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Am J Physiol Heart Circ Physiol 295: H77-H88, 2008. First published May 2, 2008; doi:10.1152/ajpheart.01355.2007
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A novel signaling pathway of ADP-ribosyl cyclase activation by angiotensin II in adult rat cardiomyocytes

Rukhsana Gul,1 Seon-Young Kim,1 Kwang-Hyun Park,1 Byung-Ju Kim,1 Se-Jin Kim,1 Mie-Jae Im,1 and Uh-Hyun Kim1,2

1Department of Biochemistry, 2Institute of Cardiovascular Research, Chonbuk National University Medical School, Jeonju, Republic of Korea

Submitted 20 November 2007 ; accepted in final form 28 April 2008


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
ADP-ribosyl cyclase (ADPR-cyclase) produces a Ca2+-mobilizing second messenger, cADP-ribose (cADPR), from NAD+. In this study, we investigated the molecular basis of ADPR-cyclase activation in the ANG II signaling pathway and cellular responses in adult rat cardiomyocytes. The results showed that ANG II generated biphasic intracellular Ca2+ concentration increases that include a rapid transient Ca2+ elevation via inositol trisphosphate (IP3) receptor and sustained Ca2+ rise via the activation of L-type Ca2+ channel and opening of ryanodine receptor. ANG II-induced sustained Ca2+ rise was blocked by a cADPR antagonistic analog, 8-bromo-cADPR, indicating that sustained Ca2+ rise is mediated by cADPR. Supporting the notion, ADPR-cyclase activity and cADPR production by ANG II were increased in a time-dependent manner. Application of pharmacological inhibitors and immunological analyses revealed that cADPR formation was activated by sequential activation of Src, phosphatidylinositol 3-kinase (PI 3-kinase)/protein kinase B (Akt), phospholipase C (PLC)-{gamma}1, and IP3-mediated Ca2+ signal. Inhibitors of these signaling molecules not only completely abolished the ANG II-induced Ca2+ signals but also inhibited cADPR formation. Application of the cADPR antagonist and inhibitors of upstream signaling molecules of ADPR-cyclase inhibited ANG II-stimulated hypertrophic responses, which include nuclear translocation of Ca2+/calcineurin-dependent nuclear factor of activated T cells 3, protein expression of transforming growth factor-β1, and incorporation of [3H]leucine in cardiomyocytes. Taken together, these findings suggest that activation of ADPR-cyclase by ANG II entails a novel signaling pathway involving sequential activation of Src, PI 3-kinase/Akt, and PLC-{gamma}1/IP3 and that the activation of ADPR-cyclase can lead to cardiac hypertrophy.

angiotensin II; calcium ion, adenosine 5'-triphosphate-ribosyl cyclase; cardiomyocytes

ADP-RIBOSYL CYCLASE (ADPR-cyclase) catalyzes production of cADP ribose (cADPR), a Ca2+-mobilizing metabolite, from nicotinamide adenine dinucleotide (NAD+) (12). cADPR is known to increase intracellular Ca2+ concentration ([Ca2+]i) by releasing Ca2+ from the intracellular stores or by Ca2+ influx through plasma membrane Ca2+ channels (15). CD38 represents the predominant ADPR-cyclase and is expressed ubiquitously (41). Studies with CD38 gene knockout mice (Cd38–/–) have indicated that this enzyme is involved in T cell activation (22), and oxytocin exocytosis that is closely related to autism (24), and acts mainly as a NAD+ glycohydrolase, thereby regulating intracellular NAD+ levels (27). An existence of other ADPR-cyclase(s) has also been suggested, since cADPR content of Cd38–/– tissues such as kidney, brain, or heart was nearly the same as that of the wild-type counterparts (34). In addition, Zn2+-sensitive and -reducing agent-insensitive ADPR-cyclases, which are characteristically different from CD38, are expressed in brain (5), heart (49), kidney (32), and arterial smooth muscle cells (6). Galione et al. (10) were the first to suggest that an extracellular stimulus results in the activation of ADPR-cyclase with an increase of intracellular cADPR levels. Subsequently, a number of studies have demonstrated that production of cADPR by ADPR-cyclases is stimulated by G protein-coupled receptors (GPCRs) such as receptors for epinephrine (3, 19), ANG II (8, 20), acetylcholine (21), and interleukin (IL)-8 (36). However, except IL-8-mediated CD38 activation in lymphokine activated killer (36) cells, the molecular mechanism of ADPR-cyclase activation by other GPCRs and subsequent cellular responses remains unknown.

The major renin-angiotensin system effector, ANG II, is an octapeptide, produced from angiotensinogen by the action of proteases, renin, and angiotensin-converting enzyme. ANG II plays a critical roles in cell growth, vascular contraction, migration, and salt-water retention (45). It also has a pathophysiological role in various cardiovascular diseases such as hypertension, cardiac hypertrophy, and congestive heart failure (7, 14, 37, 39). ANG II exerts its effects via two distinct receptor subtypes, angiotensin type 1 receptor (AT1R) and angiotensin type 2 receptor (AT2R) (7). The effects of ANG II are mainly mediated by AT1R activation (14, 37, 45) via complex interacting signaling pathways involving primary stimulation of phospholipase C (PLC) and Ca2+ mobilization and secondary activation of nonreceptor tyrosine kinase Src, phosphatidylinositol 3-kinase (PI 3-kinase)/protein kinase B (Akt), and mitogen-activated protein kinase (37). ANG II induces the [Ca2+]i rise by G protein-mediated activation of PLC that leads to phosphatidylinositol hydrolysis and formation of inositol trisphosphate (IP3) and diacylglycerol (DAG). IP3 is generally accepted to induce [Ca2+]i increase via the activation of IP3 receptor (IP3R) (2), whereas DAG can directly activate canonical transient receptor potential channel (TRPC)-3/6 that in turn slowly increases the membrane potential and concurrently activates L-type Ca2+ channel to induce Ca2+ influx (33). Increase of [Ca2+]i is augmented by cADPR through release from the intracellular stores, via ryanodine receptor (RyR) (11, 27, 30) and/or Ca2+ influx from the extracellular space through plasma membrane Ca2+ channels in a variety of cells (16, 18, 29, 34). ANG II-induced ADPR-cyclase activation has been observed in rat neonatal cardiomyocytes and afferent arterioles (8, 20). However, the signaling mechanism(s) of ADPR-cyclase activation remains elusive.

In this study, we investigated the molecular mechanism of ADPR-cyclase activation in ANG II signaling using adult rat primary cardiomyocytes, and the results showed that ADPR-cyclase is activated in a concerted way, involving Src, PI 3-kinase/Akt, and PLC-{gamma}1/IP3, resulting in upregulation of hypertrophic responses.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Reagents. ANG II, 8-bromo (Br)-cADPR, nicotinamide guanine dinucleotide (NGD+), and other chemicals were purchased from Sigma-Aldrich (St. Louis, Mo). Wortmannin, PD-123319, and xestospongin C were purchased from Calbiochem (Darmstadt, Germany). Losartan was purchased from Merck (Whitehouse Station, NJ). Anti-PLC-{gamma}1 monoclonal antibody was kindly provided by S. G. Rhee (Ewha Womans University, Seoul, Korea). Horseradish peroxidase-conjugated anti-mouse IgG, anti-goat IgG, and anti-rabbit IgG were purchased from Santa Cruz Biotechnology.

Animals. Sprague-Dawley male rats were obtained from Orientbio (Seoungnam, Korea). Specific pathogen-free 12- to 15-wk-old C57BL/6 CD38 knockout mice (Cd38–/–) and littermates (Cd38+/+) were obtained from Jackson Laboratories (Bar Harbor, ME). Animals were housed in a 12:12-h light-dark schedule with food and water ad libitum. All experimental animals used were under a protocol approved by the Institutional Animal Care and Use Committee of the Chonbuk National University Medical School. Standard guidelines for laboratory animal care were followed (23).

Preparation of cardiomyocytes. Ventricular cardiomyocytes were isolated from rats (200–220 g) and C57BL/6 Cd38+/+ and Cd38–/– mice (25–30 g) by using modification of the method described previously (49). Briefly, hearts were rapidly excised, cannulated, and subjected to retrograde perfusion on a Langendorff apparatus with Krebs-Henseleit (KH) buffer (in mmol/l: 10 HEPES, 118 NaCl, 4.7 KCl, 1.2 MgSO4, 1.2 KH2PO4, 25 NaHCO3, 10 pyruvate, 11 glucose, and 1 CaCl2, pH 7.37) for 2 min and then with Ca2+-free KH buffer for 2 min. The perfusion buffer was then changed to Ca2+-free KH buffer containing 0.5 mg/ml of collagenase type II and 0.08 mg/ml protease type IV for 25 min. The perfusate was gassed with 95% O2 and 5% CO2 and maintained at 37°C. The left ventricle was removed, chopped into small pieces, and incubated in a 50-ml Falcon tube at 37°C for 3 min with shaking, and then undigested tissue was allowed to settle down for ~1 min. Pellet containing undigested tissue was discarded, and Ca2+ concentration in the supernatants was gradually increased up to 1 mmol/l. Isolated cardiomyocytes were pelleted by centrifugation at 23 g for 1 min at room temperature and resuspended in a stabilizing buffer (pH 7.4) containing 20 mmol/l HEPES, 137 mmol/l NaCl, 4.9 mmol/l KCl, 1.2 mmol/l MgSO4, 15 mmol/l glucose, and 10 mmol/l, 2,3-butanedione monoxime, and 1 mmol/l Ca2+. The cell preparation was kept in the stabilizing buffer containing 1% BSA for ~15 min at 37°C and then washed and resuspended in medium 199 (Invitrogen, Grand Island, NY) supplemented with 2% albumin, 2 mmol/l L-carnitine, 5 mmol/l creatine, 5 mmol/l taurine, 100 IU/ml penicillin, 100 µg/ml streptomycin, and 25 µg/ml gentamycin. Cardiomyocytes were seeded on the plates coated previously with laminin for 1 h (10 µg/ml; Sigma). After incubation for 20 min at 37°C in an incubator humidified with 5% CO2 and 95% air, the medium was changed to remove round and unattached cells. This technique routinely yields 90% of cardiomyocytes retaining rod-shaped morphology. Experiments were performed on the same day or 12–16 h after isolation. Before treatment with blockers or ANG II, cardiomyocytes were incubated with a buffer containing 20 mmol/l HEPES, 137 mmol/l NaCl, 4.9 mmol/l KCl, 1.2 mmol/l MgSO4, 15 mmol/l glucose, and 1 mmol/l Ca2+ for 20 min.

[Ca2+]i measurement. Cardiomyocytes attached on laminin-coated confocal dishes were loaded with the Ca2+ indicator fluo 3-AM (3 µmol/l) (Molecular Probes, Eugene, OR) and incubated for 20 min at 37°C. Changes of [Ca2+]i in cardiomyocytes were determined at 488 nm excitation/530 nm emission by an air-cooled argon laser system (36). The emitted fluorescence at 530 nm was collected using a photomultiplier. One image every 3 s was scanned using a confocal microscope (Nikon). [Ca2+]i calculation was performed by using an equation given by Tsien et al. (45a), i.e., [Ca2+]i = Kd (F – Fmin)/(Fmax – F), where the dissociation constant (Kd) is 450 nM for fluo 3 and F is the observed fluorescence levels. Each tracing was calibrated for the maximal intensity (Fmax) by addition of 8 µmol/l ionomycin and for the minimal intensity (Fmin) by addition of 50 mmol/l EGTA at the end of each measurement.

Fluorometric assay for ADPR-cyclase. ADPR-cyclase activity was fluorometrically determined by measuring cyclic GDP-ribose (cGDPR) using NGD+ as a substrate (13). Briefly, aliquots of cardiomyocyte homogenates were incubated with NGD+ (200 µmol/l) in 0.1 mol/l sodium phosphate buffer (pH 7.2) at 37°C for 10 min. The reaction was stopped by adding 50 µl TCA (10%). The samples were centrifuged at 22,000 g for 10 min at 4°C, and the supernatant (90 µl) was diluted with 910 µl of sodium phosphate buffer (pH 7.2). Fluorescence of cGDPR produced was determined at excitation/emission wavelengths of 297/410 nm (Hitachi F-2000).

Measurement of cADPR level. Cyclic enzymatic assay was used to measure cADPR level as described previously (13). Briefly, cardiomyocytes were treated with 0.3 ml of 0.6 mol/l perchloric acid under sonication. Precipitates were removed by centrifugation at 20,000 g for 10 min at 4°C. To remove perchloric acid, aqueous sample was mixed with a solution containing 1,1,2-trichlorotrifluoroethane and tri-n-octlyamine in the ratio of 3:1. After centrifugation for 10 min at 1,500 g, the aqueous layer was collected and neutralized with 20 mmol/l sodium phosphate (pH 8). To remove contaminating nucleotides, the samples were incubated overnight with 2.5 mmol/l MgCl2 and the following enzymes in 20 mmol/l sodium phosphate buffer (pH 8.0) at 37°C: 0.44 U/ml nucleotide pyrophosphatase, 12.5 U/ml alkaline phosphatase, and 0.0625 U/ml NADase. Enzymes were removed by filtration using Centricon-3 filters. To convert cADPR to NAD+, the samples (0.1 ml/tube) were incubated with 50 µl of a cycling reagent containing 0.3 µg/ml Aplysia ADPR-cyclase, 30 mmol/l nicotinamide, and 100 mmol/l sodium phosphate (pH 8.0) at room temperature for 15 min. The samples were further incubated with the cycling reagent (0.1 ml) containing 2% ethanol, 100 µg/ml alcohol dehydrogenase, 20 µmol/l resazurin, 10 µg/ml diaphorase, 10 µmol/l riboflavin 5-phosphate, 10 mmol/l nicotinamide, 0.1 mg/ml BSA, and 100 mmol/l sodium phosphate (pH 8.0) for 2 h at room temperature. An increase in resorufin fluorescence was measured at 544 nm excitation and 590 nm emission using a fluorescence plate reader (Spectra-Max GEMINI; Molecular Devices).

Immunoblotting. Protein extraction and immunoblotting of cardiomyocytes were performed as previously described (36). Proteins (20 µg/lane) were resolved on 10% or 12% SDS-PAGE and transferred to polyvinylidene difluoride membranes (GE Healthcare, Little Chalfont, Buckinghamshire, UK). Blots were incubated overnight at 4°C with primary antibodies raised against Akt and phospho-Akt (Ser473) (1:2,500 dilution; Cell Signaling), transforming growth factor (TGF)-β1 (1:1,500 dilution; Santa Cruz Biotechnology), Src and phospho-Src (Tyr416) (1:2,500 dilution; Cell Signaling), phosphotyrosine (1:2,500 dilution; Cell Signaling), or actin (1:1,500 dilution; Cell Signaling). The blots were rinsed four times with blocking buffer and incubated with horseradish peroxidase-conjugated secondary antibodies (1:5,000 dilution of each antibody) for 1 h at room temperature. Signals were detected by using the enhanced chemiluminescence system (Bio-Rad, Munich, Germany). For quantitative measurements, blots were scanned, and density was normalized to the density of a control protein for each sample. Protein concentrations were determined using a Bio-Rad protein assay kit, and known concentrations of BSA were used as the standard.

Immunoprecipitaion. Immunoprecipitation with cardiomycyte lysates was performed as previously described (35). Briefly, after stimulation with ANG II (150 nmol/l) or ANG II plus drugs, cells were washed two times with ice-cold PBS (in mmol/l: 10 Na2HPO4, 1.7 KH2PO4, 136 NaCl, 2.6 KCl, and 1 Na3VO4, pH 7.4). Cells were then lysed in an ice-cold lysis buffer [20 mmol/l HEPES, pH 7.2, 1% Triton X-100, 10% glycerol, 100 mmol/l NaCl, 1 mmol/l EDTA, 1 mmol/l phenylmethylsulfonyl fluoride (PMSF), 1 mmol/l NaF, 1 mmol/l Na3VO4, 10 µg/ml leupeptin, 10 µg/ml pepstatin, and 10 µg/ml aprotinin]. The lysates collected were centrifuged at 22,000 g for 10 min at 4°C to remove debris. An anti-PLC-{gamma}1 antibody (5 µg) was added to the lysates (500 µg), and the mixtures were incubated overnight at 4°C, with gentle rotation, and then 50 µl of 50% protein-G agarose was added to the mixtures for an additional 2 h. The immunoprecipitates recovered by centrifugation were washed four times with the same lysis buffer. For immunoblot analysis, immunoprecipitated proteins were dissolved in 2x sample buffer, boiled for 10 min, and resolved on 8% SDS-polyacrylamide gel. The blots were probed with antibodies raised against phospho-Akt (Ser473), Akt (1:2,500 dilution), or PLC-{gamma}1 (1:1,500 dilution).

Preparation of cytosolic and nuclear fraction. Cytosolic and nuclear fractions were prepared by using modification of the method described previously (17). Briefly, the samples were homogenized in homogenization buffer containing 25 mmol/l Tris (pH 7.5), 0.5 mmol/l EDTA, 0.5 mmol/l EGTA, 1 mmol/l PMSF, 1 mmol/l dithiothreitol, 25 µg/ml leupeptin, 25 mmol/l NaF, and 1 mmol/l Na3VO4 and incubated on ice for 10 min with continuous vortexing. The homogenates were centrifuged at 22,000 g for 15 min, and the resulting supernatants were collected as cytosolic fraction. Insoluble pellets, which contain nuclei, were washed in buffer containing 10 mmol/l HEPES, 1.5 mmol/l MgCl2, 10 mmol/l KCl, 1 mmol/l dithiothreitol, 25 µg/ml leupeptin, and 1 mmol/l PMSF. Samples were centrifuged at 1,000 g for 10 min at 4°C, and the resulting nuclear pellets were resuspended in solubilizing buffer (20 mmol/l HEPES, 25% glycerol, 0.42 mol/l NaCl, 1.5 mmol/l MgCl2, 0.2 mmol/l EDTA, 0.5 mmol/l dithiothreitol, 1 mmol/l PMSF, and 1% Triton X-100) and vortexed for 15 s every 10 min for 40 min. After centrifugation at 22,000 g for 20 min, the resulting supernatants were collected and used as nuclear extracts. Protein concentration was determined using a Bio-Rad protein assay kit. Nuclear and cytosolic fractions (30 µg/lane) were subjected to immunoblotting analysis using an antibody raised against nuclear factor of activated T cells (NFAT) 3 (1:2,500 dilution; Santa Cruz Biotechnology).

[3H]leucine incorporation. Freshly isolated cardiomyocytes in 12-well plates were incubated in triplicate with 150 nmol/l ANG II in the presence or absence of blockers for 24 h. They were then incubated in the same medium with 1.0 µCi/ml [3H]leucine for 12 h. After aspiration, cells were washed with ice-cold PBS and fixed with 10% TCA for 30 min. After washing, radioactivity incorporated in the precipitate was determined by liquid scintillation counting.

Statistical analysis. All immunoreactive and phosphorylated signals were analyzed by densitometric scanning (Fuji Photo Film, Tokyo, Japan). Results were then normalized by arbitrarily setting the ratio of densitometry data of the control to 1.0. Data represent means ± SE of at least three separate experiments. Statistical comparisons were performed using one-way ANOVA followed by Scheffé's test. Statistical significance of difference between groups was determined using the Student's t-test. A value of P < 0.05 was considered to be significant.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Activation of AT1R by ANG II induces a sustained [Ca2+]i rise via IP3 and cADPR in cardiomyocytes. Treatment of cardiomyocytes with ANG II (150 nmol/l) produced a rapid initial peak and then sustained Ca2+ increase (Fig. 1A). These ANG II-evoked Ca2+ increases were completely blocked by the AT1R antagonist losartan (10 µmol/l) but not by the AT2R blocker PD-123319 (10 µmol/l) (Fig. 1, B and C). To determine second messengers involved in Ca2+ mobilization, a PLC inhibitor (U-73122), an IP3R blocker (xestospongin C), and an antagonistic cADPR analog (8-Br-cADPR) were employed. Pretreatment with U-73122 (2.5 µmol/l) or xestospongin C (2 µmol/l) completely inhibited the ANG II-mediated initial Ca2+ peak and sustained Ca2+ rise (Fig. 1, D and E). However, pretreatment with 8-Br-cADPR (100 µmol/l) blocked the sustained Ca2+ rise only but not the initial Ca2+ peak (Fig. 1F). To further ascertain the involvement of cADPR in the sustained Ca2+ increase, cardiomyocytes were exogenously stimulated with cADPR. Addition of cADPR (100 µmol/l) showed a sustained Ca2+ signal (Fig. 1G) that was completely blocked by pretreatment with 8-Br-cADPR (Fig. 1H). To clarify whether a prototype ADPR-cyclase, CD38, is involved in ANG II-induced Ca2+ signal, we assessed the ANG II-induced Ca2+ response in cardiomyocytes isolated from Cd38+/+ and Cd38–/– mice. Treatment with ANG II generated an initial peak and sustained Ca2+ rise in both Cd38+/+ and Cd38–/– cardiomyocytes (Fig. 1, I and K). Pretreatment with 8-Br-cADPR, however, had no effect on the initial peak Ca2+ signal, although it completely blocked the sustained Ca2+ signal in both Cd38+/+ and Cd38–/– cardiomyocytes (Fig. 1, J and L). These data suggest that AT1R in cardiomyoctes couples to an ADPR-cyclase but not CD38, thereby producing a sustained Ca2+ rise.


Figure 1
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  		Fig. 1. ANG II-stimulated Ca2+ increase in cardiomyocytes requires inositol trisphosphate receptor (IP3R) and ADP-ribosyl (ADPR) cyclase activation. A: representative tracings of Ca2+ response to 150 nmol/l ANG II in rat cardiomyocytes. B–F: typical response to ANG II after pretreatment with 10 µmol/l losartan (Los) (B), 10 µmol/l PD-123319 (C), 2 µmol/l xestospongin C (Xe C) (D), 2.5 µmol/l U-73122 (E), and 100 µmol/l 8-bromo (Br)-cADPR (F). G: representative tracings of Ca2+ response to 100 µmol/l cADP-ribose (cADPR) in rat cardiomyocytes. H: typical response to cADPR after pretreatment with 8-Br-cADPR. I and K: representative tracings of Ca2+ response to ANG II in Cd38+/+ and Cd38–/– cardiomyocytes. J and L: typical response to ANG II after pretreatment with 8-Br-cADPR in Cd38+/+ and Cd38–/– cardiomyocytes. M–O: representative tracings of Ca2+ response to ANG II in the absence of extracellular Ca2+ (M), after pretreatment with 20 µmol/l ryanodine (N), and 10 µmol/l nifedipine (O) in rat cardiomyocytes. Cardiomyocytes were treated with indicated concentrations of pageers and incubated at 37°C for 30 min before ANG II treatment. Arrows indicate the time to add ANG II.

 
To explore the molecular basis of cADPR-induced Ca2+ signal, we examined ANG II-mediated Ca2+ signal in the absence of extracellular Ca2+ and in the presence L-type Ca2+ channel blocker (nifedipine) or RyR blocker (ryanodine). The absence of extracellular Ca2+ (Fig. 1M) and treatment with nifedipine (10 µmol/l) (Fig. 1N) showed only initial Ca2+ peak, but not sustained Ca2+ increase. Pretreatment with high concentration of ryanodine (20 µmol/l) (Fig. 1O) resulted in a gradual decrease of the sustained Ca2+ signal. These data together suggest that AT1R activation by ANG II induces an increase of [Ca2+]i that involves an initial Ca2+ peak via Ca2+ release through IP3R and cADPR-mediated sustained Ca2+ signal via Ca2+ influx through L-type Ca2+ channel and Ca2+ release through RyR.

ADPR-cyclase activation requires IP3-mediated Ca2+ rise in ANG II signaling. The above data suggest that AT1R activation stimulated ADPR-cyclase/cADPR formation, since 8-Br-cADPR blocked the sustained Ca2+ signal. To corroborate the observation, the effect of ANG II on ADPR-cyclase activity and cADPR formation was examined in cardiomyocytes. Both ADPR-cyclase activity and cADPR levels were increased in a time-dependent manner and were sustained thereafter, showing that the ANG II-induced sustained Ca2+ increase is regulated by cADPR formation via ADPR-cyclase activation (Fig. 2, A and B). Inhibition of PLC or IP3R abolished the sustained Ca2+ rise (Fig. 1, D and E), suggesting that IP3-mediated Ca2+ rise by PLC stimulation is involved in activation of ADPR-cyclase. Therefore, the effects of PLC and IP3R blockers on ADPR-cyclase activation and cADPR formation were examined. Thus cardiomyocytes were pretreated with losartan, U-73122, or xestospongin C for 30 min, and ANG II-stimulated ADPR-cyclase activity (Fig. 2C) and cADPR formation (Fig. 2D) were assessed. All of the blockers significantly decreased both ADPR-cyclase activity and cADPR generation, indicating that ANG II-induced PLC activation via AT1R and Ca2+ mobilization through IP3R are required for the activation of ADPR-cyclase/cADPR generation.


Figure 2
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  		Fig. 2. ANG II-induced ADPR-cyclase activation requires phospholipase C (PLC)/inositol trisphosphate (IP3)-mediated Ca2+ rise. A and B: ANG II-mediated increase of ADPR-cyclase activity and cADPR production, respectively, in a time-dependent manner. C and D: ANG II stimulated ADPR-cyclase activity and cADPR production in the presence of Los, U-73122, and Xe C. Cardiomyocytes were treated with indicated concentrations of pageers and incubated at 37°C for 30 min before ANG II treatment. *P < 0.01 vs. control, #P < 0.01 vs. ANG II. Values are mean ± SE of 3 independent experiments.

 
ANG II-induced activation of ADPR-cyclase involves the PI 3-kinase/Akt pathway. An involvement of PI 3-kinase/Akt in ADPR-cyclase activation was examined using a PI 3-kinase inhibitor, wortmannin. Pretreatment of cardiomyocytes with wortmannin (100 nmol/l) completely abolished the ANG II-mediated Ca2+ increase and cADPR formation (Fig. 3, A and B), showing that PI 3-kinase activates ADPR-cyclase in ANG II signaling. Activation of PI 3-kinase by ANG II was also ensured by evaluating the level of phospho-Akt, a primary downstream target of PI 3-kinase. Thus Akt phosphorylation was evaluated by immunoblotting using anti-phospho-Akt Ser473 antibody. ANG II-induced Akt phosphorylation started within 15 s and was sustained up to 4 min, whereas total Akt levels were unaltered (Fig. 3C). Pretreatment with wortmannin blocked ANG II-induced Akt phosphorylation (Fig. 3D). However, U-73122, xestospongin C, and 8-Br-cADPR did not alter the ANG II-induced Akt phosphorylation, although losartan significantly inhibited the Akt phosphorylation (Fig. 3, D and E). These data indicate that the PI 3-kinase-Akt is involved in ANG II signaling pathway and that PLC/IP3/ADPR-cyclase/cADPR are activated downstream of PI 3-kinase/Akt.


Figure 3
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  		Fig. 3. ANG II-induced intracellular Ca2+ concentration ([Ca2+]i) increase and activation of ADPR-cyclase involves the phosphatidylinositol 3-kinase (PI 3-kinase)/protein kinase B (Akt) pathway. A and B: effect of 100 nmol/l wortmannin (Wor) on ANG II-stimulated increases in [Ca2+]i and cADPR formation, respectively. Top: representative immunoblot of ANG II-induced Akt phosphorylation in a time-dependent manner (C), ANG II-induced Akt phosphorylation in the presence of Wor and Los (D), and ANG II-induced Akt phosphorylation in the presence of U-73122, Wor, Xe C, and 8-Br-cADPR (E). Bottom: summary quantifications presented as the ratio of phosphorylated to total protein (p-Akt/Akt). P < 0.01 vs. control (*) and vs. ANG II (#). Values are means ± SE of 3 independent experiments.

 
Src is involved in ANG II-induced increase of [Ca2+]i and cADPR formation. An involvement of Src in ANG II-induced ADPR-cyclase activation was evaluated using a selective Src inhibitor, 4-amino-5-(4-chlorophenyl)-7-(t-butyl)pyrazolol[3,4-d] pyrimidine (PP2). Pretreatment of the cells with PP2 (10 µmol/l) completely inhibited both the ANG II-induced Ca2+ increase and cADPR formation (Fig. 4, A and B), indicating that Src may be involved in the initial Ca2+ rise and, in turn, PLC activation. To confirm ANG II-mediated Src activation, the level of phospho-Src (p-Src) was determined by immunoblotting using anti- p-Src Tyr416 antibody. The level of p-Src started to increase within 15 s and remained sustained up to 4 min after ANG II treatment (Fig. 4C). Pretreatment of cardiomyocytes with PP2 and losartan significantly reduced the level of p-Src. In addition, to understand the relationship between Src and PI 3-kinase in activation of ADPR-cyclase/Ca2+ signal, ANG II-induced phosphorylation of Src was determined in the presence of wortmannin. However, wortmannin did not alter the effects of ANG II on the Src phosphorylation (Fig. 4D). To evaluate the relationship between Src and PI 3-kinase/Akt, the effect of PP2 on the ANG II-induced Akt phosphorylation was also examined. Interestingly, pretreatment with PP2 significantly inhibited the ANG II-induced Akt phosphorylation (Fig. 4E), suggesting that Src and PI 3-kinase/Akt are in the same axis for activation of ADPR-cyclase.


Figure 4
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  		Fig. 4. Src phosphorylation is required for ANG II-induced PI 3-kinase/Akt activation. A and B: effects of 10 µmol/l PP2 on ANG II-stimulated increases in [Ca2+]i and cADPR formation, respectively. Top: representative immunoblot of time course of ANG II-stimulated Src phosphorylation (C), ANG II-mediated Src phosphorylation in the presence of PP2, Los, and Wor (D), and ANG II-mediated Akt phosphorylation in the presence of Wor and PP2 (E). Bottom: summary quantifications presented as the ratio of phosphorylated to total protein (p-Src/Src) or p-Akt/Akt. P < 0.01 vs. control (*) and vs. ANG II (#). Values are means ± SE of 3 independent experiments.

 
Akt and PLC-{gamma}1 form a complex, and ANG II stimulation phosphorylates Akt in the complex via PI 3-kinase. The above results indicate that activation of ADPR involves Src, PI 3-kinase/Akt, and PLC. Therefore, we examined whether ANG II activates PLC-{gamma}1-tyrosine phosphorylation of PLC-{gamma}1 by immunoprecipitating PLC-{gamma}1 with a PLC-{gamma}1 antibody, followed by Western blot analysis with a phosphotyrosine antibody. As shown in Fig. 5A, ANG II-induced PLC-{gamma}1 phosphorylation was rapid, occurring within 15 s. Pretreatment of cardiomyocytes with wortmannin blocked ANG II-induced phosphorylation of PLC-{gamma}1 (Fig. 5B), whereas, 8-Br-cADPR did not alter the ANG II-induced phosphorylation of PLC-{gamma}1 (Fig. 5B). These data indicate that PLC-{gamma}1 is involved in ANG II signaling and that it is activated downstream of PI 3-kinase/Akt but upstream of ADPR-cyclase.


Figure 5
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  		Fig. 5. ANG II stimulation of ADPR-cyclase involves upstream phosphorylation of Akt in Akt and PLC-{gamma}1 complex. Top, A: ANG II-induced tyrosine phosphorylation of PLC-{gamma}1 in a time-dependent manner. B: ANG-II-mediated tyrosine phosphorylation of PLC-{gamma}1 in the presence of Wor and 8-Br-cADPR. Immunoprecipitates of PLC-{gamma}1 were immunoblotted with anti-phospho-tyrosine (p-Tyr) antibody and then reprobed with anti-PLC-{gamma}1 antibody to demonstrate equal loading of protein in each lane. Bottom: summary quantifications presented as the ratio of phosphorylated to total protein (p-Tyr/PLC-{gamma}1). Top, C: ANG II-induced phosphorylation of Akt in Akt and PLC-{gamma}1 complex in a time-dependent manner. D: ANG-II-mediated phosphorylation of Akt in Akt and PLC-{gamma}1 complex in the presence of Wor and 8-Br-cADPR. Immunoprecipitates of PLC-{gamma}1 were blotted with anti-phospho-Akt antibody and then reprobed with antibodies against Akt and PLC-{gamma}1. Bottom: summary quantifications presented as the ratio of phosphorylated to total protein (p-Akt/PLC-{gamma}1). P < 0.01 vs. control (*) and vs. ANG II (#). Values are means ± SE of 3 independent experiments.

 
To further clarify the connection between PLC-{gamma}1 and PI 3-kinase/Akt, PLC-{gamma}1 was immunoprecipitated, and association of Akt with PLC-{gamma}1 was evaluated. The results revealed that PLC-{gamma}1 was associated with Akt and that treatment with ANG II increased the levels of phospho-Akt in the immunoprecipitates, starting as early as 30 s and remaining up to 8 min without change of the total Akt level (Fig. 5C). To further ascertain the observation that PI 3-kinase/Akt activates PLC-{gamma}1, cardiomyocytes, which had been pretreated with wortmannin or U-73122, were incubated with ANG II, and then PLC-{gamma}1 was immunoprecitated. The PLC-{gamma}1 immunoprecipitates were then immunoblotted with phospho-Akt antibody. As shown in Fig. 5D, the PI 3-kinase inhibitor wortmannin, but not the PLC inhibitor U-73122, significantly reduced the level of phospho-Akt without altering the Akt level. These results indicate that Akt and PLC-{gamma}1 exist as a complex at the basal level and that Akt in the complex is phosphorylated via PI 3-kinase, which is activated by ANG II, thereby stimulating PLC-{gamma}1.

ANG II-induced hypertrophic responses require ADPR-cyclase activation in rat cardiomyocytes. We also examined cellular responses of ADPR-cyclase activation by ANG II. Because ANG II is known to induce cardiac hypertrophy, known factors of cardiac hypertrophy were evaluated. To examine whether ANG II-induced stimulation of ADPR-cyclase activates NFAT3, which is known to be expressed in heart and involved in cardiac hypertrophy (31), nuclear and cytosolic fractions of cardiomyocytes were prepared after ANG II treatment, and they were then subjected to immunoblotting using an antibody against NFAT3. As shown in Fig. 6A, treatment of cardiomyocytes with ANG II increased the levels of NFAT3 in the nuclear fraction compared with that in the absence of ANG II (the control), whereas pretreatment with calcineurin inhibitor, cyclosporin A, losartan, wortmannin, and 8-Br-cADPR significantly reduced ANG II-induced NFAT3 nuclear translocation. These results indicate that NFAT3 translocation is downstream of ADPR-cyclase, and cADPR-mediated sustained Ca2+ signal activates NFAT3 through activation of calcineurin. ANG II also increased the level of TGF-β1 protein, whereas pretreatment with losartan, wortmannin, xestospongin C, or 8-Br-cADPR significantly reduced the protein expression (Fig. 6B), indicating that ADPR-cyclase signaling is associated with TGF-β1 expression. Finally, ANG II-induced cardiac hypertrophy was examined by determining [3H]leucine incorporation in the presence of cyclosporin A, losartan, wortmannin, xestospongin C, or 8-Br-cADPR (Fig. 6C). As expected, ANG II induced an increase of [3H]leucine incorporation, whereas treatment with the blockers significantly reduced ANG II-mediated [3H]leucine incorporation. These results indicate that ANG II-induced ADPR-cyclase activation can ultimately lead to cardiomyocyte hypertrophy.


Figure 6
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  		Fig. 6. ANG II-mediated ADPR-cyclase activation induces cardiac hypertrophy in vitro. A: representative immunoblot with summary quantifications of ANG II-stimulated nuclear factor of activated T cell (NFAT) 3 expression in nuclear (NFAT3N) and cytosolic (NFAT3C) fractions after pretreatment with cyclosporin A (CyA), Los, Wor, and 8-Br-cADPR. P < 0.01 vs. control (* and {infty}), vs. ANG II in nuclear fractions (#), and vs. ANG II in cytosolic fractions ({dagger}). B: representative immunoblot with summary quantifications of ANG II-stimulated transforming growth factor (TGF)-β1 expression after pretreatment with Los, Wor, Xe C, and 8-Br-cADPR. C: ANG II-induced [3H]leucine incorporation after pretreatment with Los, U-73122, Wor, Xe C, and 8-Br-cADPR. P < 0.01 vs. control (*) and vs. ANG II (#). Values are means ± SE of 3 independent experiments.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
To date, none of the studies has elucidated the molecular basis of ADPR-cyclase activation in ANG II signaling. Our study was set out to elucidate the molecular events that are involved in ANG II-induced ADPR-cyclase activation in adult rat cardiomyocytes and subsequent cellular responses. Our results revealed for the first time that activation of ADPR-cyclase by AT1R in adult rat cardiomyocytes was triggered by a sequential activation of Src, PI 3-kinase/Akt, PLC-{gamma}1, and IP3-mediated early Ca2+ increase (Fig. 7). Studies with cardiomyocytes isolated from Cd38+/+ and Cd38–/– indicated that the cADPR-induced Ca2+ increase was mediated by a novel ADPR-cyclase, but not CD38. Moreover, the results demonstrated that the activation of ADPR-cyclase generated not only a sustained Ca2+ signal via L-type Ca2+ channel and RyR but also enhanced hypertrophic responses that included NFAT nuclear translocation, TGF-β1 expression, and [3H]leucine incorporation.


Figure 7
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  		Fig. 7. Proposed model of the signaling pathways underlying ANG II-mediated ADPR-cyclase activation and cardiac hypertrophy. ANG II stimulation of angiotensin type 1 receptor (AT1R) leads to sequential activation of Src and PI 3-kinase, which in turn results in phosphorylation of Akt in Akt and PLC-{gamma}1 complex. Activation of PLC-{gamma}1 generates IP3, which induces Ca2+ release, on binding to its sarcoplasmic reticulum (SR) receptor, resulting in activation of ADPR-cyclase and cADPR formation. cADPR induces a sustained Ca2+ increase by Ca2+ influx and activation of ryanodine receptor (RyR) via CICR. Subsequently, the [Ca2+]i rise activates calcineurin, which causes NFAT3 nuclear translocation by its dephosphorylation, leading to cardiac hypertrophy.

 
Assessment of the characteristics of Ca2+ signals induced by ANG II showed an initial peak Ca2+ release via IP3R and then a sustained Ca2+ rise via two types of Ca2+ channels, L-type Ca2+ channel and RyR. The sustained Ca2+ rise appeared to be mediated by cADPR; a cADPR antagonist, 8-Br-cADPR, completely inhibited the sustained Ca2+ rise, but not the initial Ca2+ peak. The ANG II-induced cADPR-dependent sustained Ca2+ rise and the initial peak of Ca2+ rise were mediated by AT1R. An AT1R-specific blocker losartan completely abolished the ANG II-mediated Ca2+ signal, whereas the AT2R-specific blocker PD-123319 did not. Moreover, ANG II-induced ADPR-cyclase activity and cADPR formation were also effectively blocked by losartan. It appears that Ca2+ signal initiated by IP3 is a prerequisite for the cADPR-mediated sustained Ca2+ rise, since pretreatment of rat cardiomyocytes with the IP3R inhibitor xestospongin C completely abolished ANG II-mediated Ca2+ signals. The involvement of the IP3R-mediated Ca2+ signal in activation of CD38 by IL-8 has been observed in human LAK cells (36). In addition, the inhibition of RyR with a high concentration of ryanodine gradually reduced the sustained Ca2+ signal with time, suggesting that influx of Ca2+ through L-type Ca2+ channel induces Ca2+ release via RyR. A number of studies have shown that the primary target of cADPR to mediate Ca2+ release is RyR, which is one of the most important mechanisms of Ca2+-induced Ca2+ release (11, 28). Studies have proposed that cADPR is required for extracellular Ca2+ influx that in turn increases the amount of Ca2+ released from the intracellular store in neutrophils (16), mouse T cells (34), human LAK cells (36), and neuronal cells (18). Our findings confirmed that cADPR participates in this extracellular Ca2+ entry mediated by the activation of L-type Ca2+ channels. Although the molecular basis of activation of L-type Ca2+ channels by cADPR remains largely unknown, binding of cADPR to TRPM2 (melastatin-like transient receptor potential 2 channel) (44) and opening of L-type channel via activation of TRPC3/TRPC6 by ANG II (33) have been demonstrated. It has also been reported that ANG II-induced PI 3-kinase-dependent Akt activation leads to extracellular Ca2+ entry through L-type Ca2+ channels (45a). On the basis of observations by us and others, we propose that ANG II-mediated L-type Ca2+ channel activation via TRPCs or PI 3-kinase/Akt may be due to activation of ADPR-cyclase/cADPR formation. Previous studies have indicated that Src acts as a regulator in ANG II-stimulated Ca2+ mobilization, involving TRPC-mediated Ca2+ influx (26), and mediates AT1R signal to PI 3-kinase to induce cell proliferation, differentiation, and hypertrophy (9, 45a).

Studies on ANG II signaling in vitro and in vivo have revealed a complex, interacting signaling pathway involving Gq/Gi-PLC, G protein-independent activation of Src, PI 3-kinase/Akt, and other effectors (37). Our present study revealed that Src, PI 3-kinase/Akt, and PLC-{gamma}1 are upstream signaling molecules of ADPR-cyclase. The first line of evidence is that inhibitors of these effectors completely abolished the ANG II-mediated Ca2+ signals and the cADPR formation. A Src inhibitor, PP2, significantly reduced the levels of phospho-Akt and cADPR formation induced by ANG II, suggesting that Src is upstream of PI 3-kinase/Akt and ADPR-cyclase. Furthermore, a PI 3-kinase inhibitor, wortmannin, significantly inhibited ANG II-induced cADPR formation, suggesting that PI 3-kinase/Akt is upstream of ADPR-cyclase. ANG II-stimulated Akt phosphorylation was not inhibited by U-73122, xestospongin C, or 8-Br-cADPR, whereas a PI 3-kinase inhibitor, wortmannin, significantly inhibited ANG II-induced tyrosine phosphorylation of PLC-{gamma}1, indicating that PI 3-kinase/Akt is acting upstream of PLC. To date, a number of mechanisms have been proposed to explain the activation of PI 3-kinase/Akt induced by ANG II. Some studies demonstrated that the activation of Gq- or Gi-coupled ANG II receptors could mediate PI 3-kinase/Akt phosphorylation (38, 43), whereas other studies demonstrated that members of the Src-kinase family control PI 3-kinase/Akt activation (25). Furthermore, both PI 3-kinase and PLC-{gamma} have been shown to be involved in IP3 generation and intracellular Ca2+ release (38), and PI 3-kinase/Akt and PLC-{gamma}1 interact directly with each other (4, 35, 38). The interaction between PLC-{gamma}1 and Akt upon epidermal growth factor receptor stimulation has been also demonstrated in COS-7 cells (48). In fact, our data showed a direct interaction of Akt with PLC-{gamma}1, forming a complex at the basal state. Moreover, Akt in the complex was phosphorylated via PI 3-kinase activated by ANG II, suggesting that phosphorylation of Akt in the complex leads to PLC-{gamma}1 activation in ANG II signaling. Taken together, these findings suggest that ANG II-mediated phosphorylation of Akt via PI 3-kinase leads to PLC-{gamma}1 activation, thereby inducing an initial Ca2+ increase via IP3 formation and subsequently generating a sustained Ca2+ rise by ADPR-cyclase via cADPR formation.

Growing evidence indicates that the sustained rise of [Ca2+]i via membrane Ca2+ channels leads to cardiac hypertrophy (47). ANG II-mediated increase of [Ca2+]i through L-type Ca2+ has been shown to activate the Ca2+- and calmodulin-dependent phosphatase calcineurin, which in turn induces nuclear translocation of NFAT3 (31). NFAT transcription factors are normally hyperphosphorylated and sequestered in the cytoplasm, and these transcription factors are translocated into nucleus via calcineurin-mediated dephosphorylation upon an increase of [Ca2+]i. To understand physiological consequences of the cADPR-mediated sustained Ca2+ increase, we elucidated several hypertrophic maker proteins. Treatment of cardiomyocytes with ANG II increased the level of NFAT3 in nuclear fractions that was significantly inhibited by pretreatment with 8-Br-cADPR, cyclosporin A, losartan, and wortmannin, indicating that the ANG II-induced sustained Ca2+ rise mediated by cADPR activates calcineurin to induce NFAT3 nuclear translocation and hypertrophic responses. In support of the observation, [3H]leucine incorporation was also significantly reduced by cyclosporin A, losartan, wortmannin, or 8-Br-cADPR. TGF-β1, a founding member of a large superfamily, is upregulated in myocardium by increased workload and suffices to provoke the hypertrophic program of cardiac gene expression (40). TGF-β1 has been shown to be an important mediator of ANG II-stimulated cardiomyocyte hypertrophy (1, 42). Our present data are consistent with the concept that treatment of ANG II increases TGF-β1 expression in cardiomyocyte that may act as an autocrine/paracrine stimulus for cardiac hypertrophy. In addition, our results showed that the ANG II-induced increase of TGF-β1 protein expression was significantly decreased by pretreatment with losartan, wortmannin, xestospongin C, or 8-Br-cADPR, suggesting that ANG II-induced TGF-β1 expression requires the activation of PI 3-kinase/PLC-{gamma} and ADPR-cyclase/cADPR to induce sustained Ca2+ increase, which can ultimately stimulate cardiac hypertrophic responses.

In summary, we elucidated molecular mechanism underlying ADPR-cyclase activation in ANG II signaling and the crucial role of ADPR-cyclase in ANG II-induced hypertrophic responses in adult rat cardiomyocytes. Our findings may lead to a new therapeutic strategy of heart diseases, involving ANG II/AT1R and Ca2+ homeostasis.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
S. Y. Kim and M.-J. Im are the recipients of BK21 program of the Ministry of Education of Korea. R. Gul was supported by the foreigner support program from the Korea Research Foundation (KRF-2006-211-E00003). S.-J. Kim was supported by a grant of the Chonbuk National University Postdoctoral Fellow Program. This study was supported by Korea Research Foundation Grant KRF-2004-005-E00108 (U.-H. Kim).


   ACKNOWLEDGMENTS
 
We thank Dr. S. G. Rhee (Ewha Womans University, Seoul, Korea) for providing anti-PLC-{gamma}1 monoclonal antibody.


   FOOTNOTES
 

Address for reprint requests and other correspondence: U.-H. Kim, Dept. of Biochemistry, Chonbuk National Univ. Medical School, Keum-am dong, Jeonju, 561-182, Republic of Korea (e-mail: uhkim{at}chonbuk.ac.kr)

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.


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 GRANTS
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