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Perinatal Research Laboratories, Departments of Obstetrics/Gynecology (T.D., J.A.S., R.R.M.) and Meat/Animal Sciences (R.R.M.), University of Wisconsin-Madison, Madison, Wisconsin 53715; and Center for Perinatal Biology (L.Z.), Loma Linda University School of Medicine, Loma Linda, California 92350
Address all correspondence and requests for reprints to: Ian M. Bird, Ph.D., University Wisconsin-Madison, Department of Obstetrics and Gynecology, Perinatal Research Laboratories, 7E Meriter Hospital/Park, 202 South Park Street, Madison, Wisconsin 53715. E-mail: imbird{at}facstaff.wisc.edu
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 Top Abstract IntroductionMaterials and MethodsResults and DiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResults and DiscussionReferences |
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Although Ca2+ and members of the
mitogen-activated protein kinase (MAPK) pathway probably regulate
cPLA2 activity, as seen in other endothelial
cells (9), the role of Ca2+ in
activation of eNOS is less clear, and activation of eNOS may indeed
more closely correlate to an ability to activate extracellular
signal-regulated protein kinases 1 and 2 (ERK-1/2) for several agonists
of heptahelical or growth factor receptors (8). In initial
characterizations of eNOS, it was quickly realized that this enzyme
requires Ca2+ for activity, but an enzyme’s
requirement for Ca2+ does not always mean it is
physiologically regulated by changes in cytosolic
Ca2+. Intracellular Ca2+
typically ranges from 50–1000 nM. If the
enzyme’s requirement for Ca2+ is below or above
that range it will either be permanently activated or never activated
by Ca2+ elevation, respectively. However, studies
of proteins such as cPLA2 have shown that
alteration of protein structure through phosphorylation can result in a
dramatic increase in Ca2+ sensitivity and so a
shift of the Ca2+ dose response to the left,
which in turn results in enhanced activation even at resting cytosolic
[Ca2+]i (9).
Thus, phosphorylation alone can result in a marked increase in activity
without additional increases in free cytosolic
Ca2+. The situation with eNOS is more
complicated, and not yet fully understood. It is clear that eNOS can be
activated by Ca2+ and calmodulin (CaM) and that
subsequent complete removal of Ca2+ from the
media abolishes such activation of eNOS in vitro, but it is
also becoming increasingly clear that either inhibition of tyrosine
phosphatases (10), or shear stress
(11) can also increase eNOS phosphorylation and activity
independently of an increase in
[Ca2+]i in endothelial
cells. Recent studies in pulmonary artery endothelial cells as well as
COS-7 cells overexpressing estrogen receptor 
(ER) suggest that
eNOS can be activated by estrogen in a manner associated with ERK
activation, which in turn can be blocked by the MEK inhibitor PD98059
(12). eNOS is known to have a number of putative
phosphorylation sites, but it is not known if ERK-1 or ERK-2, or indeed
other members of the MAPK family directly regulate this enzyme’s
activity. Additional studies have shown that binding to caveolin-1 (a
scaffold protein located in caveolae) results in potent inhibition of
eNOS (13), and more detailed recent studies suggest that
caveolin-1, Ca2+/CaM and the site of eNOS
phosphorylation may all lie within the same
Ca2+/CaM binding domain of eNOS
(14). Thus at rest eNOS is bound to caveolin-1 at the
caveolae (plasma membrane invaginations rich in signaling proteins and
substrates) and is therefore inactive. Two events, however, seem able
to out-compete or destabilize the binding of eNOS to the caveolin and
phospholipid bilayer, namely competitive binding of
Ca2+/CaM formed on increases in cytosolic
Ca2+, or phosphorylation of the same domain by a
mechanism which in many cells seems to involve a
Ca2+-independent protein kinase C, acting through
a MAPK pathway (14, 15). Thus eNOS itself, and more
specifically the Ca2+/CaM binding domain, is a
possible point of convergence of Ca2+/CaM
vs. protein kinases signaling. Translocation from the
membrane/caveolae to the cytosol occurs as a result of the competitive
inhibition of binding to the caveolae, with subsequent activation of
the enzyme (16). Serine phosphorylation of eNOS by protein
kinase B, more recently known as Akt, has also recently been shown to
cause activation even at resting Ca2+ levels and
the phosphorylated form becomes insensitive to further changes in
Ca2+ (17).
In our UAEC model, AII does not stimulate Ca2+ mobilization, whereas ATP clearly stimulates Ca2+ mobilization in both NP-UAEC and P-UAEC (8). In P-UAEC alone, however, both AII and ATP also stimulate ERK-1/2 activation and both agonists stimulate a corresponding marked increase in NO production. In contrast, in NP-UAEC neither AII nor ATP evoke detectable ERK-1/2 stimulation and also fail to stimulate a significant increase in NO production, even in the face of Ca2+ elevation (8). We propose, therefore, that kinase regulation of eNOS activity occurs in our cells, and further suggest such phosphorylation is a more effective means of eNOS activation than Ca2+/CaM binding alone in UAEC. To this end, we chose to extend our former observations by first examining the effects of ATP on Ca2+, phosphoinositol production, ERK-1/2 and Akt activation in NP-UAEC and P-UAEC, and then subsequently examining the effects of BAPTA [a Ca2+ chelator and known blocker of Ca2+ mobilization in endothelial cells (16)], or U0126 [a known inhibitor of MAP kinase kinase (MEK1), the upstream kinase responsible for ERK-1/2 activation (18)] on both NO and PGI2 production. Our data suggest that eNOS activation may absolutely require ERK-1/2 signaling and that the physiologic consequence of such an event is an even greater increase in eNOS sensitivity to cytosolic Ca2+ than observed for cPLA2. Thus, the increased coupling of agonist receptors in UAEC to the ERK-1/2 pathway during pregnancy underlies the maternal adaptive response, serving to increase endothelial NO-mediated as well as PGI2-mediated vasodilation of the uterine artery and so support the growing fetus.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResults and DiscussionReferences |
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Isolation of uterine artery endothelial cells
Uterine arteries were obtained from Polypay and mixed Western
breed nonpregnant sheep (n = 4) and pregnant ewes at 120–130 days
of gestation (n = 6) during nonsurvival surgery, as described
previously (8). Procedures for animal handling and
protocols for experimental procedures were approved by the University
of Wisconsin-Madison Research Animal Care Committees of both the
Medical School and the College of Agriculture and Life Sciences and
follow the recommended AVMA guidelines for humane treatment and
euthanasia of laboratory farm animals. Briefly, primary uterine
arteries were flushed free of blood using M199 medium, before tying off
arterial branches, clamping off the larger diameter end, and inflating
with M199 containing 5 mg/ml collagenase B (Roche Molecular Biochemicals, through Roche, Indianapolis IN) and
0.5% BSA (Fraction V, Sigma) through a luerlock three way
TAP Pharmaceuticals, Inc. Digestion was allowed to proceed
at 37 C for 55 min before flushing the collagenase solution and
endothelial cell sheets from the inner surface of the vessel. Freshly
isolated cells (Passage 0) were plated to 35 mm dishes in MEM
containing 20% FBS/1% penicillin streptomycin/1% gentamycin (growth
medium; used throughout). Cells were then grown and passaged as
previously described over 12–16 days to approximately 70% confluence
in T75 flasks at which point they were passaged once more (Passage 3)
to medium containing 10% dimethylsulfoxide and frozen in liquid
nitrogen for long term storage. Cells were later recovered and grown in
T75 flasks to approximately 70% confluence and subcultured for
experimental use (Passages 4–5).
Agonist-stimulated production of NO and PGI2
Cells plated overnight in 12-well dishes were washed twice with
modified Kreb’s buffer (125 mM NaCl, 5 mM KCl,
1 mM MgSO4, 1 mM
KH2PO4, 6 mM
glucose, 25 mM HEPES, 2 mM
CaCl2, pH 7.4) before incubation for 1 h in
450 µl modified Kreb’s buffer per well. Agonists were then added as
a 50-µl volume in wells and incubation continued for a further 40 min
(or as otherwise stated). Medium was collected for assay, wells drained
thoroughly and cells solubilized in lysis buffer for protein assay. NO
production/release was measured immediately by conversion of total
nitrate and nitrite back to NO and electrochemical detection using a
Seivers model 280 NO analyzer (100 µl medium injected). Results were
calculated against a standard curve (using known quantities of
NaNO3). Samples were then stored frozen for
further assay. To determine PGI2 production, the
spontaneous breakdown product 6 keto-PGF1 was
assayed in stored medium by EIA using a commercially available EIA kit
(Cayman Chemical Co., Ann Arbor, MI), using 10 µl of medium per well,
and including quality control standards of Kreb’s buffer alone or
standards made up in Kreb’s buffer to monitor for assay drift.
Standard curves typically show correlations of
r2=0.98.
Phospholipase C activation assay
Cells maintained to passage 4 were subcultured onto 24-well
plates and culture continued in medium supplemented with 20 µCi/ml
[3H]inositol (Amersham Pharmacia Biotech, Arlington Heights, IL) for 48 h. Cells were washed
in M199 and then incubated in M199/20 mM LiCl/10
mM inositol for 15 min before cell treatment as shown. Cell
stimulation was allowed to proceed 30 min, at which time cells were
rapidly lysed with perchloric acid (final concentration 5%) exactly as
described previously (19). Cell lysates were separated by
centrifugation into aqueous products and cell membranes, and the
aqueous phase neutralized by mixing with 110% by vol of 1:1
Freon/octylamine. Water soluble [3H]inositol
phosphates and [3H]inositol separated by Dowex
chromatography as previously described (20).
[3H]phosphoinositides were extracted from the
cell membrane pellet by an acidified Bligh/Dyer procedure as previously
described (19). Results shown for dose dependency are
mean ± SE of data from quadruplicate incubations, and
four separate experiments each, with data normalized to total cell
labeling and expressed as the fold increase in inositol phosphates
labeling over unstimulated controls.
Fura-2 Ca2+ imaging studies
For cellular imaging of Ca2+ using Fura 2,
cells at passage 3 from four separate animals each were thawed and
grown to passage 4, combined, and then split 1:8 before freezing with
10% dimethylsulfoxide (Sigma) for later plating at lower
density as required. Note in test studies the data from these cells was
indistinguishable from our previously published data on pre-passage 4
cells (8). UAEC plated to low density (10–20%
confluence) on 35 mm dishes with glass coverslip windows
(Intracellular Imaging, Inc., Cincinnati, Ohio) the night
before use to allow attachment. The next day, immediately before use,
cells were loaded with 5 µM Fura 2-AM (Molecular Probes, Inc.) in Kreb’s buffer (with 2 mM
CaCl2, see above) for 45 min and cells rinsed
three times and covered with 2 ml prewarmed (37 C) Kreb’s buffer and
incubated a further 30 min to complete ester hydrolysis. Cells were
washed once more and covered with 1 ml Kreb’s buffer, and Fura 2
loading was verified by viewing at 380 nm UV excitation on a
Nikon inverted microscope (InCyt Im2, Intracellular Imaging, Inc.). A single isolated cell was then set in the field
of view and recordings commenced, using alternate excitation at 340 nm
and 380 nm at 50 msec intervals, and measuring emitted light
using a photomultiplier. From the ratio of emission at 510 nm detected
at the two excitation wavelengths, and by comparison to a standard
curve established for the same settings using buffers of known free
[Ca2+], the
[Ca2+]i was then
calculated in real time using the InCyt Im2 software on line. All
agonist additions were made in an equal volume (1 ml) of buffer to
ensure rapid mixing, and recovery of cells in fresh buffer was
routinely performed for 20 min before restimulation to avoid
desensitization.
Western blot analysis for Phospho-ERK and Phospho-Akt
UAEC were passaged (passage 4) to 60-mm dishes and maintained
for 24 h before serum withdrawal by incubation in 2.97 ml MEM (1%
penicillin-streptomycin and 1% gentamycin) with 0.01% BSA for 4
h. Cells were stimulated with agonists (30 µl vol) for the
appropriate time. In further experiments cells were preincubated with
10 µM U0126 (a dose that fully inhibited ERK-1/2
activation by basic fibroblast growth factor (bFGF); Sullivan, J.,
unpublished data) or 10 µM LY294002 (a dose that fully
inhibits EGF stimulated Akt phosphorylation; Sullivan, J., unpublished
data) for the final 20 min of the 4 h serum withdrawal before
being stimulated with 30 µM ATP for 10 min. Reactions
were terminated by addition of ice-cold PBS. Cells were immediately
washed twice in ice cold PBS, and solubilized in lysis buffer (4
mM
NaP2O7.H2O,
50 mM HEPES, 100 mM NaCl, 10 mM
EDTA, 10 mM NaF, 2 mM
Na3VO4, 1 mM
PMSF, 1% Triton X-100, 5 µg/ml leupeptin, and 5 µg/ml aprotinin)
with sonication. Solubilized protein was quantified in cell lysates by
a modified bicinchoninic acid assay procedure (Sigma).
Proteins (10 µg/lane) were then separated on 7.5% polyacrylamide
gels (100V, 2 h, Mini-Protein II, Bio-Rad Laboratories, Inc.) before transfer to Immobilon P membrane (100 V, 2
h).
For ERK-1/2 phosphorylation assays, Western analysis with the anti-active MAPK pAb (Promega Corp., Madison, WI) and HRP conjugated second antibody (HRP-linked antirabbit IgG; New England Biolabs, Inc., Beverly, MA) were used at a 1:5000 and 1:2000 dilution, respectively, to detect phosphorylated ERK-1/2. This antiactive MAPK antibody recognizes only the dually phosphorylated MAPK protein (pTpY) indicative of activity (21). To then normalize for loading, the membrane was reprobed by using anti-p42/p44 MAPK antibody diluted 1:1000 (New England Biolabs, Inc.), followed by second antibody (HRP-linked antirabbit IgG, 1:2000 dilution; New England Biolabs, Inc.). In each case, specific binding was detected by enhanced chemiluminescence reagent detection system, as described by Amersham Pharmacia Biotech, and exposed to Hyperfilm. The levels of protein expression were quantified by transmission scanning densitometry (Bio-Rad Laboratories, Inc., 670 scanning densitometer).
For Akt phosphorylation assays, membranes were blotted with anti-phospho-Akt antibody (New England Biolabs, Inc.) at 1:750 dilution and secondary HRP-linked antirabbit IgG antibody (New England Biolabs, Inc.) at 1:7500 dilution. Total Akt expression was determined with anti-Akt antibody at 1:1000 dilution and HRP-linked antirabbit IgG antibody (New England Biolabs, Inc.) at 1:2000 dilution.
Statistical analysis
Data were analyzed by one-way ANOVA or Student’s t
test, as appropriate. Data presented are the means ±
SE. Results were considered significant at the
P < 0.05 level.
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Results and Discussion |
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TopAbstractIntroductionMaterials and MethodsResults and DiscussionReferences |
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). Both ATP and bFGF
gave rise to similar magnitude responses in P-UAEC. In contrast, no NO
production was detected in response to AII, and both bFGF and ATP gave
rise to only marginal NO production, which was not significant at any
time in NP-UAEC (Fig. 1). NP-UAEC also failed to produce a significant
increase in PGI2 in response to AII or bFGF, but
retained the ability to increase production of
PGI2 in response to ATP, albeit in a diminished
fashion (Fig. 1). ATP-induced NO and PGI2
production was dose-dependent in P-UAEC (Fig. 2), with a maximal effect on both NO and
PGI2 production at 30–100 µM. In NP-UAEC,
however, there was no significant increase in NO production, whereas
PGI2 production was dose-dependently increased
but to a much lesser extent than in P-UAEC (Fig. 2).
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). The dose-response to ATP was
dissimilar to that for vasodilator production in Fig. 2
, and only rose
more dramatically at doses of 100–300 µM ATP. The
noncompetitive nature of Li+ inhibition of
inositol phosphate degradation, however, may have skewed the dose
response by potentiating the accumulation of more product as the weak
response increases (because the degree of inhibition of inositol
phosphate phosphatase is proportional to the product of
[Li+] multiplied by [inositol phosphate]).
Nevertheless, these data are intriguing because even at the maximal
dose tested there was no major discrepancy between the response
observed for NP-UAEC compared with that for P-UAEC for either AII or
ATP. Thus differences in PLC activation alone do not account for the
major differences in ATP stimulated NO production whereas they may
relate to the lesser difference observed in PGI2
production in NP-UAEC vs. P-UAEC.
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. The resting
[Ca2+]i of P and NP-UAEC
was typically below 100 nM. ATP was able to elicit a
response in more than 80% of P-UAEC and in approximately 60% of
NP-UAEC, and this response included both an acute rise within seconds
and a sustained phase lasting several minutes. The lag time of
[Ca2+]i increase was
shortened with increasing ATP concentrations (not shown), and both the
initial peak and the sustained rise showed dose-dependent increases
(Fig. 4B). The acute and particularly the sustained responses in P-UAEC
were consistently greater than the responses for NP-UAEC when assessed
as peak level (mean maximal
[Ca2+]i was 280
nM in P-UAEC and 230 nM in NP-UAEC) or total
area under the curve for 2.5 min stimulation (Fig. 4B). Although there
were no significant differences between NP-UAEC and P-UAEC responses
when assessed as -log EC50, significant
differences were observed in peak or area responses for threshold or
maximal doses of ATP (Fig. 4B). Thus, changes in
Ca2+ signaling were of a magnitude that appeared
to correlate with changes in production of PGI2
and possibly PLC activation, but not NO production in NP-UAEC
vs. P-UAEC.
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). We also found these
[Ca2+]i oscillations
occurred inside P-UAEC in the presence or absence of extracellular
Ca2+ (not shown). Our studies are insufficient to
establish the exact mechanism underlying the Ca2+
oscillation, although we clearly can conclude that extracellular influx
does not play a critical part in this response. We were, however, able
to completely block elevation of
[Ca2+]i in NP-UAEC and
P-UAEC by preloading the cells with BAPTA (Fig. 5A).
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), suggesting a rapid
and reversible mechanism of action consistent with
P2Y receptor inhibition at the plasma membrane
level.
ATP is known to act through P2Y receptors
in a number of cell types including astrocytes (25, 26)
and vascular smooth muscle cells (27, 28) to stimulate
ERK-1/2 activation as well as Ca2+ mobilization.
We have previously demonstrated that ATP can induce ERK-1/2
phosphorylation in P-UAEC in a manner that correlates with vasodilator
production (8), consistent with the observations of Chen
et al. (12), whereas others (17, 29) have recently shown Akt phosphorylation of eNOS can activate
the enzyme. The time-course of ERK-1/2 activation was examined by
stimulating P-UAEC and NP-UEAC with ATP (30 µM)
for time points ranging from 2–40 min (Fig. 6). The phosphorylation of ERK-1/2 peaked
around 5 min, i.e. after the peak Ca2+
response, but before the peak in NO and PGI2
production. The elevated level of ERK-1/2 phosphorylation then returned
to basal values by 20 min in P-UAEC. Consistent with our previous
observations (8) a weak but otherwise significant ERK-1/2
phosphorylation was detected within 5 min in NP-UAEC In contrast, Akt
phosphorylation was not observed in response to ATP in P-UAEC or
NP-UAEC at any time of stimulation (inset, Fig. 6).
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). This is consistent with the dose of
ATP that induced maximal
[Ca2+]i mobilization and
NO and PGI2 production.
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). This indicates that MEK mediates
ERK-1/2 activation in response to ATP and that this is not dependent on
or mediated indirectly by PI3-kinase. This effect of U0126 also
provides us with a useful means to study cause and effect between MEK
activation/ERK phosphorylation and vasodilator production (below). In
addition the stimulatory action of ATP (10 µM) on ERK-1/2
phosphorylation was blocked by preincubation with suramin at 30 and 100
µM (Fig. 8B) to a similar extent as the
Ca2+ responses were blocked by suramin in P-UAEC
and NP-UAEC (Fig. 5). This suggests a similar receptor/G protein
mechanism for ATP-stimulated Ca2+ mobilization
vs. ERK-1/2 phosphorylation.
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) and reduced (50%) but did
not abolish the substantial increase in ERK-1/2 phosphorylation
response to 30 µM ATP. When P-UAEC and NP-UAEC
were stimulated in the continued presence of U0126, ATP-stimulated (30
µM) increases in both NO production and
PGI2 production were abolished in P-UAEC and
ATP-stimulated PGI2 production was abolished in
NP-UAEC (Fig. 9). Similarly, when cells
were preincubated with 10 µM BAPTA, ATP (30
µM) stimulated increases in both NO production
and PGI2 production were abolished in P-UAEC and
ATP-stimulated PGI2 production was abolished in
NP-UAEC (Fig. 9). This finding suggests that in both NP-UAEC and
P-UAEC, both eNOS and cPLA2 activation are
dependent on at least resting
[Ca2+]i and at least some
basal level of MEK-ERK-1/2 activation. However, absolute dependence of
the responses on these pathways still does not indicate physiologic
sensitivity to or endocrine dose-dependent regulation by increases in
[Ca2+]i or the MEK and
ERK-1/2 signaling pathway. This question is answered more clearly,
however, by direct comparison of the dose-dependency for NO,
PGI2, Ca2+, and ERK-2
phosphorylation responses to ATP in NP-UAEC and P-UAEC (Fig. 10). Such analysis reveals that in
P-UAEC there is a direct correlation between NO production and
Ca2+ elevation (P < 0.0005) or
ERK-2 phosphorylation (P < 0.0001), and similarly
between PGI2 production and
Ca2+ elevation (P < 0.005) or
ERK-2 phosphorylation (P < 0.0001). In NP-UAEC,
however, there is no correlation between NO production and the
otherwise still detectable Ca2+ elevation,
whereas there is a correlation, albeit much reduced, between
PGI2 production and Ca2+
elevation (P < 0.005). Thus, in physiologic terms,
whereas eNOS and cPLA2 activation may be both
Ca2+ and MEK-ERK-1/2 dependent,
cPLA2 activation is more physiologically
sensitive to Ca2+ than eNOS in the absence of
dramatic ERK-1/2 activation, as observed in NP-UAEC. Both enzymes,
however, show much greater Ca2+ sensitivity in
response to dramatically increased MEK-ERK stimulation as seen in
P-UAEC. Because Akt was not stimulated by ATP in these cells, and
ATP-stimulated ERK-1/2 phosphorylation is insensitive to the PI3-kinase
inhibitor LY294002 in these cells (not shown), it also appears
activation of eNOS occurred independently of the previously described
PI3-kinase/Akt pathway in UAEC.
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Although our previous data (8) as well as our findings herein provide a molecular model for changes in UA endothelial function in pregnancy, just how this pregnancyinduced change in UAEC signaling through ERK-1/2 occurs or is regulated in vivo remains unclear. ATP as well as other agonists previously tested (8) have similar Ca2+ and ERK-mediated signaling and subsequent vasodilator production responses in P-UAEC as previously reported in bovine aorta endothelial cells (BAEC) (9), and human hand vein endothelial cells from pregnant women (30). The reduced level of Ca2+ signaling in NP-UAEC is also seen in hand vein endothelial cells from nonpregnant women (30) and, more importantly, the pregnancy-specific enhancement in Ca2+ signaling is not seen in preeclamptic pregnancy (30). Thus, while a pregnancy-specific response, the differences in Ca2+ signaling is common to both uterine/reproductive and systemic/nonreproductive vasculature. In contrast, it is the NP-UAEC alone that appear unusual in their general lack of efficient coupling of agonists to ERK-1/2 signaling at a level distal to receptor occupancy and proximal to MEK activation. This specificity of changes in ERK-1/2 signaling but not Ca2+ signaling to the uterine artery endothelium suggests that different endocrine mechanisms may control these two endothelial responses through perhaps a specific paracrine vs. general endocrine mechanism, respectively. This is also consistent with the finding that the pregnancy-induced differences in uterine artery endothelial function are as much a reflection of UA endothelial under-responsiveness in the nonpregnant state as due to enhanced responsiveness in the pregnant state.
Our finding that the sensitivity of eNOS to Ca2+ is associated with changes in ERK-1/2 signaling is also of general importance to our understanding of mechanisms of Ca2+-dependent and independent eNOS activation in many other endothelial cell types. It is now generally accepted that ERK-mediated direct phosphorylation of cPLA2 brings about a dramatic increase in Ca2+ sensitivity such that almost full activity can be achieved at near resting Ca2+ levels (9). Our data likewise suggest that in the absence of significant ERK-1/2 activity, eNOS is relatively Ca2+ insensitive and that ERK-1/2 activation leads to a dramatic increase in eNOS Ca2+ sensitivity. Our findings are thus consistent with and extend those of Chen et al. (12). In contrast, a recent study in BAEC has demonstrated bradykinin induced eNOS phosphorylation in whole cells in a manner sensitive to PD98059, an inhibitor of MEK (31), but the authors also reported that such phosphorylation resulted in inhibition of eNOS activity. However, eNOS activity was determined in an immunoprecipitate, and the assay performed in a buffer containing 1 mM CaCl2. Such a high level of Ca2+ is substantially above the normal cytosolic range of 100-1000 nM, so the changes in Ca2+ sensitivity indicated by our observations would not be apparent even if they occurred. In turn we are unable to comment from our data alone whether ERK-1/2 associated increases in eNOS activity and Ca2+ sensitivity are due to direct eNOS phosphorylation by ERK, and further studies will clearly be necessary to establish this point. The UAEC cell culture model, however, clearly gives a unique system ideally suited to pursue this fundamental question of eNOS regulation.
Thus far our attention has focused on the role of Ca2+ and ERK-1/2 in controlling vasodilator production, and how this changes in UA endothelium during pregnancy. The observation that P2Y receptors couple to Ca2+ and ERK but not Akt is not, however, unique to UA endothelium, or indeed endothelium in general, and has also been reported in astrocytes (25, 26) and coronary artery smooth muscle cells (CASMC) (27, 28). It is of particular relevance that in CASMC ATP stimulation also did not result in Akt activation, but when Akt was costimulated with insulin, a hormone elevated during pregnancy, a synergistic effect was observed on mitogenesis (28). Preliminary studies suggest that P-UAEC and NP-UAEC also respond to insulin at the level of Akt phosphorylation (Sullivan, J., unpublished data). We are currently examining the possible synergy of ATP with insulin to control UA endothelial proliferation at this time.
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Footnotes |
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Received January 25, 2001.
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References |
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TopAbstractIntroductionMaterials and MethodsResults and DiscussionReferences |
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mediates the
nongenomic activation of endothelial nitric oxide synthase by estrogen.
J Clin Invest 103:401–406[Medline]
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D. Xiao and L. Zhang Adaptation of uterine artery thick- and thin-filament regulatory pathways to pregnancy Am J Physiol Heart Circ Physiol, January 1, 2005; 288(1): H142 - H148. [Abstract] [Full Text] [PDF]
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S. M. Gifford, J. M. Cale, S. Tsoi, R. R. Magness, and I. M. Bird Pregnancy-Specific Changes in Uterine Artery Endothelial Cell Signaling in Vivo Are Both Programmed and Retained in Primary Culture Endocrinology, August 1, 2003; 144(8): 3639 - 3650. [Abstract] [Full Text] [PDF]
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I. M. Bird, L. Zhang, and R. R. Magness Possible mechanisms underlying pregnancy-induced changes in uterine artery endothelial function Am J Physiol Regulatory Integrative Comp Physiol, February 1, 2003; 284(2): R245 - R258. [Abstract] [Full Text] [PDF]
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T. Kimura, K. Ayajiki, Y. Noda, and T. Okamura Comparison of the Responses to Thrombin in Monkey Renal and Uterine Arteries Reproductive Sciences, May 1, 2002; 9(3): 146 - 151. [Abstract] [PDF]
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