ω-alkynyl lipid surrogates for polyunsaturated fatty acids: Free radical and enzymatic oxidations

Article abstract of DOI:10.1021/ja506038v

Lipid and lipid metabolite profiling are important parameters in understanding the pathogenesis of many diseases. Alkynylated polyunsaturated fatty acids are potentially useful probes for tracking the fate of fatty acid metabolites. The nonenzymatic and enzymatic oxidations of ω-alkynyl linoleic acid and ω-alkynyl arachidonic acid were compared to that of linoleic and arachidonic acid. There was no detectable difference in the primary products of nonenzymatic oxidation, which comprised cis,trans-hydroxy fatty acids. Similar hydroxy fatty acid products were formed when ω-alkynyl linoleic acid and ω-alkynyl arachidonic acid were reacted with lipoxygenase enzymes that introduce oxygen at different positions in the carbon chains. The rates of oxidation of ω-alkynylated fatty acids were reduced compared to those of the natural fatty acids. Cyclooxygenase-1 and -2 did not oxidize alkynyl linoleic but efficiently oxidized alkynyl arachidonic acid. The products were identified as alkynyl 11-hydroxy-eicosatetraenoic acid, alkynyl 11-hydroxy-8,9-epoxy-eicosatrienoic acid, and alkynyl prostaglandins. This deviation from the metabolic profile of arachidonic acid may limit the utility of alkynyl arachidonic acid in the tracking of cyclooxygenase-based lipid oxidation. The formation of alkynyl 11-hydroxy-8,9-epoxy-eicosatrienoic acid compared to alkynyl prostaglandins suggests that the ω-alkyne group causes a conformational change in the fatty acid bound to the enzyme, which reduces the efficiency of cyclization of dioxalanyl intermediates to endoperoxide intermediates. Overall, ω-alkynyl linoleic acid and ω-alkynyl arachidonic acid appear to be metabolically competent surrogates for tracking the fate of polyunsaturated fatty acids when looking at models involving autoxidation and oxidation by lipoxygenases.

Full text of DOI:10.1021/ja506038v

Article
pubs.acs.org/JACS
Open Access on 07/17/2015
ω‑Alkynyl Lipid Surrogates for Polyunsaturated Fatty Acids: Free
Radical and Enzymatic Oxidations
William N. Beavers,†,∥ Remigiusz Serwa,†,∥,⊥ Yuki Shimozu,‡ Keri A. Tallman,† Melissa Vaught,†
Esha D. Dalvie,§,# Lawrence J. Marnett,†,‡,§ and Ned A. Porter*,†,‡
A.B. Hancock Memorial Laboratory for Cancer Research, Departments of †Chemistry, ‡Biochemistry, and §Pharmacology, Vanderbilt
Institute for Chemical Biology, Vanderbilt Center in Molecular Toxicology, Vanderbilt Ingram Cancer Center, Vanderbilt University,
Nashville, Tennessee 37235, United States
S
* Supporting Information
ABSTRACT: Lipid and lipid metabolite profiling are important
parameters in understanding the pathogenesis of many diseases.
Alkynylated polyunsaturated fatty acids are potentially useful
probes for tracking the fate of fatty acid metabolites. The
nonenzymatic and enzymatic oxidations of ω-alkynyl linoleic
acid and ω-alkynyl arachidonic acid were compared to that of
linoleic and arachidonic acid. There was no detectable difference
in the primary products of nonenzymatic oxidation, which
comprised cis,trans-hydroxy fatty acids. Similar hydroxy fatty
acid products were formed when ω-alkynyl linoleic acid and ω-
alkynyl arachidonic acid were reacted with lipoxygenase
enzymes that introduce oxygen at different positions in the
carbon chains. The rates of oxidation of ω-alkynylated fatty
acids were reduced compared to those of the natural fatty acids.
Cyclooxygenase-1 and -2 did not oxidize alkynyl linoleic but efficiently oxidized alkynyl arachidonic acid. The products were
identified as alkynyl 11-hydroxy-eicosatetraenoic acid, alkynyl 11-hydroxy-8,9-epoxy-eicosatrienoic acid, and alkynyl
prostaglandins. This deviation from the metabolic profile of arachidonic acid may limit the utility of alkynyl arachidonic acid
in the tracking of cyclooxygenase-based lipid oxidation. The formation of alkynyl 11-hydroxy-8,9-epoxy-eicosatrienoic acid
compared to alkynyl prostaglandins suggests that the ω-alkyne group causes a conformational change in the fatty acid bound to
the enzyme, which reduces the efficiency of cyclization of dioxalanyl intermediates to endoperoxide intermediates. Overall, ω-
alkynyl linoleic acid and ω-alkynyl arachidonic acid appear to be metabolically competent surrogates for tracking the fate of
polyunsaturated fatty acids when looking at models involving autoxidation and oxidation by lipoxygenases.
Tracking lipids, lipid metabolites, and lipid decomposition
products in cells is a formidable task as the complexity of the
mixture challenges the most powerful analytical tools.17−20
Stable isotope derivatives of lipids have been used with some
success to track the distribution of molecular species into
different lipid classes in organelles, but detecting a minor
metabolic byproduct from thousands of different lipid species is
particularly difficult since stable isotope-labeled compounds are
often isobaric with endogenous species.21 Radiolabels have also
been used successfully in many applications, but associating a
particular radioactive chromatography fraction with the
structure of a molecular species in a complex mixture is a
challenge. Recently, an affinity labeling technique that makes
use of terminal alkynyl lipid analogues was reported.22 In this
approach, stable, but reversible, alkyne−cobalt complexes are
formed on a phosphine-modified silica to isolate alkynyl lipids.
This strategy has been used to monitor the distribution of
INTRODUCTION
■
The lipidome is a complex mixture of fatty acid and sterol
molecular species, which include the fatty acid esters of sterols,
triglycerides, and glycerophospholipids, such as the ethanol-
amines, cholines, and inositols.2 Polyunsaturated fatty acids
(PUFAs) such as linoleic acid (LA) (18:2) and arachidonic acid
(AA) (20:4) and their esters are particularly important
molecular species. These essential fatty acids are involved in a
number of consequential metabolic transformations via their
oxidation by lipoxygenase (LOX) and cyclooxygenase (COX)
enzymes.3−6 Oxidized lipids play a significant role in a number
of physiological and pathophysiological events, including
cardiovascular disease, cancer, and neurodegenerative diseases.7
The nonenzymatic peroxidation of both LA and AA by
molecular oxygen generates intermediate peroxyl free radi-
cals.8−12 Products that result from this lipid peroxidation
include peroxides and hydroperoxides, as well as secondary
electrophilic products capable of covalently modifying bio-
molecules, potentially altering their function.13−16
Received: June 16, 2014
Published: July 17, 2014
© 2014 American Chemical Society
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NMBHA in acetonitrile was added MeOAMVN in acetonitrile. After
35 h of incubation at 37 °C, BHT/PPh3 in acetonitrile was added, and
the mixture was vortexed for 2 min. Acetonitrile was removed under a
stream of argon, and the remaining material was reconstituted in 1.2%
isopropanol in hexanes with 0.1% acetic acid for HPLC-UV analysis.
MS studies of aHETE and NMR studies of aHETE were performed as
described above for aHODE.
alkynyl fatty acids into various cellular phospholipid classes
along with the subsequent lipase-catalyzed metabolism of those
lipids.23 In recent studies, terminal alkynyl analogues of 4-
hydroxy-2E-nonenal (HNE) and 4-oxo-2E-nonenal (ONE),
lipid-derived electrophiles known to modify proteins and
nucleic acids, were used to globally profile electrophile
adduction of proteins. UV-cleavable biotin azide was used to
isolate, identify, and quantify cellular protein−electrophile
adducts.16,24,25 This affinity chemistry allows protein−lipid
adducts to be concentrated and identified by standard
proteomic protocols. One shortcoming of adding electrophiles
exogenously to cells is that it does not mimic endogenous lipid
electrophile diversity, concentration, time course of generation,
and location of formation. To address these issues, we have
developed a series of ω-alkynyl PUFAs to investigate
endogenous lipid oxidation and its cellular consequences.
Tracking lipid incorporation and metabolism in cellular
systems has long been a goal for chemists, biochemists, and
biologists alike. Alkynyl analogues of lipid and cholesterol
species have been utilized to facilitate the tracking and retrieval
of these species in cells.16,22,26 During these studies, it is
assumed that the ω-alkyne confers only a minimal structural
change to its lipid analogue, resulting in nearly identical
chemical, biochemical, and biological properties. Here, we
report the oxidation chemistry for the alkynyl lipid analogues,
aAA and aLA, revealing that alkyne substitution has no effect
on fatty acid free radical autoxidation, and minimal effect on
enzymatic oxidation. This establishes that alkynyl PUFAs
provide a method to selectively study lipid distribution,
chemistry, and lipid metabolite interactions with cellular
macromolecules.
HPLC−UV/MS analysis of (a)HODEs and (a)HETEs. All HPLC
Analyses of (a)HETE and (a)HODE were carried out on a single
Beckman 5 μm ultrasphere silica column (250 mm × 4.6 mm) using
isocratic normal phase conditions (1.2% IPA in hexanes containing
0.1% acetic acid).28 Chiral HPLC analyses of HETE and HODE
methyl esters were performed on a Chiralpak AD column (250 mm ×
4.6 mm) produced by Chiral Technologies Inc., Exton, PA. aHETE
products have been eluted with 2% ethanol in hexanes, whereas
aHODE were eluted with 5% methanol in hexanes. Direct infusion MS
experiments were performed on ThermoFinnigan TSQ Quantum
triple quadrupole mass spectrometer, whereas all HPLC/MS analyses
were conducted on the same instrument coupled with a Surveyor MS
Pump and Surveyor Autosampler (for RP-HPLC) or with Waters
Alliance 2690 Separation Module (NP-HPLC). Detailed information
about solvent gradients and MS settings applied during these analyses
is given in the appropriate protocols presented below. Unless stated
otherwise, all the HPLC separations were conducted with a solvent
flow rate of 1 mL/min.
Formation and Analysis of Alkynyl F2α-Isoprostane (aF2α-
IsoP). MeOAMVN was added to a 165 mM solution of aAA in
benzene, and the mixture was incubated at 37 °C for 24 h. Solvent was
then evaporated, and the residue was treated with mixture of 1 mmol
BHT and 10 mmol P(OMe)3 in 3:1, acetonitrile/water and vortexed
for 5 min. The following solvent gradient was applied: 10% (95:5,
acetonitrile/methanol) in 2 mM ammonium acetate was held for 10
min, then ramped to 25% (95:5, acetonitrile/methanol) in 2 mM
ammonium acetate over 45 min. LC/MS/MS with negative ion mode
electrospray ionization (ESI) was used, and the important mass
spectrometer parameters were optimized for commercially available
PGF2α. The m/z transitions monitored were for a5F2α-IsoP (349 >
115), a8F2α-IsoP (349 > 127), a12F2α-IsoP (349 > 151), and a15F2α-
IsoP (349 > 193). Control oxidations of AA were performed using
analogous reaction conditions; however, slightly different analytical
conditions were applied to analyze the AA oxidation products. The
following solvent gradient was applied for AA oxidation products: 20%
(95:5, acetonitrile/methanol) in 2 mM ammonium acetate was held
for 10 min, then ramped to 40% (95:5, acetonitrile/methanol) in 2
mM ammonium acetate over 40 min. The m/z transitions monitored
were for 5F2α-IsoP (353 > 115), 8F2α-IsoP (353 > 127), 12F2α-IsoP
(353 > 151), and 15F2α-IsoP (353 > 193).
Cyclooxygenase O2 Uptake Kinetics. Quantification of cyclo-
oxygenase activity was performed in a thermostated cuvette at 37 °C
with stirring and monitored using a polarographic electrode with an
Instech System 203 oxygen monitor (Instech Laboratories Inc.,
Plymouth Meeting, PA). Substrates were solubilized in dimethyl
sulfoxide (DMSO). Activity assays were performed in 100 mM Tris
buffer containing 500 μM phenol, with hematin-reconstituted
cyclooxygenase protein. Substrate concentration was varied (1−50
μM), and maximal reaction velocity data were obtained from the linear
portion of the oxygen uptake curves. The data were analyzed by
nonlinear regression with GraphPad Prism (GraphPad, San Diego,
CA).
Crude Lipoxygenase Kinetic Parameters. LOX activity was
detected by monitoring the absorbance of the conjugated diene
product, HpETE, at 235 nm. UV assays were monitored using a
Hewlett-Packard 8453 diode array spectrophotometer equipped with a
thermostated cuvette at 25 °C, with stirring at 180 rpm. The enzyme
reactions included reaction buffer 50 mM Tris pH 7.4 with 0.03%
Tween-20 and substrate, and were initiated by the addition of enzyme.
Compounds were dissolved in acetonitrile containing 10% acetic acid
before addition to the reaction buffer; acetonitrile was kept below 1%
of the total reaction volume. Substrate concentration was varied (1−50
μM), and maximal reaction velocity data were obtained from the linear
EXPERIMENTAL SECTION
■
Materials. All reagents are from Sigma, St. Louis, MO, unless
otherwise noted. All native fatty acids and deuterated lipid metabolites
are from Cayman Chemical, Ann Arbor, MI, unless otherwise noted.
The alkynyl fatty acids, aLA and aAA, were synthesized as previously
described.26 N-methyl benzohydroxamic acid (NMBHA) has been
prepared as previously described.27 Benzene (HPLC grade) was
passed through a column of neutral alumina and stored over molecular
sieves (benzene is a carcinogen and mutagen, and should be used with
extreme care). Commercial α-tocopherol was chromatographed before
use. Diazomethane was prepared by portion-wise addition of
nitrosomethylurea into heterogeneous mixture of 40% aqueous
KOH and ethanol at 0 °C. The deep-yellow organic layer was
decanted and dried over NaOH. The dried ethereal MeN2 was used
immediately.
Formation and Analysis of Alkynyl Hydroxy Octadecadie-
noic Acid (aHODE) by Autoxidation. To a solution of aLA in
benzene was added 2,2′-azobis(4-methoxy-2,4-dimethylvaleronitrile)
(MeOAMVN) in benzene, and the mixture was incubated at 37 °C.
After 45 min, a solution of butylated hydroxytoluene (BHT) and
triphenyl phosphine (PPh3) in benzene was added, and the mixture
was vortexed for 1 min. Benzene was evaporated under a stream of
argon, and the residual material was reconstituted in 1.2% isopropanol
in hexanes with 0.1% acetic acid for HPLC-UV analysis. For direct
infusion MS studies of aHODE, eluted peaks from HPLC-UV analysis
were collected, and solvents were evaporated under a stream of argon;
then the residues were reconstituted in methanol. For NMR studies of
aHODE, corresponding peaks collected from HPLC-UV analysis were
combined, solvents were evaporated, and the residues were dried
under high vacuum for 2 h. These dried materials were reconstituted
in benzene-d6 and placed into 1.7 mm OD sample tubes for NMR
analysis.
Formation and Analysis of Alkynyl Hydroxy Eicosatetrae-
noic Acid (aHETE) by Autoxidation. To a mixture of aAA and
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portion of the absorbance curves. Rates were converted from
absorbance units/s to μM aHpETE/s using the molar absorptivity
constant of 0.027 μM−1 cm−1. The data were analyzed by nonlinear
regression with GraphPad Prism.
HETE (315.2 > 253.2), 15-HETE (319.2 > 257.2), and 15-HETE-d8
(327.2 > 264.2) for 100 ms each.
mCOX2 was diluted to a final concentration ratio of 100:1, fatty
acid/enzyme in 100 mM Tris, 500 μM phenol, 2× [mCOX2] Hematin
pH 8. The mCOX2 was incubated 5 min at 37 °C. AA and aAA were
added from 100× stocks in DMSO, and incubated 15 min at 37 °C.
The reactions were quenched and fatty acid metabolites extracted and
analyzed as detailed for the lipoxygenase enzyme full scan mode
experiment.
Monitoring the Oxidation of LA, AA, aLA, and aAA to
Completion. s15LOX1, LA, AA, aLA, and aAA were diluted to 2×
final concentration in 100 mM borate pH 9 at 25 °C. One mL fatty
acid was added to a 1 cm cuvette in a Beckman-Coulter DU-800
spectrophotometer and blanked. One mL enzyme was added, and the
reaction was monitored at 235 nm sampling every 1.5 s, until the
Δabsorbance reached 0. The data was fit to a one-phase exponential
association in GraphPad Prism to get R2 values and maximum
absorbances.
Kinetic Measurements of aLA, LA, aAA, and AA by s15LOX1.
s15LOX1, LA, AA, aLA, and aAA were all diluted to 2× final
concentration in 100 mM borate pH 9 at 25 °C. One mL fatty acid
was added to a 1 cm cuvette in a Beckman-Coulter DU-800
spectrophotometer as a blank control. One mL enzyme was added,
and the reaction was monitored at 235 nm sampling every 1.5 s in
triplicate. The slope was averaged over 10 points in the linear portion
of the curve to get the Δabsorbance per second, which was converted
to Vo using the molar extinction coefficients of 0.023 μM−1 cm−1 for
HpODE and 0.027 μM−1 cm−1 for HpETE. Kinetic parameters were
determined in GraphPad Prism using Michaelis−Menten fitting.
Enzymatic Oxidation of aLA and LA for LC/MS/MS Analysis.
Soybean 15 lipoxygenase 1 (s15LOX1) (Cayman Chemical) was
diluted to have a final concentration ratio of 100:1, fatty acid/enzyme
in 100 mM borate pH 9. Potato 5 lipoxygenase (5LOX) (Cayman
Chemical) was diluted to have a final concentration ratio of 100:1,
fatty acid/enzyme, in 100 mM phosphate pH 6.3. LA and aLA were
added from 100× stocks in DMSO, and incubated 15 min at 25 °C.
The reactions were quenched and fatty acid metabolites extracted with
ethyl acetate containing 0.5% acetic acid, PPh3, 9-HODE-d4, and
13-HODE-d4. Organic layer was dried under inert gas and dissolved
in methanol for LC/MS/MS analysis. Metabolites were analyzed on a
Thermo Finnigan TSQ Quantum with ESI source interfaced to
Surveyor MS Pumps and Surveyor Autosampler in both full scan and
SRM modes. Metabolites were separated by reverse-phase gradient
HPLC on a C18 (50 mm × 2.1 mm, 3 μm) column (Supelco, St. Louis,
MO) using 0.1% formic acid in water and 0.1% formic acid in
acetonitrile as the A and B mobile phases, respectively. Full scan
samples were separated by holding 20% B for 2 min, then ramping to
98% B over 6 min, holding 98% B for 3 min, then equilibrating to 20%
B for 3 min. Q1 was scanned in negative ion mode from 250 m/z to
380 m/z in 1 s. SRM samples were separated by holding 40% B for 0.5
min, then ramping to 98% B over 2 min, holding at 98% B for 2 min,
then equilibrating to 40% B for 2.5 min. Metabolites were detected by
SRM in negative ion mode observing the m/z transitions for a9-
HODE (291.2 > 171.2), a13-HODE (291.2 > 195.2), 9-HODE (295.2
> 171.2), 13-HODE (295.2 > 195.2), 9-HODE-d4 (299.2 > 172.2),
and 13-HODE-d4 (299.2 > 198.2) for 100 ms each.
NMR of aAA Metabolites. mCOX2 was incubated 5 min in 100
mM Tris, 500 μM phenol, and 2× [mCOX2] hematin pH 8 at 37 °C.
aAA was added, and the reaction was allowed to proceed for 15 h at 37
°C. The reaction was extracted with two volumes ethyl acetate, and the
organic layer was removed and evaporated under inert gas. The residue
was dissolved in ethanol and separated by reverse-phase HPLC on a
SUPELCOSIL C18 (150 mm × 3.0 mm, 5 μm) column, eluted with a
linear gradient with A and B buffers consisting of 0.1% acetic acid in
water and 0.1% acetic acid in acetonitrile. The gradient was held at
10% B for 10 min, then ramped to 100% B over the next 20 min, then
held at 100% B for 5 min, all at the flow rate of 1.0 mL/min. The
elution profile was monitored by absorbance at 235 nm. Peaks were
collected, dried under inert gas, and dissolved in CDCl3 for NMR
analysis. 1H and 1H−1H COSY spectra were recorded on Bruker AV-II
600 MHz spectrometer equipped with a cryoprobe. Chemical shifts are
reported in parts per million relative to the signal of residual
nondeuterated solvent.
LC/MS/MS-Based Kinetics for mCOX2. Metabolites of mCOX2
do not have an absorbance that can be used to perform kinetic
measurements. Therefore, LC/MS/MS was used to measure kinetic
parameters based on specific metabolites. mCOX2 was diluted to 25
nM in 100 mM Tris, 500 μM phenol, 50 nM hematin pH 8, and
incubated at 37 °C. AA and aAA were added from 100× stocks in
DMSO, and incubated at 37 °C. The length of incubation was
optimized to give less than 20% substrate turnover. The reactions were
quenched and fatty acid metabolites extracted with ethyl acetate
containing 0.5% acetic acid, PGE2-d4, and 13-HODE-d4. Metabolites
were separated by reverse-phase gradient HPLC on a C18 (50 mm ×
2.1 mm, 3 μm) column using 0.1% formic acid in water and 0.1%
formic acid in acetonitrile as the A and B mobile phases, respectively.
Metabolites were separated by holding 25% B for 0.5 min, then
ramping to 99% B over 2.5 min, holding at 99% B for 3 min, then
equilibrating to 25% B for 3 min. Metabolites were analyzed in
negative ion mode by SRM, monitoring the transitions for a11-HETE
(315.2 > 167.2), aPG (347.2 > 267.2), PG (351.2 > 271.2), 13-
HODE-d4 (299.2 > 198.2), PGE2-d4 (355.2 > 275.2) on an ABI/Sciex
3200 QTrap interfaced to a Shimadzu LC system. No deuterated
standard exists for 11-HETE. The signal intensity ratio between a11-
HETE and 13-HODE-d4 was observed to be similar in both full scan
MS and SRM modes, indicating that 13-HODE-d4 can be used to
quantify a11-HETE. The amount of each product formed was
converted into a concentration, and then into initial reaction rates (Vo)
using the incubation time. Kinetic parameters were determined in
GraphPad Prism using Michaelis−Menten fitting.
Mouse COX2 (mCOX2) was diluted to have a final concentration
ratio of 100:1, fatty acid/enzyme in 100 mM Tris, 500 μM phenol, 2×
[mCOX2] hematin pH 8. The mCOX2 was incubated 5 min at 37 °C.
LA and aLA were added from 100× stocks in DMSO, and incubated
15 min at 37 °C. The reactions were quenched and fatty acid
metabolites extracted and analyzed as detailed for the lipoxygenase
enzymes.
Enzymatic Oxidation of aAA and AA for LC/MS/MS Analysis.
s15LOX1 was diluted to a final concentration ratio of 100:1, fatty acid/
enzyme, in 100 mM borate pH 9. AA and aAA were diluted to 100×
stocks in DMSO. Fatty acids were separately added to enzyme
solutions, and incubated 15 min at 25 °C. The reactions were
quenched and fatty acid metabolites extracted with ethyl acetate
containing 0.5% acetic acid, PPh3, and 15-HETE-d8. The organic
layer was dried under inert gas and dissolved in methanol for LC/MS/
MS analysis. Metabolites were analyzed on a Thermo Finnigan TSQ
Quantum with ESI source interfaced to Surveyor MS Pumps and
Surveyor Autosampler in both full scan and SRM modes. Metabolites
were separated by gradient HPLC on a C18 (50 mm × 2.1 mm, 3 μm)
column using 0.1% formic acid in water and 0.1% formic acid in
acetonitrile as the A and B mobile phases, respectively. Full scan
samples were separated by holding 20% B for 2 min, then ramping to
98% B over 6 min, holding 98% B for 3 min, then equilibrating to 20%
B for 3 min. Q1 was scanned in negative ion mode from 250 m/z to
380 m/z in 1 s. SRM samples were separated by holding 40% B for 0.5
min, then ramping to 98% B over 2 min, holding at 98% B for 2 min,
then equilibrating to 40% B for 2.5 min. Metabolites were detected by
SRM in negative ion mode observing the m/z transitions for a15-
RAW264.7 Macrophage Enrichment, Activation, and Meas-
urement of aAA Metabolites. RAW264.7 macrophages (ATCC,
Manassas, VA) were plated at 10% confluence in DMEM+Glutamax
(Life Technologies, Grand Island, NY) supplemented with 10% fetal
bovine serum (Atlas Biologicals, Fort Collins, CO). Fatty acid free
BSA/aAA (6:1) was prepared as previously described.29 After 24 h,
media was replaced with serum free DMEM+Glutamax containing
aAA/BSA added to 15 μM final concentration of aAA. After 24 h, the
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Figure 1. Autoxidation products of of LA (A), aLA (B), AA (C), aAA (D). LA/aLA peaks are labeled with the hydroxyl position on either the 9 or
13 carbon, and the double bond configuration which resulted in either cis/trans (c,t) or trans/trans (t,t). AA/aAA peaks are labeled with the position
of the oxygen on either the 5, 8, 9, 11, 12, or 15 carbon. Both fatty acid pairs oxidize to similar product profiles with similar elution orders.
cells were washed with one volume DMEM+Glutamax to remove any
unincorporated aAA, and treated with DMEM+Glutamax 100 ng/
mL Kdo2-lipid A (KLA) (Avanti Polar Lipids, Alabaster, AL), prepared
as previously described.30 After 24 h, cells were scraped into media and
extracted with two volumes ethyl acetate containing 0.5% acetic acid,
PGE2-d4, and 13-HODE-d4. The organic layer was dried under an inert
gas stream and dissolved in methanol for LC/MS/MS analysis.
Metabolites were separated by reverse-phase gradient HPLC on a C18
(50 mm × 2.1 mm, 3 μm) column using 0.1% formic acid in water and
0.1% formic acid in acetonitrile as the A and B mobile phases,
respectively. Metabolites were separated by holding 25% B for 0.5 min,
then ramping to 99% B over 2.5 min, holding at 99% B for 3 min, then
equilibrating to 25% B for 3 min. Metabolites were analyzed in
negative ion mode by SRM, monitoring the transitions for a11-HETE
(315.2 > 167.2), a11-8,9-HEET (331.2 > 165.2), aPG (347.2 > 267.2),
PG (351.2 > 271.2), 13-HODE-d4 (299.2 > 198.2), and PGE2-d4
(355.2 > 275.2) on an ABI/Sciex 3200 QTrap interfaced to a
Shimadzu controller, autosampler, and HPLC pumps.
aLA produces a mixture of R- and S-HODE enantiomers,
similar to that of LA (SI Figure 2).
Contrary to the simplicity of products generated from LA
autoxidation, peroxidation of AA yielded a much more complex
mixture of products. In addition to acyclic hydroxy and
hydroperoxy products (HETEs and HpETEs) analogous to
HODEs and HpODEs, a mixture of diastereomeric isopros-
tanes (IsoPs) was produced from AA peroxidation.31−34
Oxidation of aAA under conditions that gave HpETEs as
major products was promoted by NMBHA.27 As shown in
Figure 1, characterization of peroxidation products was
performed on the aHETEs after reduction of the corresponding
hydroperoxides. The HPLC−UV elution profile for aHETEs
was similar to the profile obtained for their natural analogues.
MS analyses established the position of oxygen substitution on
the carbon chain (SI Table 1), and NMR analysis provided
information about stereoisomeric geometry (SI Figure 1). The
major HETE stereoisomers have Z,E-conjugated diene
geometry, analogous to the structure of AA-derived HETEs.
These experiments show that the elution order of aHETEs is
identical to the elution order observed for HETEs. Additionally,
oxidation of equimolar mixtures of AA and aAA generated
nearly equimolar mixtures of HETE and aHETE.
HETEs are not the only autoxidation products formed from
AA, so aAA was exposed to the radical initiator MeOAMVN in
the absence of NMBHA under conditions of oxidation and
workup that were expected to yield significant quantities of the
aF2α-IsoPs. LC/MS/MS analysis of the major (a)F2α-IsoPs
formed in this sequence are shown in SI Figure 3. Chromato-
grams of only the 5- and 15-series (a)F2α-IsoPs are presented
because these compounds are formed in a large excess
compared to the 8- and 12-regioisomers. The preference for
formation of the 5- and 15-regioisomers and the elution profiles
observed for both the natural and ω-alkynyl analogues are
consistent with a previous report.35
RESULTS
■
LA and aLA were oxidized under conditions that would allow
us to compare autoxidation rates and products. As shown in
Figure 1, four ω-alkynyl conjugated diene hydroperoxides
(alkynyl hydroperoxyoctadecadienoic acids, aHpODEs) were
produced as primary products from aLA autoxidation, a result
directly analogous to the chemistry observed with LA.11
Reduction of the hydroperoxides to the corresponding alcohols,
aHODEs, was carried out immediately after peroxidation since
the HODEs are more stable and better suited for HPLC
analysis than HpODEs. HPLC-UV analysis revealed that
parallel oxidation of equimolar amounts of aLA and LA
generated equivalent amounts of aHODEs and HODEs, with
elution orders identical to those of the natural compounds.11
The isolated products were then analyzed by ESI-MS and NMR
to confirm hydroxyl position and conjugated diene geometry
(Supporting Information [SI] Figure 1). Utilizing chiral
chromatography, it was determined that the autoxidation of
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Table 1. Kinetic Values Comparing aAA and AA for the Enzymes Ovine Cyclooxygenase 1 (oCOX1), Human Cyclooxygenase 2
(hCOX2), Human Platelet-Type 12 Lipoxygenase (plt12LOX), Porcine Leukocyte-Type 12 Lipoxgygenase (lk12LOX), Rabbit
a
Reticulocyte 15 Lipoxygenase 1 (r15LOX1), and Soybean 15 Lipoxygenase 1 (s15LOX1)
enzyme
oCOX1
substrate
product
Km (μM)
Vmax (μMs1−
)
kcat (s−1
57
(51 3)
11
)
kcat/Km (μM−1 s−1
)
Vmax/Km (s−1
)
aAA
AA
O2 cons.
6.2 0.8
(3.4 0.6)
4.5 0.7
n/a
6
9
6
n/a
(15 3)
hCOX2
aAA
AA
O2 cons.
n/a
1
2
(2.4)
n/a
1
n/a
(6.1 0.6)
7.0 0.3
(14.7 0.5)
n/a
plt12LOX
lk12LOX
r15LOX1
s15LOX1
aAA
AA
Abs 235 nm
Abs 235 nm
Abs 235 nm
Abs 235 nm
4.53 0.08
(13.3 0.3)
1.37 0.09
(13.1 0.7)
0.61 0.05
(8.6 0.4)
0.025 0.002
0.12 0.01
0.6 0.3
(9.5 0.7)
(1.4 0.7)
aAA
AA
4
1
n/a
n/a
n/a
0.3
1
(7.8 1.3)
(2 1)
aAA
AA
7
2
n/a
0.09
2
(0.4 3)
n/a
(20 3)
3.1 0.9
aAA
AA
2.5 0.2
0.8 0.2
4.3 0.2
6
1
24
2
a
All COX enzymes were assayed using oxygen electrode. LOX enzymes were assayed using absorbance at 235 nm. AA kinetic values taken from the
literature are in parentheses.1 Vmax/Km values are reported for crude enzyme preparations, while kcat/Km values are reported for purified enzymes.
Figure 2. LA, aLA, AA, and aAA were all incubated with s15LOX1 and observed at 235 nm until ΔAbs = 0. Despite having different kinetic
parameters, fatty acid pairs aLA/LA (A) and aAA/AA (B), are eventually oxidized completely.
Using alkynyl fatty acids to further probe the biochemistry of
cellular systems requires detailed knowledge of the chemistry of
enzymatic oxidation. Kinetic parameters were determined for
the transformations of aAA in the presence of several LOX and
COX enzymes by measuring alkene formation or O2
consumption, respectively. Data presented in Table 1
demonstrate small differences in kcat/KM for the alkynyl and
natural fatty acids, suggesting that aAA is an efficient substrate
for both COX-1 and COX-2. The catalytic efficiency of human
platelet-type 12-LOX in the presence of aAA was also found to
be similar to the efficiency observed for AA as a substrate. On
the other hand, porcine leukocyte-type 12-LOX, rabbit
reticulocyte 15-LOX1, and s15-LOX1 did not oxidize aAA as
efficiently as AA, illustrated by the relatively large differences in
Vmax/Km and kcat/Km values between these substrates. Despite
these differences in kinetic parameters, aAA is completely
converted by a15LOX1 when reacted for long enough times
(Figure 2).
The kinetic parameters for the transformations of aLA and
LA by s15LOX1 were also measured, and kcat/Km values were
determined to be 0.51 0.07 μM−1 s−1 and 5.2 0.6 μM−1 s−1,
respectively. These values are similar to those observed for aAA
and AA (Table 1), and like aAA, aLA is also eventually
completely reacted (Figure 2). aLA and LA give a similar
product profile of primarily (a)9-HODE or (a)13-HODE for
the enzymatic transformation by 5-LOX or s15-LOX1
respectively (SI Figures 4 and 5). a13-HODE produced from
s15LOX1 was assessed for optical purity, and determined to be
entirely the S isomer, as anticipated from the stereochemistry of
LA oxidation (SI Figure 3). Ovine COX1 generated a product
profile similar to that of mCOX2 when reacted with LA, and
neither enzyme oxygenated aLA (SI Figure 5).
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Figure 3. XIC analysis of the metabolite profiles of aAA (A) and AA (G) catalyzed by mCOX2 show very different products. The metabolism of
aAA by mCOX2 shows some aAA remaining (B) and four products corresponding to the addition of one (C), two (D), three (E), and four (F)
atoms of oxygen. The profiles in B−F are all to the same scale. The metabolism of AA by mCOX2 shows a single major product showing the
addition of three atoms of oxygen and having the same m/z as PGE2/PGD2 (K). In addition to the major metabolite and remaining AA (H),
metabolism of AA by mCOX2 shows three other products corresponding to the addition of one (I), two (J), and four (L) oxygens. These AA
products correspond to the nonalkynylated versions of the products seen when aAA is metabolized by mCOX2. The profiles in H−L are to the same
scale.
We compared the kinetics of mCOX2 oxidation of aAA
determined by O2 uptake (Table 1) to values determined by
LC/MS/MS and noticed that the product profile from aAA was
different from that of AA. As demonstrated in Figure 3, four
aAA-derived oxygenation products were identified by MS,
which correspond to the addition of one (m/z = 315.2), two
(m/z = 331.2), three (m/z = 347.2), and four atoms of oxygen
(m/z = 365.2). The product at m/z = 315.2 corresponds to
a11-HETE, and the product at m/z = 347.2 corresponds to
aPGE2/D2. One possibility for the identity of the product at m/
z = 331.2 is aHpETE; however, attempted reduction of the
hydroperoxide with either TCEP or PPh3 did not alter the peak
elution time (data not shown), indicating a chemically distinct
species from the hydroperoxide. Although the metabolite
profile of AA by mCOX2 resulted in peaks with m/z values
corresponding to the addition of one, two, three, and four
atoms of oxygen, similar to what was seen with aAA, the
intensity of the peaks displayed major differences. AA
oxygenation by mCOX2 results in a major peak at m/z =
351.2, corresponding to PGE2/D2, and a minor peak at m/z =
319.2, corresponding to a single oxygen atom incorporation.
The remaining oxygen addition peaks were very minor by
comparison, but have similar retention time and molecular
weight shifts relative to PG as was seen with aAA. Ovine COX1
generated a product profile similar to that of mCOX2 for aAA
and AA (data not shown).
To identify the metabolites depicted in Figure 3, product
peaks at m/z = 315.2 and 331.2 were isolated and analyzed via
1D and 2D NMR. The compound present at m/z = 365.2 was
not stable through the isolation process and thus was not
1
analyzed. Figure 4 shows the structure and H−1H COSY for
the peak at m/z = 315.2. It was determined that the identity of
this peak is (5Z, 8Z, 12E,14Z)-11-hydroxyeicosa-5,8,12,14-
tetraen-19-ynoic acid (alkynyl 11-hydroxyeicosatetraenoic acid,
a11-HETE). SI Table 2 shows the chemical shifts relative to
1
CDCl3 and coupling constants as determined from the H
NMR (SI Figure 6). The coupling constants for the alkene
between C12 and C13, J12,13 = 15.2 and 15.1 Hz respectively,
identify the bond as trans. Figure 5 shows the structure and
1H−1H COSY for the peak at m/z = 331.2. It was determined
that the identity of this peak is (Z)-7-(3-((3E, 5Z)-2-
Hydroxyundeca-3,5-dien-10-yn-1-yl)oxiran-2-yl)hept-5-enoic
acid (alkynyl 11-hydroxy-8,9-epoxy-eicosatrienoic acid, a11-8,9-
HEET). SI Table 3 shows the chemical shifts relative to CDCl3
1
and coupling constants as determined from the H NMR (SI
Figure 7). The coupling constants J12,13 = 15.2 and 15.1 Hz,
assign the alkene between C12 and C13 as trans. The coupling
constants for the epoxide were measured as J8,9 = 4.2 and 4.3
Hz, identifying the epoxide as cis.
Due to the distinct product profile of aAA metabolism, the
kinetic parameters for aAA were reevaluated by LC/MS/MS.
AA kinetic parameters were determined using the product
PGE2, whereas the kinetic parameters for aAA were determined
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Figure 4. 1H−1H COSY spectrum of the collected LC/MS peak with m/z = 315.2 with peaks assigned, which was identified to be that of a11-HETE.
using aPGE2 and a11-HETE. Michaelis−Menten plots for these
vitro experiments, a11-HETE was more abundant than aPG
and a11-8,9-HEET in cells. The ratio of a11-HETE to aPG and
a11-8,9-HEET is increased from the purified protein analyses
indicating that cellular and purified enzyme metabolite profiles
may have slight differences.
three kinetic experiments are found in Figure 6. The catalytic
efficiency for the formation of PGE2 by mCOX2, 1.6
0.2
μM−1 s−1, was similar to the oxygen uptake value for hCOX2
seen in Table 1. The small difference between the two can be
explained by the formation of the nonenzymatic PG
degradation product, 12-hydroxyheptadecatrienoic acid
(HHT), which accounts for approximately 20% of the total
PG signal (data not shown). The catalytic efficiencies for aAA
products were very different, however, at 0.019 0.005 μM−1
s−1 for aPG formation and 0.4 0.1 μM−1 s−1 for a11-HETE
formation. When these values are compared to the hCOX2
oxygen uptake during aAA metabolism value, 2 1 μM−1 s−1, it
can be seen that most of the oxygen consumption is due to the
formation of a11-HETE.
DISCUSSION
■
Understanding both the enzymatic and nonenzymatic metab-
olism of alkynyl fatty acids is important because lipid
oxygenation products and lipid electrophile formation has
been reported to result from both enzymatic and nonenzymatic
mechanisms.36−38 Our data indicate that aLA and LA are
kinetically equivalent substrates for free radical chain oxidation;
(a)HODEs are formed as two positional isomers with oxidation
at the 9 and 13 carbons. Additionally, the conjugated dienes are
in two different conformations, the ZE kinetic product, and the
EE thermodynamic product. Analyzing the ZE/EE product
ratios can be used as a “peroxyl radical clock” to measure
peroxidation propagation rate constants,39 further confirming
that these two substrates are equivalently oxidized.
To evaluate the potential of aAA as a tool in cellular settings,
its incorporation, release, and metabolism was tested in
RAW264.7 macrophages. BSA/aAA complexes were formed
as previously described and then were added to serum-free cell
culture medium for 24 h.29 Cells were then washed with
medium to remove any unincorporated aAA and activated with
100 ng/mL KLA for 24 h. Fatty acid metabolites were extracted
from the combined media and cells. Figure 7 shows the
quantification of a11-HETE, a11-8,9-HEET, aPG, and PG in
cells enriched with aAA and activated with KLA. The
alkynylated products were only seen at high levels in the
aAA-enriched and KLA-activated cells, which corresponds to
the conditions where levels of fatty acid release and COX2
expression are highest. Further correlating to the kinetic and in
Similarly, aAA and AA are also equivalent substrates for
autoxidation. The mechanism of HpETE and IsoP formation
has been studied in great detail and it has been established that
six major Z,E-HpETE products form with hydroperoxide
substitution at carbons 5, 8, 9, 11, 12, and 15 of the 20 carbon
eicosanoate chain.40 The IsoPs are formed as a mixture of
stereoisomers, the four sets of regioisomers identified by the
position of the allylic alcohol in the chain, 5, 8, 12, and 15.41
Each regioisomeric set of IsoPs contains eight diasteromers.
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Figure 5. 1H−1H COSY spectrum of the collected LC/MS peak with m/z = 331.2 with peaks assigned, which was identified to be that of a11-8,9-
HEET.
Quantification of the IsoP isomeric mixture has been used in
recent years as a measure of oxidative stress in vivo.8,42,43 Both
aAA and AA form the respective HpETEs and IsoPs at similar
levels. This is an important finding for setting up future lipid
oxidation studies since many disease states, including models
for cardiovascular disease and neurodegenerative diseases, are
characterized by a high level of oxidative stress.7
The kinetic values measured here indicate that with some
notable exceptions, aAA is a reasonable enzymatic substrate for
both COX and LOX classes of enzymes. Enzymatically, both
alkynyl PUFAs are metabolized by various LOX enzymes to
product profiles similar to those of the native PUFAs. Despite
the observed differences in catalytic efficiency, aAA, AA, aLA,
and LA are completely oxygenated by s15LOX1 when allowed
to react to completion (Figure 2). We hypothesize that the
reduced efficiency observed is the result of the alkyne altering
the conformation of the lipid within the LOX active site. Model
systems designed to study cellular processes all have limitations,
and the reduced enzymatic efficiency seen here may restrict the
use of this model to understand short-term enzymatic lipid
metabolism. However, in many biological settings, this reduced
enzymatic efficiency remains negligible in understanding and
tracking lipid metabolism because many studies will be looking
at changes over long time periods.
(S)-hydrogen abstraction and oxygen addition to C11 forming
the alkynyl 11-hydroperoxyl radical. The remaining reactions
proceed through two critical junctions, endoperoxide formation
and prostaglandin ring closure. The a11-hydroperoxyl radical
can be reduced by H atom transfer to form a11-HETE
(Scheme 1A), which was identified as one of the major
products by LC/MS/MS and 2D-NMR (Figure 4). When
endoperoxide formation is followed by prostaglandin ring
closure, and a final oxygenation at C15, aPGG2, the precursor
to all prostaglandins is formed (Scheme 1B). However, when
endoperoxide formation is not followed by prostaglandin ring
closure, endoperoxide homolytic cleavage results in 8,9-epoxide
and 11-alkoxyl radical formation. The alkoxyl radical can then
be terminated to give a11-8,9-HEET (Scheme 1C), which was
identified by 2D-NMR (Figure 5). The epoxide was identified
as cis due to the coupling constants for H8 and H9, which were
measured at 4.2 and 4.3 Hz, respectively. This is an interesting
observation indicating that the epoxide is formed in the enzyme
active site, before the bond between C7 and C8 can rotate.
Nonenzymatic epoxidation from endoperoxide scission would
be expected to give a 3:1 trans:cis geometry due to the free
rotation of the C7−C8 bond.44 11-8,9-HEET was first
identified when 8,9 epoxy-eicosatrienoic acid was incubated
with cyclooxygenase enzymes. The cyclooxygenase enzymes
were only able to add oxygen to C11 because the epoxide
prevented endoperoxide formation.45,46 The major mCOX2
products of aAA oxygenation we have identified are structurally
similar to previously reported COX2 variant AA metabolites.47
Schneider et al. demonstrated that mutations at Gly526 and
These data demonstrate the potential usefulness of aPUFAs
for the study of lipids in a biological setting; however, one
major finding is the differential metabolism of AA and aAA by
mCOX2 and oCOX1. All of the products in our proposed
mechanism (Scheme 1) result from the same first two steps, 13-
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Figure 7. RAW264.7 macrophages were enriched aAA, then
activated with 100 ng/mL KLA for 24 h. Metabolite levels were
measured by LC/MS/MS-SRM for the media and cells combined.
an altered product profile. One substrate that can be
investigated is the endocannabinoid 2-arachidonylglycerol (2-
AG). The crystal structure of its isomer, 1-AG, has been solved
for mCOX2, and it was revealed that it sits in the active site in
two different conformations. The structural difference in these
confirmations is a slight change in the position of the ω-tail in
the active site. Oxygenation may occur in both conformations
because abstractable hydrogens on C13 are in line with the
catalytic Tyr385, but different distances in each conformation.48
2-AG has two major products, PG-glycerol and 11-HETE-
glycerol,49 which further corroborates that there are multiple
modes of binding. These data are potentially relevant to aAA
binding in the mCOX2 active site, because they indicate that
small changes in the binding of the ω-tail has an impact on the
oxygenation and cyclization events at the center of the fatty
acid. Therefore, we hypothesize that the alkynyl tail changes the
way aAA sits in the COX2 active site, resulting in similar O2
consumption despite its altered product profile, as defined in
these studies.
On the basis of all of the in vitro oxidation, we investigated
the viability of alkynyl probes for the analysis of lipid metabolite
detection and tracking in a biological setting. RAW264.7
macrophages are the prototypical cell line used to study lipid
metabolism because their lipid chemistry has been extensively
cataloged by the Lipid MAPS Consortium (www.lipidmaps.
org). Therefore, we investigated if the in vitro mCOX2
metabolites of aAA could be measured in cultured cells and
observed a11-HETE as the major aAA metabolite in cells, with
aPG and a11-8,9-HEET also detected, but at a much lower
level. This product ratio matches the kinetic efficiencies
measured in vitro for a11-HETE and aPG formation. Many
molecules have been reported to potentiate COX2 activity in
vitro, including free fatty acids.50 It is not unreasonable to think
that many of these species are present in cells, and could
potentiate the formation of a11-HETE as was seen in our data.
11-HETE has been reported in many animal and cell models as
a COX2-derived metabolite.51−54 Additionally, it has been
reported that hydroxy fatty acid metabolites of COX2,
Figure 6. Michaelis−Menten plots and relevant kinetic parameters for
mCOX2 metabolism of AA and aAA measuring the formation of
prostaglandins (A), alkynyl prostaglandins (B), and alkynyl 11-HETE
(C).
Leu384 to larger amino acids restrict endoperoxide formation
and prostaglandin ring closure, resulting in the generation of
multiple products including 11-HpETE and PGs. Their
proposed mechanism proceeds through several intermediates
that, when terminated, will give the AA-derived products similar
to those we have identified.
While we have been unable to solve a crystal structure of
aAA in a productive conformation in the mCOX2 active site,
we can look at other substrates to corroborate the idea that
aAA may not be binding properly in the active site, resulting in
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a
Scheme 1. Proposed Mechanism of mCOX2 Oxygenation of aAA
a
All products result from the same first steps, abstraction of the 13-(S)-hydrogen and addition of molecular oxygen to the 11-carbon. Pathway A
shows the formation of a11-HETE when the reaction is terminated before 9,11-endoperoxide is formation. After endoperoxide formation, closure of
the 5-membered prostaglandin ring results in pathway B and the formation of aPGG2, the precursor for all aPGs. However, if ring closure does not
occur (pathway C), the endoperoxde can cleave, resulting in an epoxide and an 11-alkoxyl radical. The alkoxyl radical can be terminated to form a11-
8,9-HEET.
including 11-HETE,55,56 can be further oxidized by cellular
AUTHOR INFORMATION
Corresponding Author
■
dehydrogenases to oxo fatty acids.36 These oxo fatty acids are
electrophilic, reacting with nucleophilic amino acids of proteins
n.porter@vanderbilt.edu
potentially changing cellular functions. Prostaglandins are not
Present Addresses
readily converted to electrophilic species; thus, hydroxy fatty
⊥Department of Chemistry, Imperial College of London, South
acids are the most viable method to study this chemistry in
Kensington Campus, Exhibition Road, London SW7 2AZ, UK.
cells. 11-Oxoeicosatetraenoic acid, the oxidized product of 11-
#Department of Molecular Toxicology, University of California
HETE, has been detected in cells, and shown to be
antiproliferative.55,56 This avenue of exploration is potentially
Berkeley, Berkeley, CA 94720, United States.
Author Contributions
viable using aAA as a part of a COX2-mediated metabolite
study, in the appropriate context.
∥W.N.B. and R.S. contributed equally to this research.
Notes
CONCLUSION
The authors declare no competing financial interest.
■
Collectively, these studies demonstrate that aPUFAs are
metabolized similarly to native PUFAs and represent a viable
tool for studying lipid distribution, metabolism, and reactions
between lipid metabolites and cellular macromolecules in many
physiological and pathophysiological models. While there are
some caveats regarding the enzymatic metabolism of aAA,
specifically the metabolism of these surrogates by the
cyclooxygenase enzymes, the nonenzymatic metabolism is
indistinguishable from that of the native lipid species.
Therefore, aPUFAs can be used as analogues for PUFAs,
especially in cellular disease models involving high amounts of
oxidative stress resulting in high levels of lipid oxidation.
ACKNOWLEDGMENTS
■
The authors would like to thank Don Stec at the Vanderbilt
University Small Molecule NMR Center. We would also like to
thank the Vanderbilt University Mass Spectrometry Resource
Center. For editorial input, we would like to thank James
Galligan. For funding sources we would like to thank the
National Institutes of Health Grants R01 HD064727 (N.A.P.)
and R37 CA087819 (L.J.M.), the National Science Foundation
Grant CHE-1057500 (N.A.P.), and the American Heart
Association Grant 13PRE17270009 (W.N.B.).
REFERENCES
■
ASSOCIATED CONTENT
* Supporting Information
This material is available free of charge via the Internet at
http://pubs.acs.org.
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