Lysophospholipases cooperate to mediate lipid homeostasis and lysophospholipid signaling

Article abstract of DOI:10.1194/jlr.M087890

Abstract Lysophospholipids (LysoPLs) are bioactive lipid species involved in cellular signaling processes and the regulation of cell membrane structure. LysoPLs are metabolized through the action of lysophospholipases, including lysophospholipase A1 (LYPLA1) and lysophospholipase A2 (LYPLA2). A new X-ray crystal structure of LYPLA2 compared with a previously published structure of LYPLA1 demonstrated near-identical folding of the two enzymes; however, LYPLA1 and LYPLA2 have displayed distinct substrate specificities in recombinant enzyme assays. To determine how these in vitro substrate preferences translate into a relevant cellular setting and better understand the enzymes’ role in LysoPL metabolism, CRISPR-Cas9 technology was utilized to generate stable KOs of Lypla1 and/or Lypla2 in Neuro2a cells. Using these cellular models in combination with a targeted lipidomics approach, LysoPL levels were quantified and compared between cell lines to determine the effect of losing lysophospholipase activity on lipid metabolism. This work suggests that LYPLA1 and LYPLA2 are each able to account for the loss of the other to maintain lipid homeostasis in cells; however, when both are deleted, LysoPL levels are dramatically increased, causing phenotypic and morphological changes to the cells.—Wepy, J. A., James J. Galligan, P. J. Kingsley, S. Xu, M. C. Goodman, K. A. Tallman, C. A. Rouzer, and L. J. Marnett. Lysophospholipases cooperate to mediate lipid homeostasis and lysophospholipid signaling.

Full text of DOI:10.1194/jlr.M087890

Lysophospholipases Cooperate to Mediate Lipid Homeostasis and Lysophospholipid Signaling
James A. Wepy†, James J. Galligan‡, Philip J. Kingsley‡, Shu Xu‡, Michael C. Goodman†, Keri A. Tallman†,§, Carol A.
Rouzer!, Lawrence J. Marnett†,‡,§, ,1
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A. B. Hancock Jr. Memorial Laboratory for Cancer Research, Departments of †Chemistry, ‡Biochemistry, and
§Pharmacology, !Vanderbilt Institute of Chemical Biology, and Vanderbilt-Ingram Cancer Center, Vanderbilt University
School of Medicine, Nashville, Tennessee 37232-0146, United States
Contact Information for Corresponding Author: 850 A Robinson Research Building, 23rd & Pierce Ave., Nashville
37232-0146, (615) 322-0907, larry.marnett@vanderbilt.edu
Running Title: The Role of Lysophospholipases in Lysophospholipid Signaling
Abbreviations – LysoPL, Lysophospholipid; LPA, lysophosphatidic Acid; LPC, lysophosphatidylcholine; LPE,
lysophosphatidylethanolamine; LPG, lysophosphatidylglycerol; LPI, lysophosphatidylinositol; LPS,
lysophosphatidylserine; MAPK, mitogen-activated protein kinase; LYPLA1, lysophospholipase A1; LYPLA2,
lysophospholipase A2; PG-G, prostaglandin glycerol ester; PG, prostaglandin; PMSF, phenylmethylsulfonyl fluoride;
APCI, atmospheric-pressure chemical ionization; DKO, double knockout; RMSD, root-mean-square deviation; MEK,
MAPK/ERK kinase.
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ABSTRACT
Lysophospholipids (LysoPL) are bioactive lipid species involved in cellular signaling processes and the regulation
of cell membrane structure. LysoPLs are metabolized through the action of lysophospholipases, including
lysophospholipase A1 (LYPLA1) and lysophospholipase A2 (LYPLA2). A new X-ray crystal structure of LYPLA2
compared to a previously published structure of LYPLA1 demonstrated near identical folding of the two enzymes;
however, LYPLA1 and LYPLA2 have displayed distinct substrate specificities in recombinant enzyme assays. To
determine how these in vitro substrate preferences translate into a relevant cellular setting and better understand the
enzymes’ role in LysoPL metabolism, CRISPR-Cas9 technology was utilized to generate stable knockouts of Lypla1
and/or Lypla2 in neuro2A cells. Using these cellular models in combination with a targeted lipidomics approach, LysoPL
levels were quantified and compared between cell lines to determine the effect of losing lysophospholipase activity on
lipid metabolism. This work suggests that LYPLA1 and LYPLA2 are each able to account for the loss of the other to
maintain lipid homeostasis in cells; however, when both are deleted, LysoPL levels are dramatically increased, causing
phenotypic and morphological changes to the cells.
KEYWORDS
Brain Lipids, Eicosanoids, Prostaglandins, Fatty Acid, Protein Kinases/MAP Kinase, Lipidomics, Mass Spectrometry,
Fluorescence Microscopy, Neurons, X-Ray Crystallography
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INTRODUCTION
Lysophospholipids (LysoPLs) are detergent-like lipid species that play a critical role in a wide variety of cellular
signaling mechanisms in addition to maintaining the structure, shape, and fluidity of cell membranes (1-3). Each LysoPL
comprises one nonpolar acyl chain, varying in length and degree of unsaturation, and a polar glycerophosphate headgroup.
Based on the structure of the headgroup, LysoPLs belong primarily to one of six classes, including lysophosphatidic acids
(LPAs), lysophosphatidylcholines (LPCs), lysophosphatidylethanolamines (LPEs), lysophosphatidylglycerols (LPGs),
lysophosphatidylinositols (LPIs), and lysophosphatidylserines (LPSs), each with distinct biological functions dependent
on physiological location and availability of their respective cellular receptors.
LysoPLs have been shown to elicit a wide range of biological effects, including cell proliferation, intracellular
calcium mobilization, metabolic activity, inflammatory and anti-inflammatory processes, and neuritogenesis (4-20). These
effects are generally evoked through ligand-receptor interactions, but this is not always the case. For example, LPCs’ high
physiological concentrations (~200 !M in human plasma) call into question a receptor agonist role in some circumstances,
and accruing evidence supports the hypothesis they can elicit effects by altering membrane properties or interacting
directly with proteins in ways other than saturable binding to a specific site (21-24). For example, LPCs have been shown
to directly promote Ras-mediated activation of the mitogen-activated protein kinase (MAPK) signaling pathway by a
mechanism that does not involve characterized LysoPL receptors (17). In contrast, LPEs, LPGs, and LPIs have also been
shown to activate this signaling pathway, but they do so via respective receptor interactions. Regardless of mechanism,
accumulated data suggest that LysoPLs play a significant role in modulating the phosphorylation state of the
Ras/Raf/MEK/ERK cascade (16, 18, 25-27). This signaling pathway is critical for the regulation of cell cycle progression
and differentiation (28-31). Indeed, LysoPL-mediated activation of the MAPK signaling pathway has consistently been
shown to induce neuronal differentiation in neuroblastoma cells (16-19).
As LysoPLs are potent signaling molecules, their cellular levels are carefully regulated through three primary
enzymatic pathways: lysophospolipase-mediated hydrolysis, acyltransferase-mediated generation of phospholipids (PLs),
and transacylase-mediated generation of PLs. In mammalian cells, the lysophospholipase pathway predominates (32-38).
The lysophospholipases A1 and A2 (LYPLA1 and LYPLA2) are cytosolic serine hydrolases with esterase and
thioesterase activity that are partially responsible for the metabolism of LysoPLs. LYPLAs hydrolyze LysoPLs at the sn-
1, and to a lesser extent, sn-2 positions to yield a free fatty acid (FA) and a derivatized glycerophosphate that can then be
recycled by a variety of phospholipases and acyltransferases to restructure the composition of the lipid membrane in a
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process called the Lands cycle (Scheme 1) (39-42). Inhibition of these regulatory mechanisms and the associated changes
in LysoPL homeostasis have been associated with a variety of neurological diseases, including cerebral atherosclerosis,
vascular dementia, and Alzheimer’s disease (24, 43-47).
In addition to their role in the degradation of LysoPLs, LYPLAs display other activities. For example, the
monoacylglycerol and endocannabinoid, 2-arachidonoylglycerol, is oxygenated by cyclooxygenase-2 to generate
prostaglandin glycerol esters (PG-Gs) (48, 49). Our previous work has identified LYPLA2 as a PG-G hydrolase,
converting PG-Gs to free prostaglandins (PGs) (41). This suggests a role for LYPLA2 in endocannabinoid metabolism.
Furthermore, LYPLA1 and LYPLA2 are among a small pool of enzymes known to exhibit thioesterase activity on
palmitoylated proteins such as the G5" subunit of heterotrimeric G proteins and Ras (50-52). This post-translational
modification serves to alter protein conformation and/or tether a cytosolic protein to a lipid membrane with the addition of
a lipophilic acyl moiety covalently bound to cysteine residues via a thioester linkage (53-56). LYPLA1 and LYPLA2 act
to remove these acyl modifications as part of a dynamic palmitoylation process, regulating the subcellular location and
conformation of a variety of cellular proteins (50, 52, 54, 57).
Despite relatively high sequence homology (68% identical, 83% similar), LYPLA1 and LYPLA2 display
moderate substrate specificity in regard to LysoPL hydrolysis (32, 58-60). For example, whereas LYPLA1 displays
general promiscuity in regard to the different LysoPL classes, LYPLA2 is considered more specific, preferring LPCs and
LPEs. However, most of this work has been done with recombinant enzymes and exogenous substrate, despite the fact
that substrate specificity of serine hydrolases differs between in vitro and cellular settings where subcellular location,
competitive metabolism, and protein expression levels impact relevant substrate availability (61).
To understand the substrate specificity of LYPLA1 and LYPLA2 in cells, we have utilized a murine neuro2a
neuroblastoma model, in which LYPLA activity is relatively high and commonly studied (62, 63). Using CRISPR-Cas9
technology, we have generated neuro2A knockout cell lines lacking Lypla1, Lypla2, or both genes to quantify and
compare endogenous LysoPL levels in the absence of respective lysophospholipase activity. This approach offers a
physiologically relevant model to determine the roles of LYPLA1 and LYPLA2 in LysoPL hydrolysis, as well as the
consequences of the loss of their activity. Our results demonstrate that the roles played by the two enzymes in intact cells
are not necessarily reflected by the results obtained with purified protein preparations.
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MATERIALS and METHODS
Chemicals, reagents, and statistics
All reagents were purchased from Sigma Aldrich (St. Louis, MO), unless otherwise stated. PG-Gs and deuterated
PGs (PGE2-d4) were purchased from Cayman Chemicals (Ann Arbor, MI). All lysophospholipids were purchased from
Avanti Polar Lipids (Alabaster, AL). LC-MS solvents were from Fisher (Pittsburgh, PA). 16:0-d3 LPC was synthesized as
described in Supplemental Information. Recombinant enzyme was expressed and purified as described previously, using
cDNA from OriGene Technologies (Rockville, MD) (41). HIS-Select nickel affinity beads were from Sigma Aldrich.
HiPrep 16/60 Sephacryl S-200 HR was from GE Healthcare (Little Chalfont, UK). Neuro2A cells were obtained from the
American Type Culture Collection (ATCC, Manassas, VA). Cell culture reagents were purchased from Invitrogen (Grand
Island, NY). All experiments were performed twice in triplicate, unless otherwise noted, and statistical significance was
determined using one-way analysis of variance, unless otherwise noted.
Cell culture
Neuro2A cells were maintained as adherent cultures in DMEM supplemented with 10% FBS from Atlas
Biologicals (Fort Collins, CO). Cells were cultured at 37ºC, 5% CO2, grown to no more than 75% confluency, and
trypsinized to passage.
X-Ray crystallography
Recombinant human LYPLA2 (10 mg/mL) was pre-incubated with phenylmethylsulfonyl fluoride (PMSF) (10X
molar ratio). Crystals were produced by hanging drop vapor diffusion with drops containing 2 !L of enzyme-inhibitor
complex and 2 !L of reservoir solution containing 0.1 M sodium citrate (pH 5.6) and 15-30% PEG 3350. After 1~2
weeks, irregular crystals formed and were supplemented with 25% glycol for cryopreservation. Diffraction data for
LYPLA2-PMSF were collected on the Advanced Photon Source LS-CAT beamline 24-ID-E at 100 K. The data were
processed with XDS and solved to 2.70 Å resolution by molecular replacement using Phaser with the A chain of LYPLA1
(PDB ID: 1FJ2) as the search model. Two monomers were present in one asymmetric unit. Difference electron density
maps contoured at 3# showed the presence of a phenylmethylsulfonyl moiety associated with active Ser122. Data
collection and refinement statistics for each structure are listed in Supplemental Table S1. Figures were generated using
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PyMol Molecular Graphics system (Schrödinger). Atomic coordinates for LYPLA2-PMSF have been deposited in the
PDB (ID: 6BJE).
Recombinant LYPLA activity assays
Recombinant enzyme solutions (100 nM) were prepared in PBS with or without 0.5% (w/v) FA-free BSA, and
100 !L aliquots of each solution were preincubated at 37 ºC for 5 min. PGE2-G, 16:0-d3 LPC, or a mix of sn1-1 and sn2-
18:0 LPC (1.5 nmol) was added in 1 !L ethanol, and samples were vortexed, and then incubated at 37 ºC for 10 min. For
quantitation of product formation (PGE2-G and 16:0-d3 LPC), enzymatic activity was quenched by adding 1 mL of ice-
cold ethyl acetate with 0.5% (v/v) acetic acid containing either 20 ng PGE2-d4 or 1 nmol 16:0-d31 as internal standards.
Samples were vortexed, and the organic layers were collected and dried under nitrogen. To quantify PG-G hydrolysis,
extracts were resuspended in 200 !L acetonitrile and analyzed via LC-ESI-MS/MS, as described below. To quantify 16:0-
d3 LPC hydrolysis, samples were derivatized with pentafluorobenzyl (PFB) bromide based on a previously described
method with minor adaptions (64). Specifically, extracts were reconstituted in 150 !L acetonitrile, and then 200 !L of 100
nM tetrabutylammonium hydrogen sulfate in 100 nM dibasic sodium phosphate buffer was added. Following addition of
200 !L 2% (v/v) PFB bromide in chloroform, the samples were vortexed and sonicated for 45 min at 35% output using a
Virsonic Cell Disrupter (Model 16-850), and then lipids were extracted via the addition of 1 mL of 9:1 hexanes:ethanol.
The organic layer was removed, dried under nitrogen, and resuspended in 200 !L acetonitrile for analysis via LC-
atmospheric-pressure chemical ionization (APCI)-MS/MS, as described below. For quantitation of substrate remaining
(18:0 LPC), enzymatic activity was quenched by adding 100 !L of ice-cold ethanol with 0.5% (v/v) acetic acid containing
2 !M 17:0 LPC as an internal standard. Samples were immediately analyzed via LC-ESI-MS/MS, as described below, to
prevent the equilibration of sn1-18:0 and sn2-18:0 LPC isomers via acyl migration.
Generation and validation of LYPLA knockout cells using CRISPR-Cas9
CRISPR technology was utilized to generate genetic knockout cells as described by Ran, et al (65). gRNAs were
designed to target early exons of the LYPLA1 and LYPLA2 genes. gRNA oligomers (LYPLA1 gRNA:
TCCGATGCCCGCCGTTGTGC; LYPLA2 gRNA: AGCTGAGCGGGAAACGGCCG) were annealed, phosphorylated,
and ligated into digested pspCas9(BB)-2A-puro plasmid (plasmid # 62988, Addgene, Cambridge, MA). Neuro2A cells
(1.5 x 105) were suspended in 2 mL DMEM supplemented with 10% FBS and plated in 6-well plates. The following day,
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5 !g of each plasmid was combined with 10 !L lipofectamine 2000 reagent (Invitrogen) in 1 mL Opti-MEM and
incubated at room temperature for 30 min. Neuro2A culture medium was replaced with the appropriate plasmid-
lipofectamine solution, and cells were incubated at 37 ºC for 24 h. The medium was then replaced with fresh DMEM plus
10% FBS, and cells were allowed to recover for 24 h at 37 ºC prior to the addition of 0.75 !g/mL puromycin. Cells were
incubated at 37 ºC for 48 h before replacing the medium. After recovering for ~1-2 weeks, cells were pelleted,
resuspended in sorting buffer (PBS + 4% FBS), and strained to separate clumps of cells. Solutions were sorted by flow
cytometry using a 5-laser BD LSRII with a 100 !m nozzle at the Vanderbilt Medical Center Flow Cytometry Core to
isolate single cell cultures in 96-well plates for each cell line. Clones were incubated until they reached ~70% confluency
and then passaged until enough cells could be harvested for knockout validation via western blotting. Cells lacking both
Lypla1 and Lypla2 genes, or double knockout (DKO) cells, were generated by repeating this process to target the Lypla1
gene in the Lypla2-/- cells.
Western blot analysis
Protein expression was determined by western blot analysis as previously described (41, 66). Samples were
separated by SDS-PAGE. Then, proteins were transferred to a nitrocellulose membrane and blocked with Odyssey
Blocking Buffer (LI-COR, Lincoln, NE) for 1 h at room temperature, and probed with rabbit anti-LYPLA1 (1:1000 v/v,
abcam, Cambridge, UK), rabbit anti-LYPLA2 (1:1000 v/v, Vanderbilt Antibody and Protein Resource Core2), rabbit anti-
ERK1/2 (1:1000 v/v, Cell Signaling Technologies (CST), Danvers, MA), rabbit anti-phospho-ERK1/2 (1:1000 v/v, CST),
rabbit anti-MEK1/2 (1:1000 v/v, CST), rabbit anti-phospho-MEK1/2 (1:1000 v/v, CST), or goat anti-$-actin (1:5000 v/v,
Santa Cruz Biotechnologies, Santa Cruz, CA) overnight at 4 °C. Membranes were washed and incubated with IR-visible
anti-rabbit or anti-goat secondary antibodies (1:5000 v/v, LI-COR). Blots were visualized using an Odyssey IR Imager.
Assay for LYPLA activity in cell lysates
WT, Lypla1-/-, Lypla2-/-, or DKO cells (1 x 106) were plated in 8 mL DMEM with 10% FBS in 100 mm plates and
incubated at 37 ºC for 24 h prior to harvesting. Cell pellets were suspended in a buffer containing 25 mM Tris (pH 7.5),
0.1 mM EDTA, and 0.1 mM DTT. Cells were sonicated into solution via 10 x 1 s pulses at 35% output. Debris was
pelleted via centrifugation at 15,000 x g for 10 min, and soluble protein concentration in the supernatant was determined
via PierceTM BCA protein assay (Thermo Scientific, Rockford, IL). Solutions (250 !g/mL) of each enzyme were prepared,
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and 100 !L aliquots were preincubated at 37 ºC for 5 min. PGE2-G or 16:0-d3 LPC (1.5 nmol) was added in 1 !L ethanol,
and samples were vortexed and incubated at 37 ºC for 1 h. Enzymatic activity was quenched by adding 1 mL of ice-cold
ethyl acetate with 0.5% (v/v) acetic acid containing either 20 ng PGE2-d4 or 1 nmol 16:0-d31 as internal standards.
Samples were vortexed, and organic layers were collected and dried under nitrogen. FAs were derivatized as described
above, and LC-MS/MS analysis was then performed as described below.
Lysophospholipid Extraction
WT, Lypla1-/-, Lypla2-/-, or DKO cells (1 x 106) were plated in 8 mL DMEM with 10% FBS in 100 mm plates and
incubated at 37 ºC for 24 h. Cells were then scraped into 3 mL ice-cold PBS and pelleted. LysoPLs were extracted from
cell pellets using a method adapted from Zhao, et al., by resuspending in 200 !L of ice-cold methanol containing 50 nM
17:0 LPA, 250 nM 17:0 LPC, 50 nM 17:1 LPE, 50 nM 17:1 LPG, 50 nM 17:1 LPI, and 150 nM 17:1 LPS (67). Cells were
sonicated with 10 x 10 s pulses at 35% output and protein was pelleted. Methanol extracts were collected for lipid
analyses via LC-ESI-MS/MS as described below. Protein pellets were resuspended in 1 mL PBS containing 0.2% SDS,
sonicated with 10 x 10 s pulses at 35% output, and subjected to BCA assay to quantify input protein.
LC-MS/MS Analysis
For activity assays, samples were run on an LC-MS system consisting of a Shimadzu liquid chromatograph in line
with a SCIEX 3200 QTrap mass spectrometer. Analyst software (ver. 1.6.2) was used for instrument control, data
acquisition, and data processing. Quantitation was achieved via stable isotope dilution against the indicated internal
standard of PGE2-d4 or 16:0-d31. For PGE2 analysis, reconstituted samples were run in negative mode on a system
configured as described in Supplemental Tables S2A-B. For 16:0-d3 analysis, reconstituted samples were run in negative
mode on a system configured as described in Supplemental Tables S3A-B. The Luna liquid chromatography reverse phase
C18 columns were from Phenomenex (Torrance, CA).
For lipidomics experiments, samples were run on an LC-MS system consisting of a Shimadzu liquid
chromatograph in line with a SCIEX 3200 QTrap or a SCIEX 6500 QTrap mass spectrometer based on methods described
by Aaltonen, et al. and Okudaira, et al. (68, 69). This method isolates LysoPLs from other glycerolipid and
glycerophospholipid species. Analyst software (ver. 1.6.2) was used for instrument control, data acquisition, and data
processing. Quantitation was achieved via internal standard dilution against the respective unnatural 17:0 or 17:1 LysoPL
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depending on the class being investigated. In all cases, the mass spectrometer was configured in electrospray mode and
operated in multiple reaction monitoring mode. LPC species were analyzed in positive ion mode with LC parameters
described in Supplemental Table S4A. LPE, LPS, LPG, LPA, and LPI species were analyzed in negative ion mode with
LC parameters described in Supplemental Table S4B. Fragmentation patterns of all LysoPL species are described in
Supplemental Table S4C.
Immunocytochemistry
WT, Lypla1-/-, Lypla2-/-, or DKO Neuro2A cells (4 x 103) were plated in 100 !L DMEM with 10% FBS in a 96-
well plate. After 48 h, the medium was replaced with 100 !L DMEM containing 2% FBS (established conditions for
inducing neuronal differentiation (17, 70-72)) and either 2.5 !M retinoic acid, 10 !M PD98059 (an inhibitor of MEK
activation used to study neuronal differentiation (73)), 10 !M 16:0 LPA, 10 !M 16:0 LPC, or DMSO control. Cells were
cultured for 48 h to promote differentiation, based on a method adapted from Riboni, et al. (74). Cells were then washed 2
x with 100 !L PBS and fixed with the addition of 3.7% formaldehyde in PBS, followed by shaking at room temperature
for 20 min. Cells were then washed 2 x with PBS and permeabilized with 0.15% Triton X-100 in PBS for 10 min at room
temperature. Cells were again washed 2 x with PBS and blocked with Odyssey Blocking Buffer by shaking for 1 h at
room temperature. Cells were then stained with mouse anti-$III-tubulin (1:2000 v/v, abcam) for 24 h at 4 ºC while
shaking. The following day, cells were washed 3 x with TBS containing 0.1% (v/v) Tween (TBS-T) for 10 min while
shaking at room temperature, prior to the addition of AlexaFluor 488-conjugated donkey anti-mouse antibody (1:1000 v/v,
Life Technologies, Carlsbad, CA) and 4',6-diamidino-2-phenylindole (DAPI) (1:1000 v/v, Invitrogen) for 1 h while
shaking at room temperature. Cells were then washed again 3 x with TBS-T and rinsed 3 x with PBS. Stained cells were
imaged at 10x power in a 5 x 5 array, and the data were processed using a MetaXpress Micro XL automated microscope
imager, which generated a single composite image per well. Neurite outgrowth quantification was performed using the
following parameters: minimum neuron area = 120 !m2, maximum neuron width = 27 !m, minimum neuron intensity
above background = 500 gray levels, minimum nuclear width = 5 !m, maximum nuclear width = 14 !m, minimum
nucleus intensity above background = 3000 gray levels, maximum neurite outgrowth width = 4 !m, minimum neurite
outgrowth length = 10 !m, minimum neurite outgrowth intensity above background = 200 gray levels.
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RESULTS
LYPLA1 and LYPLA2 display high structural alignment
The crystal structure of apo-LYPLA1 (PDB ID: 1FJ2) suggests that the enzyme is unusual in that it lacks the
traditional cap domain found in other members of the "/$-hydrolase superfamily (75). Instead, the $4-"2 loops are
positioned as a flexible lid covering the active site serine, creating a potential hydrophobic channel for lipid binding.
Another crystal structure of LYPLA1 (PDB ID: 5SYM), and the structure of LYPLA2 (PDB ID: 5SYN), have recently
been solved with the specific inhibitors, ML348 and ML349, respectively, bound in their active sites (76). Although the
previously reported structures of LYPLA1 and LYPLA2 suggest the proteins form weak dimers, size-exclusion
chromatography indicates that both enzymes are active as monomers (data not shown).
We employed sparse matrix screening of crystallization to obtain the crystals that were used to solve the structure
of LYPLA2 covalently modified by the general serine hydrolase inhibitor, PMSF, at 2.7 Å resolution (PDB – 6BJE) (Fig.
1, Supplemental Table S1). This is the second structure of LYPLA2 to be reported, and the first with a covalent
modification of the active site Ser122. Consistent with the previously reported structure, LYPLA2-PMSF crystallized as a
dimer, though it is equally active in monomer and dimer conformations. Crystals of apo-LYPLA2 were also obtained but
failed to diffract for structure studies due to lattice defects. The data confirm that like LYPLA1, LYPLA2 lacks a cap
domain, which is replaced by the $4-"2 loops (highlighted in blue in Fig. 1) that act as a flexible lid near the active site.
The LYPLA2-PMSF structure is very similar to that of LYPLA2-ML349; however, small differences in conformation are
noted. For example, the $5-"2 loop in the two structures is present in distinct conformations (Supplemental Fig. S1), with
the LYPLA2-ML349 loop extended slightly further than that of LYPLA2-PMSF. This suggests the loop is able to move to
accommodate substrates or inhibitors binding in the channel adjacent to the active Ser122. Additionally, the modified
Ser122-PMSF demonstrates a slight shift in position compared to the unmodified Ser122 of the LYPLA2-ML349
structure, likely due to the steric bulk of the PMSF modification.
Consistent with the relatively high sequence homology of LYPLA1 and LYPLA2 (Fig. 2A), a least-squares
comparison of their coordinates reveals that the two proteins are folded in nearly identical conformations (Fig. 2B).
Superposition across all 215 aligned residues yielded a root-mean-square deviation (RMSD) of 0.878 Å, and a quality of
alignment (Q-score) of 0.838, suggesting the protein structures differ by no more than 1 Å (77). This high degree of
sequence and structural similarity suggests the two proteins may share significant overlap in substrate specificity and
hydrolytic activity.
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LYPLA1 is a PG-G hydrolase in vitro
A recent report from our laboratory identified LYPLA2 as the major PG-G hydrolase in cancer cells. At that time,
LYPLA1 was ruled out on the basis of siRNA knockdown in cells, but the enzyme’s PG-G hydrolytic activity was not
evaluated directly in vitro with purified recombinant protein (41). Therefore, we quantified both LYPLA1 and LYPLA2
activity toward a representative PG-G substrate, PGE2-G. Additionally, as both LYPLA1 and LYPLA2 have been shown
to robustly hydrolyze 16:0-LPC, we quantified hydrolytic activity toward the 16:0-LPC species as a representative
LysoPL substrate. However, as the product of LysoPL hydrolysis by LYPLAs is a glycerophosphate headgroup (too small
and polar for LC separation) and a FA (notoriously omnipresent in solvents, plastics, and glassware, as well as difficult to
ionize and fragment (78-80)) (Fig. 3A), we utilized an isotopically labeled 16:0-d3 LPC species and a PFB-Br
derivatization for these assays (64). The derivatization yields a PFB-esterified acyl chain, which has electron-capturing
properties to enable sensitive ionization and detection in APCI-MS/MS. This method allowed for the LC-MS/MS-based
quantification of a FA species by avoiding contamination and ionization issues.
As demonstrated in Fig. 3B, both enzymes were able to hydrolyze these substrates. LYPLA1 hydrolyzed PGE2-G
at a rate of 48.1 ± 0.5 !mol/min/mg compared to 93.3 ± 4.7 !mol/min/mg for LYPLA2, and LYPLA1 hydrolyzed 16:0-d3
LPC at a rate of 127 ± 8 !mol/min/mg compared to 129 ± 9 !mol/min/mg for LYPLA2. Thus, contrary to our findings in
intact cancer cells, recombinant LYPLA1 displayed substantial PG-G hydrolytic activity. However, whereas the two
enzymes hydrolyzed 16:0-d3 LPC with equivalent specific activity, LYPLA2 was significantly more active than LYPLA1
toward PGE2-G. It has been reported that BSA is able to improve hydrolysis of lipid substrates in recombinant serine
hydrolase activity assays, presumably by acting as a lipid carrier to better mimic the conditions of a cellular setting, and
we have recently shown that the presence of BSA has a significant effect on LYPLA2-mediated PG-G hydrolysis (41, 81).
Therefore, we also added BSA to the enzyme solutions to promote substrate availability and compared enzymatic activity
toward both PG-G and LysoPL substrates. In the presence of BSA, LYPLA1 hydrolyzed PGE2-G at a rate of 64.2 ± 1.7
!mol/min/mg compared to 170 ± 8 !mol/min/mg for LYPLA2, and LYPLA1 hydrolyzed 16:0-d3 LPC at a rate of 137 ±
14 !mol/min/mg compared to 137 ± 2 !mol/min/mg for LYPLA2. These results suggest that the addition of BSA
promotes the hydrolytic activity of LYPLA2 but not LYPLA1 toward PG-Gs, without significantly affecting the activity
of either enzyme toward LysoPLs.
Preliminary studies by our lab have suggested LYPLA2 prefers the sn1-isomer of PG-G and glycerolipids (41).
To determine substrate specificity of LYPLAs toward sn1- vs sn2-isomers of LysoPLs, equilibrium mixtures of sn1-18:0
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LPC (~80%) and sn2-18:0 LPC (~20%) were added to 100 nM enzyme solutions of LYPLA1, LYPLA2, or a no enzyme
control under conditions in which hydrolysis occurred faster than isomeric equilibration. The isomeric composition of the
remaining 18:0 LPC substrate was assessed immediately after quenching with acidified ethanol to prevent acyl migration,
and PBS controls were included to ensure that changes to isomeric composition were not merely a result of isomeric
equilibration. With both LYPLA1 and LYPLA2, only sn1-18:0 LPC isomer levels were significantly decreased after 10
min compared to those in the PBS controls (Fig. 3C). These data demonstrate the specificity of LYPLAs toward sn1-
LysoPLs, which is consistent with our published activity of LYPLA2 toward 1(3)-PG-Gs and 1(3)-arachidonoylglycerol.
Knockout of Lypla genes results in decreased hydrolytic activity
As our results suggested that in vitro enzymatic activity does not necessarily reflect the metabolic contribution of that
enzyme in vivo, we sought to better understand the role of LYPLA1 and LYPLA2 in LysoPL turnover in intact cells.
CRISPR-Cas9 technology was utilized to generate stable genetic knockouts of Lypla1, Lypla2, or both genes in murine
neuro2A neuroblastoma cells. Sanger sequencing was used to verify genetic editing at sites in the Lypla genes targeted by
respective gRNAs and overexpressed Cas9 protein. Genetic knockouts were validated at the protein level using western
blotting with antibodies directed toward LYPLA1 and LYPLA2 (Fig. 4A). Notably, the level of each LYPLA was
unaffected by knockout of the other isoform. The ability of the cell lines to hydrolyze the canonical PG-G and LysoPL
substrates of LYPLAs was tested using cellular lysates and the previously described hydrolytic activity assays. In addition
to avoiding palmitic acid contamination from plastics and solvents, the use of isotopically labeled 16:0-d3 LPC in this
assay was instrumental to avoid mistaking the endogenous palmitic acid present in cells for hydrolyzed product of the
exogenous substrate. This same problem did not arise in the case of PG-G hydrolysis, as neuro2A cells lack the
cyclooxygenase activity required to generate endogenous PG-Gs or PGs. Enzymatic activity of neuro2A lysates that had
been normalized for protein concentration was quantified as the percent of each substrate hydrolyzed as described in Fig.
3A. The percentages of PGE2-G hydrolyzed in WT, Lypla1-/-, Lypla2-/-, and DKO cell lysates were 50.4 ± 1.3%, 51.3 ±
0.9%, 43.0 ± 0.6%, and 32.1 ± 0.4%, respectively, while the percentages of 16:0-d3 LPC hydrolyzed in WT, Lypla1-/-,
Lypla2-/-, and DKO cell lysates were 24.2 ± 3.3%, 23.4 ± 3.5%, 18.8 ± 0.7%, and 11.2 ± 0.3%, respectively (Fig. 4B).
Similar to what we observed in our previous work using human cancer cell lines, Lypla1-/- cells exhibited the same PG-G
hydrolytic activity as WT cells, whereas the activity in Lypla2-/- cells was significantly lower than that of WT cells. In
DKO cells, PG-G hydrolysis was further decreased, suggesting that both enzymes contribute to PG-G hydrolysis and that
!
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LYPLA2 can better compensate for loss of LYPLA1 than vice versa. This same trend is observed with LysoPL
hydrolysis. Notably, however, DKO cells retained 64% and 46% of the PGE2-G and 16:0-d3 LPC hydrolytic activity of
WT cells, respectively, indicating that other enzymes are available to catalyze both reactions. It is not possible to know if
expression of these enzymes increases to compensate for the loss of LYPLA1 and/or LYPLA2 in the knockout cells. If so,
these results would underestimate the contribution of the LYPLAs to PGE2-G and 16:0-d3 LPC hydrolysis in the WT
cells.
LysoPL species are increased in DKO cells
With the decreased hydrolytic activity observed in the Lypla knockout cells, we hypothesized that basal levels of
LYPLA substrates would be consequentially increased. To test this hypothesis, we utilized an LC-MS/MS-based targeted
lipidomics approach to quantify LPA, LPC, LPE, LPG, LPI, and LPS substrates containing the following acyl moieties at
both the sn-1 and sn-2 positions of the glyceryl backbone: 16:0, 16:1, 18:0, 18:1, 18:2, 18:3, 20:4, 20:5, and 22:6.
Absolute quantification of each lipid species was normalized to the amount of protein in each sample. As the variance of
the LysoPL species were unequal between samples, and values in each sample were not normally distributed, one-way
ANOVA could not be used to determine statistical significance. Instead, samples with normal distributions (skewness
between -1 and 1) were analyzed by Welch’s t-test, and samples with abnormal distributions (skewness less than -1 or
greater than 1) were analyzed by the Wilcoxon rank-sum test. Statistically significant differences in LysoPL levels in
Lypla1-/-, Lypla2-/-, or DKO cells were compared individually to those of WT cells. Results for every identified LysoPL
species are listed in Table 1, with values that are significantly different from those of WT cells marked by boldface text.
Additionally, these values are displayed in Fig. 5A-F as absolute quantities, and in Fig. 5G-L as -fold changes normalized
to the LysoPL levels in WT cells (with WT values set to 1, represented by dotted lines).
As expected, LPCs were the most abundant of the LysoPL classes, with more prevalent species found in
quantities near 500 pmol/mg protein. LysoPLs containing saturated acyl chains of 16 and 18 carbons were generally found
at levels much higher than those containing longer or more unsaturated acyl chains. Whereas the LysoPL concentrations
of Lypla1-/- cells were mostly comparable to those of WT cells [with the exception of 20:4 LPA (1.6-fold increase) and
20:4 LPC (1.3-fold increase)], Lypla2-/- cells displayed some significant differences from WT cells in certain LysoPL
species, such as 16:0 LPC (1.5-fold increase), 18:2 LPC (1.5-fold increase), 18:3 LPC (1.8-fold increase), 16:0 LPI (2.2-
fold increase), and 18:0 LPI (1.5-fold increase). However, any differences in the single knockout cells were overshadowed
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"$

by the much greater increases in the levels of many LysoPLs in the DKO cells. This is especially notable with the LPCs,
LPEs, LPIs, and LPSs, of which most lipid species increased by 3- to 9-fold in the DKO relative to the WT cells. These
results are consistent with the lysophospholipase activity measured in cell lysates (Fig. 4B). Notably, while these large
changes in LysoPLs were observed, FA levels in the lipid extracts were quantified via PFB-derivatization, as described
above, and no significant differences were seen in the knockout cells (Supplemental Fig. S4). This is not surprising as
cellular LysoPL levels are roughly 1000-fold lower than those of free FAs; therefore, LysoPL-hydrolysis is unlikely to be
a major contributor to the levels of cellular FAs. Together, the data suggest that LYPLA2 makes a greater contribution to
LysoPL hydrolysis than LYPLA1 and/or can fully compensate for the loss of LYPLA1. However, the loss of both
enzymes appears to unmask a role for LYPLA1 that is not revealed by the single knockout cells.
Interestingly, while lipids in each of the LysoPL classes exhibited significant increases in the DKO cells, albeit at
varying magnitudes, there was no consistent pattern in the exact species (as designated by the acyl chain) most affected.
This suggests that LYPLA substrate specificity is more dependent on LysoPL class than acyl chain identity, with LPC,
LPE, and LPI species in the DKO cells increasing by >300% on average compared to WT cells. Furthermore, in contrast
to in vitro studies where LYPLA2 has been shown to prefer fully saturated LysoPL substrates, unsaturated LysoPLs of
each class in the DKO cells are significantly increased, especially in the LPC, LPE, and LPI classes (41). Indeed, the
highest –fold changes between DKO and WT cells occurred in the polyunsaturated LPCs, although these were mostly
minor species. Of course, it is possible that neither LYPLA1 nor LYPLA2 plays a direct role in changing some LysoPL
levels. It is possible that genetic deletions of multiple lipases involved in the Lands Cycle alter lipid-remodeling pathways,
leading to indirect increases of specific LysoPL species that may not necessarily be directly hydrolyzed by either enzyme.
Such remodeling might also occur through the compensatory overexpression of other enzymes in response to the deletion.
Nevertheless, these data demonstrate that the in vivo impact of a specific serine hydrolase cannot be easily predicted by in
vitro studies of its substrate selectivity, stressing the importance of cellular models in studying enzymatic activity.
LysoPL signaling pathways are overactive in DKO cells
The above data indicated that Lypla knockout leads to increases in LysoPLs, which are well-characterized
signaling molecules. As activation of the MAPK pathway is commonly associated with LysoPL signaling, we evaluated
the phosphorylation states of individual kinases within the MAPK cascade using phospho-specific antibodies and
compared them between WT and Lypla knockout cells. When normalized to total MAPK/ERK kinase (MEK)1/2,
!
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phosphorylated MEK1/2 was significantly higher in DKO than WT or either single knockout cell lines (Fig. 6A).
Additionally, when normalized to total ERK1/2, phosphorylated ERK1/2 demonstrated this same trend with significant
increases observed in DKO cells (Fig. 6B). The increased phosphorylation states of these proteins suggest that the
elevated LysoPLs in DKO cells are activating the MAPK signaling pathway.
Neuronal differentiation is increased in LYPLA knockout cells
The consequences of ERK phosphorylation have been extensively studied and are dependent on cell type. In
neuroblastoma cells, the MAPK signaling pathway is known to induce neuronal differentiation, increasing neurite
outgrowth and cell area, and initiating gap junction intracellular communication (16-19). To assess the impact of MAPK
signaling pathway activation in the context of LYPLA deficiency, we utilized immunocytochemistry and high-throughput
imaging to compare the morphology and degree of neuronal differentiation between the WT and Lypla knockout neuro2A
cells. Retinoic acid was also used as a positive control to induce differentiation in WT cells. After fixing and
permeabilizing the cells in 96-well plates, the overall shape of each cell was visualized using an antibody directed toward
!III-tubulin (green). Each cell was identified using the nuclear stain DAPI to enable cell counting and normalization
between samples (Fig. 7A). Multiple 10x magnification images were “stitched together” to create one composite image
for each well (1 replicate), and images were processed with MetaXpress software using the neurite outgrowth application.
This technology allowed us to take large-scale images of entire cell populations in 96-well plates and quickly convert
them into quantitative outputs of cell sizes and morphologies as an established method of measuring neuronal
differentiation (82, 83). Parameters quantified with this application include number of cells, cell body area, straightness of
processes, numbers of processes and branches, and a total outgrowth measurement incorporating the lengths of all
processes and branches. These parameters were normalized first to the number of cells in each sample and then to the
corresponding values of WT cells to give -fold changes (Fig. 7B-D). While the straightness of the processes of each cell
line was unchanged (data not shown), both the average cell body area and the average total outgrowth of the Lypla1-/-,
Lypla2-/-, and DKO cells were significantly increased compared to those of WT cells. Furthermore, the average number of
processes/cell was significantly increased in the DKO cells to levels comparable to that of WT cells treated with retinoic
acid. Additionally, the average total outgrowth of each LYPLA knockout was comparable to that of the WT cells treated
with retinoic acid.
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Previous studies have shown that some LysoPLs can induce or suppress neuronal differentiation (16:0 LPC and
16:0 LPA, respectively) in neuro2A cells (17). Both of these species are present in FBS, suggesting the possibility that the
conditions used to induce differentiation in the cells (a decrease in FBS content in the culture medium from 10% to 2%)
resulted in changes in their levels that contributed to the differentiation process. We confirmed the presence of ~800 nM
16:0 LPC and ~30 nM 16:0 LPA in medium containing 2% FBS, suggesting starting concentrations of ~4 µM 16:0 LPC
and ~150 nM 16:0 LPA in medium containing 10% FBS. To directly assess the effects of these lipids on Neuro2A
differentiation, we conducted experiments in which each lipid was added to differentiation medium (containing 2% FBS)
at a concentration of 1 µM. The cells were not viable in the absence of serum, precluding carrying out these experiments
under serum-free conditions. As shown in Supplemental Fig. S5, the addition of 16:0 LPC significantly increased
neuronal differentiation in each cell line, whereas the addition of 16:0 LPA had only modest effects on the average
number of processes in DKO cells. These data suggest that the relatively low concentrations of LysoPLs in the 2% FBS-
containing medium may be an external source of LYPLA substrates that mediate neuronal differentiation. However, the
fact that concentrations of these lipids are actually decreased when the serum concentration is reduced to induce
differentiation under our conditions suggests that they are unlikely to be playing a major role in the phenomena that we
observe. Thus, it is most likely that higher levels of basal neuronal differentiation observed in DKO cells, which is
comparable to those of WT cells treated with a differentiation agent, are due to increased endogenous LysoPL levels.
To further verify that the increased levels of neuronal differentiation were due to increased MAPK activation, WT
and DKO cells were treated with a MEK inhibitor, PD98059, and neurite outgrowth parameters were quantified (Fig. 8).
While outgrowth parameters in cells treated with the DMSO vehicle were consistent with basal neuronal differentiation in
each cell line (Fig. 7), the cell body area, number of processes, and total outgrowth of DKO cells treated with PD98059
were all significantly reduced. Furthermore, the average number of processes and total outgrowth of PD98059-treated
DKO cells was comparable to that of untreated WT cells (Fig. 8C-D). These data suggest that inhibiting MAPK activation
with a MEK antagonist in DKO cells counteracts the effects of increased LysoPLs on neuronal differential. The
phenotypic changes to neuritogenesis, especially the increased total outgrowth of DKO cells, are likely due to the
increased LysoPL levels and their role in the activation of the MAPK signaling cascade. If this is the case, LYPLAs are
demonstrably important in maintaining LysoPL homeostasis and modulating their effects on signaling processes in cells.
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DISCUSSION
LysoPL metabolism is a significant component of the Lands cycle, modulating the levels of potent bioactive
signaling molecules and preventing membrane disruption (39, 42). LYPLA1 and LYPLA2 are responsible for a
substantial fraction of this activity, and consequentially, for regulating the signaling effects of their respective LysoPL
substrates (32). While past work has suggested LYPLA1 and LYPLA2 display relative substrate specificity in terms of
LysoPL hydrolysis, these studies have been done using recombinant enzyme and fail to accurately portray the roles of
LYPLAs in cellular settings. To compare the activities of these two enzymes, we have taken a multifaceted approach
using in vitro and cellular assays to determine how LYPLAs maintain lysophospholipid homeostasis.
As depicted in their crystal structures, LYPLA1 and LYPLA2 are nearly identically folded and share highly
similar active sites (Fig. 2B). Accordingly, the activity of the recombinant enzymes toward the most prevalent LysoPL
substrate, 16:0-LPC, is comparable (Fig. 3B). This was tested using an isotopically labeled LysoPL species, 16:0-d3 LPC,
in order to avoid contamination by the FA products of LysoPL hydrolysis. This method allows for LC-MS/MS-based
quantitation of LysoPL hydrolysis without relying on expensive radiolabeled lipid substrates or purification steps that
reduce sensitivity. However, to determine if the activity of LYPLA1 and LYPLA2 toward other LysoPLs is analogous in
a setting where all substrates would be available, we developed cellular models to compare their activity.
We have characterized the roles of LYPLA1 and LYPLA2 in LysoPL hydrolysis in a neuroblastoma cell model in
which Lypla1, Lypla2, or both genes were deleted. The results demonstrated minimal effects of Lypla1 deletion on
LysoPL levels, although it interesting to note that the only LysoPL species that increased in Lypla1-/- cells contained 20:4,
suggesting that LYPLA1, specifically, may play a role in regulating arachidonate levels and eicosanoid biosynthesis. In
contrast, Lypla2 deletion affected a greater number and broader diversity of lipid species than Lypla1 deletion, and
substantial LysoPL elevation occurred only with knockout of both enzymes (Fig. 5). In general, the data suggest that the
two enzymes act cooperatively, with apparent compensation of one isoform for the loss of the other, although that occurs
without substantial changes in protein expression (Fig. 4A). The consistent levels of protein expression in the knockout
cells suggest the enzymes are constitutively expressed to modulate LysoPL levels. However, it is possible that the
enzymes could undergo activation via post-translational modification; both LYPLA1 and LYPLA2 are palmitoylated in
cells at conserved Cys2, LYPLA1 is acetylated at Lys224 and LYPLA2 is phosphorylated at Ser82 (57, 84, 85).
Nevertheless, it appears that LYPLA1 and LYPLA2 are able to hydrolyze the majority of LysoPLs in cells despite the
differential preferences for LysoPL classes observed in vitro. One must interpret these data with caution, however, as
!
"(

lysates from DKO cells retained substantial hydrolytic activity (Fig. 4B). The specific nature of this activity is not known,
but it almost certainly has an impact on the ultimate pattern of LysoPL changes observed in the various cell lines that we
explored.
In neuroblastoma cells lacking both Lypla1 and Lypla2, lysophospholipase activity is sufficiently compromised to
lead to increased LysoPL levels and increased activation of the MAPK signaling pathway (Fig. 5). The outcome is
initiation of neuronal differentiation (Fig. 6). Correspondingly, these data suggest that LYPLA1 and LYPLA2 are able to
modulate cell differentiation and neuritogenesis in neuroblastoma cells through their hydrolytic metabolism of LysoPLs
(Fig. 7). The role of lysophospholipase-dependent modulation of LysoPL levels in physiological models of
neurodifferentiation will be an interesting subject for further study.
In summary, we have closely compared the structures and activities of LYPLA1 and LYPLA2 and determined
that the two proteins have significant overlap in their cellular functions. Both enzymes cooperate to maintain lipid
homeostasis in cells; however, losing the activity of only one of these enzymes seems to have modest to no effect on
LysoPL levels, suggesting the counterpart lysophospholipase is able to compensate for that loss. However, when both
enzymes are knocked out, the cells are unable to regulate levels of LysoPLs, leading to aberrant LysoPL-dependent
signaling, an unregulated Lands cycle, and associated phenotypic and morphological changes to the cells. This work is
one of the first documented studies of LYPLA activity in a cellular setting and gives a clear answer to why inhibiting only
LYPLA1 or LYPLA2 seems to have little to no effect (52, 86). We have focused here on the role of these proteins in
maintaining LysoPL homeostasis, but their redundancy in substrate specificity may also apply to their other major
function, namely the thioesterase activity that results in the depalmitoylation of protein substrates (87). While relatively
few palmitoylated proteins have been identified as specific substrates for either LYPLA, it has been shown that
incomplete knockdown or inhibiting only one LYPLA is ineffective, and dual inhibition of LYPLA1 and LYPLA2 with
palmostatin B is often required to affect global protein palmitoylation (51, 52, 86). These trends are consistent with our
findings that LYPLA1 and LYPLA2 seem to share LysoPL substrates in cells and offer credence to a dual
inhibition/knockout approach to further study their physiological and pathophysiological functions.
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ACKNOWLEDGEMENTS and GRANT SUPPORT
This work was supported by National Institutes of Health Grants CA089450 and T32-ES007028, the Hancock Jr.
Memorial Cancer Center Fund, and the National Institute of General Medical Sciences Fellowship, Ruth L. Kirschstein
National Research Service Award (5F31GM120879-02). Its contents are solely the responsibility of the authors and do not
necessarily represent the official views of the National Institutes of Health.
Flow Cytometry experiments were performed in the VMC Flow Cytometry Shared Resource. The VMC Flow
Cytometry Shared Resource is supported by the Vanderbilt Ingram Cancer Center (P30 CA68485) and the Vanderbilt
Digestive Disease Research Center (DK058404).
Antibodies were generated and purified by the Vanderbilt Antibody and Protein Resource. The Vanderbilt
Antibody and Protein Resource is supported by the Vanderbilt Institute of Chemical Biology and the Vanderbilt Ingram
Cancer Center (P30 CA68485).
Imaging experiments were performed in the Vanderbilt High-Throughput Screening Core Facility with assistance
provided by Debbie Mi and Joshua Bauer. The HTS core receives support from the Vanderbilt Institute of Chemical
Biology and the Vanderbilt-Ingram Cancer Center (P30 CA68485).
Statistical analyses were performed with guidance by Yu Shyr, Ph.D. from the Department of Biostatistics at
Vanderbilt University.
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#)

FOOTNOTES
1To whom correspondence should be addressed: email – larry.marnett@vanderbilt.edu
2Polyclonal rabbit anti-LYPLA2 antibody was generated by the Vanderbilt Antibody and Protein Resource Core and can
be obtained through contact with corresponding author, Dr. Lawrence Marnett.
!
#*

TABLES
Cell Line
Lipid Species
WT
Lypla1-/-
Lypla2-/-
DKO
16:0
16:1
6.90 (3.15)
3.23 (0.74)
3.54 (0.92)
4.35 (1.49)
0.86 (0.11)
545 (86.6)
62.9 (8.52)
464 (83.9)
416 (98.5)
25.1 (4.48)
1.61 (0.30)
41.4 (10.1)
1.76 (0.46)
23.3 (3.65)
30.4 (8.19)
91.0 (44.0)
63.3 (15.3)
4.23 (1.78)
25.2 (13.4)
1.29 (0.06)
5.04 (0.38)
4.18 (1.21)
50.5 (2.44)
0.24 (0.01)
15.4 (2.63)
169 (44.2)
96.7 (19.3)
2.60 (0.43)
9.07 (3.27)
5.80 (0.76)
34.7 (5.69)
15.3 (1.65)
0.62 (0.03)
6.95 (1.82)
4.23 (1.05)
3.24 (0.96)
4.85 (0.78)
1.38 (0.24)
509 (41.8)
62.5 (6.70)
442 (48.6)
460 (34.4)
26.8 (1.86)
1.59 (0.12)
53.5 (6.34)
2.14 (0.76)
23.7 (3.19)
25.7 (6.79)
70.2 (24.6)
53.9 (12.3)
3.42 (0.99)
26.8 (11.2)
1.09 (0.16)
3.99 (0.36)
3.01 (0.37)
31.9 (4.00)
0.32 (0.08)
10.8 (3.26)
166 (119)
9.28 (1.84)
3.31 (0.53)
5.85 (2.84)
6.08 (2.02)
1.02 (0.34)
823 (77.1)
85.0 (6.32)
549 (48.9)
498 (58.0)
38.6 (2.63)
2.91 (0.27)
51.4 (13.3)
3.88 (0.61)
24.8 (6.19)
35.8 (10.5)
93.0 (40.4)
50.0 (13.2)
3.10 (1.51)
21.1 (12.5)
1.31 (0.36)
6.00 (0.66)
5.08 (0.63)
29.0 (4.40)
0.23 (0.01)
34.2 (6.33)
254 (82.1)
139 (15.6)
3.34 (0.71)
10.5 (2.43)
5.52 (0.44)
34.4 (8.78)
26.3 (6.55)
0.60 (0.08)
16.2 (6.23)
4.55 (1.18)
16.3 (14.0)
8.97 (6.54)
1.36 (0.29)
1890 (713)
243 (101)
1410 (527)
1220 (503)
154 (66.4)
9.81 (2.16)
255 (167)
16.4 (7.96)
101 (61.5)
119 (74.3)
299 (215)
155 (106)
12.3 (9.95)
94.5 (88.9)
2.31 (0.17)
13.8 (2.93)
11.0 (2.42)
50.5 (14.4)
0.37 (0.01)
86.1 (33.3)
572 (242)
255 (101)
8.63 (4.27)
31.2 (22.9)
11.5 (3.73)
80.0 (43.7)
91.0 (72.9)
1.05 (0.13)
LPA
18:0
18:1
20:4
16:0
16:1
18:0
18:1
18:2
18:3
20:4
20:5
22:6
16:0
18:0
18:1
18:2
20:4
20:5
16:0
18:0
18:1
20:4
16:0
18:0
18:1
18:2
20:4
16:0
18:0
18:1
20:4
LPC
LPE
LPG
LPI
88.6 (21.9)
2.57 (0.44)
11.1 (2.57)
5.65 (0.93)
29.9 (4.60)
19.4 (5.31)
0.61 (0.06)
LPS
Table 1. Basal LysoPL levels (pmol/mg protein) in WT, Lypla1-/-, Lypla2-/-, and DKO cells, presented as mean (standard
deviation). Boldface values indicate P<0.05, determined by Welch’s t-test or the Wilcoxon rank-sum test for samples with
normal or abnormal distributions, respectively.
!
$+