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Nitric oxide (NO) and protein S-nitrosylation (SNO) play important roles in ischemic preconditioning (IPC)-induced cardioprotection. Mitochondria are key regulators of preconditioning, and most proteins showing an increase in SNO with IPC are mitochondrial. The aim of this study was to address how IPC transduces NO/SNO signaling to mitochondria in the heart. Results: In this study using Langendorff perfused mouse hearts, we found that IPC-induced cardioprotection was blocked by treatment with either N-nitro-L-arginine methyl ester (L-NAME, a constitutive NO synthase inhibitor), ascorbic acid (a reducing agent to decompose SNO), or methyl-β-cyclodextrin (MβCD, a cholesterol sequestering agent to disrupt caveolae). IPC not only activated AKT/eNOS signaling but also led to translocation of eNOS to mitochondria. MβCD treatment disrupted caveolar structure, leading to dissociation of eNOS from caveolin-3 and blockade of IPC-induced activation of the AKT/eNOS signaling pathway. A significant increase in mitochondrial SNO was found in IPC hearts compared to perfusion control, and the disruption of caveolae by MβCD treatment not only abolished IPC-induced cardioprotection, but also blocked the IPC-induced increase in SNO. Innovation: These results provide mechanistic insight into how caveolae/eNOS/NO/SNO signaling mediates cardioprotection induced by IPC. Conclusion: Altogether these results suggest that caveolae transduce eNOS/NO/SNO cardioprotective signaling in the heart. Antioxid. Redox Signal. 16, 45–56.
Ischemic preconditioning (IPC) is an adaptive phenomenon whereby brief episodes of myocardial ischemia and reperfusion render the heart resistant to subsequent prolonged ischemic injury (23). IPC initiates a number of signaling pathways at the plasma membrane, which are transduced to the mitochondria (9, 22). Among them, nitric oxide (NO) is important in IPC-induced cardioprotection (3, 14). In addition to activating cyclic guanosine monophosphate (cGMP)/protein kinase G (PKG)-dependent signaling pathways, NO can directly modify protein sulfhydryl residues through protein S-nitrosylation (SNO), which has emerged as an important post-translational protein modification in cardiovascular signaling (17, 29) and cardioprotection (18, 34). In recent studies, we have reported that IPC results in increased SNO of a number of proteins involved in cardioprotection, and most of these SNO proteins are mitochondrial proteins (15, 33).
It has been reported that blocking the internalization of signaling molecules associated with G-protein-coupled receptors inhibits cardioprotection afforded by IPC (37). Consistent with this concept, Quinlan et al. have found that preconditioning induces the assembly of a caveolar signaling platform that migrates to mitochondria to induce mitochondrial ATP-dependent potassium channel opening (27). Caveolae are flask-like invaginations of the plasma membrane enriched with cholesterol, sphingolipids, and the marker protein, caveolin. Caveolae are also enriched in other signaling molecules and thus play an important role in signaling (24, 26). The caveolin gene family consists of three isoforms: caveolin-1, -2, and -3. The first two caveolin isoforms are co-expressed in a variety of cell types, while caveolin-3 is a muscle-specific isoform (32, 36). In cardiomyocytes, caveolin-3 acts as a scaffolding protein to concentrate and organize molecules (11, 26) such as endothelial NO synthase (eNOS) within the caveolae to mediate signal transduction (5, 6). Recent studies suggested that caveolae and caveolin-3-associated signaling play an important role in preconditioning-mediated cardioprotection (4, 10, 16, 38). For example, the gene knockout of caveolin-3 abolished IPC-induced protection (10), while the cardiac-specific overexpression of caveolin-3 led to protection that mimicked IPC (38).
Myocardial caveolae have recently been implicated in cardioprotection, but the underlying mechanistic basis has not been explored in detail. In this study, the authors provide novel data showing that caveolae transduce eNOS/NO/SNO signaling to the mitochondria, and that disruption of caveolae by MβCD treatment blocks this important cardioprotective signaling pathway. These results not only suggest that caveolae signaling complexes are crucial for IPC, but also point out a possible caveolae-mediated eNOS/NO/SNO cardioprotective signaling network between caveolae and mitochondria.
The goal of this study was to examine whether caveolae play an important role in regulating compartmentalized eNOS/NO/SNO signaling in IPC-induced cardioprotection. In the present study, methyl-β-cyclodextrin (MβCD), a cholesterol sequestering reagent, was infused into Langendorff perfused mouse hearts to disrupt caveolar structure in cardiomyocytes (4, 25). Two dimensional DyLight fluorescence difference gel electrophoresis (2D DyLight DIGE) (19, 33) was employed to determine whether there are differences in protein S-nitrosylation. Consistent with the hypothesis, we found that disruption of caveolae abolished the protection afforded by IPC and blocked IPC-mediated SNO signaling.
We showed previously that IPC leads to an increase in SNO of mitochondrial proteins (15, 33). Others have reported that loss of caveolin-3 (caveolin-3-/-) blocks protection afforded by IPC (38). Thus the aim of this study was to test the hypothesis that caveolae-dependent signaling is required for the IPC-mediated increase in SNO of mitochondrial proteins. If caveolae are required for IPC-mediated SNO of mitochondrial proteins, then disruption of caveolae with MβCD, a cholesterol sequestering agent, should block the protection provided by IPC and should also block the increase of SNO that occurs with IPC.
To first confirm that MβCD results in disruption of caveolae without a general disruption of cell structure, we used transmission electron microscopy to examine cell morphology. Transmission electron microscopy (Fig. 1A) clearly demonstrated that MβCD treatment (1mmol/L for 30min perfusion) disrupted caveolae. MβCD treatment decreased the number of caveolae per μm sarcolemma from 0.31±0.03 for perfusion control (n=4) to 0.11±0.01 for MβCD-treated hearts (n=3, p<0.01). However, MβCD treatment did not change mitochondrial morphology or the myofibrillar structure. To further confirm that MβDC lead to disruption of caveolae, we used a detergent-free Na2CO3 (pH 11) method (32) to isolate caveolae-enriched fractions. As shown in Figure 1B, the disruption of caveolae by MβCD treatment led to the disappearance of the single light-scattering buoyant caveolae-enriched fraction (fraction 5) and caused migration of caveolin-3 and other associated proteins into heavier fractions of the sucrose gradients (fractions 8–12).
Numerous studies have demonstrated that eNOS is localized in caveolae via interaction with caveolin, and this compartmentalization facilitates dynamic protein–protein interactions and signal transduction events that modify eNOS activity (5, 21, 30). Caveolin-3 has been shown to be the muscle-specific caveolin. However, our whole heart homogenate contains both myocyte and endothelial cells. To evaluate the relative proportion of eNOS bound to caveolin-1 versus caveolin-3, we performed an IP of eNOS using anti-caveolin-1/caveolin-3 vs. either anti-caveolin-1 or anti-caveolin-3 alone in total heart homogenate (Fig. 2A). IP with anti-caveolin-1/caveolin-3 will pull down both the endothelial and myocardial eNOS, whereas IP with anti-caveolin-3 should only pull down myocardial eNOS. If caveolin-1 binds both myocardial eNOS and endothelial eNOS, then anti-caveolin-1 IP should pull down higher levels of eNOS than that by anti-caveolin-3 IP, which was not observed (see Fig. 2A). The data in Figure 2A suggest a specific interaction of myocardium-derived eNOS with caveolin-3, the muscle-specific isoform of caveolin. We next wanted to determine if MβCD treatment alters the interaction of eNOS with caveolin-3 in cardiomyocytes. As shown in Figure 2B, disruption of caveolae by MβCD treatment significantly decreased the association of eNOS with caveolin-3, thus significantly attenuating the compartmentalization of eNOS signaling in the caveolae.
Perfusion of hearts with MβCD alone did not change cardiac hemodynamics (Table 1). In addition, MβCD treatment alone did not lead to changes of post-ischemic functional recovery (Table 1, Fig. 3B) or infarct size (Fig. 3C). However, MβCD treatment prior to and during IPC abolished IPC-induced cardioprotection (Table 1, Fig. 3). Thus, consistent with previous data in the literature (4), treatment of Langendorff perfused heart with MβCD to disrupt caveolae blocked IPC-induced cardioprotection. These data suggest that caveolae play an important role in mediating the cardioprotective effects of IPC (4, 10, 16, 38).
To test if caveolae are necessary for eNOS/NO/SNO signaling in IPC, we examined whether disruption of caveolae with MβCD blocks SNO signaling. Total heart homogenate fractions were subjected to 2D DyLight DIGE proteomic analysis to detect SNO in IPC±MβCD hearts. A representative 2D DyLight DIGE gel is shown in Figure 4. SNO proteins in perfusion control hearts were labeled by DyLight 488 (green), IPC hearts were labeled with DyLight 649 (red), and MβCD-treated IPC hearts were labeled with DyLight 549 (yellow). As shown in Figure 4 and Table 2, IPC significantly increased SNO signaling. MβCD treatment blocked the increase in SNO induced by IPC (overlay of DyLight 549/649), and the SNO level in MβCD-treated hearts was comparable to that in the perfusion control (overlay of DyLight 488/549). The fluorescent spots indicate SNO proteins, which were extracted and identified by mass spectrometry. Data from multiple gels (n=3) were compiled in Table 2, which shows that IPC (without MβCD) leads to an increase in SNO of proteins. The ratio of ~1:1 of (MβCD+IPC)/Perf (Table 2) in DyLight intensity (SNO content) of each protein spot suggests that the disruption of caveolae in hearts by MβCD treatment prevented IPC-induced activation of SNO signaling. These data are consistent with the hypothesis that disruption of caveolae disrupts eNOS/NO/SNO signaling.
To confirm a role for SNO in IPC, the effects of constitutive NOS inhibition with N-nitro-L-arginine methyl ester (L-NAME) or decomposing SNO with ascorbic acid were tested in the Langendorff perfused mouse heart IPC-I/R model (Fig. 5A). L-NAME (10μmol/L) treatment alone did not change pre-ischemic hemodynamics, post-ischemic recovery of LVDP, or infarct size in nonpreconditioned hearts. However, treatment with L-NAME 10min prior to and during IPC blocked IPC-induced cardioprotection (Fig. 5B). Ascorbic acid (1mmol/L) treatment did not change the pre-ischemic heart rate, but caused an ~50% decrease in pre-ischemic LVDP. We were concerned that this pre-ischemic decrease in LVDP might alter the response to ischemia; however we found that addition of ascorbic acid did not alter infarct size or recovery of function following ischemia-reperfusion in hearts in non-IPC hearts. This would suggest that the pre-ischemic decrease in LVDP is not altering infarct size. However, 5min of ascorbic acid treatment in the last cycle of IPC prior to ischemia abolished IPC-induced cardioprotection (Fig. 5C). These results suggested that NO/SNO signaling plays an essential role in IPC-induced acute cardioprotection.
We were also interested in determining if disruption of caveolae altered cardioprotective signaling that leads to activation of eNOS. Studies have shown that AKT can directly phosphorylate and activate eNOS, leading to activation of NO signaling (31, 39). Immunblot analysis using total heart homogenates (Fig. 6) showed that IPC significantly increased phosphorylation of AKT (p-AKT) at Ser 473 and phosphorylation of eNOS (p-eNOS) at Ser1177 (equivalent to Ser1176 in mouse), while treatment with MβCD blocked phosphorylation of AKT and eNOS in IPC hearts. Thus, IPC-induced AKT-eNOS cardioprotective signaling pathways are blocked by MβCD treatment.
We have demonstrated that disruption of caveolae by MβCD treatment not only abolished IPC-induced cardioprotection (Fig. 3), but also blocked eNOS signaling (Figs. 2 and and6)6) and prevented the increase in SNO induced by IPC (Fig. 4). As noted, IPC leads to an increase in SNO of a large number of mitochondrial proteins (Table 2) (15, 33). These data are consistent with the hypothesis, which has also been suggested by other recent studies (12, 20, 28), that caveolae-mediated endocytosis plays a role in regulating eNOS and delivering NO to subcellular targets (e.g., mitochondria). To further test this hypothesis, we prepared mitochondria fractions from Langendorff-perfused mouse hearts immediately after perfusion and IPC. Equal amounts of mitochondrial proteins from each heart sample were separated by 4%–12% SDS-PAGE and transferred to nitrocellulose membranes. The Ponceau S staining confirms similar protein loading (Fig. 7A). VDAC-1 was used as a mitochondrial and equal-loading marker (Fig. 7B). We saw no detectable caveolin-1 (~20kDa) in the mitochondria prepared from either perfusion control or IPC hearts. Although the mitochondria isolated from perfusion control hearts did not contain caveolin-3 (~18KDa), caveolin-3 was detected in mitochondrial samples isolated from IPC hearts. In addition, IPC significantly increased levels of eNOS in mitochondria, in parallel to the change in caveolin-3. These data support the hypothesis that IPC results in mobilization of caveolae, which transduces eNOS/NO/SNO signaling to the end effector mitochondria, leading to cardioprotection.
Both eNOS and neuronal NOS (nNOS) are constitutively expressed in cardiomyocytes but within distinct subcellular locations, i.e., eNOS is predominantly localized at caveolae in the sarcolemma (1, 7) and nNOS is mostly localized to the sarcoplasmic reticulum (1, 41). In this study, we found that perfusion with a constitutive NOS inhibitor L-NAME prior to and during IPC abolished IPC-induced protection (Fig. 5), suggesting that the activation of constitutive NOS is necessary for IPC-induced acute cardioprotection. Consistent with this finding, loss of IPC-induced cardioprotection has been found in eNOS-/- mouse hearts (35), although there are data to the contrary (8).
There are emerging data suggesting that protein S-nitrosylation plays an important role in cardioprotection (17, 18, 33, 34). Our studies have demonstrated that IPC results in an increase in SNO, and that SNO can alter protein activity and protect proteins against further oxidation (15, 33, 34). A previous study by our group reported that IPC was blocked by N-acetylcysteine, a glutathione precursor and reducing agent, suggesting a redox-sensitive mechanism is involved in the protection afforded by IPC (2). In this study, the inhibition of IPC-induced cardioprotection by ascorbic acid (Fig. 5), a reducing agent that specifically decomposes SNO, provides strong evidence showing that SNO signaling plays an important role in IPC-induced cardioprotection. Consistent with our recent studies (15, 33), the majority of proteins with an increase in SNO after IPC were mitochondrial proteins (Fig. 4 and Table 2).
In this study, we tested the hypothesis that caveolae-associated eNOS/NO is important for SNO of mitochondrial proteins in IPC hearts. In support of the hypothesis, data in the literature show that that IPC increased the formation of caveolae, and a transgenic mouse with cardiomyocyte-specific overexpression of caveolin-3 showed enhanced formation of caveolae and exhibited reduced ischemia/reperfusion (I/R) injury (38). In addition, caveolin-3-/- mice showed no protection in response to IPC (38). A recent study by Quinlan et al. (27) has shown that preconditioning the heart induces formation of signalosomes, caveolae-associated signaling platforms that interact with mitochondria to open mitochondrial ATP-dependent potassium channels (27). These data not only suggest that caveolae signaling complexes are crucial for IPC, but also point out a possible cardioprotective signaling network between caveolae and mitochondria. It has been suggested that activated eNOS could be internalized to deliver NO to subcellular targets for biological effects (12, 20, 28). In this study, we found that MβCD treatment not only abolished IPC-induced cardioprotection (Fig. 3), but also markedly reduced the association of eNOS with caveolin-3 (Fig. 2) and IPC-induced phosphorylation of eNOS (Fig. 6). Furthermore, MβCD treatment blocked the IPC-induced increase in SNO (Fig. 4, Table 2). The parallel changes in caveolar eNOS signaling and SNO in mitochondria in IPC hearts are consistent with the hypothesis that activation of eNOS and caveolae trafficking to mitochondria are important in generating mitochondrial-localized eNOS/NO/SNO signaling. The detection of eNOS/caveolin-3 in mitochondria isolated from IPC hearts (Fig. 7) provides support for this hypothesis.
As caveolae are important for many signaling pathways, the disruption of caveolae could inhibit protection by blocking any number of signaling pathways. Thus, one limitation of this study is that we only examined the role of caveolin-3/eNOS/SNO, but did not investigate other signaling pathways such as the NO/guanylyl cyclase/cGMP/PKG pathway. However, it is clear that eNOS/NO/SNO signaling is one of the signaling pathways that is blocked by disruption of caveolae. Since SNO had been previously shown to play a role in cardioprotection, it is likely that disruption of this signaling pathway contributes to loss of cardioprotection. Because of dynamic range issues, most of the proteomic methods (2D gels and mass spectrometry) are biased toward detection of high-abundance proteins (34). Thus, the prevalence of mitochondrial proteins identified as SNO proteins might be due in part to their high abundance. Our recent study using a SNO-RAC proteomic approach confirms that most of SNO proteins in IPC hearts are mitochondrial proteins (15), suggesting that mitochondria are indeed one of the major subcellular organelles targeted by SNO signaling in IPC hearts.
In summary, we found that MβCD treatment not only disrupted caveolae and association of eNOS with caveolin-3, but also blocked IPC-induced cardioprotection and the increase in SNO of mitochondrial proteins that normally occur with cardioprotection. These data are consistent with the hypothesis that caveolin-3-associated eNOS/NO trafficking between plasma membrane and mitochondria provide an important signaling pathway regulating SNO of mitochondrial proteins.
C57BL/6J male mice were obtained from Jackson Laboratories (Bar Harbor, Maine) and were between 12 and 16 weeks old at the time of experimentation. All animals were treated in accordance with National Institutes of Health guidelines and the “Guiding Principles for Research Involving Animals and Human Beings.” This study was reviewed and approved by the Institutional Animal Care and Use Committee of the National Heart Lung and Blood Institute.
Mice were anesthetized with pentobarbital and anti-coagulated with heparin. Hearts were excised quickly and placed in ice-cold Krebs-Henseleit buffer (in mmol/L: 120 NaCl, 11 D-glucose, 25 NaHCO3, 1.75 CaCl2, 4.7 KCl, 1.2 MgSO4, and 1.2 KH2PO4). The aorta was cannulated on a Langendorff apparatus and the heart was perfused in retrograde fashion with Krebs-Henseleit buffer at a constant pressure of 100cm of water at 37°C. Krebs-Henseleit buffer was oxygenated with 95% O2/5% CO2, and maintained at pH 7.4. After equilibrium perfusion or IPC (4 cycles of 5min of ischemia and 5min of reperfusion), mouse hearts were subjected to 20min of no-flow ischemia, followed by 90min of reperfusion. Drug administration is illustrated for each IPC-I/R protocol in corresponding figures. To prevent SNO breakdown, Langendorff perfusion and sample preparations were carried out in the dark.
To monitor left ventricular contractile function, a latex balloon connected to a pressure transducer was inserted into the left ventricle of a Langendorff perfused heart. The left ventricular developed pressure (LVDP) was recorded and digitized using a PowerLab system (ADInstruments, Colorado Springs, CO). We used the rate pressure product (RPP=LVDP×heart rate) as a measure of function. The post-ischemic functional recovery was expressed as percentage of preischemic RPP during the equilibrium period. For measurement of myocardial infarct size, hearts were perfused with 1% (w/v) of 2,3,5-triphenyltetrazolium chloride (TTC) after 90min of reperfusion and incubated in TTC at 37°C for 15min, followed by fixation in 10% (w/v) formaldehyde. Infarct size was expressed as the percentage of total area of cross-sectional slices through the ventricles.
Total heart homogenate was obtained by grinding a snap-frozen mouse heart into powder in liquid nitrogen, followed by homogenization with a tight-fitting glass Dounce homogenizer on ice in 1.5ml homogenate buffer containing (in mmol/L, pH7.4): 70 NaCl, 20 HEPES, 2.5 MgCl2, 1 EDTA, 0.1 neocuproine (a copper chelating agent), and 0.5% (v/v) Triton X-100 with protein protease/phosphatase inhibitors (Roche Diagnostics, Indianapolis, IN). The mixture was kept on ice for 60min with gentle vortex mixing every 15min. The sample was homogenized again on ice using a Dounce glass homogenizer. These total heart homogenates were snap frozen in liquid nitrogen and stored at −80°C.
Mitochondria were rapidly isolated by differential centrifugation according to standard procedures (40) immediately after perfusion or IPC protocol. Briefly, hearts were minced in mitochondria isolation buffer (pH 7.25, in mmol/L: 225 mannitol, 75 sucrose, 5 MOPS, 2 taurine, and 0.5 EGTA) and homogenized with a Polytron. Trypsin (0.001g/0.1g wet tissue) was added to the homogenate for 5min on ice. Digestion was stopped by adding isolation buffer containing protein protease/phosphatase inhibitors. The homogenate was centrifuged at 500 g for 5min, and the resulting supernatant was spun at 11,000 g for 5min to pellet the mitochondria. The final mitochondrial pellet was resuspended in isolation buffer with protein protease/phosphatase inhibitors. Protein content was determined using a Bradford assay.
Caveolae-enriched fractions were isolated using a protocol modified from Song et al. (32) using a discontinuous (5%/35%/45%, w/v) sucrose gradient ultracentrifugation in buffer containing (pH 6.5, in mmol/L): 25 2-(N-morpholino)ethanesulfonic acid, 150 NaCl, and 1 EDTA. After ultracentrifugation at 200,000 g for 18h at 4°C in a Beckman SW 41 swinging bucket rotor, the sucrose gradient was collected in 1ml fractions from the top to the bottom. A light-scattering band confined to the 5%–35% sucrose interface was observed and collected as the caveolae-enriched fraction (fraction 5 as shown in Fig. 1B).
Perfusion control (n=4) and MβCD-infused (n=3) hearts were examined for ultrastructural changes. Following treatment, hearts were perfused for 2min with a relaxing buffer (pH 7.2, in mmol/L: 80 potassium acetate, 10 potassium phosphate, and 5 EGTA) followed with the fixative (5% glutaraldehyde in 0.12mol/L of sodium cacodylate, pH 7.35). Hearts were dissected, samples of left ventricular myocardium were removed and put into fresh fixative for 1h at 4°C. Samples were postfixed in 1% osmium tetroxide, en bloc stained with 1% uranyl acetate, dehydrated with an ethanol series and propylene oxide, then embedded in EMbed-812 (Electron Microscopy Sciences, Hatfield PA). Thin sections were stained with uranyl acetate and lead citrate. Electron microscopy was performed on a JEM 1400 electron microscope (JEOL USA, Peabody MA) with an AMT XR-111 digital camera (Advanced Microscopy Techniques Corporation, Woburn MA). The number of caveolae/μm membrane length was counted on more than 40 cross-section images taken at 10,000X magnification for each sample.
The modified biotin switch method (13) using DyLight-maleimide sulfhydryl-reactive fluors (Pierce Biotechnology, Rockford, IL) was applied to identify SNO proteins (19, 33). Since MβCD treatment caused disruption of caveolae, the total heart homogenates rather than caveolae-enriched fractions isolated from MβCD-treated mouse hearts were used for 2D DyLight DIGE study. After the DyLight switch and labeling, equal amounts (250μg) of each sample were mixed together and subjected to 2D DyLight DIGE. Each gel was scanned at the unique excitation/emission wavelengths of each DyLight fluor using a Typhoon 9400 imager (GE Healthcare Life Sciences, Piscataway, NJ) at a resolution of 100μm. Image analysis software PG240 v2006 (Nonlinear Dynamics, Newcastle upon Tyne, UK) was used to align, detect, match, and pick spots for each DyLight image. The method selected for spot normalization was total spot volume using the scaling factor total area where the volume of each spot is divided by the total volume of all of the spots in the image multiplied by a scaling factor. Protein spots were chosen for identification with mass spectrometry if they exhibited a >1.5-fold difference volume intensity between IPC and perfusion control with a value of p<0.05. The Ettan Spot Handling Workstation (GE Healthcare Life Sciences) performed automated extraction of the selected protein spots followed by in-gel trypsin digestion. Protein identification was carried out using a MALDI-TOF/TOF instrument (4700 Proteomics Analyzer, Applied Biosystems, Carlsbad, CA) in positive ion mode with a reflector. For MS analysis, an 800–4000 m/z mass range was used with 1500 shots per spectrum. Result dependent analysis (RDA) was used for MS/MS selection. A maximum of six precursors per protein were selected with a confidence interval (C.I.) percentage of 50 or higher with a minimum signal/noise ratio of 50. In addition, a low confidence investigation (peptides not matched to top proteins) was used to allow a maximum of 5 precursors per spot with minimum signal/noise ratio of 50 selected for data-dependent MS/MS analysis. A 1-kV collision energy was used for collision-induced dissociation (CID), and 1500 acquisitions were accumulated for each MS/MS spectrum. For both MS and MS/MS analysis, the default calibration was calibrated with a 4700 mass standard peptide mix (Applied Biosystems) achieving a mass accuracy within 50 ppm. The peak-list generating software used was GPS Explorer software, set to default parameters (version 3.0, Applied Biosystems). Mascot search engine (version 2.2, Matrix Science, Boston, MA) was used for peptide and protein identifications. Swiss-Prot Protein knowledgebase database (Sprot 2010.01.19) was searched against and MS peak filtering was set for all trypsin autolysis peaks. The taxonomy selected was Mus musculus (mouse), 46,021 protein sequences. The acceptance criteria for individual MS/MS spectra had a significance threshold set to p<0.05 with expectation values <0.05 (number of different peptides with scores equivalent to or better than the result reported that are expected to occur in the database search by chance). The p value was chosen to reflect a 95% probability that the protein identification is correct. The positive identifications criteria were two peptides or more, a MS/MS C.I. of 95% or higher, a correct molecular mass and pI position, and identification from all 2D gels.
Mouse monoclonal anti-caveolin-3 and/or anti-caveolin-1 antibody (BD Biosciences, San Jose, CA) was used in the immunoprecipitation (IP) study along with Dynabeads Protein G and DynaMag™-2 (Invitrogen, Carlsbad, CA) according to the product instruction. The Dynabeads (1.5mg) were washed and resuspended in 200μl of IP buffer (phosphate-buffered saline containing 0.05% Tween-20 and protein protease/phosphatase inhibitors). Five μg of negative control mouse IgG1, anti-caveolin-1, anti-caveolin-3, or both monoclonal antibodies were added and incubated at room temperature for 30min with gentle agitation to form Dynabead-antibody complex. After washing, the total homogenate (~500μg) of each heart sample was added and the mixture was incubated at room temperature for 30min with gentle agitation. After five washes with 500μl of IP buffer, the Dynabeads were resuspended into 80μl of 1×NuPAGE LDS sample buffer with 10% (v/v) 2-mercaptoethanol, and heated for 10min at 70°C. The extract was subjected to SDS-PAGE.
Equal volumes (from caveolae fractions) or equal amounts of total heart homogenate protein were separated by 4%–12% Bis-Tris SDS-PAGE (Invitrogen) and transferred to a nitrocellulose membrane. After the transfer, the membrane was stained with Ponceau S and then washed with TBS-T composed of (pH 8.0, in mmol/L) 10 Tris, 150 NaCl, and 0.1% (v/v) Tween 20. TBS-T supplemented with 5% (w/v) nonfat dry milk was used for the blocking solution. The antibodies were diluted as follows: 1:5000 for mouse monoclonal anti-caveolin-3 antibody (#610421, BD Biosciences), 1:1000 for mouse monoclonal anti-caveolin-1 antibody (#610407, BD Biosciences), 1:200 for goat polyclonal anti-VDAC-1 (#sc-8828) and anti-eNOS (#sc-654, Santa Cruz), and 1:1000 for rabbit polyclonal antibodies including anti-phospho(Ser473)-AKT (#9271S), anti-AKT (#9272), and anti-phospho(Ser1177)-eNOS (#9570) antibodies (Cell Signaling, Danvers, MA). The corresponding IgG HRP-conjugated secondary antibodies (1:5000 dilution, Cell Signaling) were used in combination with a chemiluminescent substrate (GE Healthcare Life Sciences) according to standard procedures.
Results are expressed as mean±SE. Statistical significance was determined by Student's t test or ANOVA as required. Differences were regarded to be significant at *p<0.05, and **p<0.01.
This work was supported by the NIH Intramural Program (JS, TN, AA, PC, SE, MG, MD and EM), NIH grants 1F32HL096142 (MK), and 5R01HL039752 (CS).
No competing financial interests exist.