure C,D). It should be noted that part of the lid region (433–459) is not determined in the X-ray structure. However, after running multiple replicates of 500 ns equilibrium MD, the lid region is stabilized, as measured by the protein backbone RMSD (Figure S5).

After further investigation, we checked the lid region and noted several key differences at the C-terminal end (residues 435–457) (Figure ). First, we overlapped the highest clustered conformation of aqueous cPLA2 and fully activated cPLA2 (with Ca2+ and PIP2) (Figure A). The lid region (residues 412–457) is composed of two α-helices (α-helices 16 and 17) and a loop (residues 435–457). The helices are expected to be more stable than the loop. Fully activated form cPLA2 (with Ca2+ and PIP2) has the most stable loop (residues 435–457). Interestingly, loop 435–457 fluctuates up to 2-fold more in an aqueous environment (Figure B). Second, membrane association stabilized these fluctuations (Figure A,B). The fully activated form of cPLA2 that has bound Ca2+ with accessible PIP2 in the membrane was found to have the smallest RMSF, while aqueous cPLA2 has the largest RMSF in the 435–457 region (Figure B). Third, we noted several other conformational states of the lid region (Figure C–F) that we classified as closed, semi-open, and fully open states. Aqueous cPLA2 is in a closed state, while the fully activated form of cPLA2 that has bound Ca2+ and accessible PIP2 is in a fully open state.

3.

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Lid region dynamics. (A) Lid region conformational dynamics after membrane association. (B) RMSF of α-helix 16, α-helix 17, and loop 435–457. RMSFs for the Cα atoms were measured from the combined n = 3 replicates of 500 ns all-atom MD simulations. Superimposed top clustered conformer of each system: (C) water vs +Ca2+/+PIP2, (D) −Ca2+/+PIP2 vs +Ca2+/+PIP2, (E) −Ca2+/–PIP2 vs +Ca2+/–PIP2, and (F) +Ca2+/–PIP2 vs +Ca2+/+PIP2. RMSFs for the Cα atoms were measured from the n = 3 replicates of 500 ns all-atom MD simulations. All non-aqueous systems are membrane-associated.

cPLA2 features a deep, channel-like active site shielded by a lid region. Additionally, the lid and the areas surrounding active site residues S228 and D549 are enriched with aromatic amino acids, which favorably recognize and bind the double bonds in the 20:4 arachidonic acid tail via π–π stacking (Figure S7). The X-ray structure (Protein Data Bank entry1CJY) is presumed to represent the “closed” form of cPLA2. Although it is partially activated with Ca2+ bound in the C2 domain, the fact that a fatty acyl chain lacks direct access to the active site confirms that cPLA2 also needs further conformational changes. Upon membrane association, it appears that the active site will become more accessible, which we confirmed in our MD simulation data.

Using principal component analyses (PCAs), we detected significant movement of loop 495–540 (Figure A,B). The goal of PCA in molecular simulations of protein dynamics is to identify the most dominant and collective motions of a protein by reducing the dimensionality of the trajectory data. Here, the Cα RMSD was used in the PC calculations, and the two most dominant PCs, PC1 and PC2, were projected onto the cPLA2 structure. Interestingly, the free energy minimum of aqueous cPLA2 is adjacent to that of fully activated cPLA2 (with Ca2+ and PIP2), suggesting that the lid region can readily transition between open and closed states (Figure B). Additionally, the simulation movie revealed a lateral displacement of the loop spanning residues 495–540 (as captured by PC1). This movement appears to facilitate the exposure of allosteric lysine residues K488, K541, K543, and K544 following membrane association and activation.

4.

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Mechanistic dissection of cPLA2 activation. (A) Projection of principal components 1 and 2 onto the protein structure. The X-ray structure (gray) was superimposed in the last frame of PCs 1 and 2. (B) Probability density of all of the PCs from systems 1 (aqueous cPLA2), 2 (cPLA2/–Ca2+/–PIP2), 3 (cPLA2/+Ca2+–PIP2), 4 (cPLA2/–Ca2+/+PIP2), and 5 (cPLA2/+Ca2+/+PIP2). (C) Evolution of a phospholipid substrate tunnel inside the catalytic domain active site. The cavity volume was calculated and visualized with the CAVER3.0 web server using a value of 0.1 for the minimum probe radius. (D) cPLA2 membrane association with Ca2+. Proposed mechanism of active site opening: C2 domain association with the membrane, catalytic domain association, transition of the lid region and loop 490–544 from the closed to open state, and opening of the active site to accommodate the SAPI substrate.

In human cPLA2, the lid region is composed of 23.9% negatively charged residues (D and E), 9% positively charged residues (R and K), and 28% hydrophobic residues (Figure S8). Sequence alignment of human, monkey, rabbit, and mouse cPLA2 shows that this region shares 85–98% identity across these species, with the chicken form being an exception at 70% identity. Notably, the basic residues are 67% conserved and the acidic residues are 81.2% conserved, highlighting the strong evolutionary conservation of the lid region. For RAW264.7 macrophage membranes, which carry a net negative charge, the 23.9% acidic composition of the lid likely facilitates its repulsion and spontaneous exposure of the active site. We propose that the lid’s amphipathic nature promotes membrane association and enhances substrate recognition and binding, particularly in environments where the membrane patch exhibits a variable net charge.

The transition of cPLA2 from a partially activated form to its fully activated form is also characterized by a 3-fold increase in the active site cavity volume compared to aqueous cPLA2 (closed state) around the catalytic dyad residues, S228 and D549 (Figure C). We used the CAVER 3.0 web server to detect the cavity size around the active site (Figure C). Allosteric site residue K544 also is exposed to the cavity after membrane association. K544 exposure is not observed during simulations of aqueous cPLA2. Furthermore, with CAVER cavity detection analyses, we observed that the active site became more available and thus should allow easier access for phospholipid substrates.

We propose a mechanistic event following membrane association (Figure D). Based on our all-atom MD simulations data, we deduced that the active site opening started with (step 1) association of the C2 domain with the membrane, (step 2) catalytic domain association, (step 3) the transition of the lid region and loop 490–544 from a closed state to an open state, and (step 4) opening of the active site to accommodate the SAPI substrate.

The lid region opening of cPLA2 is an allosteric hub in a population-shifting allostery model. This model posits that the protein’s free energy landscape comprises an ensemble of conformations. In contrast to its behavior in aqueous environments, membrane-associated cPLA2 exhibits a more open lid region with a small probability of transitioning to a closed state. The additions of Ca2+ and PIP2 to the C2 domain and membrane strengthen the lid opening effect. This work adds to the important role of the membrane in fine-tuning the protein’s functional mechanisms, similar to the case of other membrane proteins such as Ras proteins (K-Ras, H-Ras, and N-Ras) and protein kinase C (PKC). −

Although our simulations are limited and may not capture phospholipid substrate SAPI or PIP2 binding, we surmised that we sampled enough events to characterize the lid region opening and loop conformational plasticity after membrane association. MD simulations helped to refine the loop structures and suggested the deeper biophysical basis of their role in cPLA2 membrane association and activation. It should be noted that simulating protein loop dynamics remains one of the challenges in membrane protein modeling.

In summary, we show how membrane-associated cPLA2 transitions from its closed to open state by opening its lid region. Ca2+ allows the stabilization of the C2 domain after cPLA2 associates with the membrane, which verifies published experimental data. Furthermore, we see how the movement of loops 495–540 preceding the allosteric site activation can be crucial for PIP2 recognition. Lastly, this more profound understanding of the cPLA2–membrane environment can help illuminate the foundation for its mechanism of interaction with the membrane and the discovery of better therapeutics for inflammatory diseases based on active and/or allosteric site inhibition of cPLA2.

Supplementary Material

Download video file (6.1MB, mp4)
Download video file (8.2MB, mp4)
jz5c00860_si_003.pdf (1.6MB, pdf)
jz5c00860_si_004.pdf (1.9MB, pdf)

Acknowledgments

The authors thank Dr. Nicolas Frederic-Lipp, Dr. Abigail Dommer, Prof. Itay Budin, Prof. J. Andrew McCammon, Dr. Varnavas Mouchlis, Dr. Daiki Hayashi, Dr. Gosia Murawska, Dr. Carla Calvó-Tusell, Dr. Lorenzo Casalino, Dr. Mohamed Shehata, Xandra Nuqui, Clare Morris, and Nicholas Wauer for helpful discussions and advice on phospholipase A2 simulations and analysis. Supercomputing resources were provided to R.E.A. by XSEDE (NSF TG-CHE060073) and ACCESS (NSF TG-CHE060063) allocations. The study was supported by National Institutes of Health Grant R35 GM139641 (E.A.D.).

Glossary

Abbreviations

AA

arachidonic acid

cPLA2

group IVA (GIVA) cytosolic phospholipase A2

ER

endoplasmic reticulum

HDX-MS

hydrogen/deuterium exchange-mass spectrometry

MD

molecular dynamics

PAPC

1-palmitoyl-2-arachidonoyl-sn-glycero-3-phosphocholine

PC

phosphatidylcholine

PCA

principal component analysis

PLA2

phospholipase A2

PI

phosphatidylinositol

PIP2

phosphatidylinositol 4,5-bisphosphate

PS

phosphatidylserine

RMSF

root-mean-square fluctuation

RMSD

root-mean-square deviation

SAPI

1-stearoyl-2-arachidonoylphosphatidylinositol

Trajectories, structures, simulation scripts, analysis scripts, and data files were uploaded to our group’s website and can be accessed via https://amarolab.ucsd.edu/data.php.

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.jpclett.5c00860.

  • MD simulations of aqueous cPLA2 (Movie S1) (MP4)

  • MD simulations of membrane-associated cPLA2 (Movie S2) (MP4)

  • Computational methods (human group IVA cPLA2 modeling, RAW264.7 macrophage endoplasmic reticulum membrane modeling, all-atom molecular dynamics simulations, and trajectory analyses), lipid composition (number of lipids present in the lipid bilayer membrane without PIP2 and with 1% PIP2 and cPLA2 secondary structure (Tables S1–S3, respectively)), and molecular dynamics simulations (distance, RMSF, RMSD, sequence alignment, and loop and lid region movement (Figures S1–S10)) (PDF)

  • Transparent Peer Review report available (PDF)

M.K.E.B., E.A.D., and R.E.A. designed the project. M.K.E.B. performed the modeling, performed the simulations, and analyzed the data. E.A.D. and R.E.A. supervised the modeling and MD simulations. M.K.E.B. wrote the initial draft of the paper. E.A.D. and R.E.A. secured the funding and resources for the project. All authors contributed to the writing and editing of the manuscript.

The authors declare no competing financial interest.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

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jz5c00860_si_003.pdf (1.6MB, pdf)
jz5c00860_si_004.pdf (1.9MB, pdf)

Data Availability Statement

Trajectories, structures, simulation scripts, analysis scripts, and data files were uploaded to our group’s website and can be accessed via https://amarolab.ucsd.edu/data.php.

Articles from The Journal of Physical Chemistry Letters are provided here courtesy of American Chemical Society

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