| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
ARTICLES |
Department of Pharmacology of the University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania 15261
Address all correspondence and requests for reprints to: Guillermo Romero, Department of Pharmacology, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania 15261. E-mail: ggr+{at}pitt.edu
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
The recent cloning and study of specific isoforms of PLD (11, 12, 13) have led to the suggestion of three main mechanisms for the regulation of PLD activity by cell surface receptors. These mechanisms are mediated by: 1) G proteins of the ADP-ribosylation factor (ARF) family; 2) G proteins of the Rac/Rho/Cdc42 family; and 3) protein kinase C (PKC) (11, 12, 13, 14, 15, 16). All of these are likely to act simultaneously in some cell systems to produce complete activation of the PLD pathway. Furthermore, the evidence suggests that these three mechanisms are not independent and that substantial cross-talk among these pathways exists (17).
Recent work has suggested that PLD plays a very important role in the proliferation of VSMC. Wilkie et al. (6) suggested in 1996 that PLD activity and mitogen activated protein kinase (MAPK) activation are both required for agonist-induced proliferation of VSMC. Furthermore, comparative analysis suggests that the VSMC hyperplasia observed in a spontaneous hypertensive rat model is associated with a significant increase in the activity of PLD in response to external agents, when compared with the levels observed in the normotensive rat (WKY) (18). These observations are significant due to the important contribution of VSMC proliferation in the development of atherosclerotic plaques and the thickening of the blood vessel wall in hypertension. Therefore, the analysis of the mechanisms by which external stimuli regulate the activity of PLD in VSMC is of significant interest. However, these mechanisms remain largely undefined. In this paper, we show that the regulation of PLD activity by angiotensin II (AngII), endothelin-1 (ET-1), platelet-derived growth factor (PDGF), and phorbol 12-myristate 13-acetate (PMA) in A10 cells, a cell model widely used to study the basic biology of VSMC, is mediated, at least in part, by proteins of the ARF family. We also show that Rho plays a minor role in the regulation of PLD activity in these cells. A general model for the activation of PLD and its functional role in VSMC biology is proposed.
|
Materials and Methods |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
ARF proteins
The following ARF proteins were used: wild-type
(wt)-ARF1, wt-ARF6, Q71L-ARF1, Q67L-ARF6, T31N-ARF1, and
T27N-ARF6. The latter are putative dominant negative mutants with
reduced affinity for GTP. These mutants were generated by
site-directed mutagenesis, cloned in the multiple cloning site of the
vector pGEFP-N1 (CLONTECH Laboratories, Inc., Palo
Alto, CA), and fused to the green fluorescent protein (GFP) of A.
victoria, as described (19). Cells were transfected with these
constructs as described elsewhere (20). The efficiency of transfection
was estimated by counting cells using DIC and fluorescence microscopy
of the same fields. Based exclusively on the ratio of green fluorescent
cells to cells detected by DIC, the transfection efficiency was better
that 50%. When the efficiency of transfection was measured in terms of
the number of viable cells (as determined by propidium iodide
exclusion), the transfection efficiency was very near 90%. The PLD
activity of ARF-GFP-transfected cells was determined approximately
48 h after transfection.
Detection of PLD isoforms
PLD1a, PLD1b, and PLD2 expression was quantitated using a
semiquantitative PCR procedure. Two 100-mm plates were used to isolate
total RNA using commercial kits (QIAGEN, Valencia,
CA). Two micrograms of total RNA were used for 1st-strand complementary
DNA (cDNA) synthesis (using a CLONTECH Laboratories, Inc.
kit). Once the cDNA was obtained, the following primers were used to
amplify PLD 1a (expected 527 bp) and 1b (expected 413 bp):
5'-CGTGAACCACAGAACCAATG-3' and 5'-TCTCACGGCAGCATCAGTAG-3'. PLD 2
was amplified (expected 467 bp) with
5'-CAGGAGGCGGTTGAGGTAAT-3' and 5'-AGTTGCACATGGAGCCAGAT-3'.
Additionally, the glyceraldehyde-3-phosphate dehydrogenase primers,
5'-TACTCCTTGGAGGCCATGTA-3' and 5'-CGTGGAGTCTACTGGCGTCT-3', were used
for quantitation purposes. The amplification cycles were designed so
that all the reactions would occur simultaneously (annealing at 60 C
for 45 sec and elongation at 72 C for 60 sec). This cycle was repeated
for a total of 30 times. Fifteen-microliter aliquots of the resulting
PCR products were loaded in each lane of a 3% agarose gel.
PLD mutants
Wild-type and catalytically inactive variants of PLD1 and PLD2
(K898R-PLD1 and K758R-PLD2) were made as previously described (21), and
fused to A. victoria green fluorescent protein by subcloning
into pEFGP-C1 (CLONTECH Laboratories, Inc.). The enzymatic
activity of the wild-type enzyme-GFP chimeras expressed in
Baculovirus was determined in vitro to confirm
that the GFP tag did not generate an inactive phenotype. Cells were
transfected with Lipofectamine, as described (21). Transfection
efficiencies were determined as described above.
Immunoblotting
ARF was detected with 2 µg/ml affinity-purified 1D9 (a
monoclonal antibody that recognizes most members of the ARF family
(22); a gift from R. Kahn). Bound antibodies were detected by
chemiluminescence.
PLD assays
Cells (60-mm plates; 70–80% confluence) were serum starved and
labeled overnight with 3H-palmitate (5 µCi/ml)
in DMEM/F12 containing 0.1% BSA. Cells were stimulated with PDGF (30
ng/ml), ET-1 (100 nM), AngII (1 µM), or PMA
(500 nM) in the presence of 0.3% ethanol for 20 min. The
reaction was stopped by addition of chloroform:methanol (1:1). The
lipid phase was extracted and developed by TLC on silica gel 60 plates
using ethyl acetate:trimethylpentane:acetic acid (9:5:2) as the
solvent. The position of major phospholipids was determined using true
standards (Avanti Biochemicals) and autoradiography. The TLC
plates were scraped, and the total amount of radioactivity associated
with each lipid species was determined by liquid scintillation
counting. The data were expressed as the number of counts associated
with the phosphatidylethanol spot normalized by the total number of
counts of lipid.
Digitonin treatment
A10 cells were serum starved, scraped gently, resuspended in
PBS, and treated with 8 µM digitonin in the presence or
absence of PDGF (30 ng/ml), ET-1 (100 nM), AngII (1
µM), or PMA (500 nM), with or without the
addition of GTP
S (100 µM) at 37 C for 15–20 min. ATP
(1 mM) and magnesium (2 mM) were included in
all the experiments. To release intracellular proteins,
digitonin-treated cells were centrifuged in a microfuge for 20 min.
Supernatants and pellets were collected separately and resolved by
SDS-PAGE. ARF proteins were detected by immunoblotting, as mentioned
above. PLD activity of digitonin-treated cells was determined using
cells that had been labeled overnight with
3H-palmitate (see above). Recombinant ARF was
isolated from bacteria that coexpress ARF1 and yeast
N-myristoyltransferase, as described (23, 24). Recombinant
ARF protein (final concentration, 8 µg/ml) was added to
cytosol-depleted cells where indicated. Human recombinant PKC
(Calbiochem; 5 µg/ml, final concentration) was included
in all assays of the effects of PMA. The choice of PKC was
arbitrary, and was included with the purpose of maximizing the effects
of PMA, because endogenous PKC is likely to leak out of the cell during
the permeabilization and centrifugation procedures.
Treatment with ARF and Rho inhibitors
Cells were treated with brefeldin A (BFA) (25 µg/ml, final) 10
min before stimulation with the agonists in serum-free medium.
C3 exotoxin treatment was done by the scrape loading method of
Malcolm et al. (25). Briefly, cells were grown in 10-cm
plates and scraped gently in 500 µl buffer [10
mM Tris (pH 7.2), 114 mM
KCl, 25 mM NaCl, 5.5 mM
MgCl2] with or without 5 µg/ml C3 transferase.
The cells were then distributed onto 6-cm plates and allowed to recover
overnight. Cells were then serum-starved and labeled with 5 µCi/ml
3H-palmitate for 17 h, as described
previously (23). The effectiveness of the treatment with C3 was
determined by in vitro ADP-ribosylation of cell extracts
after treatment with the toxin, as described by Shome et al.
(24). According to these controls, ADP-ribosylation of endogenous Rho
proteins was at least 95% complete under the conditions of the
scrape-loading assay described.
MAP kinase activity
A10 cells were treated as described above or as described in the
figure legends. Cells were washed with cold PBS and scraped into
microcentrifuge tubes in Buffer A [10 mM HEPES (pH 7.4), 2
mM EDTA, 1 mM
Na3VO4, and 1
mM phenylmethylsulfonylflouride] and then pelleted via
centrifugation. Cells were resuspended in a 0.5 ml detergent lysis
buffer [50 mM HEPES (pH 7.4), 0.1 M NaCl,
1.5% sodium cholate, 1 mM EDTA, 1 mM EGTA, 5
µg/ml leupeptin, 1 mM phenylmethylsulfonylflouride, and 1
mg/ml soybean trypsin inhibitor] and lysed for 30 min at 4 C. Cell
lysates were analyzed by SDS-PAGE, followed by immunoblotting with a
phosphospecific anti-MAPK antibody (New England Biolabs, Inc.). Immunocomplexes were detected by ECL.
Confocal microscopy
A10 cells were transfected with ARF-GFP chimeras using
Lipofectamine. The cells were examined by confocal microscopy 24 h
after transfection using a 2002 instrument (Molecular Dynamics, Inc.), equipped with an argon laser. Data were obtained
using the 488-nm band of the laser and a 530 ± 35-nm bandpass
filter. This filter was used to reduce errors caused by the background
autofluorescence associated with the use of Lipofectamine.
|
Results |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
summarize these studies. As shown,
AngII, ET-1, PDGF, and PMA considerably stimulated PLD activity (2- to
5-fold). The addition of 25 µg/ml BFA, 10 min before stimulation of
the cells with these agents, consistently inhibited the activation of
PLD. AngII, ET-1, and PDGF-dependent PLD activation was completely
blocked by BFA treatment, whereas the activation of PLD by PMA was
reduced by approximately 50% (Fig. 1).
|
). These experiments
suggest that the activation of PLD by AngII, ET-1, and PDGF is mediated
by ARF proteins and that Rho proteins do not play an essential role in
receptor- mediated activation of PLD in A10 cells.
|
Figure 3 shows the results of a
representative experiment using AngII and PDGF as agonists. As shown,
ARF leaked out of permeabilized cells during treatment (digitonin
lane), and the inclusion of GTPS, AngII, or PDGF alone during the
permeabilization procedure did not induce membrane association of ARF
proteins. Incubation with agonist and GTPS during
permeabilization resulted in a significant increase translocation
of ARF to cell membranes (Fig. 3, 5th and 7th lanes). Identical results
were obtained with ET-1 and PMA (not shown). These data show
unequivocally that the activation of ARF proteins can be regulated by
the agonists AngII, ET-1, and PDGF and by the activation of PKC by
PMA.
|
|
S, following the protocol described
above; and the activation of PLD was measured by the production of
phosphatidylethanol. Cells that had been permeabilized in the
presence of agonist and GTPS showed high levels of PLD activity
that were comparable to those observed with intact cells after
agonist stimulation (Figs. 1
, 4A, and 4C). In contrast, the PLD
activity of cells permeabilized in the presence of agonist alone or
GTPS alone was indistinguishable from that of control cells.
|
S. These cells contained very little
ARF, because of the depletion of their intracellular contents during
the permeabilization procedure (Fig 3). Figure 4, B and D, shows that the addition of
GTPS, agonist, or a solution containing both GTPS and the
corresponding agonist failed to restore PLD activity. However, when
recombinant, myristoylated human ARF1 was included in this assay, the
activity of PLD was fully recovered to levels comparable with those
obtained by the stimulation of intact cells with the corresponding
agonist.
Inhibition of receptor-mediated PLD activation by dominant negative
mutants of ARF
A10 cells were transfected with ARF proteins that had been
subcloned in pEGFP-C1 as fusion proteins containing an Aequorea
victoria GFP tag in their C-terminus. The following chimeras were
tested: wild-type ARF1-GFP, wt-ARF6-GFP, and the dominant negative
mutants T31N-ARF1-GFP and T27N-ARF6-GFP. The GFP fusion constructs were
selected to facilitate the estimation of the transfection efficiency,
using phase contrast and fluorescent microscopy.
The data obtained are shown in Fig. 5.
Additional controls were run using the empty vector, and the PLD
activity of these cells was indistinguishable from that of
untransfected cells. The overexpression of wt-ARF1-GFP and wt-ARF6-GFP
produced a small (but significant) increase in agonist-dependent PLD
activity. More important, the expression of the dominant negative
mutants T31N-ARF1-GFP and T27N- ARF6-GFP abolished the
activation of PLD by AngII, ET-1, and PDGF. In contrast, the dominant
negative ARF1 mutant had no significant effects on PMA-dependent PLD
activity, whereas the dominant negative ARF6 mutants reduced the
effects of PMA by about 40%. These results show that AngII, ET-1, and
PDGF activate PLD by a mechanism mediated primarily by the activation
of ARF proteins. In contrast, PMA activates PLD by a mechanism that is
only partially dependent on the activation of ARF.
Both ARF dominant negative mutants inhibited agonist-mediated PLD
activation in A10 cells. However, the intracellular distribution of
ARF6 has been reported to be very different from that of other members
of the ARF family (28, 29). Because PLD1 and PLD2 have been found
confined to cell membranes (11, 12), we studied the distribution of
wt-ARF1-GFP, wt-ARF6-GFP, T31N-ARF-GFP, and T27N-ARF6-GFP by confocal
microscopy. Additional controls included the mutants Q71L-ARF1-GFP and
Q67L-ARF6-GFP, two mutants with reduced GTPase activity, that
have been described to act as dominant activated mutants in other
systems (22, 28). The results are shown in Fig. 6. Wild-type-ARF1 and the dominant
activated mutant Q71L-ARF1 were both distributed between the Golgi and
the cytosol. Q71L-ARF1-GFP also localized to some punctate
intracellular structures or on the plasma membrane, although the
fraction of the chimera in these regions was small. In contrast,
T31N-ARF1-GFP was mainly cytosolic. All ARF6 chimeras were found
associated with small intracellular vesicles. However, a significant
fraction of the wild-type and GTPase-deficient chimeras were also found
on the plasma membrane, in regions that seem to be membrane ruffles
(see Fig. 6).
|
. A
short acute preexposure to BFA blocked the activation of MAP kinase by
either PDGF or AngII. However, addition of PA reversed the inhibitory
effects of BFA, suggesting a role for PLD-dependent generation of
phosphatidate in the modulation of the MAP kinase cascade by
extracellular signals in A10 cells.
|
. A10 cells expressed primarily PLD1b
and PLD2, whereas the expression of PLD1a was barely detectable. Also,
as shown, the predominant PLD expressed in A10 cells is PLD2.
|
shows clearly that the
overexpression of the catalytically inactive K758R-PLD2 completely
abolished PDGF- and AngII-dependent PLD activity, whereas the
overexpression of the corresponding PLD1 mutant had no apparent
effects. These data demonstrate that the activation of PLD by PDGF and
AngII in A10 cells is primarily mediated by PLD2. |
Discussion |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Two different PLD genes have been identified in mammals (11, 12). One of these, PLD1, codes for two different gene products generated by alternative splicing (30). PLD1a and PLD1b are activated in vitro by small GTPases of the ARF and Rho families and by direct interaction with PKC (31). Although the sensitivity of PLD2 to regulatory stimuli is lost during purification (12), the PLD activity of crude extracts prepared from cells that overexpress this protein are sensitive to ARF stimulation (32). The linkage between receptor-mediated activation of PLD and the activation of PLD1 and PLD2 has not been examined in detail. In particular, the study of the relative roles of ARF, Rho, and PKC requires a significant amount of attention.
We have recently developed a two-pronged approach to study the role of small GTPases in the regulation of PLD activity in HIRcB fibroblasts (23, 24). This approach is based on a combination of permeabilization/reconstitution assays and on the use of dominant negative mutants of these GTPases. Using this approach, we have recently shown that ARF plays a major role in the modulation of PLD activity by insulin and PDGF in fibroblasts (23, 24), thus proving that the regulation of PLD by receptor tyrosine kinases is primarily mediated by ARF. Some evidence linking ARF to the activation of PLD by G protein-linked receptors has also been reported (33, 34). Here, we extend these studies to mitogenic agents that regulate the proliferation of VSMC.
We also investigated the effects of ARF1 and ARF6 dominant negative
mutants on the activation of PLD. ARF6 was chosen because it is the
only member of the ARF family abundantly found on the plasma membrane
(28, 29). ARF1 was selected because: 1) it is the most abundant of the
ARF proteins in mammalian cells; and 2) it is representative of the
remaining members of the family, in that it exists in a steady-state
distribution between cytosolic and membrane-bound forms. Our data show
that the overexpression of both ARF1 and ARF6 dominant negative mutants
inhibits agonist- dependent PLD activation. This is consistent with
previous data from our laboratory (24). Therefore, the question of
which ARF mediates this signaling pathway remains undecided.
Examination of cells that express ARF-GFP chimeras failed to provide
conclusive results. ARF6 was found on the plasma membrane or on
intracellular vesicles, a distribution that corresponds closely to that
of mammalian PLDs (11, 12, 13). However, although most of the ARF1-GFP
chimeras were found localized to the Golgi and cytosol (Fig. 6), a
small fraction was visible in the plasma membrane and small
intracellular vesicles. Furthermore, agonist treatment failed to
produce a noticeable displacement of either ARF1-GFP or ARF6-GFP to the
plasma membrane. This result seems to be at odds with the observed
complete displacement of endogenous ARF proteins to the membrane in the
permeabilization experiments shown in Fig. 3. Our interpretation of
this apparent discrepancy is that, when ARF1 and ARF6 are
overexpressed, then the concentrations of these proteins are so high
that only a very small fraction of the protein is activated and
displaced to the membrane. Because the cytosolic protein is also
intensely fluorescent, the displacement of a small fraction of the
ARF-GFP chimeras is very difficult to examine using conventional
live-cell confocal microscopy techniques.
Significantly, both ARF dominant negatives were found to be equally effective in the inhibition of PLD activation by ET-1, AngII, and PDGF. The mechanism by which these dominant negative mutants inhibit PLD activity is likely to result from the competition with endogenous ARF proteins for a limited number of ARF activators (guanine nucleotide exchange factors, or GEFs). The fact that both ARF dominant negatives are effective in the blockade of the activation of PLD suggests that these GEFs cannot distinguish between ARF1 and ARF6. Several ARF-GEFs have been described in recent years (35, 36, 37, 38, 39). They are characterized by their homology to the yeast protein Sec7, which is itself an ARF-GEF (35). Many of these ARF-GEFs are not sensitive to BFA in vitro (36, 37). However, our data show clearly that BFA inhibits the activation of PLD in A10 cells. The ARF-GEFs ARNO and GRP-1 have been shown to be recruited to the plasma membrane by a mechanism that seems to involve the activation of PI3 kinase (40, 41). However, the activation of ARF by these GEFs in vitro does not seem to be BFA sensitive (37). There are two possible explanations for these discrepancies: either the GEFs that mediate the activation of ARF by cell surface receptors in A10 cells are BFA sensitive, or the activation of these GEFs by cell surface receptors is BFA sensitive. Our laboratory is presently working on the resolution of this issue. Finally, the dominant negative mutants of ARF1 and ARF6 had very small effects on the activation of PLD by phorbol esters. This suggests that ARF proteins do not mediate the effects of PKC on PLD. Because PKC has been shown to activate directly PLD1 (16, 17), PKC-dependent pathways for the activation of PLD may bypass completely the ARF signaling pathway.
Our data show that PDGF and AngII activate PLD2, rather than PLD1. This
observation is particularly interesting, considering that the activity
of PLD2 in vitro is independent of ARF (12); and thus PLD2
was thought to be an unlikely candidate for receptor-sensitive PLD
activity. Contrary to this expectation, we find that the catalytically
inactive variant of PLD2, but not PLD1, functions as a dominant
negative and blocks PDGF- and AngII-dependent activation of PLD,
suggesting that PLD2 mediates receptor-responsive PLD activity in A10
cells. Interestingly, this further suggests that PLD2 is regulated by
ARF in vivo, as follows from the fact that the PDGF- and
AngII-dependent PLD activity in this model requires ARF activation.
Recent evidence from Sung et al. (32) supports this model.
Although immunopurified PLD2 was found to be unresponsive to
ARF, PLD2 overexpressed in COS-7 cells was activated by ARF preloaded
with GTPS, suggesting that PLD2 may be regulated by ARF in
vivo. Furthermore, a PLD2 mutant lacking the N-terminal 308 amino
acids displays both a reduced in vitro activity and
ARF-sensitive activity. These results are consistent with a model for
ARF-mediated PLD2 activation in response to cell surface receptors.
Finally, it is important to consider the role of PLD activation in the biology of VSMC cells. Several pieces of work have strongly suggested that PLD activation is essential for the proliferative responses of VSMC cells to a variety of stimuli (6, 7, 8, 9, 10). However, the mechanism by which PLD modulates proliferation is not clearly understood. Recent work by Rizzo et al. (21) has shown that the activation of the MAP kinase pathway by cell surface receptors is BFA sensitive and requires the generation of PA, but that PA alone is insufficient to activate the MAPK cascade. This seems to be a direct consequence of the effects of PA on the catalysis of the recruitment of Raf to membrane compartments (21). This function of PA was originally described in fibroblasts. Our data demonstrate a similar phenomenon in A10 cells. Furthermore, in agreement with our previous data, PLD activation alone is insufficient to activate the MAPK cascade in A10 cells (21). This is evidenced by the fact that ET-1 was a very poor activator of MAPK phosphorylation, although the activation of PLD by ET-1 was comparable with that observed upon AngII and PDGF treatment. This suggests that the different effects of ET-1, PDGF, and AngII in the phosphorylation of MAPK probably reflect differences in the mechanisms of activation of Ras or Raf-1 and that this mechanisms are either not present or not fully active in A10 cells.
Taken altogether, our observations suggest a working model for the mechanism of activation and function of PLD in A10 cells. Agonists stimulate the activation of ARF proteins by a mechanism that is likely mediated by ARF-GEFs related to ARNO. Activated, GTP-bound ARF interacts with PLD and promotes its activation. Once activated, PLD hydrolyzes phosphatidylcholine to choline and phosphatidate. The latter, in conjunction with Ras, catalyzes the activation of Raf-1, thus initiating the events that lead to the activation of MAP kinase and cell proliferation. The general aspects of this model are strongly supported by our data, but additional studies are required, to validate its details.
|
Footnotes |
|---|
2 Recipient of an Independent Investigator Award from NIDDK.
Received October 27, 1999.
|
References |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
-diacylglycerol kinase activation is an essential
step in IL-2-mediated lymphocyte proliferation. J Biol Chem 271:10334–10340 and ß receptors in vascular smooth muscle cells. J
Biol Chem 269:30546–30552. J Biol Chem
272:3860–3868
This article has been cited by other articles:
 |
|
 |
|
  K. J. Allahdadi, L. C. Duling, B. R. Walker, and N. L. Kanagy Eucapnic intermittent hypoxia augments endothelin-1 vasoconstriction in rats: role of PKC{delta} Am J Physiol Heart Circ Physiol, February 1, 2008; 294(2): H920 - H927. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 Y. Ozawa and H. Kobori Crucial role of Rho-nuclear factor-{kappa}B axis in angiotensin II-induced renal injury Am J Physiol Renal Physiol, July 1, 2007; 293(1): F100 - F109. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 H. Ohtsu, H. Suzuki, H. Nakashima, S. Dhobale, G. D. Frank, E. D. Motley, and S. Eguchi Angiotensin II Signal Transduction Through Small GTP-Binding Proteins: Mechanism and Significance in Vascular Smooth Muscle Cells Hypertension, October 1, 2006; 48(4): 534 - 540. [Full Text] [PDF]
|
|
|
|
 |
|
 Z. Yang, L. D. Asico, P. Yu, Z. Wang, J. E. Jones, C. S. Escano, X. Wang, M. T. Quinn, D. R. Sibley, G. G. Romero, et al. D5 dopamine receptor regulation of reactive oxygen species production, NADPH oxidase, and blood pressure Am J Physiol Regulatory Integrative Comp Physiol, January 1, 2006; 290(1): R96 - R104. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J.-H. Parmentier, Z. Pavicevic, and K. U. Malik ANG II stimulates phospholipase D through PKC{zeta} activation in VSMC: implications in adhesion, spreading, and hypertrophy Am J Physiol Heart Circ Physiol, January 1, 2006; 290(1): H46 - H54. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
B. T. Andresen, K. Shome, E. K. Jackson, and G. G. Romero AT2 receptors cross talk with AT1 receptors through a nitric oxide- and RhoA-dependent mechanism resulting in decreased phospholipase D activity Am J Physiol Renal Physiol, April 1, 2005; 288(4): F763 - F770. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 F. Li and K. U. Malik Angiotensin II-Induced Akt Activation through the Epidermal Growth Factor Receptor in Vascular Smooth Muscle Cells Is Mediated by Phospholipid Metabolites Derived by Activation of Phospholipase D J. Pharmacol. Exp. Ther., March 1, 2005; 312(3): 1043 - 1054. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Z. Yang, L. D. Asico, P. Yu, Z. Wang, J. E. Jones, R.-k. Bai, D. R. Sibley, R. A. Felder, and P. A. Jose D5 dopamine receptor regulation of phospholipase D Am J Physiol Heart Circ Physiol, January 1, 2005; 288(1): H55 - H61. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
B. T. Andresen, G. G. Romero, and E. K. Jackson AT2 Receptors Attenuate AT1 Receptor-Induced Phospholipase D Activation in Vascular Smooth Muscle Cells J. Pharmacol. Exp. Ther., April 1, 2004; 309(1): 425 - 431. [Abstract] [Full Text]
|
|
|
|
 |
|
 D. N. Robertson, M. S. Johnson, L. O. Moggach, P. J. Holland, E. M. Lutz, and R. Mitchell Selective Interaction of ARF1 with the Carboxy-Terminal Tail Domain of the 5-HT2A Receptor Mol. Pharmacol., November 1, 2003; 64(5): 1239 - 1250. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. A Frias, F. Dubouloz, M. C Rebsamen, and U. Lang Prostacyclin production in rat aortic smooth muscle cells: role of protein kinase C, phospholipase D and cyclooxygenase-2 expression Cardiovasc Res, November 1, 2003; 60(2): 438 - 446. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. Mitchell, D. N. Robertson, P. J. Holland, D. Collins, E. M. Lutz, and M. S. Johnson ADP-ribosylation Factor-dependent Phospholipase D Activation by the M3 Muscarinic Receptor J. Biol. Chem., September 5, 2003; 278(36): 33818 - 33830. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. A. Foster and L. Xu Phospholipase D in Cell Proliferation and Cancer Mol. Cancer Res., September 1, 2003; 1(11): 789 - 800. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
B. T. Andresen, J. J. Linnoila, E. K. Jackson, and G. G. Romero Role of EGFR Transactivation in Angiotensin II Signaling to Extracellular Regulated Kinase in Preglomerular Smooth Muscle Cells Hypertension, March 1, 2003; 41(3): 781 - 786. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L. Xu, P. Frankel, D. Jackson, T. Rotunda, R. L. Boshans, C. D'Souza-Schorey, and D. A. Foster Elevated Phospholipase D Activity in H-Ras- but Not K-Ras-Transformed Cells by the Synergistic Action of RalA and ARF6 Mol. Cell. Biol., January 15, 2003; 23(2): 645 - 654. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Souchet, E. Portales-Casamar, D. Mazurais, S. Schmidt, I. Leger, J.-L. Javre, P. Robert, I. Berrebi-Bertrand, A. Bril, B. Gout, et al. Human p63RhoGEF, a novel RhoA-specific guanine nucleotide exchange factor, is localized in cardiac sarcomere J. Cell Sci., January 2, 2002; 115(3): 629 - 640. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J.-H. Parmentier, M. M. Muthalif, A. T. Nishimoto, and K. U. Malik 20-Hydroxyeicosatetraenoic Acid Mediates Angiotensin II-Induced Phospholipase D Activation in Vascular Smooth Muscle Cells Hypertension, February 1, 2001; 37(2): 623 - 629. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
B. T. Andresen, E. K. Jackson, and G. G. Romero Angiotensin II Signaling to Phospholipase D in Renal Microvascular Smooth Muscle Cells in SHR Hypertension, February 1, 2001; 37(2): 635 - 639. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. R. Dana, C. Eigsti, K. L. Holmes, and T. L. Leto A Regulatory Role for ADP-ribosylation Factor 6 (ARF6) in Activation of the Phagocyte NADPH Oxidase J. Biol. Chem., October 13, 2000; 275(42): 32566 - 32571. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J.-H. Parmentier, M. M. Muthalif, A. E. Saeed, and K. U. Malik Phospholipase D Activation by Norepinephrine Is Mediated by 12(S)-, 15(S)-, and 20-Hydroxyeicosatetraenoic Acids Generated by Stimulation of Cytosolic Phospholipase A2. TYROSINE PHOSPHORYLATION OF PHOSPHOLIPASE D2 IN RESPONSE TO NOREPINEPHRINE J. Biol. Chem., May 4, 2001; 276(19): 15704 - 15711. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
L.-A. Denmat-Ouisse, C. Phebidias, P. Honkavaara, P. Robin, B. Geny, D. S. Min, S. Bourgoin, M. A. Frohman, and M.-N. Raymond Regulation of Constitutive Protein Transit by Phospholipase D in HT29-cl19A Cells J. Biol. Chem., December 21, 2001; 276(52): 48840 - 48846. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Endocrinology | Endocrine Reviews | J. Clin. End. & Metab. |
| Molecular Endocrinology | Recent Prog. Horm. Res. | All Endocrine Journals |
