|
|
|
|
|
|
|
|
|||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | |||||
First published online 11 December 2007
doi: 10.1242/jcs.011692
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Research Article |
1 Center for Lung Biology, The University of South Alabama College of Medicine, Mobile, AL 36688, USA
2 Department of Pharmacology, The University of South Alabama College of Medicine, Mobile, AL 36688, USA
3 Department of Cell Biology and Neuroscience, The University of South Alabama College of Medicine, Mobile, AL 36688, USA
* Author for correspondence (e-mail: tstevens{at}jaguar1.usouthal.edu)
Accepted 24 September 2007
 |
Summary |
|---|
 Top Summary IntroductionResultsDiscussionMaterials and MethodsReferences |
|---|
II spectrin within this membrane domain. Although constitutive PDE4D4 activity limits cAMP access to the bulk cytosol, inhibiting its activity permits cAMP to access a cytosolic domain that is rich in microtubules, where it promotes protein kinase A (PKA) phosphorylation of tau at Ser214. Such phosphorylation reorganizes microtubules and induces interendothelial cell gap formation. Thus, spectrin-anchored PDE4D4 shapes the physiological response to cAMP by directing it to barrier-enhancing effectors while limiting PKA-mediated microtubule reorganization.
Key words: Adenylyl cyclase, Lung, Permeability
|
Introduction |
|---|
|
TopSummaryIntroductionResultsDiscussionMaterials and MethodsReferences |
|---|
; Mehta and Malik, 2006; Moore et al., 1998). Endothelial barrier strength relies on adenylyl cyclase production of cAMP just beneath the plasma membrane. Whereas increases in membrane cAMP enhance barrier function, decreased membrane cAMP impairs barrier function and increases permeability (Cioffi et al., 2002; Moore et al., 1998; Stevens et al., 1995). Even though cAMP is a readily diffusible molecule, cells maintain a membrane-to-cytosol cAMP diffusion gradient, suggesting its free diffusion into the whole cell compartment is hindered (Rich et al., 2000; Rich et al., 2001a; Willoughby et al., 2006). Although cellular organelles such as the endoplasmic reticulum may impede cAMP diffusion, selective targeting of cAMP to barrier-enhancing membrane microdomains is better achieved through spatial and temporal regulation of cAMP availability (Baillie et al., 2005; Barnes et al., 2005; Brunton, 2003; Conti et al., 2003; Houslay and Adams, 2003; Jin et al., 1998). PDEs hydrolyze cAMP to 5'AMP, and in so doing terminate cAMP signaling. PDEs thereby channel signaling to relevant effectors by degrading cAMP before it accesses physiologically inappropriate cellular domains.
Maintenance of the membrane-to-cytosol cAMP gradient limits cAMP access to targets that are potentially barrier disruptive. Indeed, production of cAMP outside the membrane domain disrupts, rather than strengthens, the endothelial cell barrier (Fischmeister, 2006; Sayner and Stevens, 2006; Sayner et al., 2006; Sayner et al., 2004). The bacterium Pseudomonas aeruginosa uses a type III secretion system to inject the exotoxin ExoY into endothelial cells, where ExoY acts as an adenylyl cyclase within the cytosol. ExoY-dependent cAMP synthesis disrupts the endothelial cell barrier, and increases lung permeability (Sayner et al., 2004). Similarly, heterologous expression of a soluble chimeric mammalian adenylyl cyclase (sAC) into pulmonary microvascular endothelial cells (PMVECs) results in cytosolic cAMP production. Activation of this sAC disrupts PMVEC monolayers, similarly to the actions of ExoY (Sayner et al., 2006). These findings collectively illustrate that the physiological actions of cAMP depend largely upon where it is made, and where it acts within the cell. Thus, under normal circumstances, endothelial cells must possess regulatory mechanisms that compartmentalize cAMP to domains that strengthen the endothelial cell barrier.
PDEs probably fulfil this regulatory role by controlling cAMP availability. Although endothelial cells express PDE4 (Creighton et al., 2003; Stevens et al., 1999; Zhu et al., 2004; Zhu et al., 2005), and possibly PDE3 (Suttorp et al., 1996; Suttorp et al., 1993) and PDE7 (Zhu et al., 2005), identity of the PDE isoform/splice variant that is responsible for maintaining membrane cAMP that strengthens barrier function is unknown. Our present findings reveal that the D4 splice variant of the PDE4 phosphodiesterase family PDE4D4, is expressed in PMVEC membranes. We show that PDE4D4 colocalizes with non-erythroid spectrin, a key component of the cytoskeletal and cell-tethering machinery (De Matteis and Morrow, 2000; Goodman and Zagon, 1986), near regions of cell-cell contact in PMVECs. Inhibiting membrane PDE4D4 activity increases the range of cAMP signaling and results in microtubule reorganization and endothelial cell barrier disruption. Thus, we report that PDE4D4 compartmentalizes cAMP, which is necessary to control PMVEC barrier integrity.
|
Results |
|---|
|
TopSummaryIntroductionResultsDiscussionMaterials and MethodsReferences |
|---|
). Our present studies therefore used caveolin-containing membranes to determine whether adenylyl cyclase (AC) and PDE activities reside within the same membrane fraction. Calcium concentrations reflecting those present under basal (100 nM) and stimulated (10 µM) conditions were used to confirm that membrane AC activity is inhibited by Ca2+, consistent with previous studies showing that AC6 activity dominates in this cell type. Increasing the Ca2+ concentration from 100 nM to 10 µM inhibited 97% of AC activity in PMVEC membranes (66.5±3.1 vs 4.6±0.2 pmol cAMP/mg protein/minute) compared with 22% inhibition of adenylyl cyclase activity in pulmonary artery endothelial cell (PAEC) membranes (18.5±3.3 vs 14.3±0.2 pmol cAMP/mg protein/minute) (Fig. 1A). Since PMVECs possess a high rate of cAMP-to-5'AMP turnover because of type 4 phosphodiesterase (PDE4) activity (Creighton et al., 2003; Stevens et al., 1999), our next studies examined the effect of rolipram (PDE4-specific inhibitor) on cAMP accumulation. In PMVEC membranes, rolipram (10 µM, EC100) increased cAMP content by 330% (78.7±0.6 vs 258.2±8.0 pmol cAMP/mg protein/minute), but had no effect on cAMP accumulation (33.4±2.8 vs 32.7±3.6 pmol cAMP/mg protein/minute) in PAEC membranes (Fig. 1B).
|
Phosphodiesterase activity studies indicated that PMVECs and PAECs possess similar rates of whole-cell cAMP hydrolysis (37.0±1.8 vs 38.0±1.6 pmol cAMP hydrolyzed/minute x107 cells, respectively; Fig. 1C). Total cAMP hydrolysis rates were also similar in PMVEC and PAEC membranes (12.2±0.2 vs 10.5±0.4 pmol cAMP hydrolyzed/minute/mg protein, respectively). However, rolipram inhibited 75% of cAMP PDE activity in PMVEC membranes (12.2±0.2 vs 3.4±0.1 pmol cAMP hydrolyzed/minute/mg protein) compared with 20% inhibition in PAEC membranes (10.5±0.4 vs 8.2±1.0 pmol cAMP hydrolyzed/minute/mg protein). Collectively, these studies indicate greater rolipram-sensitive PDE4 and Ca2+-inhibitable AC6 activities colocalize to PMVEC membranes than PAEC membranes.
PDE4D4 is expressed in PMVECs
Nearly 20 isoforms and splice variants of the PDE4 phosphodiesterase subfamily have been identified thus far. Although all PDE4 isozymes have closely related kinetic properties and are sensitive to rolipram inhibition, they vary in subcellular localization (Beard et al., 1999; Brunton, 2003; Georget et al., 2003; Houslay and Adams, 2003; Jin et al., 1998; Jurevicius et al., 2003; Karpen and Rich, 2001; Willoughby et al., 2006). Our next studies sought to identify the specific PDE4 responsible for localizing cAMP hydrolyzing activity to PMVEC membranes. Western blot analysis of the 30-40% sucrose gradient membranes using a pan PDE4 antibody revealed enrichment of a 115 kDa band in PMVEC membranes (Fig. 2A). Of the four genes that encode PDE enzymes, Pde4d encodes the largest number of proteins, including nine known splice variants, which are grouped into long [contain upstream conserved region (UCR) 1 and UCR2] and short forms (lack UCR1). Thus, the various lengths of the protein products are indicative of individual splice variants (Fig. 2B). Based on the molecular mass of the band in our western analysis (Fig. 2A), we identified the PDE4D splice variant as PDE4D4; these results were confirmed using pan PDE4, PDE4D (data not shown) and PDE4D4 (Fig. 2C) antibodies. PDE4D4-specific expression was verified by RT-PCR (Fig. 2D).
|
). Non-erythroid spectrin is composed of II and βII subunits that form a heterotetramer (De Matteis and Morrow, 2000; Goodman and Zagon, 1986). A single SH3 domain is located on each II spectrin subunit (Fig. 3A). Thus, co-immunoprecipitation studies were performed using antibodies to spectrin to determine whether PDE4D4-spectrin interaction occurs in lung endothelium. In whole-cell lysates, II spectrin co-immunoprecipitated with a 115 kDa PDE4 protein in PMVECs, but not in PAECs (Fig. 3B). These results are consistent with the 115 kDa PDE4 band obtained using 30-40% sucrose gradient membranes (see Fig. 2A). Since it is possible that other PDE4 proteins could be interacting with spectrin in lung endothelium, full-length PDE4D4 was overexpressed in PAECs and the co-immunoprecipitation studies were repeated. A 115 kDa band was only resolved in PAECs expressing PDE4D4, confirming that PDE4D4 and not other PDE4 isoforms expressed in PAECs interacts with II spectrin (Fig. 3B).
|
Confocal microscopy was performed to examine localization of PDE4 and spectrin in intact cells. PDE4 and spectrin colocalized at sites of cell-cell borders in PMVECs (Fig. 4A, top panel). By contrast, PDE4 and spectrin were more randomly dispersed on PAEC membranes and did not intensely colocalize (Fig. 4A, bottom panel).
|
We next determined whether PDE4D41-166 inhibited PDE4 activity. Membrane cAMP-PDE activity was reduced 
30% in PDE4D41-166-expressing PMVECs (9 pmol/minute/mg protein), compared with controls (13 pmol/minute/mg protein). Rolipram further reduced cAMP-PDE activity to 3.5 pmol/minute/mg protein (Fig. 4E). These findings are consistent with RT-PCR and western blot data, illustrating that PMVECs express PDE4B3, PDE4D4, PDE4D5 and PDE4A5 (data not shown). Rolipram inhibits all PDE4 isozymes, whereas expression of the PDE4D41-166 peptide reduces only a fraction of the total PDE4 activity, particularly in PMVECs.
Inhibiting PDE activity with rolipram or IBMX increases whole-cell cAMP. To determine whether PDE4D41-166 increases whole-cell cAMP, radioimmunoassay studies were performed. Basal cAMP levels were twofold higher in PMVECs expressing PDE4D41-166 compared with (control) mock-infected cells (120±14 pmol/106 cells vs 60±4 pmol/106 cells, respectively) (Fig. 4F). Forskolin stimulation of adenylyl cyclase increased cAMP to less than 150 pmol/106 cells in controls, and to more than 250 pmol/106 cells in PDE4D41-166 expressing PMVECs.
To address more rigorously whether PDE4D41-166 specifically inhibits PDE4D4 activity and not the activities of other PDEs, we infected PAECs with the retrovirus expressing PDE4D41-166, selected the cells to homogeneity and examined whole-cell PDE activity. II spectrin does not typically interact with PDE4 in PAECs (see Fig. 3B). However, when PDE4D4 was expressed in these PAECs, it co-immunoprecipitated with II spectrin (Fig. 4G). Rolipram inhibited approximately 40% of cAMP hydrolysis in (control) mock-infected cells, and approximately 50% of cAMP hydrolysis in PDE4D41-166-expressing PAECs (Fig. 4H). These findings indicate that PDE4D41-166 does not nonspecifically inhibit PDE4 (or PDE7) activity, and suggest that it functions as a dominant negative enzyme.
PDE4D4 inhibition potentiates cAMP signaling
Inhibition of PDE4D4 activity increases cAMP, and should also potentiate cAMP signaling. To determine whether such a potentiation of cAMP signaling occurs in cells expressing the catalytically inactive PDE4D4 peptide, PMVECs were fractionated into membrane and cytosolic fractions and PKA activity was assessed. Expression of the catalytically inactive PDE4D4 peptide increased both membrane and cytosolic PKA activity, and significantly potentiated activation of PKA by forskolin (Fig. 5).
|
; Sayner et al., 2004). The P. aeruginosa toxin ExoY is a bacterial adenylyl cyclase, which, when introduced into PMVECs, appears to localize with the microtubule-organizing center and/or microtubules (Sayner et al., 2004). We therefore questioned whether inhibition of PDE4D4 activity promotes cAMP signaling and influences microtubule architecture. PKA activity is an important determinant of microtubule architecture. Microtubules are stabilized by tau, and by other microtubule-assembly proteins. Multiple phosphorylation sites are prominent on tau. Phosphorylation dissociates tau from microtubules, and promotes microtubule disassembly (Schneider et al., 1999). Of the tau phosphorylation targets, only two represent consensus PKA phosphorylation sites, including Ser214 and Ser409 (Andorfer and Davies, 2000; Godemann et al., 1999; Jicha et al., 1999; Kyoung Pyo et al., 2004; Schneider et al., 1999; Zheng-Fischhofer et al., 1998). Ser409 is present on long forms of tau (tau-39 and tau-40), and is not found on the short form that is expressed in endothelium (tau-37; Fig. 6). Thus, we examined whether tau-Ser214 (consensus sequence...RTPSL...) is constitutively phosphorylated in PMVECs, and whether forskolin stimulation augments its phosphorylation. In control experiments, tau-Ser214 was not constitutively phosphorylated, and forskolin treatment did not increase phosphorylation (Fig. 6A). However, in cells expressing the catalytically inactive PDE4D4 peptide, tau-Ser214 was constitutively phosphorylated, and forskolin substantially increased its phosphorylation. tau-Ser262 is phosphorylated by multiple different protein kinases, including microtubule-affinity-regulating kinase, calmodulin-dependent kinase and calcium-dependent protein kinase, in addition to PKA. tau-Ser262 was not a PKA target in either control cells or cells expressing PDE4D41-166 (Fig. 6A, lower panel). Thus, tau-Ser214 represents a PKA target that is normally protected from phosphorylation by PDE4D4 activity. Inhibition of PDE4D4 activity allows the cAMP signal to result in tau-Ser214 phosphorylation.
|
Since phosphorylated tau reduces microtubule formation, studies were performed to assess whether phosphorylated tau-Ser214 was associated with depolymerized tubulin. As anticipated, immunoprecipitation of β-tubulin and immunoblotting for phosphorylated tau-Ser214 resolved phosphorylated tau-Ser214 in both control and PDE4D41-166-expressing cells (Fig. 6B). Forskolin increased tau-Ser214 phosphorylation in the depolymerized pool of β-tubulin, although the magnitude of this effect was greater in PDE4D41-166-expressing cells. Forskolin-induced tau-Ser214 phosphorylation was due to PKA activation, as the PKA antagonist H89 abolished the increase in tau-Ser214 phosphorylation (Fig. 6C). These findings are compatible with the idea that PKA phosphorylation of tau-Ser214 dynamically regulates microtubule architecture.
To further probe the interaction between microtubule architecture and tau-Ser214 phosphorylation, microtubule architecture was evaluated by confocal fluorescence microscopy. Forskolin treatment of control PMVEC cultures revealed no significant microtubule redistribution. However, forskolin treatment of PDE4D41-166-expressing cells realigned microtubules from a network distribution into elongated bundles (Fig. 6D). These changes in microtubule architecture paralleled a loss of cell-cell apposition.
PDE4D4 activity is a critical component of PMVEC barrier function
If PDE4D4 cAMP-hydrolyzing activity at the membrane is necessary for maintenance of endothelial cell-cell apposition and barrier integrity, then inactivating PDE4D4 in PMVECs using the catalytically inactive PDE4D4-GFP peptide should result in a loss of endothelial cell-cell interaction and cause gap formation. Microtubule reorganization accompanies endothelial cell gap formation and is, indeed, sufficient to produce gap formation (Birukova et al., 2006; Birukova et al., 2005; Birukova et al., 2004a; Birukova et al., 2004c; Petrache et al., 2003; Verin et al., 2001). Confluent control, vector control and PDE4D4-GFP-expressing PMVEC monolayers were monitored over time in the presence and absence of 1 µM forskolin. Images were taken every 2 minutes for 1 hour using time-lapse microscopy. In control and vector control monolayers, 1 µM forskolin did not cause a loss of PMVEC cell-cell contact (Fig. 7, top and bottom panels). However, in PDE4D4-GFP-expressing PMVECs, forskolin treatment initiated the formation of intercellular gaps that appeared 15 minutes after application; gaps did not reseal by the end of the experiment at 60 minutes (Fig. 7, second panel). Areas of gap formation in the second panel are expanded in the third panel. Formation of gaps in response to minimal stimulation of cAMP synthesis reveals a physiological role for PDE4D4 in the dynamic regulation of cell-cell apposition in PMVECs.
|
|
Discussion |
|---|
|
TopSummaryIntroductionResultsDiscussionMaterials and MethodsReferences |
|---|
; Jurevicius et al., 2003; Ostrom et al., 2001; Rybin et al., 2000). Rich and co-workers (Rich et al., 2000; Rich et al., 2001a) have indicated that cells maintain tenfold greater cAMP levels at the membrane compared with the bulk cytosol. Phosphodiesterase activity contributes to the membrane-to-cytosol cAMP gradient, by hydrolyzing cAMP to 5-AMP as it diffuses from its site of synthesis (Barnes et al., 2005; Willoughby et al., 2006). We now report that in PMVECs PDE4D4 activity is necessary to direct cAMP to its barrier-enhancing targets and to prevent activation of PKA within the cytosol that phosphorylates tau-Ser214, reorganizes microtubules and disrupts the PMVEC barrier.
PMVECs exhibit a high cAMP turnover rate that is attributable to rolipram-sensitive PDE4 activity (Creighton et al., 2003; Stevens et al., 1999). Given the considerable molecular diversity within the PDE4 enzyme family, we sought to resolve putative splice variants that may account for membrane-associated PDE4 activity. Global PCR analysis and western blotting using pan PDE4, PDE4D and PDE4D4 antibodies all resolved the presence of PDE4D4 in PMVEC membranes. Phosphodiesterase 4D4 was not abundantly expressed in PAEC membranes. PDE4D4 interacts with spectrin, especially along cell-cell borders.
The membrane distribution of PDE4D4 has important functional implications. Although PDEs exhibit Michaelis-Menton kinetics, PDE4 possesses a high Km of approximately 1-4 µmol (Rich et al., 2001a; Rich et al., 2001b). This high Km allows cAMP to achieve high membrane-associated concentrations. Inhibiting the activity of PDE4D4 would then be predicted to increase membrane cAMP and, perhaps most importantly, allow cAMP to access intracellular sites that are not normally accessible. In PMVECs expressing PDE4D41-166, basal and forskolin-stimulated cAMP concentrations were increased and PKA activation was potentiated. Such potentiation of PKA activation was prominent in the cytosolic fraction, which typically exhibited limited PKA activity. Increased cytosolic PKA activity phosphorylated tau-Ser214, and depolymerized and reorganized microtubules. Thus, PDE4D4 normally prevents cAMP from disrupting microtubule organization.
Microtubules regulate endothelial cell barrier strength (Birukova et al., 2006; Birukova et al., 2005; Birukova et al., 2004a; Birukova et al., 2004c; Petrache et al., 2003; Verin et al., 2001). In our studies, cytosolic PKA activation, tau-Ser214 phosphorylation and microtubule reorganization paralleled intercellular gap formation. These findings are consistent with prior work from other labs, which demonstrated that microtubule disruption is sufficient to increase permeability, whereas microtubule stabilization (i.e. with taxol) prevents inflammatory agonists from increasing permeability (Birukova et al., 2006; Birukova et al., 2005; Birukova et al., 2004a; Birukova et al., 2004c; Petrache et al., 2003; Verin et al., 2001). Indeed, inflammatory agonists activate intracellular signals that reorganize microtubule architecture, partly contributing to loss of endothelial barrier strength. Calcium transitions are associated with microtubule reorganization, as is the activation of Rho-Rho kinase. However, inflammatory mediators such as thrombin decrease, rather than increase, intracellular cAMP (Cioffi et al., 2002; Stevens et al., 1995), and thus cAMP signaling mechanisms have not been incriminated as a mechanism by which inflammatory agonists reorganize microtubules and increase permeability. Indeed, typical membrane-delimited cAMP fluctuations support microtubule architecture (Birukova et al., 2004b).
However, there is an emerging appreciation that cytosolic cAMP transitions occur during mitosis, and may contribute to microtubule reorganization at the spindle poles (Zippin et al., 2003). Although most mammalian adenylyl cyclases (AC1-AC9) are transmembrane proteins, one mammalian adenylyl cyclase, AC10, exhibits a cytosolic distribution. AC10 was first described in germ line cells; although we now know it is ubiquitously expressed (Buck et al., 1999; Chen et al., 2000; Hess et al., 2005; Stessin et al., 2006; Wu et al., 2006; Zippin et al., 2003; Zippin et al., 2004; Zippin et al., 2001). It localizes to intracellular organelles such as mitochondria and the nucleus, and to cytoskeletal structures like the microtubule-organizing center, i.e. centrosome (Zippin et al., 2003). At each of these sites, AC10 appears to co-associate with other members of the cAMP-signaling system, including PDEs, PKA and Epac (Diviani and Scott, 2001). The function of such discrete AC10-dependent cAMP pools is still poorly understood, but centrosome-associated AC10 activity has been implicated in control of microtubule organization during cell division (Zippin et al., 2003). Taken together with our present findings, PDE4D4 may protect microtubule structures from membrane-derived cAMP signaling in quiescent cells. However, inhibition of PDE4D4, or generation of cAMP from within the cytosol, may activate PKA and phosphorylate tau necessary for microtubule reorganization. Future studies will have to be completed to better resolve these possibilities.
The idea that selectively inhibiting membrane PDE4D4 activity results in a cAMP rise that disrupts the barrier stands in contrast to whole animal, isolated organ, and cultured endothelial cell studies using non-selective PDE inhibitors. In these prior studies PDE4 inhibition reduced or prevented the permeability increase that was caused by various mediators of lung injury (Adkins et al., 1992; Moore et al., 1998; Seibert et al., 1992). Part of the difference among these prior studies and our present results relates to the cell type studied. Whereas pan inhibition of PDE activity using 3-isobutyl-1-methylxanthine or PDE4 inhibition using rolipram strengthen barrier function in endothelial cells isolated from conduit blood vessels (e.g. aorta, pulmonary artery and umbilical vein), rolipram increases susceptibility to edema formation in the microcirculation of the lungs (Wu et al., 2005). Thus, PMVECs partition cAMP to membrane domains in a unique way, contributing to the phenotypic distinction between conduit and microvascular endothelial cells.
Perhaps most importantly, our findings indicate that PDE4D4 expressed in endothelial cells fulfils a unique physiological role in PMVECs by limiting the cAMP signal to barrier enhancing effectors or by protecting barrier disrupting effectors from activation. Our results are generally compatible with emerging evidence that PDE4D isozymes are discretely located within the cell to restrict cAMP to functional microdomains. Lehnart and colleagues (Lehnart et al., 2005) have recently shown that PDE4D3 is colocalized with the ryanodine receptor in the heart. Normal function of PDE4D3 is required for proper heart function, because loss of this enzyme results in prolonged Ca2+ release from the sarcoplasmic reticulum during excitation-contraction coupling, and contributes to development of congestive heart failure.
In summary, our data provide novel insight into the molecular composition and spatial organization of the cAMP-compartmentalizing machinery in lung microvascular endothelium. We identify a critical role for spectrin-anchored PDE4D4 in steering cAMP to barrier-enhancing membrane domains and preventing activation of cytosolic PKA. Such PKA activation phosphorylates tau-Ser214, reorganizes microtubules and disrupts the PMVEC barrier.
|
Materials and Methods |
|---|
|
TopSummaryIntroductionResultsDiscussionMaterials and MethodsReferences |
|---|
). Pepstatin A and leupeptin were from Peninsula Laboratories. Antibodies for PDE4, PDE4D, and PDE4D4 were obtained from Fab-Gennix (Shreveport, LA). Caveolin 1, βII spectrin, and HRP antibodies were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Antibody for II spectrin was purchased from Biomol (Plymouth Meeting, PA). Antibody for βII spectrin was a kind gift from Steve Goodman, University of Texas, Dallas, TX. GFP antibody was purchased from BioVision (Mountain View, CA). Secondary antibodies, conjugated to HRP or Alexa fluor fluorescent probes, were purchased from Invitrogen/Molecular Probes (Eugene, OR). Antibodies for β tubulin, polymerized (SMI62) and depolymerized (SMI61) microtubules were obtained from Covance (Berkeley, CA). The antibody for phosphorylated tau at Ser214 and Ser262 was purchased from Biosource (Camarillo, CA). Unless otherwise noted, all other materials and reagents were obtained from Sigma (St Louis, MO).
Isolation and culture of rat PMVECs and PAECs
Cells were isolated and cultured in DMEM supplemented with 10% FBS and penicillin-streptomycin using a method previously described by Stevens and colleagues (King et al., 2004; Stevens et al., 1999).
Preparation of cell membrane extractions
Pre-confluent PMVECs and PAECs were washed and collected in cold PBS. The cell pellets were re-suspended in TME-PI buffer (20 mM Tris-HCl, 5 mM MgCl2, 0.5 mM EDTA, pH 7.4), with protease inhibitors: 10 µM TLCK, 2 µM leupeptin, 2 µM pepstatin A, 10 µM benzamidine, 2000 U aprotinin/ml) and homogenized on ice. After centrifugation at 20,000 g for 20 minutes at 4°C, the membrane pellet (particulate fraction) was obtained by decanting and discarding the supernatant (cytosolic fraction) from the homogenate. The total membrane pellet or membranes isolated from the 30-40% sucrose gradient were resuspended in Triton lysis buffer (Cell Signaling, 20 mM Tris-HCl, pH 7.5, with 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 2.5 mM sodium pyrophosphate, 1 mM β-glycerophosphate, 1 mM Na3VO4, 1 µg/ml leupeptin and 1% Triton X-100) and extracted at 4°C for 15 minutes. The extracts were centrifuged as before and the supernatant used as the membrane fraction. The membrane extraction was used for PDE activity, western blot and immunoprecipitation studies.
Preparation of plasma membranes from 30-40% sucrose gradients
Sucrose-gradient-purified membranes were obtained using a previously described method (Stevens et al., 1999).
PDE activity measurement
PDE activity from 30-40% sucrose gradient fractions or whole membrane extractions was measured using 0.25 µM 3H-cAMP as the substrate. Rolipram (10 µM, EC100) was used as the PDE4-selective inhibitor (Creighton et al., 2003).
Measurement of adenylyl cyclase activity
In vitro determination of adenylyl cyclase activity was performed on isolated membranes as previously described (Stevens et al., 1999) with the following modifications. Reactions were allowed to proceed for 20 minutes at 30°C before the addition of pharmacological agents. Data are presented as pmol [32P]cAMP formed/mg protein/minute.
Measurement of cAMP concentration
Assessment of cAMP content was performed using standard radioimmunoassay (Biomedical Technologies, Stoughton, MA). Cells were seeded onto 2-cm2 24-well plates at 40,000 cells/ml, and grown to confluency over 3-4 days. Experiments were conducted in DMEM with physiological extracellular calcium concentrations and pH balanced to 7.4, with osmolality of 285-305 mosM. Agonists were added for the times indicated, cells were lysed and reactions stopped with 1 M NaOH. After assessment of cAMP concentrations, the results were standardized to cell counts (106 cells).
PKA assay
MESACUP Protein Kinase Assay ELISA system (Upstate, Lake Placid, NY) was used to measure PKA activity. Briefly, cells treated with or without forskolin (1 µM, 30 minutes) were stopped on ice, washed with cold PBS, and collected in cell lysate buffer (50 mM Tris-HCl, pH 7.5, 5 mM EDTA, 10 mM EGTA, 50 mM 2-mercaptoethanol, 1 mM PMSF, 10 mM benzamidine). After centrifugation at 20,000 g at 4°C for 40 minutes, the cytosolic (supernatant) and membrane (pellet resuspended in cell lysate buffer) fractions were used to perform kinase reactions on microwell strips coated with the phosphorylated form of the substrate peptide (RFARKGSLPQKNV). For in situ kinase activity, the reaction system contained 0.1 mM ATP, 0.5 mM EDTA, 1 mM EGTA, 3 mM MgCl2, and 5 mM 2-mercaptoethanol in 25 mM Tris-HCl, pH 7.0. For total activity, 2 µM cAMP was added to the reaction mixture. The reaction was initiated by addition of cell extractions and incubated at 25°C for 15 minutes. The reaction was stopped by H3PO4 (10%) for ELISA assay (OD490 nm).
Immunoprecipitation and western blot analysis
Whole cell membrane extractions or sucrose gradient derived membrane extractions in Triton Lysis Buffer were incubated with antibody and Ez-view Red Protein A Affinity Gel at 4°C for 12 hours. The gels were washed with the same incubation buffer and used for western blot assays. The protein samples from cell membrane extractions, sucrose gradient extractions, or immunoprecipitated proteins from the affinity gel were dissolved in SDS buffer for loading onto 7% precast Novex gels (Invitrogen). Secondary antibodies conjugated to AP were used to visualize the proteins.
Immunocytochemistry
Cells were seeded onto 12 mm coverslips and grown to confluency over 4-5 days. The cells were fixed with cold methanol for 1 minute and rehydrated in PBS + 5% milk for 15 minutes, then permeabilized using 0.1% Triton X-100 in PBS for 5 minutes. Cells were incubated overnight (4°C) with antibody to βII spectrin at 1:250 dilution in PBS + 5% milk. Cells were rinsed with fresh PBS + 5% milk + 0.1% Triton X-100 for 5 minutes, followed by PBS + 5% milk for 15 minutes, then the secondary antibody was incubated with the cells at 1:500 in PBS + 5% milk for 1 hour. Following a rinse with PBS + 5% milk + 0.1% Triton for 5 minutes then PBS + 5% milk for 15 minutes, antibody to PDE4 was added in PBS + 5% milk at 1:100 for 1 hour. The secondary antibody for PDE4 was applied in the same manner as for spectrin. After a final rinse in PBS the cells were mounted onto glass microscope slides using Dako (Carpintaria, CA) fluorescent mounting medium. Cells were imaged using a Leica TCs SP2 confocal microscope fitted with a 63x oil-immersion objective and Leica Microsystems Confocal Software Version 2.61.
Retroviral constructs
To generate a retrovirus that encoded a fusion between N-terminal portion of the rat PDE4D4 and EGFP, a fragment of rat PDE4D4 gene encoding the first 166 aa was amplified with primers PDE4D4ratF (gcggaattcgccaccatggaggcagagggcagcagcgtgc) and PDE4D4ratR (cgcagatctgctggtcatgggatccaag-ggactcc) using a pcDNA3 plasmid containing the full-length PDE4D4 rat sequence as template (accession number AF031373). EcoRI and BglII sites (italicized and underlined) were incorporated into primers to facilitate subsequent manipulations. The PCR product was digested with EcoRI and BglII and ligated to pMA1254 (EGFP containing adenovirus shuttle plasmid, unpublished data), which was digested with EcoRI and BamHI. The resulting plasmid, pMA1853, contains in-frame PDE4D4-EGFP fusion. This fusion was extracted from pMA1853 with EcoRI and XhoI and ligated to similarly digested pMA1629rc, a Moloney Murine Leukemia virus-based retroviral vector that encodes puromycin resistance (unpublished data). The construct obtained this way, pMA1870, was used to generate retroviral supernatants using Phoenix Ampho packaging cell line (a kind gift of Gary Nolan, Stanford University, Palo Alto, CA). To stably transfect the PDE4D4-GFP construct into PAECs and PMVECs, cells were rinsed with PBS, trypsinized, and transferred to 1.7 ml centrifuge tubes. Following gentle centrifugation (250 g for 5 minutes), the trypsin was aspirated and the cell pellet resuspended in the retroviral supernatant and incubated at 37°C for 1 hour prior to plating as for normal cell culture. Puromycin, at 5 µg/ml for PAECs and 20 µg/ml for PMVECs, was added to normal growth medium to select for fusion protein expression. Selection was considered complete after 3 days' incubation with antibiotic.
Cytosolic and microtubule-enriched extractions
Cells were collected in cold TME buffer (20 mM Tris-HCl, pH 7.5, 5 mM magnesium acetate and 1 mM EDTA) and centrifuged at 20,000 g at 4°C for 20 minutes. The cytosolic (supernatant) fraction was used to perform western blot and immunoprecipitation. For microtubule-enriched extraction, cells collected in cold TME buffer were kept on ice for 30 minutes to facilitate polymerized microtubule release to cytosolic fractions before centrifugation.
Immunocytochemistry for microtubule and phosphorylated tau
Cells were directly fixed in cold methanol and kept in –20°C for 5 minutes. After washing with PBS, cells were permeabilized using 1% Triton X-100 in PBS for 15 minutes at room temperature and blocked in 3% BSA for 1 hour. Antibodies for β tubulin and phosphorylated tau-Ser214 were added to the cells for 1 hour, incubated with secondary antibody conjugated with FITC or TRITC, and evaluated by confocal microscopy.
Gap studies
Cells were plated onto 25 mm coverslips and grown to confluency. Experiments were performed in Kreb's buffer with 2 mM calcium. Images were taken at 2 minute intervals for 60 minutes using an Olympus IX70 microscope and a 40x oil immersion lens. Diagnostics Instruments' (Sterling Heights, MI) Spot software version 4.0.9 was used to acquire and edit image sequences.
Statistical analysis
As appropriate, one-way ANOVA or unpaired Student's t-test was used to determine statistical significance between groups. Values represent means ± s.e.m. Data were presented using GraphPad Prism version 3.00 for Windows, GraphPad Software San Diego, CA. Values of P<0.05 were considered significant.
|
Acknowledgments |
|---|
|
Footnotes |
|---|
|
References |
|---|
|
TopSummaryIntroductionResultsDiscussionMaterials and MethodsReferences |
|---|
Adamson, R. H., Liu, B., Fry, G. N., Rubin, L. L. and Curry, F. E. (1998). Microvascular permeability and number of tight junctions are modulated by cAMP. Am. J. Physiol. 274, H1885-H1894.[Medline]
Adkins, W. K., Barnard, J. W., May, S., Seibert, A. F., Haynes, J. and Taylor, A. E. (1992). Compounds that increase cAMP prevent ischemia-reperfusion pulmonary capillary injury. J. Appl. Physiol. 72, 492-497.
Andorfer, C. A. and Davies, P. (2000). PKA phosphorylations on tau: developmental studies in the mouse. Dev. Neurosci. 22, 303-309.[CrossRef][Medline]
Baillie, G. S., Scott, J. D. and Houslay, M. D. (2005). Compartmentalisation of phosphodiesterases and protein kinase A: opposites attract. FEBS Lett. 579, 3264-3270.[CrossRef][Medline]
Barnes, A. P., Livera, G., Huang, P., Sun, C., O'Neal, W. K., Conti, M., Stutts, M. J. and Milgram, S. L. (2005). Phosphodiesterase 4D forms a cAMP diffusion barrier at the apical membrane of the airway epithelium. J. Biol. Chem. 280, 7997-8003.
Beard, M. B., O'Connell, J. C., Bolger, G. B. and Houslay, M. D. (1999). The unique N-terminal domain of the cAMP phosphodiesterase PDE4D4 allows for interaction with specific SH3 domains. FEBS Lett. 460, 173-177.[CrossRef][Medline]
Birukova, A. A., Birukov, K. G., Smurova, K., Adyshev, D., Kaibuchi, K., Alieva, I., Garcia, J. G. and Verin, A. D. (2004a). Novel role of microtubules in thrombin-induced endothelial barrier dysfunction. FASEB J. 18, 1879-1890.
Birukova, A. A., Liu, F., Garcia, J. G. and Verin, A. D. (2004b). Protein kinase A attenuates endothelial cell barrier dysfunction induced by microtubule disassembly. Am. J. Physiol. Lung Cell Mol. Physiol. 287, L86-L93.
Birukova, A. A., Smurova, K., Birukov, K. G., Usatyuk, P., Liu, F., Kaibuchi, K., Ricks-Cord, A., Natarajan, V., Alieva, I., Garcia, J. G. et al. (2004c). Microtubule disassembly induces cytoskeletal remodeling and lung vascular barrier dysfunction: role of Rho-dependent mechanisms. J. Cell. Physiol. 201, 55-70.[CrossRef][Medline]
Birukova, A. A., Birukov, K. G., Gorshkov, B., Liu, F., Garcia, J. G. and Verin, A. D. (2005). MAP kinases in lung endothelial permeability induced by microtubule disassembly. Am. J. Physiol. Lung Cell Mol. Physiol. 289, L75-L84.
Birukova, A. A., Adyshev, D., Gorshkov, B., Bokoch, G. M., Birukov, K. G. and Verin, A. D. (2006). GEF-H1 is involved in agonist-induced human pulmonary endothelial barrier dysfunction. Am. J. Physiol. Lung Cell Mol. Physiol. 290, L540-L548.
Brunton, L. L. (2003). PDE4: arrested at the border. Sci. STKE 2003, PE44.[Medline]
Buck, J., Sinclair, M. L., Schapal, L., Cann, M. J. and Levin, L. R. (1999). Cytosolic adenylyl cyclase defines a unique signaling molecule in mammals. Proc. Natl. Acad. Sci. USA 96, 79-84.
Chen, Y., Cann, M. J., Litvin, T. N., Iourgenko, V., Sinclair, M. L., Levin, L. R. and Buck, J. (2000). Soluble adenylyl cyclase as an evolutionarily conserved bicarbonate sensor. Science 289, 625-628.
Cioffi, D. L., Moore, T. M., Schaack, J., Creighton, J. R., Cooper, D. M. and Stevens, T. (2002). Dominant regulation of interendothelial cell gap formation by calcium-inhibited type 6 adenylyl cyclase. J. Cell Biol. 157, 1267-1278.
Conti, M., Richter, W., Mehats, C., Livera, G., Park, J. Y. and Jin, C. (2003). Cyclic AMP-specific PDE4 phosphodiesterases as critical components of cyclic AMP signaling. J. Biol. Chem. 278, 5493-5496.
Creighton, J. R., Masada, N., Cooper, D. M. and Stevens, T. (2003). Coordinate regulation of membrane cAMP by Ca2+-inhibited adenylyl cyclase and phosphodiesterase activities. Am. J. Physiol. Lung Cell Mol. Physiol. 284, L100-L107.
De Matteis, M. A. and Morrow, J. S. (2000). Spectrin tethers and mesh in the biosynthetic pathway. J. Cell Sci. 113, 2331-2343.[Abstract]
Diviani, D. and Scott, J. D. (2001). AKAP signaling complexes at the cytoskeleton. J. Cell Sci. 114, 1431-1437.[Abstract]
Fischmeister, R. (2006). Is cAMP good or bad? Depends on where it's made. Circ. Res. 98, 582-584.
Georget, M., Mateo, P., Vandecasteele, G., Lipskaia, L., Defer, N., Hanoune, J., Hoerter, J., Lugnier, C. and Fischmeister, R. (2003). Cyclic AMP compartmentation due to increased cAMP-phosphodiesterase activity in transgenic mice with a cardiac-directed expression of the human adenylyl cyclase type 8 (AC8). FASEB J. 17, 1380-1391.
Godemann, R., Biernat, J., Mandelkow, E. and Mandelkow, E. M. (1999). Phosphorylation of tau protein by recombinant GSK-3beta: pronounced phosphorylation at select Ser/Thr-Pro motifs but no phosphorylation at Ser262 in the repeat domain. FEBS Lett. 454, 157-164.[CrossRef][Medline]
Goodman, S. R. and Zagon, I. S. (1986). The neural cell spectrin skeleton: a review. Am. J. Physiol. 250, C347-C360.[Medline]
Head, B. P., Patel, H. H., Roth, D. M., Lai, N. C., Niesman, I. R., Farquhar, M. G. and Insel, P. A. (2005). G-protein-coupled receptor signaling components localize in both sarcolemmal and intracellular caveolin-3-associated microdomains in adult cardiac myocytes. J. Biol. Chem. 280, 31036-31044.
Hess, K. C., Jones, B. H., Marquez, B., Chen, Y., Ord, T. S., Kamenetsky, M., Miyamoto, C., Zippin, J. H., Kopf, G. S., Suarez, S. S. et al. (2005). The "soluble" adenylyl cyclase in sperm mediates multiple signaling events required for fertilization. Dev. Cell 9, 249-259.[CrossRef][Medline]
Houslay, M. D. and Adams, D. R. (2003). PDE4 cAMP phosphodiesterases: modular enzymes that orchestrate signalling cross-talk, desensitization and compartmentalization. Biochem. J. 370, 1-18.[CrossRef][Medline]
Jicha, G. A., Weaver, C., Lane, E., Vianna, C., Kress, Y., Rockwood, J. and Davies, P. (1999). cAMP-dependent protein kinase phosphorylations on tau in Alzheimer's disease. J. Neurosci. 19, 7486-7494.
Jin, S. L., Bushnik, T., Lan, L. and Conti, M. (1998). Subcellular localization of rolipram-sensitive, cAMP-specific phosphodiesterases. Differential targeting and activation of the splicing variants derived from the PDE4D gene. J. Biol. Chem. 273, 19672-19678.
Jurevicius, J., Skeberdis, V. A. and Fischmeister, R. (2003). Role of cyclic nucleotide phosphodiesterase isoforms in cAMP compartmentation following beta2-adrenergic stimulation of ICa,L in frog ventricular myocytes. J. Physiol. 551, 239-252.
Karpen, J. W. and Rich, T. C. (2001). The fourth dimension in cellular signaling. Science 293, 2204-2205.
King, J., Hamil, T., Creighton, J., Wu, S., Bhat, P., McDonald, F. and Stevens, T. (2004). Structural and functional characteristics of lung macro- and microvascular endothelial cell phenotypes. Microvasc. Res. 67, 139-151.[CrossRef][Medline]
Kyoung Pyo, H., Lovati, E., Pasinetti, G. M. and Ksiezak-Reding, H. (2004). Phosphorylation of tau at THR212 and SER214 in human neuronal and glial cultures: the role of AKT. Neuroscience 127, 649-658.[CrossRef][Medline]
Lehnart, S. E., Wehrens, X. H., Reiken, S., Warrier, S., Belevych, A. E., Harvey, R. D., Richter, W., Jin, S. L., Conti, M. and Marks, A. R. (2005). Phosphodiesterase 4D deficiency in the ryanodine-receptor complex promotes heart failure and arrhythmias. Cell 123, 25-35.[CrossRef][Medline]
Mehta, D. and Malik, A. B. (2006). Signaling mechanisms regulating endothelial permeability. Physiol. Rev. 86, 279-367.
Moore, T. M., Chetham, P. M., Kelly, J. J. and Stevens, T. (1998). Signal transduction and regulation of lung endothelial cell permeability. Interaction between calcium and cAMP. Am. J. Physiol. 275, L203-L222.[Medline]
Ostrom, R. S., Gregorian, C., Drenan, R. M., Xiang, Y., Regan, J. W. and Insel, P. A. (2001). Receptor number and caveolar co-localization determine receptor coupling efficiency to adenylyl cyclase. J. Biol. Chem. 276, 42063-42069.
Petrache, I., Birukova, A., Ramirez, S. I., Garcia, J. G. and Verin, A. D. (2003). The role of the microtubules in tumor necrosis factor-alpha-induced endothelial cell permeability. Am. J. Respir. Cell Mol. Biol. 28, 574-581.
Rich, T. C., Fagan, K. A., Nakata, H., Schaack, J., Cooper, D. M. and Karpen, J. W. (2000). Cyclic nucleotide-gated channels colocalize with adenylyl cyclase in regions of restricted cAMP diffusion. J. Gen. Physiol. 116, 147-161.
Rich, T. C., Fagan, K. A., Tse, T. E., Schaack, J., Cooper, D. M. and Karpen, J. W. (2001a). A uniform extracellular stimulus triggers distinct cAMP signals in different compartments of a simple cell. Proc. Natl. Acad. Sci. USA 98, 13049-13054.
Rich, T. C., Tse, T. E., Rohan, J. G., Schaack, J. and Karpen, J. W. (2001b). In vivo assessment of local phosphodiesterase activity using tailored cyclic nucleotide-gated channels as cAMP sensors. J. Gen. Physiol. 118, 63-78.
Rybin, V. O., Xu, X., Lisanti, M. P. and Steinberg, S. F. (2000). Differential targeting of beta -adrenergic receptor subtypes and adenylyl cyclase to cardiomyocyte caveolae. A mechanism to functionally regulate the cAMP signaling pathway. J. Biol. Chem. 275, 41447-41457.
Sayner, S. and Stevens, T. (2006). Soluble adenylate cyclase reveals the significance of compartmentalized cAMP on endothelial cell barrier function. Biochem. Soc. Trans. 34, 492-494.[CrossRef][Medline]
Sayner, S. L., Frank, D. W., King, J., Chen, H., VandeWaa, J. and Stevens, T. (2004). Paradoxical cAMP-induced lung endothelial hyperpermeability revealed by Pseudomonas aeruginosa ExoY. Circ. Res. 95, 196-203.
Sayner, S. L., Alexeyev, M., Dessauer, C. W. and Stevens, T. (2006). Soluble adenylyl cyclase reveals the significance of cAMP compartmentation on pulmonary microvascular endothelial cell barrier. Circ. Res. 98, 675-681.
Schneider, A., Biernat, J., von Bergen, M., Mandelkow, E. and Mandelkow, E. M. (1999). Phosphorylation that detaches tau protein from microtubules (Ser262, Ser214) also protects it against aggregation into Alzheimer paired helical filaments. Biochemistry 38, 3549-3558.[CrossRef][Medline]
Seibert, A. F., Thompson, W. J., Taylor, A., Wilborn, W. H., Barnard, J. and Haynes, J. (1992). Reversal of increased microvascular permeability associated with ischemia-reperfusion: role of cAMP. J. Appl. Physiol. 72, 389-395.
Stessin, A. M., Zippin, J. H., Kamenetsky, M., Hess, K. C., Buck, J. and Levin, L. R. (2006). Soluble adenylyl cyclase mediates nerve growth factor-induced activation of Rap1. J. Biol. Chem. 281, 17253-17258.
Stevens, T., Nakahashi, Y., Cornfield, D. N., McMurtry, I. F., Cooper, D. M. and Rodman, D. M. (1995). Ca (2+)-inhibitable adenylyl cyclase modulates pulmonary artery endothelial cell cAMP content and barrier function. Proc. Natl. Acad. Sci. USA 92, 2696-2700.
Stevens, T., Creighton, J. and Thompson, W. J. (1999). Control of cAMP in lung endothelial cell phenotypes. Implications for control of barrier function. Am. J. Physiol. 277, L119-L126.[Medline]
Suttorp, N., Weber, U., Welsch, T. and Schudt, C. (1993). Role of phosphodiesterases in the regulation of endothelial permeability in vitro. J. Clin. Invest. 91, 1421-1428.[Medline]
Suttorp, N., Ehreiser, P., Hippenstiel, S., Fuhrmann, M., Krull, M., Tenor, H. and Schudt, C. (1996). Hyperpermeability of pulmonary endothelial monolayer: protective role of phosphodiesterase isoenzymes 3 and 4. Lung 174, 181-194.[Medline]
Thompson, W. J., Ashikaga, T., Kelly, J. J., Liu, L., Zhu, B., Vemavarapu, L. and Strada, S. J. (2002). Regulation of cyclic AMP in rat pulmonary microvascular endothelial cells by rolipram-sensitive cyclic AMP phosphodiesterase (PDE4). Biochem. Pharmacol. 63, 797-807.[CrossRef][Medline]
Verin, A. D., Birukova, A., Wang, P., Liu, F., Becker, P., Birukov, K. and Garcia, J. G. (2001). Microtubule disassembly increases endothelial cell barrier dysfunction: role of MLC phosphorylation. Am. J. Physiol. Lung Cell Mol. Physiol. 281, L565-L574.
Willoughby, D., Wong, W., Schaack, J., Scott, J. D. and Cooper, D. M. (2006). An anchored PKA and PDE4 complex regulates subplasmalemmal cAMP dynamics. EMBO J. 25, 2051-2061.[CrossRef][Medline]
Wu, K. Y., Zippin, J. H., Huron, D. R., Kamenetsky, M., Hengst, U., Buck, J., Levin, L. R. and Jaffrey, S. R. (2006). Soluble adenylyl cyclase is required for netrin-1 signaling in nerve growth cones. Nat. Neurosci. 9, 1257-1264.[CrossRef][Medline]
Wu, S., Cioffi, E. A., Alvarez, D., Sayner, S. L., Chen, H., Cioffi, D. L., King, J., Creighton, J. R., Townsley, M., Goodman, S. R. et al. (2005). Essential role of a Ca2+-selective, store-operated current (ISOC) in endothelial cell permeability: determinants of the vascular leak site. Circ. Res. 96, 856-863.
Zheng-Fischhofer, Q., Biernat, J., Mandelkow, E. M., Illenberger, S., Godemann, R. and Mandelkow, E. (1998). Sequential phosphorylation of Tau by glycogen synthase kinase-3beta and protein kinase A at Thr212 and Ser214 generates the Alzheimer-specific epitope of antibody AT100 and requires a paired-helical-filament-like conformation. Eur. J. Biochem. 252, 542-552.[Medline]
Zhu, B., Kelly, J., Vemavarapu, L., Thompson, W. J. and Strada, S. J. (2004). Activation and induction of cyclic AMP phosphodiesterase (PDE4) in rat pulmonary microvascular endothelial cells. Biochem. Pharmacol. 68, 479-491.[CrossRef][Medline]
Zhu, B., Strada, S. and Stevens, T. (2005). Cyclic GMP-specific phosphodiesterase 5 regulates growth and apoptosis in pulmonary endothelial cells. Am. J. Physiol. Lung Cell Mol. Physiol. 289, L196-L206.
Zippin, J. H., Levin, L. R. and Buck, J. (2001). CO (2)/HCO (3)(–)–responsive soluble adenylyl cyclase as a putative metabolic sensor. Trends Endocrinol. Metab. 12, 366-370.[CrossRef][Medline]
Zippin, J. H., Chen, Y., Nahirney, P., Kamenetsky, M., Wuttke, M. S., Fischman, D. A., Levin, L. R. and Buck, J. (2003). Compartmentalization of bicarbonate-sensitive adenylyl cyclase in distinct signaling microdomains. FASEB J. 17, 82-84.
Zippin, J. H., Farrell, J., Huron, D., Kamenetsky, M., Hess, K. C., Fischman, D. A., Levin, L. R. and Buck, J. (2004). Bicarbonate-responsive "soluble" adenylyl cyclase defines a nuclear cAMP microdomain. J. Cell Biol. 164, 527-534.
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati 
Twitter What's this?
Related articles in JCS:
This article has been cited by other articles:
|
|
 |
|
  N. Prasain, M. Alexeyev, R. Balczon, and T. Stevens Soluble adenylyl cyclase-dependent microtubule disassembly reveals a novel mechanism of endothelial cell retraction Am J Physiol Lung Cell Mol Physiol, July 1, 2009; 297(1): L73 - L83. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
B. Zhu, L. Zhang, M. Alexeyev, D. F. Alvarez, S. J. Strada, and T. Stevens Type 5 phosphodiesterase expression is a critical determinant of the endothelial cell angiogenic phenotype Am J Physiol Lung Cell Mol Physiol, February 1, 2009; 296(2): L220 - L228. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
