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Enzymes and Receptors of Prostaglandin Pathways with Arachidonic Acid-derived Versus Eicosapentaenoic Acid-derived Substrates and Products

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Dietary fish oil containing omega 3 highly unsaturated fatty acids has cardioprotective and anti-inflammatory effects. Prostaglandins (PGs) and thromboxanes are produced in vivo both from the omega 6 fatty acid arachidonic acid (AA) and the omega 3 fatty acid eicosapentaenoic acid (EPA). Certain beneficial effects of fish oil may result from altered PG metabolism resulting from increases in the EPA/AA ratios of precursor phospholipids. Here we report in vitro specificities of prostanoid enzymes and receptors toward EPA-derived, 3-series versus AA-derived, 2-series prostanoid substrates and products. The largest difference was seen with PG endoperoxide H synthase (PGHS)-1. Under optimal conditions purified PGHS-1 oxygenates EPA with only 10% of the efficiency of AA, and EPA significantly inhibits AA oxygenation by PGHS-1. Two- to 3-fold higher activities or potencies with 2-series versus 3-series compounds were observed with PGHS-2, PGD synthases, microsomal PGE synthase-1 and EP1, EP2, EP3, and FP receptors. Our most surprising observation was that AA oxygenation by PGHS-2 is only modestly inhibited by EPA (i.e. PGHS-2 exhibits a marked preference for AA when EPA and AA are tested together). Also unexpectedly, TxA(3) is about equipotent to TxA(2) at the TP alpha receptor. Our biochemical data predict that increasing phospholipid EPA/AA ratios in cells would dampen prostanoid signaling with the largest effects being on PGHS-1 pathways involving PGD, PGE, and PGF. Production of 2-series prostanoids from AA by PGHS-2 would be expected to decrease in proportion to the compensatory decrease in the AA content of phospholipids that would result from increased incorporation of omega 3 fatty acids such as EPA.
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Enzymes and Receptors of Prostaglandin Pathways with
Arachidonic Acid-derived Versus Eicosapentaenoic
Acid-derived Substrates and Products
*
}
Received for publication, April 16, 2007, and in revised form, May 11, 2007 Published, JBC Papers in Press, May 22, 2007, DOI 10.1074/jbc.M703169200
Masayuki Wada
‡1
, Cynthia J. DeLong
‡1
, Yu H. Hong
, Caroline J. Rieke
, Inseok Song
, Ranjinder S. Sidhu
,
Chong Yuan
, Mark Warnock
§
, Alvin H. Schmaier
§¶
, Chieko Yokoyama
,EmerM.Smyth**, Stephen J. Wilson**,
Garret A. FitzGerald**, R. Michael Garavito
‡‡
, De Xin Sui
‡‡
, John W. Regan
§§
, and William L. Smith
‡2
From the
Department of Biological Chemistry and
§
Department of Internal Medicine, University of Michigan,
Ann Arbor, Michigan 48109, the
Department of Medicine, Case Western Reserve University, Cleveland, Ohio 44106,
the
21st Century Center of Excellence Program, Department of Cellular Physiological Chemistry, Tokyo Medical and Dental
University, 1-5-45, Yushima, Tokyo-113-8549, Japan, the **Institute for Translational Medicine and Therapeutics, University of
Pennsylvania, Philadelphia, Pennsylvania 19104, the
‡‡
Department of Biochemistry and Molecular Biology, Michigan State
University, East Lansing, Michigan 48109, and the
§§
Department of Pharmacology and Toxicology, University of Arizona,
Tucson, Arizona 85721
Dietary fish oil containing
3 highly unsaturated fatty acids
has cardioprotective and anti-inflammatory effects. Prostaglan-
dins (PGs) and thromboxanes are produced in vivo both from
the
6 fatty acid arachidonic acid (AA) and the
3 fatty acid
eicosapentaenoic acid (EPA). Certain beneficial effects of fish oil
may result from altered PG metabolism resulting from increases
in the EPA/AA ratios of precursor phospholipids. Here we
report in vitro specificities of prostanoid enzymes and receptors
toward EPA-derived, 3-series versus AA-derived, 2-series pros-
tanoid substrates and products. The largest difference was seen
with PG endoperoxide H synthase (PGHS)-1. Under optimal
conditions purified PGHS-1 oxygenates EPA with only 10% of
the efficiency of AA, and EPA significantly inhibits AA oxygen-
ation by PGHS-1. Two- to 3-fold higher activities or potencies
with 2-series versus 3-series compounds were observed with
PGHS-2, PGD synthases, microsomal PGE synthase-1 and EP1,
EP2, EP3, and FP receptors. Our most surprising observation
was that AA oxygenation by PGHS-2 is only modestly inhibited
by EPA (i.e. PGHS-2 exhibits a marked preference for AA when
EPA and AA are tested together). Also unexpectedly, TxA
3
is
about equipotent to TxA
2
at the TP
receptor. Our biochemical
data predict that increasing phospholipid EPA/AA ratios in cells
would dampen prostanoid signaling with the largest effects
being on PGHS-1 pathways involving PGD, PGE, and PGF. Pro-
duction of 2-series prostanoids from AA by PGHS-2 would be
expected to decrease in proportion to the compensatory
decrease in the AA content of phospholipids that would result
from increased incorporation of
3 fatty acids such as EPA.
North American and Western European diets have relatively
high levels of
6 fatty acids (e.g. linoleic acid (1, 2)). As a result,
the most common highly unsaturated fatty acid is the C20
6
fatty acid, arachidonic acid (AA).
3
AA is present mainly at the
sn2-position of membrane phospholipids. Humans ingesting
fish oil enriched in
3 fatty acids show increased amounts of
eicosapentaenoic acid (EPA) in their membrane phospholipids
and an approximately corresponding decrease in the level of
AA. The ratio of
3 EPA/
6 AA in tissue phospholipids from
human populations averages less than 0.1 (1, 2) but can be
increased to almost 0.7 with palatable diets enriched in fish oil
(3, 4). An increased dietary intake of fish oil is cardioprotective,
anti-inflammatory, and anti-carcinogenic (2, 5–14).
The molecular basis for the health benefits of dietary fish oil
is almost surely multifactorial. For example,
3 fatty acids
attenuate responses of T-cells (15) and macrophages (16) to
agents working through cell surface receptors perhaps by
changing the composition of membrane microdomains (17,
18). One
3 fatty acid, docosahexaenoic acid (DHA), has been
shown to be essential in the development and maintenance of
neuronal functions including visual acuity. This may also be
related to the ability of DHA to change the physical properties
of membranes in a way that facilitates rhodopsin signaling (17,
19–22). Anti-arrhythmic effects of
3 fatty acids may relate to
their stabilizing effect on cardiac cell membranes and inhibi-
tion of the fast, voltage-dependent sodium and L-type calcium
currents (12). Nonesterified polyunsaturated fatty acids, partic-
ularly EPA, can also influence transcription acting through per-
oxisomal proliferator-activated receptors and sterol response
* This work was supported in part by National Institutes of Health (NIH) Grant
GM68848 (to W. L. S.), NIH HL56773 (to R. M. G.), and NIH NSRA HL075993
(to C. J. D.) and by a postdoctoral fellowship from the Heart and Stroke
Foundation of Canada (to R. S. S.). The costs of publication of this article
were defrayed in part by the payment of page charges. This article must
therefore be hereby marked advertisement in accordance with 18 U.S.C.
Section 1734 solely to indicate this fact.
}
This article was selected as a Paper of the Week.
1
These authors contributed equally to this work.
2
To whom correspondence should be addressed: Dept. of Biological Chem
-
istry, University of Michigan Medical School, 5301 Medical Science
Research Bldg. III, 1150 W. Medical Center Dr., Ann Arbor, MI 48109-0606.
Tel.: 734-647-6180; Fax: 734-764-3509; E-mail: smithww@umich.edu.
3
The abbreviations used are: AA, arachidonic acid; PG, prostaglandin; PGDS,
PGD synthase; mPGES-1, microsomal PGE synthase-1; PGHS, prostaglandin
endoperoxide H synthase; COX, cyclooxygenase; EPA, eicosapentaenoic
acid; DHA, docosahexaenoic acid; PRP, platelet-rich plasma; Tx, thrombox-
ane; TxAS, thromboxane A synthase; HHTrE, 12(S)-hydroxy-5,8,10-hepta-
decatrienoic acid; HHTE, 12(S)-hydroxy-5,8,10-heptadecatetraenoic acid;
hu, human; mu, murine; ov, ovine; IP, inositol phosphate; YPD, yeast
extract/peptone/dextrose; YEL, yeast extract-sodium lactate; DMEM, Dul-
becco’s modified Eagle’s medium; PBS, phosphate-buffered saline;
Ni-NTA, nickel-nitrilotriacetic acid; MES, 4-morpholineethanesulfonic acid.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 282, NO. 31, pp. 22254 –22266, August 3, 2007
© 2007 by The American Society for Biochemistry and Molecular Biology, Inc. Printed in the U.S.A.
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element binding protein-1c, major transcription factors con-
trolling lipid metabolism (2, 23, 24). Other proteins that can be
activated directly by polyunsaturated fatty acids, and thus,
whose activities might be altered by changes in EPA/AA ratios
include protein kinase C (25), NADPH oxidase (26), and a two-
pore domain K
channel (27). Polyunsaturated fatty acids such
as AA can promote apoptosis but the mechanism is not known
(28).
Finally, the eicosanoid pathways for lipid mediator forma-
tion; including the cyclooxygenase pathways, the 5-, 12-, and
15-lipoxygenase pathways, the P450 epoxygenase pathways,
and non-enzymic oxidative pathways; are influenced by
changes in EPA/AA ratios (29 –35). Anti-thrombotic, anti-in-
flammatory, and anti-carcinogenic effects of
3 fatty acids
could result, at least in part, from their ability to attenuate the
synthesis of specific eicosanoids and/or to alter the nature of
the eicosanoid products formed or to serve as precursors of
novel products such as isoprostanes and resolvins (32, 33,
35–38).
Prostanoids are synthesized via the cyclooxygenase pathway,
most commonly from AA, in response to various hormones and
physical stimuli (29). The pathway involves three stages: (a)
mobilization of AA from membrane phospholipids by cytosolic
phospholipase A
2
(cPLA
2
) sometimes in conjunction with
secretory sPLA
2
s; (b) conversion of AA to the prostaglandin
endoperoxide PGH
2
by prostaglandin endoperoxide H syn
-
thase-1 or -2 (PGHS-1 or -2) also known as cyclooxygenase-1 or
-2 (COX-1 or -2); and (c) isomerization of PGH
2
to a “2-series”
product, PGD
2
, PGE
2
-, PGF
2
, PGI
2
, or thromboxane A
2
(TxA
2
), by specific synthases. Newly formed PGs exit cells and
function primarily through G-protein-coupled receptors on
neighboring or parent cells to elicit responses. Because PGs act
at or near their sites of synthesis and are rapidly metabolized,
they are considered to be “local” hormones. Importantly, EPA
can serve as a substrate for PG formation generating “3-series”
PG products including PGD
3
, PGE
3
, PGF
3
, PGI
3
, and TxA
3
.
There is only limited biochemical information available on
the specificities of the enzymes and receptors of the prostanoid
pathways with EPA-derived versus AA-derived substrates and
products. Here we report studies that address this topic.
EXPERIMENTAL PROCEDURES
Materials—U46619,
17
U46619, PGI
2
, PGI
3
, iloprost,
SQ29548, AA, EPA, HHTrE, lipocalin PGD synthase (L-PGDS),
and hematopoietic (H) PGDS were purchased from Cayman
Chemicals (Ann Arbor, MI). [
3
H]SQ29548 was purchased from
PerkinElmer Life Sciences. [1-
14
C]AA and [1-
14
C]EPA were
from American Radiolabeled Chemicals. [
3
H]myoinositol and a
cAMP assay kits were from Amersham Biosciences. SQ22536
was from Biomol. Cell culture materials were purchased from
Invitrogen. Human fibrinogen,
-thrombin, and
IIa-thrombin
were purchased from Hematologic Technologies, Inc. Collagen
was obtained from Chronolog Corp. Complete protease inhib-
itor was from Roche Applied Science. BCA protein reagent was
from Pierce. Restriction enzymes were from New England Bio-
labs, Inc. Ni-NTA was from Qiagen. All other materials were
purchased from Fisher Scientific.
Expression, Purification, and Assay of PGHSs—Hexahisti-
dine-tagged (His
6
) ovine (ov) PGHS-1, murine (mu) murine
PGHS-2, and human (hu) PGHS-2 were expressed in S21 insect
cells and purified through Ni-NTA chromatography essentially
as described previously (39 41). His
6
-muPGHS-1 was
expressed also in insect cells but was unstable following Ni-
NTA chromatography so in the experiment using this enzyme,
the supernatant from centrifugation of solubilized cell pellets
was used for COX assays. Oxygen electrode assays for COX
activity were performed as detailed in previous reports (39
41). COX assays of purified enzymes utilizing radio thin layer
chromatography assays of PGH
2
or PGH
3
formation were
performed using [1-
14
C]AA and [1-
14
C]EPA as described
previously (41).
Preparation of Platelet-rich Plasma (PRP)—Platelets were
obtained from normal human donors who had not taken med-
ication during the 2 weeks prior to donation. Whole blood was
drawn into 3.8% sodium citrate (1:9; citrate:blood). The blood
was centrifuged at 180 g for 10 min at room temperature, and
PRP was transferred to a new tube. The remaining blood was
centrifuged at 1000 g for 10 min at room temperature to
obtain PRP. For aggregation studies using PRP, the platelet
count was determined on a Coulter counter (Model Z; Coulter
Electronics, Hialeah, FL) and adjusted with HEPES-Tyrode’s
buffer (137 m
M NaCl, 3 mM KCl, 12 mM NaHCO
3
, 0.34 mM
Na
2
HPO
4
, 14.7 mM HEPES, 0.35% dextrose, and 0.35% bovine
serum albumin, pH 7.4) to 2.2 to 2.5 10
8
platelets/ml. For
preparation of washed platelets, human platelets in PRP were
separated from plasma by gel filtration over Sepharose 2B col-
umns in HEPES-Tyrode’s buffer. The peak tubes were pooled,
and the platelet count was adjusted to 2.5 10
8
platelets/ml
before proceeding with platelet aggregation studies. Washed
platelets (400
l) were placed in a cuvette in the aggregometer
and stirred at 37 °C. The integrity of the washed platelets was
tested by their ability to be activated by collagen (1–5 mg/ml)
and
-thrombin (3 nM).
PGDS Assays—PGDS activity was determined essentially as
described previously (42, 43). First, PGH
2
or PGH
3
were pre
-
pared from 18
M [1-
14
C]AA or [1-
14
C]EPA, respectively, by
incubation for 20 s at room temperature with purified His
6
-
muPGHS-2 (30 unit) in 100
lof0.1M Tris-Cl, pH 8.0, contain-
ing2m
M phenol, 20
M hematin, and
-globulin (1 mg/ml).
PGH
2
/PGH
3
isomerization to PGD
2
/PGD
3
was initiated by the
addition of either lipocalin or hematopoietic PGDS (0.04 unit)
premixed with 0.1 m
M GSH, followed by incubation for 40 s at
room temperature. Reactions were quenched by adding 500
l
of diethylether/methanol/0.2
M citric acid (30:4:1). After vor-
texing for 10 s, the reaction mixture was centrifuged at 1000
g 10 min at 4 °C. An aliquot of organic extract (100
l) was
separated by thin layer chromatography on a silica gel plate
in ethyl acetate/2,2,4-trimethylpentane/acetic acid/water
(110:50:20:100). Regions of the plates migrating with PGD,
PGH, and other products were scraped into vials and radioac-
tivity quantified by liquid scintillation counting. One unit of
PGDS enzyme represents 1
mol of PGD
2
/min at 25 °C in 100
m
M Tris-HCl, pH 8.0, containing 1 mM GSH, 1 mg/ml
-glob-
ulin, and 40
M PGH
2
.
PG Enzymes and Receptors
AUGUST 3, 2007 VOLUME 282 NUMBER 31 JOURNAL OF BIOLOGICAL CHEMISTRY 22255
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Expression, Purification, and Assay of Human Microsomal
PGE Synthase-1 (hu mPGES-1)—The human PGES cDNA
(Invitrogen) was amplified using High Fidelity PCR kit (Invitro-
gen) with the 5-primer (with BspHI) GAA TTC ATC ATG
ATC CCT GCC CAC AGC CTG GTG A and the 3-primer
(with HindIII and His
6
-tag) CAT CCA AGC TTG TCA GTG
GTG GTG GTG GTG GTG CAG GTG GCG GGC CGC AAC.
The PCR product was purified with a QIAquick PCR purifica-
tion kit (Qiagen). The pRMGsp expression vector
4
was digested
by AflIII and XhoI and the amplified PGES PCR product was
digested with BspHI and HindIII. The digested DNA was iso-
lated by electrophoresis on a 1% agarose gel, and the DNA band
was purified with a QIAquick gel purification kit (Qiagen). The
digested and purified expression plasmid and His
6
-hu
mPGES-1 insert were ligated with T4 DNA ligase. The ligation
sample was transformed into DH5-
competent cells, and plas-
mids from positive colonies were sequenced to confirm the
expression construct pRMGsp-PGES-His
6
.
The pRMGsp-PGES-His
6
DNA with the aid DNA pAL9
(digested by PstI) were transformed into freshly made Schizos-
accharomyces pombe competent cells. After selection of posi-
tive colonies on MAA plates, the potential transformants were
screened twice on yeast extract/peptone/dextrose medium
(YPD)/G418 plates (containing G418 at 20
g/ml). Positive sin-
gle colonies were then grown in yeast extract-sodium lactate
(YEL) medium (with 10
g/ml G418) to make glycerol stocks,
which were stored at 80 °C.
For expression of His
6
-hu mPGES-1 in S. pombe,5mlofYEL
medium (with 10
g/ml G418) was inoculated with 200
lofa
glycerol stock culture and shaken for 24 h at 32 °C. The culture
was transferred into 50 ml of fresh YEL/G418 medium and
incubated at 32 °C for 48 h with shaking until the A
600
was
10–12. The culture was then transferred into 1 liter of YPD
medium (with 100
g of G419/ml) and incubated at 32 °C with
shaking until the A
600
exceeds 20 (48 60 h). The cells were
harvested by centrifugation at 3000 g for 15 min at 4 °C, and
the cell pellet was stored at 80 °C.
After completely thawing the cell paste in ice water, 5 ml of
lysis buffer (15 m
M Tris-HCl, 250 mM sucrose, 0.1 mM EDTA, 1
m
M reduced glutathione, pH 8.0) was added for per gram of cell
pellet. The resuspended cell pellet was lysed using an Emulsi-
Flex-C3 (at 20,000 –25,000 p.s.i. with two passes). The cell
lysate was centrifuged at 8000 g for 20 min at 4 °C. The mem-
brane-containing supernatant fraction was then centrifuged at
200,000 g for1hat4°C.Themembrane pellet was resus-
pended in loading buffer (50 m
M sodium phosphate, 300 mM
NaCl, 10% glycerol, pH 8.0) and dodecyl maltoside (Anatrace)
was added to a final concentration of 1%. The membrane frac-
tion was stirred for1hat4°Candthen centrifuged at 200,000
g for 1 h at 4 °C. Imidazole was added to the supernatant frac-
tion to a final concentration of 10 m
M, and the mixture was
loaded onto an Ni-NTA (Qiagen) column equilibrated with
loading buffer supplemented with 10 m
M imidazole; all column
buffers contained 0.05% dodecyl maltoside. After loading, the
column was washed with loading buffer containing 22 m
M
imidazole. The bound His
6
-hu mPGES-1 was eluted using load
-
ing buffer containing 200 m
M imidazole. Elution fractions con-
taining His
6
-hu mPGES-1 were pooled and concentrated using
a 30 molecular weight cutoff Centricon spin concentrator.
Microsomal Preparations of TxA Synthase (TxAS)—Mouse
(mu) TXAS was expressed in Sf21 insect cells as described pre-
viously (44, 45). The cells from a 250-ml culture were harvested
after 4 days, collected, and washed twice with ice-cold phos-
phate-buffered saline (PBS) and stored at 80 °C. Cell pellets
were thawed on ice and resuspended in 100 m
M Tris-HCl pH
7.4, 1 m
M EDTA and 1 Complete protease inhibitor. Cells
were disrupted by sonication and centrifuged at 10,000 g for
10 min at 4 °C. The supernatant was then centrifuged at
100,000 g for1hat4°C.Theresulting pellet was homoge-
nized in 10 m
M Tris-HCl, pH 7.4, 0.1 mM EDTA, and 20% glyc-
erol using a Dounce homogenizer and the protein concentra-
tion measured. The protein was used immediately for in vitro
synthesis of TxA
2
or TxA
3
.
PGE (EP), PGF (FP), and TxA/PGH (TP) Receptor Binding
HEK cell lines expressing various human PGE (EP2, EP3 (EP3II
isoform (46)), EP4 (47, 48)), PGF (FP; (49)), PGI (IP; (50)), and
TxA/PGH (TP; (51, 52)) receptors were maintained in Dulbec-
co’s modified Eagle’s medium (DMEM) containing 10% heat
inactivated fetal bovine serum, 250
g/ml Geneticin, 200
g/ml
hygromycin, 100
g/ml gentamicin and maintained at 37 °C
with 5% CO
2
. The cells were grown to 60% confluence and
harvested from five 100-mm tissue culture dishes by scraping
into the medium and centrifuged at 100 g for 5 min. The cell
pellets were washed once with ice-cold PBS, harvested, and
stored at 80 °C until membranes were prepared for competi-
tive binding assays.
The PCR was used to amplify the coding domain of the
huEP1 receptor (nucleotides 1–1209; GenBank
TM
accession
number L22647 (53)) from human kidney cDNA. The product
encoding the huEP1 was purified by agarose gel electrophoresis
and cloned into the EcoRV site of pcDNA3 to yield
huEP1/pcDNA3. The sequence of huEP1 in huEP1/pcDNA3
was verified by DNA sequencing. huEP1/pcDNA3 encoding the
huEP1 was transiently transfected into HEK293 cells using
Lipofectamine 2000 according to the recommendation of the
manufacturer, and cells were harvested 30 h post-transfection.
Cell pellets were stored at 80 °C and membranes prepared
from these cells were used to determine the relative affinities of
PGE
2
versus PGE
3
.
Membranes were prepared from HEK293 cells essentially as
described by Ungrin et al. (54). Briefly, cell pellets were thawed
on ice and resuspended in Buffer A (10 m
M HEPES/KOH, pH
7.4, with 1 m
M EDTA and 1 Complete protease inhibitor),
disrupted by sonication, and centrifuged at 10,000 g for 10
min at 4 °C. The supernatant was centrifuged at 100,000 g for
1 h at 4 °C. The pellet was homogenized in Buffer A, and ali-
quots of the suspended protein (50 –100
g) were used imme-
diately for binding assays.
Binding assays were performed in 200
lof10mM MES, pH
6.0, 1 m
M EDTA and 10 mM MgCl
2
. Binding isotherms were
performed for [
3
H]PGE
2
,[
3
H]PGF
2
, or the TPA antagonist
[
3
H]SQ29548 to estimate K
d
values for the different receptors
with the cognate 2-series PG ligand. A concentration of
3
H-la
-
4
D. Sui and R. M. Garavito, unpublished results.
PG Enzymes and Receptors
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beled ligand corresponding to the K
d
value for each receptor
was then used in competition binding assays with PGE
2
versus
PGE
3
, PGF
2
versus PGF
3
, or U46619 versus
17
U46619. Non
-
specific binding was determined in the presence of 10
M unla-
beled ligand. Samples were incubated at 30 °C for 1 h and then
filtered through Whatman GF/C glass filters. The filters were
washed three times with cold MES buffer (without EDTA) and
radioactivity measured by liquid scintillation counting. Recep-
tor binding data were analyzed by nonlinear regression in Ori-
gin. Statistical analyses were performed using Student’s t test
and/or ANOVA.
cAMP Assays—HEK cell lines expressing EP2 and EP4 recep-
tors that had been grown as described above in 6-well plates
were treated for 15 min at 37 °C with fresh DMEM containing
50 m
M isobutylmethylxanthine. Cells were then treated with
various concentrations of PGE
2
or PGE
3
for an hour. The treat
-
ments were terminated by scrapping cells into 0.5 ml of TE (50
m
M Tris-HCl, pH 7.5, containing 4 mM EDTA) and boiling for 8
min. After centrifuging the lysates, 50 ml of the supernatant
(from about 10
5
cells) was used for cAMP analysis using an
Amersham Biosciences cAMP assay system kit following the
instructions of the manufacturer. The samples were quantified
by scintillation counting and values for cAMP were calculated
from the cAMP standard curve.
Assays of Inositol Phosphates (IPs)—Assay of IPs was per-
formed by measuring receptor induced production of [
3
H] IPs
as described previously (55). Briefly, HEK293 cells expressing
EP3 or FP receptors were grown in 24-well plates as described
above and labeled by incubating overnight with 1
Ci of
[
3
H]myoinositol (Amersham Biosciences) per ml of DMEM.
The cells were treated with 10 m
M LiCl for 15 min prior to
adding various concentrations of PGE
2
/PGE
3
or PGF
2
/PGF
3
for 1 h. Assays were terminated by adding 3 ml of chloroform/
methanol/water (1:1:1) to each well. The whole cell lysates were
collected and centrifuged, and the resulting aqueous phase was
applied to a Dowex AG1-X8 anion exchange column (Bio-Rad)
to remove unincorporated [
3
H]myoinositol. The IPs were
eluted with 0.2
M ammonium formate, 0.1 M formic acid and
quantified by liquid scintillation counting.
Measurements of Intracellular Ca
2
—Intracellular Ca
2
concentrations were measured as described previously by
Fisher et al. (56). Nontransfected HEK293 cells and huEP1 plas-
mid-transfected HEK293 cells were incubated with 1
M Fura-
2/AM (Invitrogen) for 15 min before adding various concentra-
tions of PGE
2
or PGE
3
. The Ca
2
signals were measured using a
Shimadzu RF-5301 PC spectrofluorometer.
Platelet Aggregation—Aggregation assays were performed on
a Chronolog dual-channel aggregometer at 37 °C. The assays
were carried out with 360
l of undiluted human PRP, and the
compounds to be tested were added to obtain a final volume of
400
l. The assay was measured for 3 min following platelet
activation. Only PRP that was responsive to 20 40 n
M
IIa-
thrombin was used for experiments. Variable amounts of each
aggregatory compound were added to PRP to determine the
threshold concentration for platelet aggregation: U46619 or
17
U46619 (0.1 nM to 2
M) and collagen (1–2
g/ml). In some
experiments, inhibition of aggregation induced by 2
M
U46619 was examined using different amounts of anti-aggre-
gatory compounds: iloprost (0.1–10 n
M) and PGI
2
or PGI
3
(10
p
M to 2
M); flurbiprofen was added at 100
M. All compounds
were diluted from a stock solution in organic solvent into PBS,
pH 7.5, prior to the collection of blood, except for PGI
2
and
PGI
3
, which were prepared by removing a solid chemical stock
vial from 80 °C (stored for less than 1 week) and dissolving in
PBS less than 15 min before adding to PRP. The stability of each
prostacyclin was checked by comparing the anti-aggregatory
activity of 1 n
M at the end of the experiment to that of the
beginning of the experiment.
Platelet Aggregation by TxA
2
Versus TxA
3
—An aliquot of
PRP (370
l) was placed into a cuvette in the aggregometer
while stirring at 37 °C. The TxA
2
and TxA
3
were prepared just
before adding to PRP. In an Eppendorf tube, AA or EPA was
added to a final concentration of 5
M in reaction buffer (100
m
M Tris-HCl, pH 7.4, 1 mM phenol, and 10
M heme) contain-
ing 260 units of COX-2 to initiate the synthesis of PGH
2
or
PGH
3
. The sample was vortexed for 20 s. Then, 30
lofTxA
microsomal protein (500
g) was added to produce TxA
2
or
TxA
3
in a final volume of 100
l. The mixture was vortexed for
10 s, at which time 30
l was removed and added to the 370 ml
of PRP. Immediately, prior to the experiments using PRP, par-
allel reactions were performed with
14
C-labeled AA or EPA and
the products quantified by radio thin layer chromatography as
described above. This provided an accurate estimate of the var-
ious products being added in each case to the PRP.
RESULTS
Specificities of PGHS-1 and PGHS-2 with AA and EPA—As
shown in Fig. 1, purified ovPGHS-1 and muPGHS-2 oxygenate
AA with comparable catalytic efficiencies at concentrations of
1
M AA where reasonably precise O
2
electrode measure
-
ments of enzyme activity can be performed. ovPGHS-1 is essen-
tially inactive with EPA while muPGHS-2 can use EPA with
about 30% of the efficiency of AA in the range of 1–100 m
M.
5
A
solubilized preparation of muPGHS-1 expressed in baculovirus
showed qualitatively similar results to those shown for
ovPGHS-1 (data not shown); muPGHS-1 was unstable in our
hands, and so we were unable to analyze purified enzyme.
Although ovPGHS-1 was not active with low concentrations of
EPA, significant activity (10% of that with AA) was observed
when 15
M 15-hydroperoxyeicosatetraenoic acid was added to
the reaction mixtures (data not shown). Purified huPGHS-2
also showed results very similar to those in Fig. 1 for
muPGHS-2 (data not shown) when tested under essentially
identical enzyme and substrate conditions.
Results similar to those illustrated in Fig. 1 have been
reported by Kulmacz and co-workers using ovPGHS-1 (57, 58).
Moreover, the results with purified and semipurified enzymes
are consistent with studies comparing the utilization of AA ver-
sus EPA by microsomal huPGHS-1 and huPGHS-2 (59), where,
under optimal conditions, huPGHS-1 is 5% as active with EPA
as with AA while huPGHS-2 is 25–30% as active with EPA as
5
Critical micelle concentrations for AA and EPA determined in 0.1 M sodium
phosphate, pH 7.6. using fluorescence assays (i.e. with 1 m
M N-phenyl-1-
naphthylamine) were approximately 62 and 210 mM, respectively.
PG Enzymes and Receptors
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AA. It also is clear that EPA can be oxygenated by PGHS-1 in
intact cells in a manner that is peroxide dependent (60).
EPA and AA have similar K
m
values with PGHS-1 and
PGHS-2 (Fig. 1 and (57, 59)), and so EPA would be expected to
compete with AA for oxygenation. EPA/AA competition has
been shown previously with PGHS-1 (61), and the results in Fig.
2 confirm these findings. Half-maximal inhibition occurs with
equimolar AA versus EPA. Essentially identical results were
also obtained with a solubilized preparation of His
6
-
muPGHS-1. AA is about a 10 times better substrate than EPA
for ovPGHS-1 in vitro, and as shown in Fig. 3, inhibition of
oxygenation reflects primarily inhibition of AA oxygenation. A
5-fold excess of EPA caused 40% inhibition of [1-
14
C]AA oxy
-
genation by ovPGHS-1. This result is similar but not identical
to that of Fig. 2, which shows about 75% inhibition at these
concentrations of AA plus EPA. As expected, [1-
14
C]EPA was a
poor substrate for PGHS-1; however, EPA oxygenation was
augmented slightly by the presence of AA. Again, this is prob-
ably because hydroperoxide is being generated when AA is
present along with EPA in the reaction mixtures and hydroper-
oxides potentiate EPA oxygenation (59, 60, 62).
In contrast to the results obtained with PGHS-1, EPA was a
relatively poor inhibitor of AA oxygenation by PGHS-2 (Fig. 2).
For example, at equimolar AA and EPA concentrations, the
first point at which there was a statistically significant decrease
in the rate of oxygenation with muPGHS-2, there was only a
10% decrease in O
2
consumption and even with a 5-fold excess
of EPA there was less than a 20% decrease in oxygenase activity.
Based on the kinetic constants for muPGHS-2 (Fig. 1) and
huPGHS-2 for AA and EPA tested individually, one would
expect about a 35% lower oxygenation rate with 20
M AA plus
20
M EPA and a 60% decrease in the rate with 20
M AA plus
100
M EPA. To examine this inconsistency, we incubated puri-
fied enzymes with [1-
14
C]AA or [1-
14
C]EPA with and without
unlabeled competing substrate and measured the formation of
radioactive PGH
2
or PGH
3
(plus HHTE (12(S)-hydroxy-5,8,10-
heptadecatetraenoic acid) a degradation product of PGH
3
)
using radio thin layer chromatography (Fig. 3). When 20
M
unlabeled AA was added to reaction mixtures containing 20
M
[1-
14
C]EPA, oxygenation of EPA by PGHS-2 was inhibited by
70–90% (Fig. 3). In contrast, with 20
M [1-
14
C]AA and 100
M
unlabeled EPA there was only a modest inhibition (10%) of
AA oxygenation (Fig. 3, lower panel).
Thus, EPA acts as an effective inhibitor of AA oxygenation by
PGHS-1 but not PGHS-2, and indeed PGHS-2 shows a marked
and unanticipated preference for AA when presented with a
mixture of AA and EPA. The basis for the uneven competition
between AA and EPA with PGHS-2 is not clear. It may involve
FIGURE 1. COX activities of ovPGHS-1 (left panel) and muPGHS-2 (right panel) with AA and EPA. Specific activities were measured using purified His
6
-
tagged native ovPGHS-1 (8
g) or His
6
-tagged muPGHS-2 (ca. 8
g). Assays were performed on an O
2
electrode using standard COX assays with the indicated
concentrations of substrates.
5
Shown in this figure are results obtained with fatty acid substrate concentrations of 1–5
M; however, K
m
and V
max
values were
determined using fatty acid substrate concentrations of 1–100
M. Circles, AA; squares, EPA. Each data point represents a total of four assays involving two
separate enzyme preparations, and error bars represent mean S.D.
FIGURE 2. Inhibition of COX activities of ovPGHS-1, muPGHS-2, and
huPGHS-2 with AA and EPA. Enzyme activity was measured using a stand-
ard COX oxygen electrode assay with 6 8
gofHis
6
-tagged purified pro
-
teins, 20
M AA, and the indicated concentrations of EPA as described under
“Experimental Procedures” (39). Duplicate samples were assayed, and exper-
iments with three different enzyme preparations yielded similar results. Error
bars represent mean S.D.
PG Enzymes and Receptors
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half of sites activity with PGHS-2 (40). In the case of PGHS-2
but not PGHS-1, binding of certain fatty acids to one COX site
may facilitate oxygenation of AA bound to the other site.
In brief, our results with PGHSs show that (a)AAisan
equally good substrate for PGHS-1 and PGHS-2; (b) EPA is a
poorer substrate than AA for both PGHS-1 and PGHS-2 and a
particularly poor substrate for PGHS-1; (c) EPA is an efficient
inhibitor of AA oxygenation by PGHS-1 but not PGHS-2; and
(d) in the presence of EPA, PGHS-2 shows a marked preference
for AA.
Specificities of Lipocalin PGDS, Hematopoietic PGD Synthase
(hPGDS), and Microsomal PGES-1 (mPGES-1) toward PGH
2
Versus PGH
3
—Table 1
shows data obtained in estimating the
specificities of hPGDS, lPGDS, and mPGES-1 with PGH
2
versus
PGH
3
. PGH
2
and particularly PGH
3
are unstable, and so they
were generated in situ quantitatively from AA or EPA using an
excess of purified PGHS-2 and then a PGDS or PGES was added
immediately and the reactions continued for 20 40 s. PGH
3
was found to be less stable than PGH
2
, so it was necessary to add
more EPA than AA in generating the endoperoxides so that the
PGH
2
and PGH
3
concentrations were about the same when a
PGDS or PGES was added. The reactions were terminated
before 20% of the PGH was consumed, and the reactions were
performed with amounts of enzyme that provided approxi-
mately linear product formation with time. Somewhat different
V
max
and K
m
values have been reported for each of the various
PGD and PGE synthases we tested (63– 68). Because of this and
the technical difficulties associated with multiple assays with
unstable substrates and limited amounts of enzymes, we
elected to use an endoperoxide substrate concentration in the
range of 5
M for all of our assays, because as noted earlier,
5
5
M PGH
2
or PGH
3
would likely be as high a concentration as
would be encountered by a PGDS or PGES in an intact cell.
With all these provisos, the human versions of H-PGDS,
L-PGDS, and mPGES-1 were all more than 3-fold less active
with PGH
3
than with PGH
2
.
Quantitative data comparing the specificities of various pros-
tanoid biosynthetic enzymes with AA versus EPA derived sub-
strates is summarized in Table 2.
PGE and PGF Receptor Specificities—Membranes from HEK
cell lines expressing various PGE (EP) and PGF (FP) receptors
were used to determine the relative affinities of 2- versus 3-se-
ries PGE and PGF (Table 3). Except for the EP4 receptor, the
affinity of each receptor was significantly greater for the 2-se-
ries than the 3-series PGs. The most dramatic difference was
with the FP receptor, which had a 78-fold higher affinity for
PGF
2
than PGF
3
.
Fig. 4 shows the potencies of 2- versus 3-series PGs in elicit-
ing second messenger formation via the EP and FP receptors.
Differences were significant with EP1, EP2, EP3, and FP recep-
tors as determined by ANOVA for EP1 (p 0.001), EP2 (p
FIGURE 3. Oxygenation of [1-
14
C]EPA and [1-
14
C]AA in the presence and
absence of unlabeled AA or unlabeled EPA. Radio thin layer chromatogra-
phy assays were performed as described under “Experimental Procedures”
(41). The indicated substrates were mixed with 0.5
g(12 units) of the
purified His
6
-tagged PGHSs and the reactions continued for 30 s. Products
were extracted, separated, and visualized by autoradiography. The thin layer
plates were subsequently scraped and the amounts of radioactivity associ-
ated with the substrates and products determined by scintillation counting
and used to compute the relative rates indicated in the figure.
TABLE 1
Specificities of human hematopoietic and lipocalin PGD synthases and microsomal PGE synthase-1 toward PGH
2
versus PGH
3
Human hematopoietic PGD synthase (H-PGDS; 0.04 unit (1
g/assay)) and lipocalin PGDS (L-PGDS; 0.04 unit (18
g/assay)) PGD synthases were from Cayman Chemical
Co. Purified, solubilized His
6
mPGES-1 was expressed and purified as indicated under “Experimental Procedures” and 0.67–1
g used for the assays presented in the table.
1-
14
CPGH
2
or 1-
14
CPGH
3
was prepared by incubation of 18
M 1-
14
CAA or 1-
14
CEPA for 20 or 40 s. Hematopoietic or lipocalin PGDS or mPGES-1 was then added
and the incubations performed for 40 s for PGDSs or 20 sec for mPGES-1 under conditions in which the rate of conversion to product (PGD or PGE) was approximately
linear with time and added enzyme. 1-
14
CPGH
2
and 1-
14
CPGH
3
were generated in situ with purified muPGHS-2 or huPGHS-2 and H-PGDS, L-PGDS, or PGES was
added to initiate the reactions in a final volume of 0.1 ml. Products were extracted, separated at 4 °C by thin-layer chromatography, and quantified by scintillation counting.
Values in parentheses are numbers normalized for the indicated starting PGH
2
concentration.
PG synthase PGH
2
PGH
3
PGD
2
or PGE
2
PGD
3
or PGE
3
PGD
2
/PGD
3
or PGE
2
/PGE
3
nmol nmol nmol
H-PGDS (1
g) 8.0 4.4 (8.0) 0.51 0.063 (0.11) 4.6
H-PGDS (1
g) 3.8 3.8 0.26 0.043 6.0
H-PGDS (1
g) 3.8 4.7 (3.8) 0.21 0.035 (0.028) 7.5
L-PGDS (18
g) 3.8 4.7 (3.8) 0.056 0.021 (0.017) 3.3
mPGES-1 (0.67
g) 6.0 3.2 (6.0) 0.13 0.021 (0.039) 3.2
mPGES-1 (1.0
g) 4.4 4.5 0.069 0.025 2.7
PG Enzymes and Receptors
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0.005), EP3 (p 0.0014), and FP (p 0.0001); ANOVA indi-
cated no difference with the EP4 receptor (p 0.054). In all
cases the differences in potencies were less than the differences
in binding affinities. However, it is important to note that, with
the possible exception of the EP4 receptor, 3-series PGs were
partial agonists. Quantitative data on receptor potencies are
summarized in Table 4.
TP
Receptor Specificity toward U46619 and
17
U46619
Using membranes from HEK293 cells expressing the huTP
receptor, we determined the equilibrium dissociation constant
for the TP
antagonist [
3
H]SQ29548 to be 9
M. Equilibrium
competition binding assays with the PGH
2
/TxA
2
and PGH
3
/
TxA
3
analogues U46619 and
17
U46619, respectively, were
used to measure displacement of 5
M [
3
H]SQ29548 (Fig. 5
).
The IC
50
values for U46619 and
17
U46619 were identical (i.e.
200 and 210 n
M, respectively).
Comparison of U46619 and
17
U46619 Activation of Platelet
Aggregation—PRP from human donors was treated with
U46619 or
17
U46619 in amounts ranging from 0.1 to 2
M to
measure the potency of each compound in platelet aggregation
(Fig. 6). The threshold concentrations ranged from 0.5 to 0.8
M for U46619 and from 0.7 to 1
M for
17
U46619 for platelets
from four donors. For individual donors, the relative potencies
of the compounds were similar, with the threshold concentra-
tion of
17
U46619 consistently around 1.2-fold greater than
that of U46619. To confirm the specificities of the analogues for
the TP receptor, the platelets were incubated first with TP
antagonist SQ29548 (1
M) before addition of the diene or
triene analogue (2
M) or of collagen (2
g/ml). SQ29548 com-
pletely blocked aggregation by either U46619 or
17
U46619 and
inhibited collagen-induced aggregation by 50% (data not
shown).
Effects of TxA
2
Versus TxA
3
on Platelet Aggregation—Previ
-
ous studies had suggested that TxA
3
was essentially inactive in
platelet aggregation (69, 70), while our results with
17
U46619
suggested that TxA
3
would be pro-aggregatory. This assess
-
ment raised the possibility that the
17
U46619 analogue
behaves differently than authentic TxA
3
. Because TxAs have
very short half lives, we developed a system for synthesizing
TxA
2
or TxA
3
, which could then be added immediately to
platelets. In brief, AA or EPA were treated with excess
huCOX-2 to convert the fatty acids quantitatively to their
respective endoperoxides. Excess microsomal TxAS was then
added to quantitatively convert PGH
2
or PGH
3
to TxA
2
or
TxA
3
, respectively; TxAS is reported not to discriminate
between PGH
2
and PGH
3
(71). An aliquot of the reaction mix
-
ture was immediately added to PRP and platelet aggregation
was monitored. The amounts of the various products formed
from AA and EPA by huPGHS-2 and TxAS were monitored in
parallel reactions using [1-
14
C]AA or [1-
14
C]EPA.
A representative experiment is shown in Fig. 7. Reaction 2,
with a concentration of 78 n
M TxA
2
, induced irreversible aggre
-
gation. However, Reaction 4, which contained 45 n
M TxA
3
,
caused only a small reversible aggregation. When added to
TABLE 2
Specificities of PG biosynthetic enzymes with AA- versus EPA-derived substrates
Enzyme
AA-derived substrates EPA-derived substrates
Ref.
K
m
Rel. rates K
m
Rel. rates
cPLA
2
k
cat
/K
m
0.5
Est. k
cat
/K
m
0.5
74–76
sPLA
2
Kinetic values are highly context dependent; mechanism of reaction does not permit
discrimination among acyl groups.
79–81
ovPGHS-1 12
M 31 units/mg No activity without added
hydroperoxide; 10% of activity
with hydroperoxide; K
m
similar
to that of AA
57, 59, 83, 87;
Fig. 1
muPGHS-2 7.6
M 32 units/mg 4.6
M 9.2 units/mg 57, 59; Fig. 1
H-PGDS 0.5 m
M k
cat
21 s
1
17% activity with
5
M PGH
3
vs. PGH
2
63, 64; Table 1
L-PGDS 14
M k
cat
50 s
1
30% activity with
5
M PGH
3
vs. PGH
2
65, 66; Table 1
mPGES-1 17
M K
cat
50 s
1
30% activity with
5
M PGH
3
vs. PGH
2
67, 68; Table 1
mPGES-2 28
M 3.3
mol/min/mg ND 120
cPGES 14
M 190
mol/min/mg ND 109
PGFS Several enzymes proteins catalyzing the formation of PGF from PGH
2
have been
reported; it is not clear which are physiologically important.
103, 104
PGI synthase 30
M k
cat
5s
1
About the same activity with
PGH
3
and PGH
2
45, 71
TxA synthase 22
M k
cat
27 s
1
About the same activity with
PGH
3
and PGH
2
71
TABLE 3
EP and FP receptor specificities for PGE
2
vs. PGE
3
and PGF
2
vs. PGF
3
Membranes were prepared from HEK293 cell lines that stably express the human EP2, EP3, EP4, and FP receptors essentially as described by Ungrin et al. (54) as detailed
under “Experimental Procedures.”
Ligand
IC
50
10
9
M for ligand binding to receptor
EP1 EP2 EP3 EP4 FP
PGE
2
or PGF
2
15 6.2
a
5.3 0.86
a
7.7 1.6
a
4.9 1.4 2.3 0.70
a
PGE
3
or PGF
3
110 31 20 5.3 37 8.7 17 11 180 110
Relative affinities (PG
3
vs.PG
2
)
7.3 3.8 4.8 3.5 78
a
Denotes significant difference between 2- and 3-series as determined by Student’s t test. All binding assays were performed with duplicate samples with at least three different
membrane preparations.
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human PRP, PGH
3
was found to isomerize to PGD
3
with a t
1
2
of
90 s, unlike PGH
2
which has a t
1
2
of 350 s. PGD
3
is also reported
to be somewhat more potent at inhibiting aggregation than
PGD
2
(70), both of which cause an increase in cyclic AMP. To
block the putative inhibitory effects of any PGD
3
,1
M adeny
-
late cyclase inhibitor SQ22536 was added to the PRP prior to
the addition of Reaction 5. This unmasked an aggregatory effect
equal to that of the TxA
2
reaction (Reaction 2).
Both PGH
2
and PGH
3
are ligands of the TP receptor. How
-
ever, neither the diene nor the triene endoperoxide produced in
the PGHS-2-only reactions 1 and 3 (Fig. 7) was generated at
sufficient concentrations to induce irreversible aggregation.
Thus, residual endoperoxide in the reactions containing TxAS
could not be responsible for the irreversible aggregation seen in
Reactions 2 or 5.
Thromboxane and HHTrE are reported to be produced in
equimolar amounts by TxAS (71). However, in our in vitro reac-
tion, HHTrE and HHTE were produced at approximately twice
the concentration of their respective thromboxanes. The effect
of HHTrE on platelet aggregation was investigated by adding 1
M HHTrE to PRP alone or prior to the addition of either 2
M
U46619 or 2
g/ml collagen. HHTrE neither induced nor inhib-
ited platelet aggregation (results not shown). Likewise, up to
100
M malondialdehyde, another side product of the TxAS
reaction, had no effect on platelet aggregation (results not
shown). The results of studies with the TPa receptor and plate-
let aggregation suggest that TxA
2
or TxA
3
are approximately
equipotent.
Comparison of PGI
2
and PGI
3
as Inhibitors of Platelet
Aggregation—PGI
2
or PGI
3
(0.1–2
M) was added to PRP,
immediately followed by 2
M U46619. Preliminary experi-
ments with several donors were performed to optimize exper-
imental conditions, including the stabilization of the prostacy-
clins and determination of the approximate threshold
concentrations of each compound, before proceeding to per-
form dose response measurements with three donors (Fig. 8).
The initial slope of each curve in Fig. 8 was measured and
expressed as the percent inhibition of aggregation versus PGI
concentration. The average IC
50
values were 0.92 0.28 nM
and 1.30 0.18 nM for PGI
2
and PGI
3
, respectively. Thus, the
potencies of PGI
2
and PGI
3
in inhibiting platelet aggregation
are approximately the same confirming earlier results (69).
DISCUSSION
The goal of the studies reported here was to compare the
specificities and potencies of PG biosynthetic enzymes and
receptors toward AA-derived, 2-series versus EPA-derived,
3-series substrates and products. We reason that this new infor-
mation will contribute to understanding whether any of the
reported beneficial health effects of dietary
3 fish oil fatty
acids are mediated through PG pathways. Our biochemical
results along with those of others are summarized in Fig. 9 and
Tables 2 and 4.
FIGURE 4. Potencies of 2- versus 3-series PGs in eliciting second messen-
ger formation by various EP receptors and the FP receptor. HEK cells
expressing the indicated EP1, EP2, EP3, EP4, and FP receptors were used to
measure changes in cAMP or IP formation, or Ca
2
mobilization with the
indicated concentrations of PGE
2
, PGE
3
, PGF
2
or PGF
3
. Details of the exper
-
imental protocols are presented under “Experimental Procedures.” All assays
were performed in duplicate or triplicate with at least three cell preparations
and data analyzed using ANOVA.
PG Enzymes and Receptors
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PhospholipasescPLA
2
is the key phospholipase involved
in AA release in most PG forming cells (72). Previous studies
have shown that cPLA
2
exhibits specificity toward AA and
EPA esterified at the 2-position of phospholipids in comparison
to other 2-position acids such as linoleate and oleate (73–76).
Moreover, although EPA- and AA-containing phospholipids
are equally good substrates for cPLA
2
, DHA-containing phos
-
pholipids are essentially inactive with cPLA
2
(76–78). Certain
sPLA
2
forms can also participate in PG biosynthesis (79, 80).
Because of the nature of the interaction with its substrates,
sPLA
2
does not discriminate among 2-position acyl groups
(81). In short, neither cPLA
2
nor sPLA
2
appears to differentiate
between the acyl chains of AA versus EPA.
PGHS-1 and PGHS-2—PGHS-1 and PGHS-2 both exhibit
specificity toward AA versus EPA. As reported previously by
others and us, both enzymes have very similar K
m
and V
max
values with AA (57, 59). PGHS-2 oxygenates EPA at about 30%
of the rate of AA. Purified PGHS-1 is not active with EPA unless
an exogenous hydroperoxide is added to the reaction mixture.
This is a consequence of the higher hydroperoxide requirement
of PGHS-1 versus PGHS-2 (57, 82, 83). PGHS-1 present in
platelets cells does oxygenate exogenously supplied EPA albeit
at a low rate in the presence of alkyl hydroperoxides (60). It is
not clear whether the hydroperoxide concentration in cells is
usually sufficient to support EPA oxygenation or whether there
are differences in hydroperoxide concentrations among cell
types (57, 84).
The behavior of PGHS-1 with AA plus EPA is consistent with
the kinetic properties of the enzyme determined with AA and
EPA individually; thus, EPA is a reasonably good inhibitor of
AA oxygenation by PGHS-1 as was originally reported by Lands
and co-workers (61). A comparison of the crystal structures of
EPA and AA with PGHS-1 suggests that EPA prefers to bind in
a catalytically incompetent conformation in the PGHS-1
cyclooxygenase site and competes with AA for binding
(85–87).
PGHS-1 mediated biological events include platelet aggrega-
tion and parturition (88, 89) and certain types of acute inflam-
mation (79, 90, 91). Cellular events involving PGHS-1 may be
dampened when EPA/AA ratios in phospholipids are
increased. At an EPA/AA ratio of 1.0, one would expect that
there would be 50% less AA to be mobilized from phospholipids
by cPLA
2
and that PGHS-1 would function at only 50% of max
-
imal efficiency because of inhibition of AA oxygenation by EPA;
however, this may well be an oversimplification because the
concentrations of enzymes, receptors, and substrates in intact
cells are unknown.
One of our most surprising observations was that PGHS-2
preferentially oxygenates AA when EPA and AA are tested
together. The results observed when PGHS-2 is mixed with
EPA plus AA cannot be explained based on the simple kinetic
properties of PGHS-2 with AA or EPA individually. The bio-
chemical basis for the selectivity of PGHS-2 for EPA versus AA
when the substrates are together may relate to the half of sites
activity of the enzyme (40). One possibility is that EPA binds
one of the two cyclooxygenase sites of the PGHS-2 dimer and
elicits an allosteric effect on the other cyclooxygenase site caus-
ing it to preferentially bind and oxygenate AA. If this is true and
also applicable to any fatty acid, it could explain why PGHS-2
can preferentially oxygenate AA at low substrate concentra-
tions when AA represents a small part of the available fatty acid
pool in cells (92–94). A situation like this could occur in so-
called late phase PG synthesis when an sPLA
2
is the operative
phospholipase (95).
PGs are importantly involved in inflammation (79, 91,
96–98), and in this context PGHS-2 is the most important
PGHS isoform (99, 100). Based on our biochemical studies, a
decrease in the formation of 2-series PGs via PGHS-2 would be
FIGURE 5. Binding of U46619 and
17
U46619 to the huTP
receptor. An
equilibrium competition binding assay was performed for U46619 and
17
U46619 versus [
3
H]SQ29548 (5
M) as the radioligand. The average K
d
and
B
max
values for [
3
H]SQ29548 binding to TPa/HEK293 microsomes from three
experiments was 10.2 3.1 n
M and 3800 980 fmol/mg of protein, respec-
tively. The average IC
50
values for U46619 and
17
U46619 were 200 and 210
n
M, respectively, from an average of three experiments depicted in the figure.
TABLE 4
PG receptors and their affinities or potencies with AA- vs. EPA-derived PGs
Receptor EC
50
2-series PG
EC
50
3-series PG
Second messenger Cell/Tissue Ref.
nM nM
DP1 109 64 G
s
, cAMP
Platelets 70, 111
DP2 7 8 G
i
,Ca
2
Eosinophil 111
EP1 17 34
a
G
q
,Ca
2
EP1 HEK cell Table 3, Fig. 4; PGE
3
is partial agonist
EP2 4.3 11
a
G
s
, cAMP
EP2 HEK cell Table 3, Fig. 4; PGE
3
is partial agonist
EP3 62 190
a
G
i
, reduced cAMP, IP increases
EP3 HEK cell Table 3, Fig. 4; PGE
3
is partial agonist
EP4 0.58 3.5 G
s
, cAMP
EP4 HEK cell Table 3, Fig. 4; PGE
3
is partial agonist
FP 14 67
a
G
q
,Ca
2
FP HEK cell Table 3, Fig. 4; PGF
3
is partial agonist
IP 0.92 1.3 G
s
, cAMP
Platelets Fig. 8
TP 650 for U44619 850 for 17-U46619 G
q
,Ca
2
Platelets Fig. 6
a
Denotes significant difference. ANOVA (p 0.05).
PG Enzymes and Receptors
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by guest on September 23, 2015http://www.jbc.org/Downloaded from
expected to occur only to the extent that AA levels in phospho-
lipids were decreased by
3 fatty acids supplanting AA.
PGD, PGE, PGF, PGI, and TxA Synthases—There are seven
different synthases reportedly involved in the conversion of PG
endoperoxides to what are considered to be the biologically
active PGs. Including the results from the present studies, there
are now data on the specificities toward PGH
2
versus PGH
3
for
H-PGDS, L-PGDS, mPGES-1, PGIS, and TXAS (Fig. 9 and
Table 2). Still lacking is information on cPGES (101) and
mPGES-2 (101, 102) and various putative NADPH-dependent
and GSH-dependent PGFSs (103, 104).
PGDSs and mPGES-1 are about
one third as active with PGH
3
as
PGH
2
. mPGES-1 does play a role in
inflammation (67, 96, 97, 105, 106),
and in principle, elevated levels of
EPA could increase PGH
3
produc
-
tion and decrease PGH
2
formation,
and the net effect would be
decreased formation of PGE
2
with
less than a corresponding increase
in PGE
3
. However, PGHS-2 appears
to be the most relevant enzyme in
inflammation and, as discussed
above, increases in EPA have a rela-
tively modest effect on the forma-
tion of PGH
2
. PGE formation can
occur via PGHS-1 and mPGES-1
(107). There are functions such as salt and water metabolism in
the kidney that involve these two enzymes, and renal PGE
2
synthesis is diminished with no detectable production of PGE
3
in rats fed diets having elevated levels of fish oil (108).
PGIS and TXAS are reported to be similarly reactive with
PGH
2
and PGH
3
(45, 71). This suggests that any effects of
changes in tissue EPA/AA levels on PGI and TxA formation
would occur primarily at the level of PGHSs and not PGIS or
TXAS.
Prostanoid Receptors—There are nine G-protein-linked PG
receptors. Previous comparisons of receptor specificities for
the 2- versus 3-series PGs had been performed for the DP1 (70),
DP2 (110, 111), EP1 (54), TP (69), and IP (69, 112) receptors. In
all cases except for the TP receptor, there was little or no dif-
ference in the potencies of the 2- versus 3-series PGs. We per-
formed both binding measurements and measurements of
receptor potencies for all of the human receptors except the IP
and DP receptors. In the case of the IP receptor, we analyzed
potencies of purified PGI
2
versus PGI
3
using human platelets.
The EP1, EP2, and EP3 receptors bound less well and were
less responsive to PGE
3
than PGE
2
. As recently reviewed by
Narumiya and coworkers (113), each of these receptor subtypes
participate in a large number of functions each of which has the
potential to be affected by increased tissue EPA/AA levels.
However again, it should be noted that functions most likely to
be affected are those that would be mediated via PGHS-1 and
mPGES-1.
The FP receptor is known to be involved in parturition. Mice
lacking cPLA
2
(114, 115), PGHS-1 (88, 89), or the FP receptor
(116) have failures of parturition. Interestingly, this is also a
characteristic of essential fatty acid deficiency that can be over-
come with omega-6 but not omega-3 fatty acids (117–119).
This could be accounted for by the low activity of PGHS-1 with
EPA and the low potency of PGF
3
with the FP receptor. It is
not clear what enzyme is responsible for PGF
2
formation in
vivo, so we did not examine the PGF synthases that have been
described (103) for their specificities toward PGH
2
versus
PGH
3
.
An unexpected observation in our studies of PG receptors
was that TxA
3
is almost as active as TxA
2
with the TP receptor.
Earlier studies indicating that TxA
3
is inactive in platelet aggre
-
FIGURE 6. Comparison of potencies of U46619 and
17
U46619 for platelet aggregation. Human PRP
(2.25 10
8
platelets per 0.4 ml) was treated with various concentrations of either U46619 or
17
U46619 as
indicated, and platelet aggregation as indicated by the change in light transmission was recorded on an
aggregometer. Shown is a representative result of four different donors. The concentration at which irrevers-
ible aggregation occurred for
17
U46619 was 1.2 times higher than for U46619 with platelets from each donor.
FIGURE 7. Platelet aggregatory properties of AA- versus EPA-derived
COX-2 and TXAS products. Enzyme reactions were initiated by adding 5
M
AA (Reactions 1 and 2) or EPA (Reactions 3, 4, and 5) to a reaction mixture
containing 750 units of purified His
6
-tagged huPGHS-2. A microsomal
huTXAS preparation (540 mg of protein) (Reactions 2, 4, and 5) or microsomal
buffer (Reactions 1 and 3) was added and the sample vortexed for an addi-
tional 10 s. An aliquot (30
l) of each reaction mixture was immediately added
to PRP and platelet aggregation was measured. The adenylate cyclase inhib-
itor SQ22536 was added to the PRP 1 min prior to addition of the reaction
mixture for Reaction 5. To calculate the concentrations of products from the
COX-2/TXAS reactions that were added to PRP, [1-
14
C]AA, or [1-
14
C]EPA was
used in place of the unlabeled substrate. The radiolabeled products were
separated by TLC and the bands corresponding to fatty acid, HETE, (12-hy-
droxyheptadecatrienoic acid), HHT, PGH, and TxB were scraped and quanti-
fied by liquid scintillation counting and the final concentration of each prod-
uct (n
M) in the PRP was calculated. The concentration of TxA
2
or TxA
3
was
corrected for degradation to TxB
2
or TxB
3
during the 10 s incubation as
described under “Experimental Procedures.”
PG Enzymes and Receptors
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gation (69) were probably compromised by the formation of
PGD
3
from PGH
3
and PGD
3
being a potent anti-aggregatory
compound (70). In general, PGH
3
appears to be significantly
less stable than PGH
2
in the aqueous systems used for our
enzyme assays; PGH
3
is rapidly converted to HHTE and malon
-
dialdehyde whereas spontaneous
conversion of PGH
2
to the homolo
-
gous products is relatively slow.
This is apparent in Fig. 3 where
there is an accumulation of HHTE
but not HHTrE.
To the extent that we have dis-
cussed our biochemical data in the
context of the biological changes
seen with dietary fish oil, we have
assumed simple linear relation-
ships based on K
m
, V
max
, and EC
50
values for the various enzymes and
receptors. All of these values were
obtained under optimal in vitro
conditions. Obviously, what occurs
in vivo cannot yet be predicted
with any certainty because the
ratios of enzymes and receptors
to substrates and agonists in-
volved in PG signaling may well be
different in vivo (89). There may
also be other eicosanoid media-
tors, including those derived from
omega-3 fatty acids that are importantly involved in PG
signaling (34, 35).
Acknowledgments—We thank Dr. Stephen C. Fischer and Dan Foster
for guidance in making Ca
2
measurements and Dr. Nisha Palackal
of Cayman Chemical Company for help with PGDS assays. We thank
Dr. William E. M. Lands for his advice and encouragement during the
course of these studies.
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PG Enzymes and Receptors
22266 JOURNAL OF BIOLOGICAL CHEMISTRY VOLUME 282 NUMBER 31 AUGUST 3, 2007
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William L. Smith
Garavito, De Xin Sui, John W. Regan and
Wilson, Garret A. FitzGerald, R. Michael
Yokoyama, Emer M. Smyth, Stephen J.
Warnock, Alvin H. Schmaier, Chieko
Ranjinder S. Sidhu, Chong Yuan, Mark
Hong, Caroline J. Rieke, Inseok Song,
Masayuki Wada, Cynthia J. DeLong, Yu H.
Substrates and Products
Eicosapentaenoic Acid-derivedVersus
Pathways with Arachidonic Acid-derived
Enzymes and Receptors of Prostaglandin
Lipids and Lipoproteins:
doi: 10.1074/jbc.M703169200 originally published online May 22, 2007
2007, 282:22254-22266.J. Biol. Chem.
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... The increase in MUFA content was in response to the loss of membrane fluidity originated by the drop of PUFA double bonds; by upregulating the activity of the Stearoyl-CoA-Desaturase, cells were able to partially counterbalance the drop in membrane fluidity through the synthesis of MUFA. The slight increase in ω-3 content was probably due to both the relatively high amount of ω-3 fatty acid supplied by the FBS and to the competition for the same enzymes by ω-6 and ω-3 fatty acid synthetic pathways [47,48] (pp. 77-80, pp. ...
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... Notably, the high levels of EPA in the retinal and ON tissues prove the successful incorporation and critical role of EPA in protecting the aforementioned tissues. EPA was shown to compete with AA and inhibit AA oxygenation in vitro, thus inhibiting the inflammatory pathway which involves prostaglandin (PG)-D, PGE, and PGF (McCappin et al., 1987;Wada et al., 2007). ...
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The purpose of this study was to evaluate the neuroprotective effects of omega-3 polyunsaturated fatty acid (ω3-PUFA) supplementation in a mouse model of OPA1-associated autosomal dominant optic atrophy (ADOA). The blood level of arachidonic acid (AA) and eicosapentaenoic acid (EPA) served to adjust the treatment dosage (AA/EPA = 1.0–1.5). Eight-month-old mice were allocated to four groups (n = 20/group): the ω3-PUFA-treated Opa1enu/+, untreated Opa1enu/+, ω3-PUFA-treated wild-type and untreated wild-type groups. Treated mice received the ω3-PUFAs, EPA and docosahexaenoic acid (DHA; 5:1 ratio) by daily gavage for 4 months based on the measured AA/EPA ratio. Blood, retina and optic nerve (ON) fatty acid levels were determined by gas chromatography, and the retina and ON were histologically examined. Western blotting and/or immunohistochemistry was performed to analyse retinal mediators involved in Opa1-mutation-mediated apoptosis, inflammation and oxidative stress. Increased EPA and reduced AA levels were primarily observed predominantly in the blood and retinal tissues, and a similarly high EPA level tended to be observed in the ONs of ω3-PUFA-treated mice. Retinal ganglion cell and ON axonal densities were higher in both mouse strains upon ω3-PUFA treatment than in the corresponding untreated groups. Caspase-3 expression analysis showed fewer apoptotic retinal cells in both groups of treated mice. Decreases in inflammatory microglia and astrocytes activation and proapoptotic Bcl-2-associated X protein (Bax) expression were noted in the treated groups, with no difference in the antioxidant superoxide dismutase-2 expression. ω3-PUFA supplementation had neuroprotective effects on the retinas of Opa1enu/+ and wild-type mice via blockade of microglia and astrocytes activation and suppression of Bax and caspase-3. Our findings indicated that inhibition of oxidative stress may not be involved in ω3-PUFA-mediated neuroprotection. These novel findings support the use of ω3-PUFAs as a beneficial therapy in the occurrence of ADOA, posing the basis for future clinical trials to confirm these observations.
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The ω3 polyunsaturated fatty acids (PUFAs) are known to have beneficial effects on health and diseases, and hence their intake is encouraged. However, it remains unknown as to how ω3 PUFAs affect female reproduction processes, in which ω6 PUFA-derived prostaglandin (PG) E2 and PGF2α play crucial roles. We therefore compared female reproductive performance between ω3 PUFA-biased linseed oil diet-fed (Lin) mice and ω6 PUFA-biased soybean oil diet-fed (Soy) mice. In Lin mice, the uterine levels of arachidonic acid (AA) and eicosapentaenoic acid (EPA) were 0.42 fold and 16 fold of those in Soy mice, respectively, with the EPA/AA ratio being 0.7 (vs 0.02 in Soy mice). Lin mice showed no alterations in any of the fertility indexes, including luteolysis and parturition. The uterine PG synthesis profiles of Lin mice were similar to those of Soy mice, but the levels of PGF2α and PGE2 were 50% of those in Soy mice, as a result of the increased EPA/AA ratio. PGF3α and PGE3 were undetectable in the uterine tissues of Soy and Lin mice. Interestingly, in Lin mice, ‘luteolytic’ PGF2α synthesis was considerably maintained even in the ω6 PUFA-reduced condition. These results suggest the existence of an elaborate mechanism securing PGF2α synthesis to a level that is sufficient for triggering luteolysis and parturition, even under ω6 PUFA-reduced conditions.
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Little evidence exists for stored prostaglandins in tissues (Jouvenaz, Nugteren, Beerthuis and Van Dorp, 1970) and a capacity of a tissue to release prostaglandins seems to reflect principally the capacity for biosynthesis of these compounds. Since prostaglandins represent a diverse family of oxygenated derivatives formed from certain polyunsaturated fatty acids, the Fig. 3.1 Formation and degradation of prostaglandins. biosynthesis of prostaglandins consists of many steps that involve a variety of different enzymes and cofactors. Inhibition of prostaglandin synthesis may thus occur by blocking any of the several processes that are involved. A brief summary of some of the metabolic events leading to the different prostaglandins is given in Figure 3.1. Because of the diversity of these processes and the stimuli and mediators that appear to modify these reactions it is expected that a wide variety of chemical agents will diminish the formation of prostaglandins in vivo. We will indicate some of the events of prostaglandin biosynthesis known to be influenced by chemical agents, and will suggest a logical framework for comparing the relative inhibitory activity of these different agents.
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Since the first AHA Science Advisory “Fish Consumption, Fish Oil, Lipids, and Coronary Heart Disease,”1 important new findings, including evidence from randomized controlled trials (RCTs), have been reported about the beneficial effects of omega-3 (or n-3) fatty acids on cardiovascular disease (CVD) in patients with preexisting CVD as well as in healthy individuals.2 New information about how omega-3 fatty acids affect cardiac function (including antiarrhythmic effects), hemodynamics (cardiac mechanics), and arterial endothelial function have helped clarify potential mechanisms of action. The present Statement will address distinctions between plant-derived (α-linolenic acid, C18:3n-3) and marine-derived (eicosapentaenoic acid, C20:5n-3 [EPA] and docosahexaenoic acid, C22:6n-3 [DHA]) omega-3 fatty acids. (Unless otherwise noted, the term omega-3 fatty acids will refer to the latter.) Evidence from epidemiological studies and RCTs will be reviewed, and recommendations reflecting the current state of knowledge will be made with regard to both fish consumption and omega-3 fatty acid (plant- and marine-derived) supplementation. This will be done in the context of recent guidance issued by the US Environmental Protection Agency and the Food and Drug Administration (FDA) about the presence of environmental contaminants in certain species of fish. ### Coronary Heart Disease As reviewed by Stone,1 three prospective epidemiological studies within populations reported that men who ate at least some fish weekly had a lower coronary heart disease (CHD) mortality rate than that of men who ate none.3–6⇓⇓⇓ More recent evidence that fish consumption favorably affects CHD mortality, especially nonsudden death from myocardial infarction (MI), has been reported in a 30-year follow-up of the Chicago Western Electric Study.7 Men who consumed 35 g or more of fish daily compared with those who consumed none had a relative risk of death from CHD of 0.62 and a relative risk of nonsudden death from MI of 0.33. In an …
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1. Chicks were reared to 3 weeks in a germ-free or conventional environment on diets containing 50% raw or heated soya-bean meal. Body-weights and pancreas weights were determined and the proteolytic and amylolytic enzymes in the pancreas were measured. In one experiment the contents of the intestinal tract were analysed for their content of nitrogenous compounds. 2. The growth of chicks given raw soya was depressed below that of controls given heated soya, and the growth depression was significantly greater in the conventional compared with the germ-free environment. 3. The pancreases of chicks given raw soya were significantly larger than those given heated soya. The degree of enlargement was similar in both environments. The concentration of α-amylase was lower in the enlarged pancreases, but the total amount per pancreas was similar to that in the normal pancreases. Conversely, the concentration of proteolytic enzymes was the same or greater in the enlarged pancreases, which had significantly greater total content of proteases than did the normal pancreases. 4. There was evidence of impaired protein digestion in the birds given the raw soya diets. Their gut contents contained larger amounts of insoluble matter, with higher nitrogen content, and the soluble fraction contained more nitrogen present as ‘protein’, with correspondingly less ‘peptide’ and ‘amino acid’, than did similar samples from birds given heated soya. No differences were observed in this respect between germ-free and conventional birds. 5. It was concluded that the gut microflora exacerbated the growth-depressing effect on chicks of raw soya meal. Possible explanations are advanced.