| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
RECEPTORS AND SIGNAL TRANSDUCTION
1Department of Integrative Biology and Pharmacology, University of Texas Health Science Center at Houston, Houston, Texas; 2Department of Pharmacology, College of Medicine and Center for Lung Biology, University of South Alabama, Mobile, Alabama; 3Department of Physiology and Pharmacology, Oregon Health and Science University, Portland, Oregon; and 4Department of Gynecology and Obstetrics, Division of Reproductive Biology, Stanford University Medical Center, Stanford, California
Submitted 15 March 2006 ; accepted in final form 28 July 2006
 |
ABSTRACT |
|---|
 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
G protein signaling; protein kinase A; phosphodiesterase; A-kinase anchoring protein; CNG channel
When using these approaches one must be keenly aware of their advantages and disadvantages, and pick the sensor best suited to measure a specific signal. Two critical factors are the sensor's kinetics and buffering capacity (33). We have previously taken advantage of the rapid kinetics and low buffering capacity of CNG channel-based sensors to examine the subcellular compartmentalization of cAMP signals. We observed that 1) cAMP concentrations near the plasma membrane were >10-fold higher than total cAMP levels in C6–2B glioma cells and human embryonic kidney (HEK)-293 cells stably overexpressing adenylyl cyclase (AC) type 8; 2) cAMP accumulation at the plasma membrane was resistant to dialysis in the whole-cell, patch-clamp configuration; and 3) the wash-in of cAMP from the patch pipette to the channels was anomalously slow (31). In addition, we developed a compartmental model of cAMP signaling that was able to describe all of these results. The model predicted that in response to a stimulus, kinetically distinct cAMP signals could be observed within cells, and that cyclic nucleotides produced from distinct pools of cyclase (e.g., particulate and soluble guanylyl cyclase) would not have the same efficacy in activating CNG channels, even if similar total cellular cyclic nucleotide levels were reached. We have subsequently provided evidence supporting both predictions (28, 32). As part of the former study we found that prostaglandin E1 (PGE1) triggers a transient cAMP signal near the plasma membrane of these cells, and presented evidence that the decline in the signal was primarily due to a PGE1-induced stimulation of PDE activity.
Here, we have used CNG channel-based sensors to more closely examine the underlying mechanisms responsible for transient cAMP signals near the plasma membrane. Our results demonstrate that PKA-mediated stimulation of PDE4 activity underlies the decline in transient cAMP signals, and that the subcellular localization of PKA via A kinase anchoring proteins (AKAPs) is critical in the regulation of these signals. We used these data to develop a plausible kinetic model of near-membrane cAMP signals. The simulations demonstrate that stimulation of PDE4 activity is sufficient to cause the observed decline in transient cAMP signals. Furthermore, based on these simulations we propose that high local concentrations of PKA serve to buffer cAMP, contributing to its slow spatial spread within cells (23, 28, 31, 40). Resolving the kinetics of the near-membrane cAMP signals in this relatively simple cellular system should help us better understand the physiological consequences of PDE4 regulation and the mechanisms by which cAMP is localized to different cellular compartments.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
60% confluence in 100-mm dishes for infection with the C460W/E583M CNG-channel-encoding adenovirus construct (multiplicity of infection 10 plaque-forming units/cell) (34). Two hours postinfection, hydroxyurea was added to the cell media at 1 mM final concentration to inhibit viral replication. Twenty-four hours postinfection cells were detached with phosphate-buffered saline containing 0.03% EDTA, resuspended in serum-containing media, and assayed within 12 h. All experiments were conducted at room temperature, 20–22°C. Unless otherwise stated all reagents were purchased from Sigma.
Measurement of PDE activity.
Cyclic AMP PDE activity was measured according to the method of Thompson and Appleman (46) as detailed previously (16). Briefly, after incubation for 1–5 min with or without PGE2 (1 µM), forskolin (50 µM), or isoproteronol (10 µM), cells were harvested and homogenized in ice-cold hypotonic buffer (in mM), 20 Tris·HCl, pH 8.0, 50 NaF, 1 EDTA, 0.2 EGTA, 10 Na2PO4, 5 
-mercaptoethanol, and a protease inhibitor cocktail [leupeptin, 0.5 µg/ml; aprotinin, 4 µg/ml; benzamidine, 50 mM; pepstatin, 0.7 µg/ml; soybean trypsin inhibitor, 10 µg/ml; and PMSF, 10 µg/ml (freshly added before use)] using an all-glass homogenizer. In some experiments, homogenates were centrifuged at 14,000 g for 15 min and the supernatant (soluble) and pellets (particulate fraction) were assayed separately. Aliquots of the homogenates or of the fractionated extracts were assayed for PDE activities with 1 µM [3H]cAMP as a substrate. PDE4 activity was defined as the fraction of cAMP PDE activity inhibited by 10 µM rolipram (a PDE4 inhibitor). Protein concentrations were determined using the Bio-Rad protein assay (Bio-Rad Laboratories, Hercules, CA) with BSA as a standard. Experiments were repeated at least three times.
Western blot analysis. After incubation with the indicated substances, cells were homogenized in ice-cold hypotonic buffer and the protein concentration was determined. Samples (40 µg protein) were boiled in Laemmli buffer (25), subjected to electrophoresis on an 8% SDS-PAGE gel, and blotted onto Immobilon-P transfer membrane (Millipore, Bedford, MA). Membranes were blocked in TBS-0.1% Tween 20 containing 5% nonfat milk. The phosphorylated and total cAMP response element binding (CREB)/activating transcription factor (ATF) proteins were detected using mouse monoclonal (Upstate Biotechnology) and rabbit polyclonal antibodies (Cell Signaling), respectively and visualized by use of ECL detection reagents (Amersham Pharmacia Biotech).
Measurement of total cellular cAMP levels. HEK-293 cells were plated at 33% confluence in 12-well plates and assayed 24–48 h later. Cells were washed and assayed in a solution containing (in mM), 145 NaCl, 4 KCl, 10 HEPES, 10 D-glucose, 1 MgCl2, 1 CaCl2, pH 7.4. Additions were made from 100x stock solutions. Reactions were terminated by addition of 1 N HCl (0.1 N HCl final) and plates were incubated on ice for 15 min, after which the cells were scraped from the well. Cellular cAMP levels were measured using enzyme immunoassays (Direct Cyclic AMP Enzyme Immunoassay Kit, Assay Designs). Sample cAMP concentrations were calculated from standard curves. Data are presented as means ± SE, performed in triplicate.
Monitoring near-membrane cAMP signals in cell populations.
Cyclic AMP signals were monitored in cell populations as described previously (32–34). Briefly, we took advantage of the Ca2+ permeability of CNG channels comprised of the rat olfactory channel 
subunits, CNGA2 (14), and measured the rate of Ca2+ influx to monitor changes in cAMP levels. In this assay, an increase in local cAMP concentration causes activation of CNG channels and a subsequent increase in the rate of Ca2+ entry (12, 31–34). Changes in the rate of Ca2+ influx in response to stimuli reflect changes in the cAMP levels. We used the fluorescent indicator fura-2 to monitor Ca2+ influx in cell populations. Cells were loaded with 4 µM fura-2 AM (the membrane-permeant form, Calbiochem) at room temperature, 20–22°C, for 30–40 min, in MEM supplemented with 20 mM HEPES, pH 7.4. Cells were washed twice, then resuspended in a solution containing (in mM), 145 NaCl, 11 D-glucose, 10 HEPES, 4 KCl, 1 CaCl2, and 1 MgCl2, pH 7.4 (3–4 x 106 cells/3 ml buffer solution), and assayed using a PTI DeltaScan-1 spectrofluorimeter (Photon Technology International). PGE1, PGE2, rolipram, and the PKA inhibitor H89 (Calbiochem) were added to a stirred cuvette from concentrated DMSO stocks, with final concentrations as indicated (final DMSO concentrations were 
0.2%). The mixing time was estimated to be on the order of 5 s. Fluorescence was measured at an excitation wavelength of 380 nm and an emission wavelength of 510 nm. Under these conditions Ca2+ influx caused a decrease in fluorescence (
F), which was expressed relative to the prestimulus fluorescence (F0) to correct for variations in dye concentration, and to allow for comparison of results on different batches of cells. Data were sampled at either 5 or 10 Hz and filtered at 0.5 or 1 Hz. All data are representative of at least four experiments.
To ensure that extracellular application of H89 did not alter the Ca2+ handling properties of HEK-293 cells, we examined the effects of a 10 min pretreatment with 10 µM H89 on thapsigargin-induced changes in intracellular Ca2+ levels. Pretreatment with H89 did not significantly change the Ca2+ response induced by 1 µM thapsigargin in nominally Ca2+-free conditions (Ca2+ release from internal stores), or in the presence of 1 mM extracellular Ca2+ (Ca2+ release from internal stores and subsequent Ca2+ entry). Thus, it is unlikely that H89 significantly altered the Ca2+ handling properties of the cells.
Monitoring near-membrane cAMP signals in single cells.
Single-cell cAMP measurements were made using the whole-cell, patch-clamp technique. Recordings were made using an HEKA EPC10 patch-clamp amplifier. To ensure adequate voltage control, pipette resistance was limited to 4 M
and averaged 2.6 ± 0.2 M (n = 43). Voltage offsets were zeroed with the pipette in the bath solution; no additional corrections were made for the liquid junction potential difference. Experiments with a series resistance-induced error in excess of 5 mV were discarded. After achieving whole-cell configuration, the preparation was allowed to equilibrate for at least 10 min to ensure sufficient time for dialysis of compounds from the patch pipette into the cell. Current records were typically sampled at 10 kHz and filtered at 2 kHz and stored on a PC. Currents were recorded during 400-ms steps to a membrane potential of +20 mV from a holding potential of 0 mV. The pipette solution contained (in mM) 140 KCl, 0.5 MgCl2, 10 HEPES, 5 Na2ATP, 0.5 Na2GTP, pH 7.4; the bath solution contained (in mM) 140 NaCl, 4 KCl, 10 D-glucose, 10 HEPES, and either 0.1 or 10 MgCl2, pH 7.4. PGE1 and rolipram (Calbiochem) were added to control solutions from concentrated DMSO stocks (final DMSO concentrations 0.2%), with final concentrations as indicated. PKI (Calbiochem), stearated (St)-Ht31 (Promega), H89, and St-Ht31P control peptide (Promega) were aliquoted as 1,000x stock solutions and stored at –20°C. Solutions were applied using the SF-77B fast-step solution switcher (Warner Instruments). The mechanical switch time was 1–2 ms. The time required to exchange the extracellular solution was measured by applying a 140 mM KCl solution to a depolarized cell (+50 mV) and monitoring changes in current through endogenous voltage-gated K+ channels; for each experiment, it was less than 100 ms. The bulk solution within the bath chamber was changed within 20 s using a custom-built, gravity-driven perfusion system.
Data analysis and mathematical simulations. All data were analyzed using custom scripts written in the MATLAB programming environment (MathWorks) and statistical analysis was performed using SigmaPlot (v.9; Systat Software) using Student's t-test. Electrophysiological data were converted to formats compatible with MATLAB software using a custom script provided by Bruxton. Simulations were performed using the fourth-order Runge-Kutta solver in the MATLAB programming environment.
|
RESULTS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
Effects of H89 on PDE4 and AC activity.
To determine the extent to which prostaglandins induce activation of PDE4, we exposed HEK-293 cells to vehicle or 1 µM PGE2 for different times and then measured PDE4 activity in either the crude homogenate, or in the soluble and particulate fractions. PDE4 activity was determined as the rolipram-sensitive PDE activity. PGE2 treatment triggered a two- to threefold increase in PDE4 activity in both the soluble and particulate fractions of the cells (Fig. 1, A and B) as well as in the homogenate (data not shown). An increase in activity was detected as early as 1 min after PGE2 addition (Fig. 1, A and B). The increase in PDE4 activity was recovered in the pellet after immunoprecipitation with PDE4D-selective antibodies (data not shown), suggesting that activation of this isoenzyme plays a major role in this G protein-coupled receptor (GPCR)-dependent regulation of cAMP signals. Pretreatment of cells for 10 min with the PKA inhibitor H89 reduced the PGE2-induced stimulation of PDE4 activity in a dose-dependent manner (Fig. 1C). Half-maximal inhibition of PGE2-stimulated PDE activity occurred following pretreatment with 4 µM H89, and complete inhibition occurred at 10 µM H89. Similar results were observed following 10 µM isoproterenol-induced or 100 µM forskolin (an AC activator)-induced stimulation of cAMP production (data not shown).
|
40 nM (17), and could potentially inhibit other protein kinases. However, under these conditions we were unable to estimate the intracellular H89 concentrations. To ensure that H89 was indeed inhibiting PKA, we tested the dose dependence of H89 inhibition of CREB phosphorylation in the same preparation. Under these experimental conditions, H89 inhibited CREB phosphorylation with a similar dose dependence as it inhibited PDE4 stimulation (Fig. 1D). In control experiments, pretreatment with 1–30 µM H85, an analog of H89 that does not inhibit PKA, had little or no effect on prostaglandin-induced stimulation of PDE4 activity or CREB phosphorylation (data not shown).
We next examined the effects of H89 and rolipram on forskolin-, PGE1-, and PGE2-induced total cellular cAMP accumulation using enzyme immunoassays (Fig. 2). In response to vehicle alone (5 min) 0.9 ± 0.3 pmol/well of cAMP was measured, and there was no significant increase in cAMP levels following 10 min pretreatment with 10 µM H89 alone (open bars), pretreatment with 10 µM rolipram alone (cross-hatched bars) or H89 and rolipram (hatched bars). Stimulation with a subsaturating concentration of forskolin (1 µM, 5 min) triggered no significant increases in total intracellular cAMP, whereas PGE1 and PGE2 (1 µM, 5 min) triggered significant increases (3-fold over vehicle alone). Pretreatment with H89, rolipram, or H89 + rolipram triggered significant increases in forskolin- and prostaglandin-induced cAMP accumulation. There were no significant differences between the prostaglandin-induced cAMP accumulation observed following pretreatment with rolipram alone vs. H89 + rolipram, indicating that the primary effect of H89 was the inhibition of prostaglandin-induced stimulation of PDE4 activity. This is consistent with our previous work that demonstrated the rate of total cAMP accumulation is nearly linear in the presence of PDE inhibitors (see figure 1B of Ref. 32).
|
Figure 3A shows a typical response to 0.1 and 1 µM PGE1 in populations of HEK-293 cells (3–4 x 106 cells per cuvette) expressing C460W/E583M channels. Addition of PGE1 (added at 1 min, arrow) triggered an initial influx of Ca2+ through C460W/E583M channels, followed by a slower reduction in Ca2+ levels. Little or no response was observed in cells not expressing C460W/E583M channels. Pretreatment with 10 µM rolipram (3 min), a selective PDE4 inhibitor, prevented the decline in the responses triggered by either 0.1 or 1 µM PGE1 (Fig. 3B). Ten minute pretreatment with 10 µM H89 dramatically reduced the decline in the PGE1-induced responses (Fig. 3C). We determined that slopes of the PGE1-induced responses following 10 min pretreatment with 10 µM H89 and 3 min pretreatment with 10 µM rolipram were not significantly different than slopes of the PGE1-induced responses following pretreatment with rolipram alone (Fig. 3D). In control experiments, pretreatment with 10 µM H85 had little or no effect on prostaglandin-induced cAMP transients (data not shown). Also, pretreatment with 10 µM H89 had little or no effect on the time course or amplitude of Ca2+ influx in response to extracellular application of 100 µM pCPT-cGMP (a membrane permeant cGMP analog, data not shown), indicating that pretreatment with 10 µM H89 had little or no effect on CNG channel activity. These data are consistent with the hypothesis that PKA-mediated stimulation of PDE4 activity is primarily responsible for the decay in the cAMP response.
|
40 µM). Figure 4A shows a typical response of an HEK-293 cell expressing C460W/E583M channels to 1 µM PGE1. In this cell, PGE1 triggered a transient increase in outward current through C460W/E583M channels. This response is consistent with the PGE1-induced responses that we previously reported (32). Pretreatment of cells with 10 µM rolipram largely prevented the decline in the response (Fig. 4, B and D). Addition of rolipram following a PGE1-induced transient response triggered a large increase in current through CNG channels (Fig. 4C). These data demonstrate that the exposure of cells to PGE1 does not trigger cAMP accumulation to levels that saturate CNG channels, and are consistent with calibrated measurements of cAMP concentration near CNG channels, 0.7 ± 0.4 µM (32). To assess the extent to which rolipram prevented the decline in the cAMP response, we measured the percentage current remaining 3 min after peak current. In cells in which vehicle (0.1% DMSO) was added to the patch pipette solution 26 ± 6% of the peak current remained, whereas in cells that were pretreated with 10 µM rolipram, 92 ± 6% of the peak current remained (Fig. 4D).
|
2.5-fold higher than the KI for PKA, and significantly lower than the KI for other kinases), the rise in PGE1-induced responses was similar to that observed in control cells, whereas the decay was markedly less pronounced (compare Fig. 5A with Fig. 4A). At 200 nM H89 the decay was even less pronounced (Fig. 5B). Addition of H89 to the patch pipette increased the percentage of current remaining in a dose-dependent manner (a maximum of 68% in the presence of 0.5 or 1 µM H89, Fig. 5C). To further demonstrate the role of PKA-mediated feedback control of cAMP signals we used the highly specific PKA inhibitor PKI, KI 2.3 nM (6). We introduced PKI into the cell via the patch pipette because it does not readily cross the plasma membrane. In the presence of either 5 or 20 nM PKI, the decay of the responses was dramatically blunted (Fig. 6, A and B), with 81 ± 6% and 85 ± 4% current, respectively, remaining 3 min after the peak response (Fig. 6C). Taken together, the results presented thus far strongly indicate that PKA-mediated stimulation of PDE4 activity is required for the decline in the PGE1-triggered transient cAMP response.
|
|
|
 | (1) |
 | (2) |
 | (3) |
 | (4) |
is the time constant for the reduction in the synthesis rate. This reduction in cAMP synthesis may be due feedback regulation of AC activity or a small amount of receptor desensitization. The equations describing PKA activity are given below.
 | (5) |
 | (6) |
 | (7) |
 | (8) |
 | (9) |
 | (10) |
 | (11) |
 | (12) |
 | (13) |
 | (14) |
 | (15) |
|
|
3-fold higher than the free cAMP concentration. This occurs because the free cAMP levels are significantly higher than the K1/2 of PKA (200 nM), even after 300 s. Note that the total cAMP concentration near the plasma membrane would be higher if the PKA concentration were increased (due to the higher buffering capacity discussed below). We next used the model to examine the effects of PDE and PKA inhibitors (Fig. 9, D–F and G–I, respectively). In the presence of 10 µM rolipram, a competitive PDE4 inhibitor, cAMP levels quickly increase, saturating CNG channels (Fig. 9D). The model predicts that total cAMP levels near the plasma membrane reach 20 µM in 5 min, 8-fold higher than control conditions. This is higher than the threefold increase in total cellular cAMP measured using enzyme immunoassays (Fig. 2). Factors that may contribute to these differences, and were not incorporated into the simulations, include mechanisms that segregate near-membrane and cytosolic cAMP signals (see Refs. 31, 32 for details), and incomplete equilibration of rolipram across the plasma membrane. Simulations using (intracellular) rolipram concentrations of 1–2 µM are consistent with the experimental observations presented in Fig. 2. In the presence of 20 nM PKI, a noncompetitive PKA inhibitor, cAMP levels reach a steady plateau in 5 min. This response demonstrates the importance of PKA-mediated stimulation of PDE4 activity in shaping the cAMP signal. Although basal levels of PDE4 activity are sufficient to prevent saturation of CNG channels (Fig. 9G) and to offset the rate of cAMP synthesis, they are not sufficient to cause the observed decline in cAMP levels.
|
had relatively small effects on the transient cAMP signals, primarily lowering peak and final cAMP levels (Fig. 10, E and F). This should not be surprising given that PKA-mediated stimulation of PDE4 activity has such profound effects on cAMP signals in this system. It is likely that processes such as receptor desensitization will have a greater impact on the time course of cAMP signals in systems in which stimulation of PDE activity does not dominate the response.
|
|
DISCUSSION |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
A study by DiPilato et al. (8) used fluorescent cAMP indicators based on EPAC to examine the dynamics of cAMP signals in HEK-293 cells. The EPAC-based sensors were targeted to different regions of the cell and cAMP responses to 10 µM isoproterenol, 10 µM PGE1, or 50 µM forskolin were observed. The agonist-induced cAMP responses reached steady plateaus within 4 min, whether the sensors were targeted to the plasma membrane or diffusely distributed to the cytosol. Interestingly, they observed a slow spatial spread of cAMP, consistent with our previous results (28, 31, 32). The differences in the kinetics of the observed responses may have occurred because the membrane localized EPAC sensors traffic to different regions of the plasma membrane than CNG channels, and thus measure a different subcellular pool of cAMP. Another possibility is that the HEK-293 cells used in the study by DiPilato et al. were inherently different. Consistent with the latter possibility, we routinely observe transient isoproterenol-induced cAMP signals using either CNG channel-based sensors or enzyme immunoassays (data not shown); however, the responses presented by DiPilato et al. reached a steady plateau, regardless of the subcellular localization of the EPAC-based sensor. Experiments need to be conducted in the same cells to reconcile potential differences between the responses monitored with different cAMP sensors.
Recently, Rochais et al. (37, 38) used CNG channels to examine cAMP PDE activity in rat cardiac myocytes. They found that exposure of cardiac myocytes to 100 nM isoproterenol triggered a transient cAMP response; however, the decline of the response was slower than those measured in HEK-293 cells. Extracellular application of H89 increased isoproterenol-induced and L-858051 (an analog of forskolin)-induced cAMP signals, as measured by CNG channels, but did not alter isoproterenol- or L-858051-induced stimulation of total cellular PDE3 or PDE4 activities. While the basis for the H89 effects is not fully understood, they were likely due, at least in part, to a PKA-mediated increase in PDE3 or 4 activity at the plasma membrane. They also found that cAMP signals triggered by different hormones were regulated by distinct subsets of PDE families present. The latter observation further implicates compartmentalization as the primary mechanism for the specificity of cAMP signals within cardiac myocytes.
The high resolution measurements of near-membrane cAMP signals presented here have allowed us to develop a plausible mathematical model of the cellular mechanisms underlying these responses. This model can be used to better understand cAMP signals measured in other systems. We have used the model to investigate the roles of PKA, PDE4, and processes such as receptor desensitization in shaping cAMP signals, by altering the concentration (PKA) or kinetics (PDE4 and receptor desensitization) of relevant enzymes. Two interesting predictions can be drawn from these simulations. First, processes such as receptor desensitization may not contribute significantly to the time course of transient cAMP signals if PKA-mediated stimulation of PDE4 activity is present. It remains to be determined if regulation of PDE activity and receptor desensitization act in concert to regulate cAMP signals, or if one process typically dominates the time course of the response. Second, the model predicts that nonuniform concentrations of PKA may buffer cAMP to different extents in different subcellular compartments. Beautiful examples of nonuniform PKA distribution caused by AKAPs are presented in Refs. 26 and 50. AKAPs also help to generate nonuniform concentrations of other proteins, including PDE4 (10, 48). High levels of localized cAMP buffers (e.g., 5 µM PKA) and PDE4 would allow cAMP signals to activate PKA without activating EPAC or native CNG channels. In this scenario, EPAC and CNG channels would be activated after prolonged stimulation of AC (after the buffers had been saturated), or when PDE activity is inhibited. This hypothesis is particularly intriguing given the recent discovery of a signaling complex that contains mAKAP, PKA, PDE4D3, and EPAC1 (9). The measurement of local PKA concentration, as well as the concentration of other proteins within signaling modules, is an important area of future research. Such data are necessary if we are to determine whether localized PKA concentrations do indeed contribute to the segregation of cAMP-mediated cellular responses, and to understand how information is transmitted through the cAMP signaling pathway.
|
GRANTS |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
|
ACKNOWLEDGMENTS |
|---|
|
FOOTNOTES |
|---|
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
|
REFERENCES |
|---|
|
TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
2. Beavo JA, Bechtel PJ, Krebs EG. Activation of protein kinase by physiological concentrations of cyclic AMP. Proc Natl Acad Sci USA 71: 3580–3583, 1974.
3. Bolger GB, Erdogan S, Jones RE, Loughney K, Scotland G, Hoffmann R, Wilkinson I, Farrell C, Houslay MD. Characterization of five different proteins produced by alternatively spliced mRNAs from the human cAMP-specific phosphodiesterase PDE4D gene. Biochem J 328: 539–548, 1997.
4. Brunton LL, Hayes JS, Mayer SE. Functional compartmentation of cAMP and protein kinase in heart. Adv Cyclic Nucleotide Res 14: 391–397, 1981.[Web of Science][Medline]
5. Carr DW, Stofko-Hahn RE, Fraser ID, Bishop SM, Acott TS, Brennan RG, Scott JD. Interaction of the regulatory subunit (RII) of cAMP-dependent protein kinase with RII-anchoring proteins occurs through an amphipathic helix binding motif. J Biol Chem 266: 14188–14192, 1991.
6. Cheng HC, Kemp BE, Pearson RB, Smith AJ, Misconi L, Van Patten SM, Walsh DA. A potent synthetic peptide inhibitor of the cAMP-dependent protein kinase. J Biol Chem 261: 989–992, 1986.
7. Corbin JD, Sugden PH, Lincoln TM, Keely SL. Compartmentalization of adenosine 3':5'-monophosphate and adenosine 3':5'-monophosphate-dependent protein kinase in heart tissue. J Biol Chem 252: 3854–3861, 1977.
8. DiPilato LM, Cheng X, Zhang J. Fluorescent indicators of cAMP and Epac activation reveal differential dynamics of cAMP signaling within discrete subcellular compartments. Proc Natl Acad Sci USA 101: 16513–16518, 2004.
9. Dodge-Kafka KL, Soughayer J, Pare GC, Carlisle Michel JJ, Langeberg LK, Kapiloff MS, Scott JD. The protein kinase A anchoring protein mAKAP coordinates two integrated cAMP effector pathways. Nature 437: 574–578, 2005.[CrossRef][Medline]
10. Dodge KL, Khouangsathiene S, Kapiloff MS, Mouton R, Hill EV, Houslay MD, Langeberg LK, Scott JD. mAKAP assembles a protein kinase A/PDE4 phosphodiesterase cAMP signaling module. EMBO J 20: 1921–1930, 2001.[CrossRef][Web of Science][Medline]
11. Doskeland SO, Ogreid D. Characterization of the interchain and intrachain interactions between the binding sites of the free regulatory moiety of protein kinase I. J Biol Chem 259: 2291–2301, 1984.
12. Fagan KA, Schaack J, Zweifach A, Cooper DMF. Adenovirus encoded cyclic nucleotide-gated channels: a new methodology for monitoring cAMP in living cells. FEBS Lett 500: 85–90, 2001.[CrossRef][Web of Science][Medline]
13. Fraser ID, Tavalin SJ, Lester LB, Langeberg LK, Westphal AM, Dean RA, Marrion NV, Scott JD. A novel lipid-anchored A-kinase Anchoring Protein facilitates cAMP-responsive membrane events. EMBO J 17: 2261–2272, 1998.[CrossRef][Web of Science][Medline]
14. Frings S, Seifert R, Godde M, Kaupp UB. Profoundly different calcium permeation and blockage determine the specific function of distinct cyclic nucleotide-gated channels. Neuron 15: 169–179, 1995.[CrossRef][Web of Science][Medline]
15. Gray PC, Scott JD, Catterall WA. Regulation of ion channels by cAMP-dependent protein kinase and A-kinase anchoring proteins. Curr Opin Neurobiol 8: 330–334, 1998.[CrossRef][Web of Science][Medline]
16. Hansen G, Jin S, Umetsu DT, Conti M. Absence of muscarinic cholinergic airway responses in mice deficient in the cyclic nucleotide phosphodiesterase PDE4D. Proc Natl Acad Sci USA 97: 6751–6756, 2000.
17. Hidaka H, Watanabe M, Tokumitsu H. Search for functional substrate proteins of protein kinases and their selective inhibitors. Adv Second Messenger Phosphoprotein Res 24: 485–490, 1990.[Web of Science][Medline]
18. Hofmann F, Bechtel PJ, Krebs EG. Concentrations of cyclic AMP-dependent protein kinase subunits in various tissues. J Biol Chem 252: 1441–1447, 1977.
19. Houge G, Steinberg RA, Ogreid D, Doskeland SO. The rate of recombination of the subunits (RI and C) of cAMP-dependent protein kinase depends on whether one or two cAMP molecules are bound per RI monomer. J Biol Chem 265: 19507–19516, 1990.
20. Houslay MD, Baillie GS. beta-Arrestin-recruited phosphodiesterase-4 desensitizes the AKAP79/PKA-mediated switching of beta2-adrenoceptor signalling to activation of ERK. Biochem Soc Trans 33: 1333–1336, 2005.[CrossRef][Web of Science][Medline]
21. Houslay MD, Sullivan M, Bolger GB. The multienzyme PDE4 cyclic adenosine monophosphate-specific phosphodiesterase family: intracellular targeting, regulation, and selective inhibition by compounds exerting anti-inflammatory and antidepressant actions. Adv Pharmacol 44: 225–342, 1998.
22. Huang LJ, Taylor SS. Dissecting cAMP binding domain A in the RI subunit of cAMP-dependent protein kinase. Distinct subsites for recognition of cAMP and the catalytic subunit. J Biol Chem 273: 26739–26746, 1998.
23. Jurevicius J, Fischmeister R. cAMP compartmentation is responsible for a local activation of cardiac Ca2+ channels by -adrenergic agonists. Proc Natl Acad Sci USA 93: 295–299, 1996.
24. Jurevicius J, Skeberdis VA, Fischmeister R. Role of cyclic nucleotide phosphodiesterase isoforms in cAMP compartmentation following 2-adrenergic stimulation of ICa,L in frog ventricular myocytes. J Physiol 551: 239–252, 2003.
25. Laemmli UK. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227: 680–685, 1970.[CrossRef][Medline]
26. Mongillo M, McSorley T, Evellin S, Sood A, Lissandron V, Terrin A, Huston E, Hannawacker A, Lohse MJ, Pozzan T, Houslay MD, Zaccolo M. Fluorescence resonance energy transfer-based analysis of cAMP dynamics in live neonatal rat cardiac myocytes reveals distinct functions of compartmentalized phosphodiesterases. Circ Res 95: 67–75, 2004.
27. Nikolaev VO, Bünemann M, Hein L, Hannawacker A, Lohse MJ. Novel single chain cAMP sensors for receptor-induced signal propagation. J Biol Chem 279: 37215–37218, 2004.
28. Piggott LA, Hassell KA, Berkova Z, Morris AP, Silberbach M, Rich TC. Natriuretic peptides and nitric oxide stimulate cGMP synthesis in different cellular compartments. J Gen Physiol 128: 3–14, 2006.
29. Ponsioen B, Zhao J, Riedl J, Zwartkruis F, van der Krogt G, Zaccolo M, Moolenaar WH, Bos JL, Jalink K. Detecting cAMP-induced Epac activation by fluorescence resonance energy transfer: Epac as a novel cAMP indicator. EMBO Rep 5: 1176–1180, 2004.[CrossRef][Web of Science][Medline]
30. Reeves ML, Leigh BK, England PJ. The identification of a new cyclic nucleotide phosphodiesterase activity in human and guinea-pig cardiac ventricle. Implications for the mechanism of action of selective phosphodiesterase inhibitors. Biochem J 241: 535–541, 1987.[Web of Science][Medline]
31. Rich TC, Fagan KA, Nakata H, Schaack J, Cooper DMF, Karpen JW. Cyclic nucleotide-gated channels colocalize with adenylyl cyclase in regions of restricted cAMP diffusion. J Gen Physiol 116: 147–161, 2000.
32. Rich TC, Fagan KA, Tse TE, Schaack J, Cooper DMF, Karpen JW. A uniform extracellular stimulus triggers distinct cAMP signals in different compartments of a simple cell. Proc Natl Acad Sci USA 98: 13049–13054, 2001.
33. Rich TC, Karpen JW. Cyclic AMP sensors in living cells: what signals can they actually measure? Ann Biomed Eng 30: 1088–1099, 2002.[CrossRef][Web of Science][Medline]
34. Rich TC, Tse TE, Rohan JG, Schaack J, Karpen JW. In vivo assessment of local phosphodiesterase activity using tailored cyclic nucleotide-gated channels as cAMP sensors. J Gen Physiol 118: 63–77, 2001.
35. Richter W, Conti M. The oligomerization state determines regulatory properties and inhibitor sensitivity of type 4 cAMP-specific phosphodiesterases. J Biol Chem 279: 30338–30348, 2004.
36. Richter W, Jin SL, Conti M. Splice variants of the cyclic nucleotide phosphodiesterase PDE4D are differentially expressed and regulated in rat tissue. Biochem J 388: 803–811, 2005.[CrossRef][Web of Science][Medline]
37. Rochais F, Abi-Gerges A, Horner K, Lefebvre F, Cooper DMF, Conti M, Fischmeister R, Vandecasteele G. A specific pattern of phosphodiesterases controls the cAMP signals generated by different Gs-coupled receptors in adult rat ventricular myocytes. Circ Res 98: 1081–1088, 2006.
38. Rochais F, Vandecasteele G, Lefebvre F, Lugnier C, Lum H, Mazet JL, Cooper DMF, Fischmeister R. Negative feedback exerted by cAMP-dependent protein kinase and cAMP phosphodiesterase on subsarcolemmal cAMP signals in intact cardiac myocytes: an in vivo study using adenovirus-mediated expression of CNG channels. J Biol Chem 279: 52095–52105, 2004.
39. Saucerman JJ, Brunton LL, Michailova AP, McCulloch AD. Modeling beta-adrenergic control of cardiac myocyte contractility in silico. J Biol Chem 278: 47997–48003, 2003.
40. Sayner SL, Alexeyev M, Dessauer CW, Stevens T. Soluble adenylyl cyclase reveals the significance of cAMP compartmentation on pulmonary microvascular endothelial cell barrier. Circ Res 98: 675–681, 2006.
41. Sette C, Conti M. Phosphorylation and activation of a cAMP-specific phosphodiesterase by the cAMP-dependent protein kinase. Involvement of serine 54 in the enzyme activation. J Biol Chem 271: 16526–16534, 1996.
42. Sette C, Iona S, Conti M. The short-term activation of a rolipram-sensitive, cAMP-specific phosphodiesterase by thyroid-stimulating hormone in thyroid FRTL-5 cells is mediated by a cAMP-dependent phosphorylation. J Biol Chem 269: 9245–9252, 1994.
43. Smith SB, White HD, Siegel JB, Krebs EG. Cyclic AMP-dependent protein kinase I: cyclic nucleotide binding, structural changes, and release of the catalytic subunits. Proc Natl Acad Sci USA 78: 1591–1595, 1981.
44. Steinberg SF, Brunton LL. Compartmentation of G protein-coupled signaling pathways in cardiac myocytes. Annu Rev Pharmacol Toxicol 41: 751–773, 2001.
45. Sutherland EW. Studies on the mechanism of hormone action. Science 177: 401–408, 1972.
46. Thompson WJ, Appleman MM. Multiple cyclic nucleotide phosphodiesterase activities from rat brain. Biochemistry 10: 311–316, 1971.[CrossRef][Medline]
47. Willoughby D, Wong W, Schaack J, Scott JD, Cooper DMF. An anchored PKA and PDE4 complex regulates subplasmalemmal cAMP dynamics. EMBO J 25: 2051–2061, 2006.[CrossRef][Web of Science][Medline]
48. Wong W, Scott JD. AKAP signalling complexes: focal points in space and time. Nat Rev Mol Cell Biol 5: 959–970, 2004.[CrossRef][Web of Science][Medline]
49. Xiang Y, Naro F, Zoudilova M, Jin SL, Conti M, Kobilka B. Phosphodiesterase 4D is required for beta2 adrenoceptor subtype-specific signaling in cardiac myocytes. Proc Natl Acad Sci USA 102: 909–914, 2005.
50. Zanolo M, Pozzan T. Discrete microdomains with high concentration of cAMP in stimulated rat neonatal cardiac myocytes. Science 295: 1711–1715, 2002.
This article has been cited by other articles:
|
|
 |
|
  S. Togo, X. Liu, X. Wang, H. Sugiura, K. Kamio, S. Kawasaki, T. Kobayashi, R. F. Ertl, Y. Ahn, O. Holz, et al. PDE4 inhibitors roflumilast and rolipram augment PGE2 inhibition of TGF-{beta}1-stimulated fibroblasts Am J Physiol Lung Cell Mol Physiol, June 1, 2009; 296(6): L959 - L969. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S. P. Adderley, E. A. Dufaux, M. Sridharan, E. A. Bowles, M. S. Hanson, A. H. Stephenson, M. L. Ellsworth, and R. S. Sprague Iloprost- and isoproterenol-induced increases in cAMP are regulated by different phosphodiesterases in erythrocytes of both rabbits and humans Am J Physiol Heart Circ Physiol, May 1, 2009; 296(5): H1617 - H1624. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 V. De Arcangelis, D. Soto, and Y. Xiang Phosphodiesterase 4 and Phosphatase 2A Differentially Regulate cAMP/Protein Kinase A Signaling for Cardiac Myocyte Contraction under Stimulation of {beta}1 Adrenergic Receptor Mol. Pharmacol., November 1, 2008; 74(5): 1453 - 1462. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 H. Liu, J. A. Enyeart, and J. J. Enyeart ACTH Inhibits bTREK-1 K+ Channels through Multiple cAMP-dependent Signaling Pathways J. Gen. Physiol., August 1, 2008; 132(2): 279 - 294. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
W. Xin, T. M. Tran, W. Richter, R. B. Clark, and T. C. Rich Roles of GRK and PDE4 Activities in the Regulation of {beta}2 Adrenergic Signaling J. Gen. Physiol., March 31, 2008; 131(4): 349 - 364. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Visit Other APS Journals Online |
