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Originally published In Press as doi:10.1074/jbc.M304396200 on September 12, 2003

J. Biol. Chem., Vol. 278, Issue 48, 48154-48161, November 28, 2003
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Localization of Functional Endothelin Receptor Signaling Complexes in Cardiac Transverse Tubules*

Valentin G. Robu{ddagger}, Emily S. Pfeiffer{ddagger}, Seth L. Robia{ddagger}, Ravi C. Balijepalli§, YeQing Pi{ddagger}, Timothy J. Kamp{ddagger}§, and Jeffery W. Walker{ddagger}

From the Departments of {ddagger}Physiology and §Medicine, University of Wisconsin, Madison, Wisconsin 53706

Received for publication, April 28, 2003 , and in revised form, September 8, 2003.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Endothelin-1 (ET-1) is an autocrine factor in the mammalian heart important in enhancing cardiac performance, protecting against myocardial ischemia, and initiating the development of cardiac hypertrophy. The ETA receptor is a seven-transmembrane G-protein-coupled receptor whose precise subcellular localization in cardiac muscle is unknown. Here we used fluorescein ET-1 and 125I-ET-1 to provide evidence for ET-1 receptors in cardiac transverse tubules (T-tubules). Moreover, the ETA receptor and downstream effector phospholipase C-{beta}1 were co-localized within T-tubules using standard immunofluorescence techniques, and protein kinase C (PKC)-{epsilon}-enhanced green fluorescent protein bound reversibly to T-tubules upon activation. Localized photorelease of diacylglycerol further suggested compartmentation of PKC signaling, with release at the myocyte "surface" mimicking the negative inotropic effects of bath-applied PKC activators and "deep" release mimicking the positive inotropic effect of ET-1. The functional significance of T-tubular ET-1 receptors was further tested by rendering the T-tubule lumen inaccessible to bath-applied ET-1. Such "detubulated" cardiac myocytes showed no positive inotropic response to 20 nM ET-1, despite retaining both a nearly normal twitch response to field stimulation and a robust positive inotropic response to 20 nM isoproterenol. We propose that ET-1 enhances myocyte contractility by activating ETA receptor-phospholipase C-{beta}1-PKC-{epsilon} signaling complexes preferentially localized in cardiac T-tubules. Compartmentation of ET-1 signaling complexes may explain the discordant effects of ET-1 versus bath applied PKC activators and may contribute to both the specificity and diversity of the cardiac actions of ET-1.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Signaling networks involving protein kinase C (PKC)1 have become a major focus in investigations of the onset and treatment of heart disease. PKC is normally activated by the lipid messenger diacylglycerol produced in response to neurohumoral stress factors such as angiotensin II, opiates, and endothelin-1 (ET-1). ET-1 profoundly influences many aspects of myocardial function, including ventricular contractility, intracellular Ca2+ handling, myocyte survival, and myocyte growth (1, 2). These effects are mediated at least in part by G-protein-coupled ETA receptors located on ventricular myocytes, which mobilize diacylglycerol and activate the {epsilon}-isoform of PKC (3–8). Studies of transgenic mice with altered PKC-{epsilon} levels have also linked this signaling pathway with contractile regulation, cardioprotection during ischemia, and the onset of hypertrophy (9, 10).

L-type Ca2+ channels represent a key target of PKC in cardiac myocytes (6, 11, 12). In addition to its well established role in excitation-contraction coupling, the L-type channel may also play a central role in regulating myocyte survival and growth. L-type Ca2+ channels are preferentially localized in the periodic deep membrane invaginations of ventricular myocytes known as transverse tubules (T-tubules) (13). This specialized membrane compartment promotes functionally relevant interactions between cardiac sarcolemma containing L-type Ca2+ channels and the sarcoplasmic reticulum containing ryanodine receptors (14, 15). This membrane architecture develops as ventricular myocytes mature in parallel with development of the electrical and regulatory properties of adult myocytes (16). Moreover, T-tubules can degenerate in cardiac disease, resulting in compromised Ca2+ mobilization, particularly deep within the myocyte cross-section (17). Cardiac T-tubules display a unique composition of membrane lipids and proteins (e.g. high in phosphoinositides (18), certain ion channels (19–22), and cytoskeletal proteins (23)), but many proteins such as dystrophin (23), ankryin B (24), sodium/calcium exchangers (25), and caveolin-3 (26) are more evenly distributed among surface sarcolemma and T-tubules.

ET-1 stimulates contractility and L-type Ca2+ channel function in cardiac myocytes in a PKC-dependent manner (5, 6). Paradoxically, other activators of PKC, notably phorbol esters, inhibit these same processes (5, 27). To better understand this dichotomy, we employed a combination of techniques including photorelease of a caged compound, fluorescence imaging, and functional analyses of cardiac myocyte contractility to provide support for the hypothesis that PKC can stimulate or inhibit cardiac function depending upon where PKC is activated. Moreover, we present evidence that functional ETA receptors and the associated signaling proteins PLC-{beta}1 and PKC-{epsilon} are preferentially co-localized within cardiac T-tubules, where they mediate a positive inotropic response. We further suggest that PKC signaling mechanisms localized to T-tubules may not be readily mimicked by bath application of cell-permeable PKC activators such as phorbol esters. Some of this work has been presented in preliminary form (28).


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Materials—[125I]ET-1 and ET-1 were from Bachem (King of Prussia, PA), collagenase was from Worthington, ETA receptor mouse polyclonal antibody was from Transduction Laboratories (Lexington, KY), and PLC-{beta}1 rabbit polyclonal antibody was from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). Wheat germ agglutinin (WGA) Alexa-660 conjugate and Alexa-488 signal amplification kit for fluorescein probes were obtained from Molecular Probes, Inc. (Eugene, OR). Unless otherwise noted, all other reagents were from Sigma.

Rat Cardiac Myocyte Preparation—All experiments were performed on freshly isolated ventricular myocytes from adult male Sprague-Dawley rats prepared as described previously (5). Myocytes were maintained in 1 mM Ca2+ Ringer's solution (125 mM NaCl, 5 mM KCl, 2 mM NaH2PO4, 5 mM sodium pyruvate, 1.2 mM MgSO4, 11 mM glucose, 1 mM CaCl2, 25 mM HEPES, pH 7.4) at 21–23 °C for up to 6 h after isolation. Twitch measurements were performed as described (5). For immunofluorescence imaging, myocytes were permeabilized with 100 µg/ml saponin for 5 min at 20 °C in relaxing solution of the following composition: 100 mM KCl, 1 mM MgCl2, 2 mM EGTA, 4.5 mM ATP, 10 mM imidazole, pH 7.0. Cells were washed and resuspended in relaxing solution containing 2% bovine serum albumin to reduce nonspecific binding of applied probes.

Caged Dioctanoylglycerol Synthesis—The bromohydroxymethyl coumarin (BHC) chromophore was synthesized and purified as described (29). The BHC chromophore was coupled through its aliphatic hydroxyl group to dioctanoylglycerol (diC8) to give BHC-caged diC8 (structure in Fig. 1B). A detailed synthesis and characterization will be presented elsewhere. Briefly, to 2 ml of dry acetonitrile was added 90 µmol of BHC, 100 µmol of 4-nitrophenyl chloroformate and 180 µmol of 4-dimethylaminopyridine. After 7 h at room temperature under nitrogen, 100 µmol of diC8 (Avanti Polar Lipids, Alabaster, AL) and 200 µmol of 4-dimethylaminopyridine were added in 2 ml of dry chloroform. Twelve hours later, the reaction was quenched by the addition of 25 ml of 15% citric acid, extracted with chloroform, dried over MgSO4, and purified by silica gel flash chromatography in methylene chloride/ethyl acetate. Final purification was accomplished by reverse phase HPLC in acetonitrile/H2O/trifluoroacetic acid. The structure of BHC-caged diC8 was confirmed by 1H NMR (CDCl3; coumarin protons: {delta}7.8 (s, 1H), 7.0 (s, 1H), 6.4 (s, 1H), 5.2 (s, 2H); diC8 protons: {delta}6.1 (m, 1H), 4.6–4.1 (m, 4H), 2.3 (m, 2H), 1.6 (m, 6H), 1.3 (m, 16H), 0.9 (m, 6H)). Matrix-assisted laser desorption ionization-time of flight mass spectrometry (theoretical 641, found 641.4) and UV-visible spectroscopy (in ethanol: {epsilon} = 21,000 M-1 cm-1 at {lambda}max = 330 nm; in 0.1 M Tris, pH 7.4, 50% ethanol: {epsilon} = 15,000 M-1 cm-1 at {lambda}max = 380 nm). The photochemical quantum yield for BHC-caged diC8 was 0.4 by direct comparison with {alpha}-carboxyl-caged diC8 (with a quantum yield of 0.2) using a continuous wave xenon arc lamp filtered to pass 350–400-nm light followed by HPLC analysis as described (30).



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  		FIG. 1.
Twitch responses of individual rat ventricular myocytes to PKC activators. A, control (untreated) and 10 min after perfusion with 10 nM ET-1 or 20 µM diC8. ET-1 caused a 90% increase, whereas diC8 caused a 29% decrease in twitch amplitude. B, structure of BHC-caged diC8, and two-photon image of a myocyte containing 200 µM BHC-caged diC8. C, representative myocyte twitches in control and 10 min after two-photon photorelease of diC8 in deep (left) versus surface (right) optical sections. Deep photolysis caused a 67% increase, whereas surface photolysis caused a 30% decrease in twitch amplitude. The arrowheads indicate diastolic and systolic cell length. The far right panel shows recovery after a complete loss of twitch amplitude by increasing stimulation voltage by 50%. Similar results were obtained in three separate experiments. Recordings were made in 1 mM Ca2+ Ringer's solution, 22 °C, 0.4-Hz field stimulation.

 
Peptide Synthesis and Fluorescence Labeling—ET-1 was synthesized on an ABI 432A automated peptide synthesizer using Fmoc (N-(9-fluorenyl)methoxycarbonyl) chemistry and a Wang resin. The crude peptide was purified by reverse phase HPLC, folded, and oxidized as described (31). Quantities were determined by mass and by UV absorbance using {epsilon} = 6,000 M-1 cm-1 at 270 nm. Synthetic ET-1 was indistinguishable from commercial ET-1 by matrix-assisted laser desorption ionization-time of flight mass spectrometry (theoretical 2492, found 2492) and by its ability to compete with 125I-ET-1 for receptor sites on rat or dog ventricular membranes. Amine-directed N-hydroxysuccimide fluorophores were coupled to ET-1 via lysine 9 in 50% dimethyl formamide and aqueous 0.1 M sodium carbonate, pH 9.3, as described (31). Fluorescein ET-1 contained an {epsilon}-aminocaproate linker between lysine 9 and fluorescein, which facilitated retention of biological activity. Fluorescein ET-1 displayed an IC50 of 2.4 ± 0.4 nM in the 125I-ET-1 competition assay and at 10 nM gave a normal positive inotropic response in the myocyte twitch assay.

Two-photon Microscopy—Myocytes were imaged with a Zeiss inverted microscope equipped with a x40 oil immersion 1.4 numerical aperture lens and a pulsed titanium-sapphire laser (32). The laser was tuned to 780 nm for experiments involving BHC-caged diC8 and to 960 nm for experiments involving fluorescein ET-1. Myocytes were loaded with 200 µM BHC-caged diC8 in Ringer's solution containing 0.1% Me2SO and then perfused to remove Me2SO. Imaging was carried out by raster scanning the laser beam under the control of Bio-Rad software at a laser power of 3–10 milliwatts. For photolysis of BHC-caged diC8, the laser power was increased to 20–25 milliwatts, and the selected cell dimensions were scanned 10–30 times. Twitch amplitudes initiated by standard electrical field stimulation (40 V, 0.4 Hz) were monitored before and after photolysis by a 30-ms line scan of the myocyte edge.

The amount of diC8 generated during two-photon photolysis was estimated to be in the range of 1–5 fmol/cell. This is based on earlier one-photon photolysis studies of {alpha}-carboxyl-caged diC8 in which the effective threshold dose (i.e. minimum required for an effect on twitches) of photoreleased diC8 was measured to be 2–3 fmol/cell or 1 diC8/300 cellular phospholipids (30). BHC-caged diC8 possesses a larger molar absorptivity and a 2-fold higher photochemical quantum yield than {alpha}-carboxyl-caged diC8, so it was used at a 4-fold lower initial concentration. Using a standard xenon arc lamp for photolysis, the twitch responses to 800 µM {alpha}-carboxyl-caged diC8 and 200 µM BHC-caged diC8 were similar, including a threshold at a similar exposure time. The two-photon photolysis experiments described here showed the same threshold phenomenon (i.e. no response if the scan time or laser power were reduced by 3-fold or more), consistent with photorelease of diC8 in the 1–5-fmol range.

Western Blotting—Rat myocyte extracts were prepared by mixing freshly isolated myocytes for 30 min on ice with cold radioimmune precipitation buffer of the following composition: 9.1 mM Na2HPO4, 1.7 mM NaH2PO4, 150 mM NaCl, 1 mM sodium orthovanadate, 1% Igepal Ca-630, 0.5% sodium deoxycholate, 0.1% SDS, 10 µg/ml phenylmethylsulfonyl fluoride, and 30 µl/ml aprotinin. Myocytes were passed through a 21-gauge needle, incubated 30 min on ice in radioimmune precipitation buffer with freshly added 10 µg/ml phenylmethylsulfonyl fluoride, and centrifuged at 10,000 x g for 10 min at 4 °C. The supernatant was electrophoresed on 12% SDS-polyacrylamide gels and then transferred to nitrocellulose membranes. Nonspecific sites were blocked for 1 h at room temperature with Blotto (150 mM NaCl, 20 mM Tris, pH 7.4, 1% (v/v) Tween 20, 5% powdered milk), and Western analysis was carried out with enhanced chemiluminescence detection as described (33).

Immunofluorescence Labeling—Myocytes skinned in 100 µg/ml saponin were washed extensively and then incubated with primary antibody (diluted 1:50 or 1:100) overnight at 4 °C. Following extensive washing with 2% bovine serum albumin in relaxing solution, myocytes were incubated with Alexa-conjugated secondary antibodies for 1 h at room temperature. Highly cross-adsorbed Alexa-488 goat anti-rabbit IgG and Alexa-568 goat anti-mouse IgG were used at 1:100 dilutions. After extensive washing, images were acquired with a Bio-Rad MRC 1024 laser-scanning confocal microscope equipped with an argon/krypton laser controlled by 24-bit LaserSharp software. For double labeling, 50 µg/ml WGA Alexa-660 conjugate was added together with primary antibody. For fluorescein-ET-1 binding, rat cardiac cells were incubated with 10 nM fluorescein-ET-1 overnight at 4 °C. After extensive washing, an Alexa 488-based signal amplification kit for fluorescein (Molecular Probes) was used according to the manufacturer's instructions.

Canine Left Ventricular Membrane Fractionation—Sarcolemmal, T-tubular, and dyadic membrane fractions were prepared as previously described (34). Briefly, 30–35 g of canine left ventricle was homogenized, and a total membrane fraction (free of myofilaments and soluble proteins) was obtained by differential centrifugation. This crude membrane fraction was loaded onto discontinuous sucrose density gradients composed of 21, 31, 40, and 55% (w/v) sucrose and centrifuged for 2 h at 141,000 x g. Membranes were recovered from three distinct interfaces (Fraction I, 10/21% sucrose; Fraction II, 21/31% sucrose; Fraction III, 31/40% sucrose), diluted, pelleted at 141,000 x g, and stored at -80 °C.

Binding Studies—Radioligand-binding assays were performed in triplicate. 50 µg of membranes were incubated with 0.025–1 nM [125I]ET-1 for 1 h at 32 °C in 0.2 ml containing 10 mM MgCl2, 2 mM EGTA, 50 mM Tris, pH 7.3, 0.0015% aprotinin, 2% bovine serum albumin. Preincubation in 1 µM unlabeled ET-1 was used to assess nonspecific [125I]ET-1 binding. Membrane-bound ligand was separated from free ligand by rapid vacuum filtration over GF/C filters on a 24-well harvester (Brandel, Gaithersburg, MD) with three washes of ice-cold incubation buffer. Data were fit to a single-site binding model by nonlinear regression analysis using SigmaPlot software.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Inotropic Responses to ET-1 and Diacylglycerol—Twitch responses after treatment of rat ventricular myocytes with ET-1 and diacylglycerol are illustrated in Fig. 1. Perfusion of electrically paced myocytes with 10 nM ET-1 increased twitch amplitude by 60–90% over 10 min (Fig. 1A). Perfusion of 20 µM diC8 did not mimic this positive inotropic response but instead showed two distinct types of inhibitory response: a 30–40% decrease in twitch amplitude and an abrupt loss of excitability (Fig. 1A). A BHC-caged diC8 compound (structure in Fig. 1B) was then used to photorelease diC8 in different subdomains of myocytes. First, two-photon microscopy showed that the intrinsically fluorescent BHC-caged diC8 compound was incorporated into virtually all intracellular membranes of ventricular myocytes (Fig. 1B). Photorelease of BHC-caged diC8 was then accomplished in selected optical sections by raster scanning the cell with the two-photon laser at a 3-fold higher laser power. The x-y dimensions of the scan were matched to the cell dimensions by the laser scanning software. The z dimension was dictated by the point spread function of the titanium sapphire laser (measured to be <=2 µm through the 40 x 1.4 numerical aperture objective) (32).

"Deep photolysis" was accomplished by scanning an optical section at mid-cell depth (typically a plane with visible nuclei; estimated distance from cell surface >= 4 µm), giving rise to a positive inotropic response (Fig. 1C). This response developed within 10 min after uncaging. In contrast, "surface photolysis" was performed by centering the upper membrane surface (the surface not in contact with the glass floor) within the scanned optical plane (estimated distance from cell surface <= 1 µm). In this case, twitch amplitudes were inhibited within 5 min of uncaging (Fig. 1C), sometimes changing quite abruptly from twitch to twitch as observed with perfusion of diC8. This sudden loss of twitch amplitude induced by diC8 was not due to nonspecific cell toxicity, because it could be reversed by washing or by increasing stimulation voltage (Fig. 1C). These data show that generation of diC8 at the surface sarcolemma produced a fundamentally different effect (twitch inhibition or negative inotropy) than did generation of diC8 within deeper membrane compartments (positive inotropy). Importantly, only the deep photolysis protocol mimicked the response to 10 nM ET-1.

Localization of ET-1 Receptors in Rat Myocytes—To explore localization of receptors capable of binding ET-1, rat myocytes were stained with biologically active fluorescein-ET-1 (Fig. 2A). An anti-fluorescein immunological amplification kit (see "Experimental Procedures") was employed to obtain this pattern, reflecting a relatively low ET-1 receptor density in these cells. Fluorescein-ET-1 binding to receptors gave a striated pattern, which was greatly reduced by preincubation of myocytes with unlabeled ET-1 (Fig. 2B) or with the ETA receptor-selective antagonist BQ123 (not shown). The pronounced striation pattern is reminiscent of T-tubule staining and provides support for a preferential localization of active ET-1 receptors (capable of binding ET-1) in T-tubules compared with non-T-tubular surface sarcolemma.



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  		FIG. 2.
Localization of ETA receptors in T-tubules of rat myocytes. A, two-photon image of a myocyte treated with 10 nM fluorescein-ET-1 and then with Alexa 488 anti-fluorescein antibody. Bar, 10 µm. B, two-photon image of a myocyte pretreated with 1 µM unlabeled ET-1 and then with 10 nM fluorescein-ET-1 and Alexa 488 anti-fluorescein antibody. C, Western blot demonstrating that the anti-ETA antibody recognizes a single protein of Mr = 50,000. Left lane, rat testis extract. Right lane, rat myocyte extract. D, confocal image of a myocyte immunostained with a polyclonal anti-ETA receptor antibody. Scale bar, 10 µm. E, same cell stained with wheat germ agglutinin Alexa 660. F, merged images of D and E with yellow/orange indicating regions of overlap and green indicating WGA alone (arrow).

 
A more traditional immunofluorescence approach was then used to examine the localization of the ETA receptor subtype on rat ventricular myocytes using confocal microscopy. Fig. 2C shows that the commercial polyclonal antibody used for receptor localization recognized a single band on a Western blot of rat myoycte extracts with the expected molecular weight for ETA receptor (Mr 50,000). Fig. 2D illustrates the confocal fluorescence pattern observed in a rat myocyte stained with this anti-ETA receptor antibody. Like the fluorescein ET-1 probe, the ETA receptor antibody gave a striated pattern. An epitope peptide for this antibody was not available because the antibody was raised against a 120-amino acid domain on the receptor. However, no staining was observed in anti-ETA receptor antibody experiments with secondary antibodies alone (not shown). Fluorescent WGA, a marker of glycoproteins on the extracellular membrane leaflet, strongly stained surface sarcolemma and T-tubules (Fig. 2E). Co-staining of myocytes with anti-ETA and WGA revealed similar striated patterns with overlap (and hence co-localization) predominantly in the T-tubules (Fig. 2F).

125I-ET-1 Binding to Cardiac Receptors—A quantitative ET-1 binding assay was employed to determine the density and affinity of ET-1 receptors in cardiac myocytes and cardiac membrane extracts. The assay was used to characterize synthetic fluorescein ET-1 and commercial ET-1 receptor antagonists used in our experiments and afforded another opportunity to examine the subcellular localization of ET-1 binding sites.

Receptors on the surface of cardiac myocytes were characterized by their EC50 values in a "competition assay," where unlabeled ligands competed with 125I-ET-1 binding. In isolated rat ventricular myocytes, we found an EC50 of 170 ± 11 pM for unlabeled ET-1 (commercial and synthetic ET-1 were similar), an EC50 of 2.4 ± 0.4 nM for fluorescein ET-1, and for receptor antagonists BQ123 (ETA-selective) and BQ788 (ETB-selective) EC50 values of 7 and 800 nM, respectively. Nonlinear least squares analysis of direct 125I-ET-1 binding in rat ventricular myocytes revealed an apparent Kd of 610 ± 174 pM and a Bmax of 2 ± 1 x 105 receptors/myocyte.

For localization of 125I-ET-1 binding sites by subcellular fractionation, dog heart homogenates were used instead of rat heart homogenates to obtain sufficient membrane material for fractionation. Dog heart appeared to be a reasonable surrogate, since dog ventricular myocytes displayed similar 125I-ET-1 binding properties to rat ventricular myocytes (EC50 = 180 ± 50 pM; 105 ET-1 receptors/myocyte). Dog heart membranes were obtained by homogenization of whole hearts, followed by sucrose density centrifugation to separate membranes into three fractions by buoyant density (33). Fraction I at the 21/31% sucrose interface was enriched in surface sarcolemma as indicated by a high oubain-sensitive Na+/K+-ATPase activity (Fig. 3A). Fraction II at the 31/40% sucrose interface was enriched in T-tubules as indicated by low oubain-sensitive Na+/K+-ATPase activity and a high density of dihydropyridine binding sites. Fraction III at the 40/55% sucrose interface contained dyads and sarcoplasmic reticulum and was relatively low both in Na+/K+-ATPase activity and dihydropyridine binding. As shown in Fig. 3A, specific 125I-ET-1 binding was detected in all three membrane fractions, but ET-1 receptor binding was highest in Fraction II enriched in T-tubules. Nonlinear least squares analysis of Fraction II binding over a range of 125I-ET-1 concentrations revealed a single class of sites with a Kd of 820 ± 190 pM and Bmax of 479 ± 53 fmol/mg (Fig. 3B). Competition over a range of unlabeled ET-1 concentrations gave an EC50 value of 178 ± 69 pM, and the majority of 125IET-1 binding was displaced by the ETA receptor antagonist BQ123 with an EC50 of 4 nM. Overall, the binding properties exhibited by Fraction II were consistent with literature values for the ETA receptor subtype (35, 36). Moreover, the Kd and EC50 values for ET-1 binding were similar in Fractions I–III, indicating that the same receptor subtype was present in all three fractions (Fig. 3B), but receptor density was highest in the T-tubule fraction. Thus, subcellular fractionation of myocytes was consistent with a preferential localization of ET receptors in cardiac T-tubules, a membrane compartment high in dihydropyridine binding sites (L-type Ca2+ channels) and low in Na+/K+-ATPase activity (Fig. 3).



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  		FIG. 3.
125I-ET-1 binding in dog heart membranes. A, three subcellular fractions (I–III) obtained from sucrose density gradients were analyzed for specific 125I-ET-1 binding, 3H-PN-200–110 binding (an L-type Ca2+ channel ligand used as a marker for T-tubules), and oubain-sensitive Na+/K+-ATPase activity (a marker for surface sarcolemma). Data from five dog hearts are presented as mean ± S.E. after normalizing to the fraction with the highest value. Hg, total homogenate. B, 125I-ET-1 binding to each fraction over a range of concentrations. Fraction II displayed the most 125I-ET-1 binding sites at all concentrations, and EC50 values were similar in each fraction (I, 1,200 ± 280 pM; II, 800 ± 95 pM; III, 890 ± 420 pM).

 
Localization of PLC-{beta}1 and PKC-{epsilon}—ETA receptors are classical G{alpha}q-coupled receptors capable of activating PLC-{beta}1 and PKC in a variety of cell types (1–8). These downstream effectors of ETA receptors were examined to establish whether they too are present in cardiac T-tubules. First, rat myocytes were stained with a polyclonal antibody to PLC-{beta}1, and confocal images showed clear staining of both T-tubules and surface sarcolemma (Fig. 4A). Staining was eliminated upon preincubation of primary antibody with its epitope peptide (Fig. 4B). To confirm antibody specificity, Western blots were carried out on rat myocyte extracts and probed with the same anti-PLC-{beta}1 antibody. Two bands were recognized by the antibody, and both were abolished by epitope peptide pretreatment (Fig. 4C). The upper band had the appropriate molecular weight for PLC-{beta}1, and the lower band was a likely proteolytic fragment. Importantly, this finding places a subpopulation of PLC-{beta}1 molecules, a recognized downstream effector of ETA receptors, also within cardiac T-tubules.



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  		FIG. 4.
Localization of PLC-{beta}1 in T-tubule and surface sarcolemma. A, confocal image of a rat myocyte decorated with a polyclonal anti-PLC {beta}1 antibody. Scale bar, 10 µm. B, preincubation of anti-PLC-{beta}1 with its epitope peptide blocked all staining. C, Western blot of a myocyte extract probed with the same anti-PLC-{beta}1 antibody. Bands at 150 and 100 kDa (lanes 1 and 2) were eliminated by pretreatment with the epitope peptide (lane 3).

 
PKC is known to redistribute to distinct cellular compartments in the time domain of seconds to minutes (7, 8, 33, 37). Therefore, a green fluorescent protein fusion construct of PKC-{epsilon} (PKC-{epsilon}-GFP) described previously (37) was used to examine the localization of this kinase under different conditions. Fig. 5A illustrates T-tubular staining by PKC-{epsilon}-GFP under the same conditions used for immunolocalization of PLC-{beta}1. The striated pattern for PKC-{epsilon}-GFP binding required preactivation of the kinase with a precise stoichiometric amount of PMA prior to incubation with saponin-skinned cardiac cells. T-tubular binding by PKC-{epsilon}-GFP was readily reversible upon washing and was partially blocked by preincubation of myocytes with unlabeled PKC-{epsilon}. Overall, these observations are consistent with the concept that PKC-{epsilon} translocates dynamically from the cytosol to the T-tubules in response to diacylglycerol production in the T-tubule membrane (37). Local formation of diacylglycerol would in turn be under the control of the ETA receptor-PLC-{beta}1 complex.



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  		FIG. 5.
Localization of PKC-{epsilon}-GFP in T-tubules and surface sarcolemma. A, confocal image of a saponin-permeabilized myocyte (not treated with PMA) and decorated with PKC-{epsilon}-GFP (preactivated with PMA). Bar, 10 µm. B, confocal image of a saponin-permeabilized myocyte first treated with 100 nM PMA and subsequently stained with PKC-{epsilon}-GFP (not preactivated with PMA). C, representative rat myocyte twitch responses before and 10 min after 100 nM PMA was applied in the bath. PMA caused a 29% decrease in twitch amplitude. D, summary of the effects of 100 nM PMA on twitch amplitude in seven myocytes.

 
Effects of PMA on Myocyte Twitches and PKC-{epsilon}-GFP Localization—Fig. 5, C and D, illustrates the functional effects of the widely used cell-permeant PKC activator PMA. Rat myocytes were paced by field stimulation at 0.4 Hz and then perfused with 100 nM PMA. A negative inotropic response characterized by a 30–40% decrease in twitch amplitude developed within 5 min under these conditions. This is similar to the response to bath-applied diC8 but quite different from the response to ET-1 (see Fig. 1A). To determine how such bath application of PMA might influence PKC redistribution, saponin-skinned myocytes were first incubated with 100 nM PMA and then with PKC-{epsilon}-GFP. Remarkably, PKC-{epsilon}-GFP accumulated primarily in the surface sarcolemma under these conditions (Fig. 5B). Note that the difference between Fig. 5, A and B, is only the order of addition of PMA and PKC-{epsilon}-GFP to the myocytes. In Fig. 5A, PMA and PKC-{epsilon}-GFP were mixed together first and then applied to saponin-skinned myocytes. In Fig. 5B, PMA was added to the saponin-skinned myocytes first, and then PKC-{epsilon}-GFP (not preactivated with PMA) was added 5 min later. This seemingly minor difference in experimental protocol resulted in T-tubular staining in one case and surface sarcolemmal accumulation in another (Figs. 5, A versus B). Surface sarcolemmal staining of PKC-{epsilon}-GFP persisted for >=20 min, at which time faint staining of T-tubules near the junction with the surface membrane became apparent. These effects of PMA on myocyte twitches and PKC-{epsilon}-GFP localization are consistent with the suggestion that PKC activation at myocyte surface membranes is associated with inhibitory or negative inotropic effects.

Effects of ET-1 on Detubulated Myocytes—Rat ventricular myocytes were detubulated by a recently developed osmotic shock procedure that effectively "seals off" cardiac T-tubules (21). The technique renders the T-tubule lumen inaccessible and permits a test of the hypothesis that ET-1 signaling depends upon access to the T-tubule lumen. Fig. 6A shows that T-tubules were no longer accessible to WGA or to fluorescent 40 nm microspheres (Molecular Probes) after the osmotic treatment. With regard to control twitch responses (no agonist), average twitch amplitudes and twitch durations were not significantly different in myocytes after detubulation compared with untreated myocytes. Twitch amplitudes normalized to resting cell length were 8.4 ± 1.1% (n = 8) intact myocytes and 7.1 ± 0.7% (n = 7) detubulated myocytes. Times to 90% relaxation were 363 ± 113 ms (n = 8) in intact myocytes and 426 ± 146 ms (n = 7) in detubulated myocytes. p values in unpaired t tests comparing intact versus detubulated myocytes were not significant for any control twitch parameter.



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  		FIG. 6.
Response of detubulated rat myocytes to ET-1 and Iso. A, confocal images of intact and detubulated (De-tub) myocytes stained with WGA (top) and 40-nm fluorescent latex microspheres (bottom). B, representative twitch traces in a single detubulated myocyte before and 10 min after the addition of 20 nM ET-1. The same myocyte was exposed to 20 nM Iso (after washing out ET-1). ET-1 caused no change, whereas Iso increased twitch amplitude by 72%. C, representative twitch traces in a single intact (not detubulated) myocyte before and 10 min after the addition of 20 nM ET-1. ET-1 caused a 60% increase in twitch amplitude. D, summary of twitch amplitudes in intact and detubulated myocytes. Data are presented as mean ± S.E. from 3–5 myocytes after normalization to a control (no agonist) twitch. *, p < 0.5 versus control.

 
Illustrated in Fig. 6B are the effects of detubulation on the myocyte twitch response to 20 nM ET-1. Despite the relatively normal twitch response following detubulation, the positive inotropic response to 20 nM ET-1 was no longer observed. At least one other signaling and regulatory process appeared to be operating, however, since treatment with 20 nM Iso to activate {beta}-adrenergic receptors initiated a robust positive inotropic response (Fig. 6B). Myocytes not subjected to detubulation gave a normal positive inotropic response to 20 nM ET-1 with a 66 ± 17% (n = 4) increase in twitch amplitude.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The results of this study demonstrate preferential localization of ETA receptors capable of binding ET-1 within cardiac T-tubules, where they co-localized with two probable downstream effectors: PLC-{beta}1 and PKC-{epsilon}. Such a "localized cascade" (illustrated in Fig. 7) is fully compatible with the properties of classical G{alpha}q/PLC-{beta}1-coupled receptors (38, 39) and places the machinery for regulated formation of diacylglycerol in the vicinity of L-type Ca2+ channels. Cardiac L-type channels are located in T-tubule membranes, where they are closely associated with ryanodine receptors of the sarcoplasmic reticulum (12–16). Ca2+ ions flowing through the L-type channel during action potentials provide the "trigger" for a much larger mobilization of Ca2+ from the sarcoplasmic reticulum. Up- or down-regulation of trigger Ca2+ leads directly to up- or down-regulated intracellular Ca2+ transients and, in turn, changes in contraction strength (12). ET-1 is among a number of diacylglycerol-mobilizing agonists that modulate ventricular contractility in part via regulation of L-type Ca2+ channel activity (12), but until now the subcellular organization of ET-1 receptors and downstream effectors in relation to L-type Ca2+ channels was unknown.



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  		FIG. 7.
Model of PKC activation in cardiac T-tubules and surface sarcolemma. "T-tubule signaling" involves a local receptor-effector signaling complex. A key feature of this arrangement is elevation of diacylglycerol (DAG) in the T-tubule and recruitment of PKC to the T-tubule in response to local agonist binding. "Surface signaling" initiated by bath-applied diC8 or PMA or by surface-localized PLC-{beta}1 occurs outside of the T-tubule compartment. Whether surface PLC-{beta}1 is associated with G-protein coupled receptors (R) is currently unknown (?).

 
There is growing evidence that complexes specialized for cyclic AMP/protein kinase A signaling are concentrated near cardiac T-tubules (reviewed in Ref. 38). Many related signaling proteins, including {beta}-receptors, G{alpha}s, adenylate cyclase, phosphodiesterase, protein kinase A, and protein phosphatase 2A, appear to be anchored at the T-tuble on scaffolding proteins (40), which provide for highly localized cyclic AMP production (41). Evidence that other receptors and associated signaling cascades reside in cardiac T-tubules is more sparse. One recent report identified an ErB2 tyrosine kinase receptor in cardiac T-tubules that is targeted in an autoimmune disease (42). Another recent report used antibodies to multiple epitopes on endothelin A and B receptors to demonstrate their presence in the nuclei of rat ventricular myocytes (43). Whereas the focus of this work was to characterize functional nuclear endothelin receptors, immunostaining of receptors in surface sarcolemma including T-tubules was apparent (43). Our data provide evidence that endothelin receptors in the T-tubules are capable of binding fluorescein ET-1 and that they mediate the positive inotropic response to ET-1. Co-localization of endothelin receptors with known effector proteins suggests local signaling in the immediate vicinity of the T-tubule. Signaling cascades featuring long range diffusion of second messengers (e.g. diacylglycerols, cis-unsaturated fatty acids) or translocation of signaling proteins (e.g. G-protein subunits, phospholipase A2, or phospholipase C-{gamma}) from the surface sarcolemma to the T-tubules are not needed to account for the data.

The biological relevance of preferential T-tubular localization of ETA receptor/diacylglycyerol/PKC signaling is also suggested by our results. Localized photorelease of diacylglycerol demonstrated that this lipid messenger promoted fundamentally different physiological responses at deep sites within the myocyte (>=4 µm from cell surface) compared with surface sites (<=1 µm from cell surface). We cannot conclude with certainty that deep photolysis only activates T-tubular PKC and that surface photolysis only activates (non-T-tubular) plasma membrane PKC, but only the deep photolysis protocol mimicked the slowly developing positive inotropic response seen with 10 nM ET-1. Consistent with this correlation between location and function, bath application of the potent PKC activator PMA inhibited myocyte twitches and promoted PKC-{epsilon}-GFP accumulation preferentially in the surface plasma membrane. Finally, myocyte detubulation experiments served as a direct test of the hypothesis that ET-1 is a positive inotrope in rat myocytes because it activates T-tubular PLC-{beta}1 and generates diacylglycerol there. ET-1 had no detectable physiological effects on detubulated myocytes (no positive inotropy and no negative inotropy), consistent with the vast majority of ET-1 receptors and associated signaling complexes being inaccessible in detubulated myocytes. Interestingly, detubulated myocytes retained their positive inotropic response to the {beta}-adrenergic agonist isoproterenol despite considerable evidence that {beta}-receptor signaling complexes are also present within the T-tubule (38–41). However, it is likely that {beta}-receptors are also expressed on surface sarcolemma outside of T-tubules (34). Establishing whether responses to isoproterenol in detubulated and intact myocytes differ in some fundamental way requires further investigation.

Overall, the data provide insight into the paradox recognized several years ago in mammalian ventricular tissues that agonists coupled to PKC signaling pathways are often positive inotropes, whereas cell-permeant PKC activators are often negative inotropes (5). PMA at the relatively high concentration of 100 nM has become widely used in studies of PKC regulation of cardiac function (2) and many other cell types. Bath application of PMA although it is cell-permeant may not give the experimenter adequate control over the subcellular location of PKC activation. PMA is a relatively hydrophobic phorbol ester that probably partitions rapidly and efficiently into the first membranes it encounters (i.e. surface rather than T-tubule or intracellular membranes). This interpretation is supported by the observation that PMA promoted preferential accumulation of PKC-{epsilon}-GFP in the surface sarcolemma under conditions similar to those in which PMA promoted negative inotropy. Surprisingly, surface accumulation of GFP-tagged PKC-{epsilon} did not redistribute to a significant extent into intracellular membranes even after 20 min or more. A similar robust and stable translocation of PKC to surface sarcolemma has been observed in cultured adult ventricular myocytes expressing GFP-tagged PKC-{delta} in response to bath-applied PMA.2 It should not be surprising then that bath application of 100 nM PMA does not adequately mimic the positive inotropic response to ET-1. In addition to PKC isoform specificity being lost (27), the use of PMA may result in PKC being recruited to inappropriate subcellular sites.

A general model of the type presented in Fig. 7 may provide insight into other apparent inconsistencies in the literature. For example, it has been recognized that agonists that stimulate G-protein-coupled receptors and phosphoinositide turnover do not always lead to the same changes in cardiac function (5, 39). For example, angiotensin II and norepinephrine promote positive inotropy (like ET-1), whereas acetylcholine, adenosine, and opiates cause negative inotropic responses. The presence of PLC-{beta}1 immunoreactivity in the surface membrane of myocytes suggests the intriguing possibility that G-protein coupled receptors not localized in T-tubules may employ phosphoinositide hydrolysis but give rise to negative inotropic responses. M2 muscarinic receptors have recently been localized predominantly to surface sarcolemma in ventricular cells (19), and these M2 receptors mediate a negative inotropic response. Significant species variability in responses to diacylglycerolmobilizing agonists has also been noted (39), possibly reflecting species differences in receptor expression and localization.

Two subtypes of the ET receptor are known, ETA receptor and ETB receptor, each derived from a separate gene (1). Molecular and pharmacological analyses reveal that functional proteins from both genes are expressed in mammalian ventricular tissue, but the density of ETA receptors is higher by 4–9-fold (35, 36, 44, 45). The polyclonal ET receptor antibody used here was specific for ETA receptors, and fluorescent ET-1 binding was blocked by the ETAR antagonist BQ-123. Moreover, the majority of the 125I-ET-1 binding in rat and dog myocytes and membrane fractions was displaced by BQ-123. Thus, the ET-1 receptors under investigation here were most likely the ETA receptor subtype. Considering the lower density of ETB receptors and the sensitivity limits of the imaging techniques employed, the subcellular distribution of ETB receptors in cardiac myocytes remains an open question. The precise mechanism of sequestering ETA receptors within cardiac T-tubules is also unclear. The cytoskeletal protein ankyrin B appears to have a critical role in maintaining the functional integrity of cardiac T-tubules (24), but its role in receptor anchoring in the heart has not been investigated. Specific interactions of receptors and signaling molecules within caveolae suggest that these microdomains may also play a role in receptor localization (38).

In conclusion, four lines of evidence indicate that location plays a major role in dictating the nature of functional responses to diacylglycerol/PKC in cardiac cells. First, confocal imaging of receptors and signaling proteins, particularly those responsible for generating and subsequently responding to diacylglycerol, placed ETA receptor signaling machinery preferentially in cardiac T-tubules. Subcellular fractionation reinforced this interpretation by demonstrating co-migration of ET receptors with L-type Ca2+ channels. Second, localized photorelease of caged diacylglycerol demonstrated that diacylglycerol location was a critical determinant of the nature of physiological response. Third, myocytes responded to ET-1 with a reproducible positive inotropic response, whereas detubulated cells did not respond to ET-1. And fourth, PMA strongly translocated PKC-{epsilon}-GFP to surface membranes and inhibited myocyte twitch contractions.

As illustrated in the model of Fig. 7, a consistent picture of PKC signaling in cardiac cells begins to emerge. G-protein-coupled ETA receptors localize preferentially to cardiac T-tubules, giving rise to a local elevation of diacylglycerol following activation of PLC-{beta}1. This leads to recruitment of PKC isoform(s) to T-tubules and ultimately gives rise to positive inotropy. A likely mechanism of this inotropic action of PKC is via stimulation of L-type Ca2+ channels, a major regulatory point in cardiac excitation-contraction coupling (12). The isoform of PKC that most likely participates in T-tubule signaling, PKC-{epsilon}, possesses unique domains that direct its docking to specific anchoring proteins such as receptors for activated C kinase, thought to reside near cardiac Z-lines/T-tubules (38, 46). We propose that activation of ETA receptors expressed on the lumen of the T-tubules of cardiac myocytes promotes a positive inotropic response by a local formation of diacylglycerol, recruitment of PKC, and phosphorylation of L-type Ca2+ channels. Activation of other G{alpha}q/PLC-{beta}1-coupled receptors may give rise to functional consequences more akin to the effect of PMA by being localized on surface sarcolemma. In light of these observations, it will be important to determine how ET-1/PKC-{epsilon} signaling and their physiological consequences differ in diseased ventricular tissue in which T-tubule morphology and function may be compromised (17, 24, 34).


   FOOTNOTES
 
* This work was supported by National Institutes of Health Grant P01 04573 (to J. W. W. and T. J. K.) and Fellowships from the American Heart Association (to V. G. R., S. L. R., and Y. Q. P.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

To whom correspondence should be addressed: Dept. of Physiology, 1300 University Ave., Madison, WI. Tel.: 608-262-6941; E-mail: jwalker{at}physiology.wisc.edu.

1 The abbreviations used are: PKC, protein kinase C; ET-1, endothelin-1; PLC, phospholipase C; diC8, dioctanoylglycerol; PMA, phorbol 13-myristate-12-acetate; WGA, wheat germ agglutinin; BHC, bromohydroxymethylcourmarin; Iso, isoproterenol; GFP, enhanced green fluorescent protein; CDCl3, deuterated chloroform; HPLC, high pressure liquid chromatography. Back

2 M. Kang and J. W. Walker, unpublished results. Back


   ACKNOWLEDGMENTS
 
We gratefully acknowledge Dr. John G. White, Kevin Eliseri, and Dr. Victoria Frolich for assistance with two-photon microscopy and Lance Rodenkirch for assistance with confocal microscopy.



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 DISCUSSION
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