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. 2010 Jun 7;588(Pt 15):2905–2917. doi: 10.1113/jphysiol.2010.192617

Regulation of myocyte contraction via neuronal nitric oxide synthase: role of ryanodine receptor S-nitrosylation

Honglan Wang 1, Serge Viatchenko-Karpinski 1, Junhui Sun 2, Inna Györke 1, Nancy A Benkusky 3, Mark J Kohr 1, Héctor H Valdivia 3, Elizabeth Murphy 2, Sandor Györke 1, Mark T Ziolo 1
PMCID: PMC2956906  PMID: 20530114

Abstract

The sarcoplasmic reticulum (SR) Ca2+ release channel (ryanodine receptor, RyR2) has been proposed to be an end target of neuronal nitric oxide synthase (NOS1) signalling. The purpose of this study is to investigate the mechanism of NOS1 modulation of RyR2 activity and the corresponding effect on myocyte function. Myocytes were isolated from NOS1 knockout (NOS1−/−) and wild-type mice. NOS1−/− myocytes displayed a decreased fractional SR Ca2+ release, NOS1 knockout also led to reduced RyR2 S-nitrosylation levels. RyR2 channels from NOS1−/− hearts had decreased RyR2 open probability. Additionally, knockout of NOS1 led to a decrease in [3H]ryanodine binding, Ca2+ spark frequency (CaSpF) and a rightward shift in the SR Ca2+ leak/load relationship. Similar effects were observed with acute inhibition of NOS1. These data are indicative of decreased RyR2 activity in myocytes with NOS1 knockout or acute inhibition. Interestingly, the NO donor and nitrosylating agent SNAP reversed the depressed RyR2 open probability, the reduced CaSpF, and caused a leftward shift in the leak/load relationship in NOS1−/− myocytes. SNAP also normalized Ca2+ transient and cell shortening amplitudes and SR fractional release in myocytes with NOS1 knockout or acute inhibition. Furthermore, SNAP was able to normalize the RyR2 S-nitrosylation levels. These data suggest that NOS1 signalling increases RyR2 activity via S-nitrosylation, which contributes to the NOS1-induced positive inotropic effect. Thus, RyR2 is an important end target of NOS1.

Introduction

Nitric oxide (NO) is an important regulator of cardiac contractility. NO within cardiac myocytes is endogenously produced via two constitutively expressed isoforms: endothelial NO synthase (NOS3) and neuronal NO synthase (NOS1). Signalling via NOS1 and NOS3 is compartmentalized and each isoform modulates cardiac function differently (Barouch et al. 2002; Ziolo et al. 2008). NOS3 is localized to the caveolae and blunts the response to β-adrenergic stimulation, while NOS1 is localized to the sarcoplasmic reticulum (SR) and enhances contraction (Barouch et al. 2002; Wang et al. 2008b; Ziolo et al. 2008). That is, NOS1 knockout (NOS1−/−) or acute inhibition has been found to decrease contraction (Wang et al. 2008a), and attenuate the force–frequency response (Khan et al. 2003; Wang et al. 2008a). However, limited data exist on the mechanism(s) of how NOS1 modulates contractility.

The SR Ca2+ release channel (ryanodine receptor, RyR2) is an essential excitation–contraction coupling protein. Upon RyR2 opening, Ca2+ is expelled from the SR (i.e. Ca2+ transient) leading to myocyte contraction (Bers, 2002). RyR2, which has been shown to be endogenously S-nitrosylated (Xu et al. 1998), has been implicated as a target for NOS1 (Barouch et al. 2002; Gonzalez et al. 2007a; Lim et al. 2008). Studies have demonstrated that NO donors can increase the open probability of RyR2 channels through a direct effect via S-nitrosylation of cysteine residues (Stoyanovsky et al. 1997; Xu et al. 1998). We further demonstrated that NO is able to modulate β-adrenergic stimulated RyR2 activity (measured as Ca2+ spark frequency) in intact cardiac myocytes (Ziolo et al. 2001). In this study, we used the NO donor, and S-nitrosylating agent, S-nitroso-N-acetylpenicillamine (SNAP) to test the hypothesis that NOS1 signalling increases S-nitrosylation levels of RyR2 leading to increased RyR2 activity and enhanced contraction.

In order to precisely define the regulation of NOS1 signalling on RyR2 activity, we used five different ways to measure activity. We directly measured open probability in purified RyR2 channels reconstituted into lipid bilayers. We also performed the [3H]ryranodine binding assay, an indirect measure of open probability. In intact cardiac myocytes, a more physiological approach, we measured Ca2+ sparks, the SR Ca2+ leak/load relationship, and fractional release (Ca2+ transient amplitude/SR Ca2+ load).

Methods

Isolation of ventricular myocytes

As previously described (Wang et al. 2008a), ventricular myocytes were isolated from male and female (4–5 months) NOS1 knockout (NOS1−/−) mice and their corresponding wild-type (WT, C57BL/6J) mice (Jackson Laboratories, Bar Harbor, ME, USA), which were anaesthetized with sodium pentobarbital (i.p. injection of 50 mg kg−1). Briefly, the heart was mounted on a Langendorff apparatus and perfused with modified minimal essential medium (MEM; Sigma, St Louis, MO, USA, 37°C, bubbled with 95% O2–5% CO2). Blendzyme type IV (0.077 mg ml−1) (Roche Applied Science, Indianapolis, IN, USA) was then added to the perfusate. After 7–20 min, the heart was taken down, the ventricles minced, and myocytes dissociated by trituration. Subsequently the myocytes were filtered, centrifuged and resuspended in MEM containing 200 μm Ca2+. Myocytes were used within 6 h after isolation. All the animal protocols and procedures were performed in accordance with National Institutes of Health guidelines and approved by the Institutional Laboratory Animal Care and Use Committee at The Ohio State University. They comply with the policies and regulations of The Journal of Physiology (Drummond, 2009).

Simultaneous measurement of Ca2+ transients and cell shortening

Functional measurements were performed as previously described (Kohr et al. 2008). Briefly, myocytes were loaded at room temperature with Fluo-4 AM (10 μm, Molecular Probes, Eugene, OR, USA) for 30 min, and then 30 min were allowed for intracellular de-esterification. The instrumentation used for cell fluorescence measurements was an epifluorescence system (Cairn Research Ltd, Faversham, UK). [Ca2+]i was measured by Fluo-4 epifluorescence with excitation at 480 ± 20 nm and emission at 535 ± 25 nm. The illumination field was restricted to collect the emission of a single cell. Data were expressed as ΔF/F0, where F is the fluorescence intensity and F0 is the intensity at rest. Simultaneous measurement of shortening was also performed using an edge detection system (Crescent Electronics, Sandy, UT, USA). Data were expressed as the precentage of resting cell length (%RCL). Measurements were performed at room temperature.

We performed experiments examining the effects of S-methyl-l-thiocitrulline (SMLT)/SNAP on Fluo-3 (pentapotassium salt) properties using a spectrometer (LS55, Perkin Elmer Inc., Waltham, MA, USA). We observed that SMLT/SNAP produced less than 1.5% variability on our measurements. Thus, we believed that SMLT/SNAP does not affect the Ca2+ affinity and /or fluorescence properties of the dye.

Measurement of SR Ca2+ load

SR Ca2+ load was measured by rapid application of 10 mm caffeine for 10 s. The amplitude of the caffeine-induced Ca2+ transient was used as an index of SR Ca2+ load (Trafford et al. 1999). Measurements were performed at room temperature.

DyLight switch method for S-nitrosothiol detection

All procedures were performed in the dark to prevent the light-induced cleavage of S-nitrosothiols (SNOs). The snap-frozen hearts were ground into powder in liquid N2 followed by Dounce glass homogenization in 1.5 ml HEN buffer (0.25 m Hepes-NaOH, pH 7.7, 1 mm EDTA, 0.1 mm neocuproine) containing 0.3 m sucrose and protease inhibitor (cocktail tablet by Roche). After spinning at 1000 g to remove the debris, the supernatant was saved as the total homogenate. Total homogenate (200 μg) of each heart was treated with 0 or 10 μm SNAP for 5 min at 24°C in the dark. Free SNAP was removed by −20°C acetone precipitation. The pellets were resuspended into 100 μl HEN buffer containing 2% SDS and 20 mm methyl methanethiolsulfonate, incubated at 50°C for 30 min with gentle vortex every 5 min. After blocking free thiols, the samples were subjected to −20°C acetone precipitation to remove free MMTS, then resuspended into 75 μl HEN buffer with 1% SDS, 1 mm ascorbate, and 1 mm DyLight-maleimide 800 (Pierce Biotechnology, Inc., Rockford, IL, USA) for modified biotin switch using DyLight-maleimide (Sun et al. 2007). The DyLight-labelled samples were resuspended into 60 μl of 1× sample buffer containing 10%β-mercaptoethanol, and incubated at 37°C for 15 min in the dark. Twenty microlitres of each mixture was loaded onto 12 wells of NuPAGE Novex 4–12% Bis-Tris gel (Invitrogen). After SDS-PAGE, the gels were rinsed with de-ionized H2O and scanned at 800 nm using Li-COR Odyssey for DyLight-maleimide 800 fluorescence/SNO signal, then were transferred to PVDF membrane at 4°C overnight for anti-RyR2 (MA3-916, Affinity BioReagents, Golden, CO, USA; 1:1000 dilution) Western blot.

Single RyR2 channel recording

Heavy SR microsomes were isolated by differential centrifugation from NOS1−/− (10 mice) and their corresponding WT (10 mice) hearts. Single RyR2s were reconstituted by fusing SR microsomes into planner lipid bilayers as previously described (Lukyanenko et al. 1996). SR vesicles were added into one side of the bilayer (defined as cis). The other side was defined as trans (virtual ground). Channel incorporation was performed in solutions containing (in mm): 350 CsCH3SO3, 0.02 CaCl2, and 20 Hepes (pH 7.4), on the cis (cytosolic) side of the bilayer, and 20 CsCH3SO3, 0.02 CaCl2, and 20 Hepes (pH 7.4) on the trans (luminal) side of the bilayer. The free [Ca2+] in our solutions was measured with a Ca2+-selective electrode. After incorporation, the trans CsCH3SO3 was adjusted to 350 mm. Single channel data were sampled during short (400 ms) repetitive steps to +40 mV from 0 mV. Single channel parameters (open probability, mean open time, mean close time and unitary current amplitude) were assessed with 400–2000 repetitive voltage steps. Single channel currents were recorded with an Axopatch 200A patch-clamp 8 amplifier (Molecular Devices, Sunnyvale, CA, USA). Data acquisition and analysis were performed by using pCLAMP 9.2 software (Molecular Devices).

[3H]Ryanodine binding assay

[3H]Ryanodine binding was performed as described (Jiang et al. 2002). The Ca2+ dependence of [3H]ryanodine binding was determined in total homogenates in medium containing 20 mmol l−1 Hepes (pH 7.4), 200 mmol l−1 KCl, 1 mmol l−1 EGTA, and CaCl2 to give a range of free [Ca2+] from pCa 8 to pCa 3 (Ca2+-EGTA constants taken from MaxChelator http://www.stanford.edu/~cpatton/webmaxcS.htm). [3H]Ryanodine binding was normalized to RyR2 expression (measured via Western blot). RyR2 antibody was purchased from Affinity BioReagents.

Assessment of Ca2+ sparks

Intracellular Ca2+ imaging was performed using an Olympus Fluoview 1000 laser scanning confocal microscope equipped with Olympus oil objective (60×, 1.4 NA). To measure Ca2+ sparks in intact myocytes, cells were incubated with Fluo-3 AM (50 μm, Molecular Probes, Eugene, OR, USA) for 25 min and then 20 min was allowed for de-esterification. Fluo-3 was excited at the 488 nm line of an argon ion laser, and fluorescence was acquired at >510 nm in the line-scan mode of the confocal system at the rate of 2.5 ms per scan. Solution for sparks recordings (in mm): 140 NaCl, 5.4 KCl, 2.0 CaCl2, 1 MgCl2, 10 Hepes, and 5.6 glucose, pH 7.3. The magnitude of fluorescence signals was quantified in terms of ΔF/F0, where F0 is baseline fluorescence and ΔF=F−F0. Ca2+ spark parameters were quantified with a detection/analysis computer algorithm.

Measurement of SR Ca2+ leak

SR Ca2+ leak was measured as the tetracaine (RyR2 inhibitor)-induced shift in diastolic [Ca2+]i (Shannon et al. 2002; Knollmann et al. 2006; Picht et al. 2007). Myocytes were loaded at 22°C with Fluo-4 AM (10 μm, Molecular Probes, Eugene, OR, USA) for 30 min, washed out, and then an additional 30 min was allowed for intracellular de-esterification. Then the cells were stimulated for 1 min in normal Tyrode solution. After reaching steady state, the solution was rapidly switched to 0 Na+, 0 Ca2+ Tyrode solution (Na+ was replaced by Li+) plus 1 mm tetracaine (Sigma) for 30 s. Afterwards, the solution was rapidly switched to 0 Na+, 0 Ca2+ Tyrode solution for 20 s. Tetracaine, by blocking RyR2 and therefore SR Ca2+ leak, caused a decrease in diastolic [Ca2+]i. The shift in diastolic [Ca2+]i upon the removal of tetracaine was used as a measure of RyR2-dependent SR Ca2+ leak. Since SR Ca2+ leak is also dependent upon SR Ca2+ load, SR Ca2+ load of each cell was measured. Measurements were performed at room temperature.

Solution and drugs

Normal Tyrode solution consisted of (in mm): 140 NaCl, 4 KCl, 1 MgCl2, 1 CaCl2, 10 glucose, 5 Hepes, 1 l-arginine, pH 7.4 adjusted with HCl. S-Methyl-l-thiocitrulline (SMLT, 10 μm, Calbiochem, La Jolla, CA, USA), a specific NOS1 inhibitor (Narayanan & Griffith, 1994), and SNAP (1 and 10 μm, Sigma), a NO donor and S-nitrosylating agent, were prepared fresh each experimental day. Myocytes were pre-incubated with SMLT (30 min) or SNAP (20 min).

Statistics

Results were expressed as means ±s.e.m. Statistical significance (P < 0.05) was determined by ANOVA (followed by the Newman–Keuls test) for multiple groups. Student's t test for unpaired data were used for comparison between two groups.

Result

NOS1−/− myocytes display decreased SR Ca2+ fractional release and RyR2 S-nitrosylation levels

Myocytes were isolated from NOS1−/− and WT mouse hearts. Shown in Fig. 1A are representative Ca2+ transient traces from a WT myocyte and NOS1−/− myocytes. Consistent with our previous findings (Wang et al. 2008a), NOS1 knockout myocytes had decreased Ca2+ transient amplitude (0.44 ± 0.02 vs. 0.26 ± 0.03 ΔF/F0, P < 0.05) and SR Ca2+ load (0.56 ± 0.03 vs. 0.87 ± 0.04 ΔF/F0, P < 0.05) compared to WT myocytes. In addition, we observed a decreased SR Ca2+ fractional release (twitch Ca2+ transient amplitude/SR Ca2+ load) with NOS1 knockout (WT: 0.49 ± 0.02, NOS1−/−: 0.37 ± 0.03, P < 0.05 vs. WT; Fig. 1A). We confirmed that the effects observed in the NOS1−/− myocytes were due to deletion of NOS1, and not an adaptive response, by acutely inhibiting NOS1 in WT myocytes with 10 μm SMLT. Similar to NOS1 knockout, acute NOS1 inhibition also decreased Ca2+ transient amplitude (0.31 ± 0.02 ΔF/F0, P < 0.05 vs. WT) and SR Ca2+ load (0.73 ± 0.04 ΔF/F0, P < 0.05 vs. WT) compared to WT myocytes. Moreover, acute NOS1 inhibition resulted in decreased SR Ca2+ fractional release (WT+SMLT: 0.35 ± 0.02, P < 0.05 vs. WT; Fig. 1A).

Figure 1. NOS1 knockout or inhibition decreased SR Ca2+ fractional release and RyR2 S-nitrosylation levels.

Figure 1

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A, representative twitch and caffeine-induced Ca2+ transient traces (left: WT, right: NOS1−/−). B, summary data (means ±s.e.m.) of SR Ca2+ fractional release (measured as twitch Ca2+ transient amplitude/SR Ca2+ Load) in WT myocytes (open bar), NOS1−/− myocytes (grey bar) and acute inhibition with SMLT in WT myocytes (black bar). C, RyR2 S-nitrosylation levels measured by the DyLight switch method. Representative blots show S-nitrosothiols (SNO) (top) and total RyR2 (bottom). Summary data (mean ±s.e.m.) of the ratio of S-nitrosothiols and total RyR2, *P < 0.05 vs. WT. n= 39–65 cells/9–15 hearts. n= 4 hearts each for S-nitrosylation measurements.

We also tested if RyR2 from NOS1−/− hearts had decreased S-nitrosylation levels. As shown in Fig. 1C, knockout of NOS1 resulted in significantly decreased RyR2 S-nitrosylation levels (normalized to total RyR2) (WT: 1.7 ± 0.1 arbitrary units (A.U.); NOS1−/−: 1.0 ± 0.2 A.U.; P < 0.05). These data together (i.e. decreased SR Ca2+ fractional release and decreased RyR2 S-nitrosylation levels) suggest that knockout of NOS1 may lead to decreased RyR2 activity.

NOS1 knockout decreases open probability of single RyR2 channels

We directly measured RyR2 activity by investigating the effect of NOS1 knockout on open probability of RyR2 channels. RyR2 channels were isolated from WT and NOS1−/− hearts and incorporated into planar lipid bilayers for single channel recordings. Shown in Fig. 2A are representative traces from a WT and NOS1−/− RyR2 channel. Summary data (Fig. 2B) show that RyR2 channels from NOS1−/− hearts displayed a 3.1-fold reduction in RyR2 open probability compared to WT (Po: 0.11 ± 0.02 vs. 0.34 ± 0.05, P < 0.05). The decrease in Po was due to a shortened mean open time (WT: 3.5 ± 0.5 ms vs. NOS1−/−: 1.3 ± 0.1 ms, P < 0.05), and a trend toward a prolonged mean close time (WT: 7.3 ± 2.0 ms vs. NOS1−/−: 11.4 ± 1.7 ms, P= 0.15). There was no difference in amplitude (WT: 22 ± 4 pA vs. NOS1−/−: 24 ± 6 pA, not significant (NS)). Thus, our data indicate that NOS1 knockout results in decreased activity of RyR2 channels.

Figure 2. NOS1 knockout decreased RyR2 activity.

Figure 2

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A, representative traces from a WT (left) and NOS1−/− (right) single RyR2 channel. B, summary data (mean ±s.e.m.) of open probability in WT (open bar) and NOS1−/− (grey bar) RyR2 channels. *P < 0.05 vs. WT. n= 10–14 channels for each group. C, [3H]ryanodine binding in WT (black) and NOS1−/− (grey) at different Ca2+ concentrations normalized to total RyR2.

NOS1 knockout decreases Ca2+ dependence of [3H]ryanodine binding

Due to the limitation of using a single, unbuffered Ca2+ concentration in our single RyR2 channels measurements, we performed the [3H]ryanodine binding assay. As shown in Fig. 2C, in the range of pCa 7 to pCa 3, NOS1 knockout decreased [3H]ryanodine binding to RyR2, confirming a reduced RyR2 open probability in NOS1−/− hearts under various, buffered Ca2+ concentrations.

NOS1 knockout or inhibition reduces Ca2+ sparks

We further investigated the effects of NOS1 on RyR2 activity in a more physiologically relevant manner. We measured Ca2+ sparks in intact myocytes isolated from WT and NOS1−/− hearts. Shown in Fig. 3A are representative images of Ca2+ sparks from a WT myocyte and NOS1−/− myocyte. NOS1−/− myocytes had significantly reduced Ca2+ spark frequency and amplitude compared to WT myocytes (frequency: 1.6 ± 0.1 vs. 2.2 ± 0.3 sparks (100 μm)−1 s−1; amplitude: 0.56 ± 0.04 vs. 0.72 ± 0.03 ΔF/F0; both P < 0.05; Fig. 3B). These Ca2+ spark data are consistent with our single channel recordings and [3H]ryanodine binding assay (i.e. reduced RyR2 activity with NOS1 knockout). Acute NOS1 inhibition resulted in similar effects on Ca2+ sparks as NOS1 knockout. That is, SMLT reduced Ca2+ spark frequency and amplitude (1.6 ± 0.2 sparks (100 μm)−1 s−1; 0.57 ± 0.03 ΔF/F0, both P < 0.05 vs. WT; Fig. 3). NOS1 knockout or acute inhibition had no effect on Ca2+ spark half-rise time (WT, 11.5 ± 0.4 ms; NOS1−/−, 12.1 ± 0.4 ms; WT+SMLT, 11.8 ± 0.3 ms; all NS), however NOS1 knockout or acute inhibition prolonged the Ca2+ spark half-decay time (WT, 28.3 ± 0.3 ms; NOS1−/−, 35.2 ± 0.9 ms, P < 0.05 vs. WT; WT+SMLT, 30.9 ± 1.2 ms, P < 0.05 vs. WT). Interestingly, our previous study showed that NOS1−/− myocytes had slower [Ca2+]i decline due to hypophosphorylation of phospholamban (Wang et al. 2008a), which could explain this slower half-decay time (Gomez et al. 1996). Our previous data have shown that SMLT had no effect on Ca2+ transient and cell shortening amplitude nor SR Ca2+ load in NOS1−/− myocytes (Wang et al. 2008a). We further show that SMLT does not affect Ca2+ spark frequency or amplitude in NOS1−/− myocytes (amplitude: 0.59 ± 0.06 ΔF/F0, frequency: 1.6 ± 0.4 sparks (100 μm)−1 s−1, NS vs. without SMLT). These data further suggest that NOS1 signalling enhances RyR2 activity, with knockout or inhibition resulting in decreased activity of RyR2.

Figure 3. NOS1 knockout or inhibition reduced Ca2+ spark frequency and amplitude.

Figure 3

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A, representative images of Ca2+ sparks from WT (top), NOS1−/− (middle) and WT+SMLT (bottom) myocytes. B, summary data (means ±s.e.m.) of Ca2+ spark frequency (top) and amplitude (bottom) in WT (open bar), NOS1−/− (grey bar) and WT+SMLT (black bar) myocytes. *P < 0.05 vs. WT. n= 219–447 sparks/16–25 cells/4 hearts.

NOS1 knockout or inhibition shifts the SR Ca2+ leak–load relationship

SR Ca2+ leak was also utilized to measure RyR2 activity in intact myocytes. In WT and NOS1−/− myocytes, we measured the tetracaine (RyR2 inhibitor)-induced shift in diastolic [Ca2+]i. Shown in Fig. 4A are the representative traces in a NOS1−/− myocyte and WT (± SMLT) myocytes. Our summary data (Fig. 4B) demonstrated that NOS1−/− myocytes have a significantly decreased SR Ca2+ leak compared to WT myocytes (0.014 ± 0.004 vs. 0.033 ± 0.005 ΔF/F0, P < 0.05). Acutely inhibiting NOS1 in WT myocytes with SMLT also resulted in decreased SR Ca2+ leak (0.016 ± 0.006 ΔF/F0, P < 0.05 vs. WT). Since the SR Ca2+ leak is strongly dependent on the SR Ca2+ load, the SR Ca2+ load was also measured in each cell. Figure 4B (right panel) shows that NOS1 knockout and inhibition decreased SR Ca2+ load, consistent with our previous data (Wang et al. 2008a) (WT: 0.92 ± 0.09 ΔF/F0; NOS1−/−: 0.65 ± 0.14 ΔF/F0, P= 0.08 vs. WT; WT+SMLT: 0.60 ± 0.12 ΔF/F0, P < 0.05 vs. WT). Then, we plotted the SR Ca2+ leak–load relationship. Figure 4C shows that all three groups displayed the normal relationship between SR Ca2+ leak and SR Ca2+ load (i.e. the higher the SR Ca2+ load the greater the SR Ca2+ leak). However, knockout or inhibition of NOS1 induced a rightward shift in this relationship, which is indicative of decreased RyR2 activity. A previous study has shown that backflux from the SR Ca2+ pump also contributes to the SR Ca2+ leak (Shannon et al. 2002). However, for this study we used the tetracaine-induced shift in diastolic [Ca2+]i as the measure of SR Ca2+ leak, which specifically examined the RyR2-dependent leak (Shannon et al. 2002). Thus, in our five distinct methods to measure RyR2 activity, we consistently found impaired RyR2 activity with NOS1 knockout or inhibition.

Figure 4. NOS1 knockout or inhibition shifts the SR Ca2+ leak–load relationship.

Figure 4

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A, representative traces of the SR Ca2+ leak/load protocol from a WT myocyte (left), NOS1−/− myocyte (middle) and WT+SMLT myocyte (right). B, summary data (means ±s.e.m.) of SR Ca2+ leak (left) and SR Ca2+ load (right) in WT, NOS1−/− and WT+SMLT myocytes. C, the SR Ca2+ leak–load relationship in WT (continuous line), NOS1−/− (grey line) and WT+SMLT (dashed line) myocytes. *P < 0.05 vs. WT. n= 20–35 cells/5–6 hearts.

SNAP reversed the reduced activity of NOS1−/− RyR2 channels

Next we repeated our experiments in the presence of the NO donor SNAP. We first investigated the effects of SNAP (1 μM) on single RyR2 channel activity. Shown in Fig. 5A (left) are representative traces from a NOS1−/− RyR2 channel in the presence of SNAP. Summary data (Fig. 5A, right) show that SNAP increased RyR2 channel open probability from NOS1−/− hearts (Po: 0.28 ± 0.05, P < 0.05 vs. NOS1−/−). SNAP had a small affect on WT RyR2 Po (Po: 0.36 ± 0.1, NS vs. WT). Moreover, there was now no difference in open probability in NOS1−/− RyR2 channels compared to WT RyR2 channels. The increase in Po with SNAP in NOS1−/− RyR2 channels was due to a prolonged mean open time (4.1 ± 1.3 ms, P < 0.05 vs. NOS1−/−). There was also a trend that SNAP shortened the mean close time (7.4 ± 1.1 ms, P= 0.08 vs. NOS1−/−). There was no affect on amplitude (21 ± 4 pA, NS vs. NOS1−/−). As with Po, there was no effect of SNAP on mean open time (4.5 ± 1.3 ms, NS vs. WT), or mean close time (6.3 ± 0.7 ms, NS vs. WT) in WT RyR2 channels.

Figure 5. SNAP reversed the decreased RyR2 activity in NOS1−/− and/or SMLT-treated myocytes.

Figure 5

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A, left, representative traces from a NOS1−/− RyR2 channel in the presence of SNAP. Right, summary data (mean ±s.e.m.) of open probability in WT (open bar, data from Fig. 2), NOS1−/− (dark grey bar, data from Fig. 2) and NOS1−/−+SNAP (light grey bar) RyR2 channels. *P < 0.05 vs. WT, **P < 0.05 vs. NOS1−/−. n= 8–14 channels for each group. B, summary data (means ±s.e.m.) of the effects of SNAP on Ca2+ spark frequency (left) and amplitude (right) in WT and NOS1−/− myocytes. n= 52–111 sparks/6–12 cells/2 hearts. *P < 0.05 vs. control. C, the SR Ca2+ leak–load relationship in WT (left), NOS1−/− (middle) and SMLT-treated (right) myocytes (± SNAP). n= 16–19 cells/4–6 hearts.

We also investigated the effects of SNAP (1 μm) on Ca2+ sparks. Figure 5B shows that SNAP only increased Ca2+ spark frequency without changing amplitude in WT myocytes (1.9 ± 0.1 vs. 2.6 ± 0.2 sparks (100 μm)−1 s−1, P < 0.05; 0.80 ± 0.03 vs. 0.77 ± 0.03 ΔF/F0, NS). However, SNAP significantly increased both Ca2+ spark frequency and amplitude in NOS1−/− myocytes (1.6 ± 0.3 vs. 2.2 ± 0.3 sparks (100 μm)−1 s−1, P < 0.05; 0.59 ± 0.03 vs. 0.71 ± 0.06 ΔF/F0, P < 0.05). Additionally, SNAP increased the NOS1−/− myocyte spark frequency and amplitude to levels that were similar to WT. These data indicate that SNAP can reverse the blunted Ca2+ spark frequency and amplitude observed in NOS1−/− myocytes.

The effects of SNAP (1 μm) were also tested in our SR Ca2+ leak/load protocol. SNAP had little effect on leak in WT myocytes, but increased the leak in myocytes with NOS1 knockout or inhibition (WT: 0.016 ± 0.002 vs. 0.020 ± 0.002 ΔF/F0, NS; NOS1−/−: 0.010 ± 0.002 vs. 0.029 ± 0.004 ΔF/F0, P < 0.05; WT+SMLT: 0.010 ± 0.001 vs. 0.019 ± 0.004 ΔF/F0, P < 0.05). Our data also show that 1 μm SNAP caused a leftward shift in the SR Ca2+ leak–load relationship in WT+SMLT and NOS1−/− myocytes (Fig. 5C). In fact, SNAP normalized the SR Ca2+ leak–load relationship in myocytes with NOS1 knockout or acute inhibition to WT myocytes. These SR Ca2+ leak/load data are consistent with our Ca2+ spark data. Thus, taken together, our Ca2+ spark and SR Ca2+ leak/load data indicate that SNAP is able to reverse the blunted RyR2 activity.

SNAP reserved the depressed contraction in NOS1−/− myocytes and increased RyR2 S-nitrosylation

In order to further confirm the finding that NOS1 knockout or inhibition results in decreased RyR2 activity and correspondingly depressed cardiac contraction, we simultaneously measured Ca2+ transients and cell shortening (± SNAP) in myocytes isolated from WT and NOS1−/−. Shown in Fig. 6A are representative Ca2+ transient and cell shortening traces. As we have previously shown (Wang et al. 2008a), NOS1 knockout or inhibition significantly decreased Ca2+ transient and cell shortening amplitudes. Incubation of myocytes with SNAP (1 μm) increased Ca2+ transient and cell shortening amplitudes in all three groups (Fig. 6B). However, SNAP had much greater effects in NOS1−/− and SMLT-treated myocytes (Ca2+ transient amplitude, WT: 26 ± 12%Δ from control, NOS1−/−: 71 ± 24%Δ from control, WT+SMLT: 89 ± 19%Δ from control; cell shortening amplitude, WT: 36 ± 48%Δ from control, NOS1−/−: 188 ± 120%Δ from control, WT+SMLT: 133 ± 68%Δ from control). In fact, our data indicate that SNAP normalized the depressed Ca2+ transient and cell shortening amplitude in NOS1−/− and acutely inhibited myocytes (Fig. 6B, NS vs. WT). Interestingly, SNAP also enhanced the SR Ca2+ fractional release in myocytes with NOS1 knockout or inhibition (WT: 0.56 ± 0.05, NOS1−/−: 0.46 ± 0.03, WT+SMLT: 0.46 ± 0.04, both P < 0.05 vs.–SNAP). Further, there was no difference in the fractional release between WT and NOS1−/− (or SMLT-treated) myocytes (Fig. 6C, NS vs. WT). These data suggest that the depressed RyR2 activity contributes to the reduced cardiac contraction observed in NOS1−/− and SMLT-treated myocytes.

Figure 6. SNAP reversed the depressed cardiac function in NOS1−/− and SMLT-treated myocytes via RyR2 S-nitrosylation.

Figure 6

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A, representative shortening (top) and Ca2+ transient (bottom) traces from a WT (left), NOS1−/− (middle) and SMLT-treated (right) myocytes (± SNAP). Note, scale bars are for all traces. B, summary data (means ±s.e.m.) of the effects of SNAP on Ca2+ transient and cell shortening amplitudes. C, summary data (means ±s.e.m.) of the effects of SNAP on SR Ca2+ fractional release. D, RyR2 S-nitrosylation levels with SNAP treatment. Representative blots show S-nitrosothiols (SNO) (top) and total RyR2 (bottom). Summary data (means ±s.e.m.) of the ratio of S-nitrosothiols and total RyR2. *P < 0.05 vs. control, #P < 0.05 vs. WT. n= 15–37 cells/4–7 hearts. n= 4 hearts each for S-nitrosylation measurements.

We also pretreated cardiac homogenates with SNAP (10 μm). As shown in Fig. 6D, SNAP significantly increased S-nitrosylation levels of both WT and NOS1−/− RyR2 (WT, 2.4 ± 0.1 A.U.; NOS1−/−, 2.0 ± 0.1 A.U., P < 0.05 vs.– SNAP). As with contraction, SNAP had a much larger effect in increasing RyR2 S-nitrosylation levels in NOS1−/−vs. WT RyR2 (104 ± 31%vs. 40 ± 11%Δ from control). In addition, SNAP also normalized the S-nitrosylation levels of RyR2 in NOS1−/−vs. WT (Fig. 6D, NS). These data indicate that the effect of SNAP to reverse the cardiac function is via, in part, increased RyR2 S-nitrosylation levels.

Discussion

Our data show that SR Ca2+ fractional release and RyR2 S-nitrosylation levels are decreased in NOS1−/− myocytes, which suggested that RyR2 activity may be reduced. By using four other methods to confirm, we observed decreased RyR2 activity resulting from the knockout of NOS1. Our data demonstrated that RyR2 channels isolated from NOS1−/− hearts have decreased open probability and decreased Ca2+-dependent [3H]ryanodine binding. In intact NOS1−/− myocytes, we observed a decrease in Ca2+ spark frequency/amplitude and a rightward shift in the SR Ca2+ leak–load relationship. We observed similar effects when we acutely inhibited NOS1 in WT myocytes with SMLT. Interestingly, the NO donor SNAP, also a S-nitrosylating agent, reversed the decreased RyR2 open probability, Ca2+ spark frequency/amplitude, and the SR Ca2+ leak/load relationship in myocytes with NOS1 knockout or with acute inhibition. SNAP also normalized the depressed contractile amplitude in myocytes with NOS1 knockout or acute inhibition. This was associated with an increase in SR Ca2+ fractional release and RyR2 S-nitrosylation levels. Thus, we are the first to show that NOS1 signalling increases RyR2 activity via S-nitrosylation contributing to its positive inotropic effect.

NOS1 signalling and myocyte contraction

It is well established that NOS1 signalling is able to regulate basal and β-adrenergic stimulated cardiac function in both normal and disease states (Ashley et al. 2002; Barouch et al. 2002; Khan et al. 2003; Zhang et al. 2008; Ziolo et al. 2008). For example, NOS1−/− hearts and myocytes exhibit a blunted force–frequency response, decreased contraction, and slowed [Ca2+]i decline. However, limited data exist on the mechanism of how NOS1 modulates cardiac contraction. Phospholamban (PLB) has been demonstrated to be an important end target for NOS1 regulation of cardiac function (Wang et al. 2008a; Zhang et al. 2008). That is, myocytes with NOS1 knockout or with acute inhibition had decreased PLB serine-16 phosphorylation, which is the cAMP-dependent protein kinase (PKA) site. Interestingly, activation of PKA via stimulation of the β-adrenergic pathway was able to normalize the depressed PLB phosphorylation levels in NOS1−/− myocytes. This also normalized the [Ca2+]i decline rates. However, the β-adrenergic stimulated [Ca2+]i transient and shortening amplitudes were still blunted in NOS1−/− myocytes. These data suggest an additional protein target(s) exists. Indeed, our present data showed that myocytes with NOS1 knockout or inhibition displayed decreased RyR2 activity (Figs 2–4). It should be noted that measuring RyR2 activity in lipid bilayers may not be equivalent to RyR2 activity within intact cardiac myocytes. For example, the influence of regulatory factors including intracellular soluble ligands, kinases, phosphatases and auxiliary proteins is likely to be disrupted in the bilayer RyR2 preparation. Another limitation of our RyR2 lipid bilayer experiments is using a single, unbuffered [Ca2+]. Ideally, RyR2 measurements should be done under conditions similar to those in situ. However, matching conditions of bilayer and myocyte experiments is difficult for several reasons. RyR2 Po near 100 nm, which corresponds to resting Ca2+ levels in myocytes, is extremely low, which precludes the acquisition of a sufficient number of events required for meaningful analysis. Thus, we measured [3H]ryanodine binding using various buffered Ca2+ concentrations. Further, we performed additional experiments examining RyR2 activity in intact myocytes (i.e. Ca2+ sparks, SR Ca2+ leak–load relationship, and fractional release). In the intact myocyte where the regulatory factors are present, RyR2 activity was still decreased with NOS1 knockout or inhibition. Thus, besides phospholamban, RyR2 is also an end target of NOS1 signalling.

One study had suggested that increased diastolic Ca2+ leak from RyR2 is, in part, responsible for the slower relaxation and attenuated force–frequency response in NOS1−/− hearts (Khan et al. 2003). They also directly measured SR Ca2+ leak in NOS1−/− myocytes and reported enhanced RyR2-dependent SR Ca2+ leak compared to WT myocytes (Gonzalez et al. 2007a). However, the myocytes used in this study exhibited spontaneous activity. As a result of this spontaneous activity, the physiological effects of NOS1 signalling on RyR2 may be altered. Since our experimental conditions were different and our myocytes did not exhibit spontaneous activity, we may not have seen the increased SR Ca2+ leak. Under our experimental conditions, we consistently observed reduced RyR2 activity using five distinct methods. Thus, this reduced RyR2 activity may contribute to the blunted cardiac contraction exhibited in myocytes with NOS1 knockout or inhibition. Indeed, the SR Ca2+ fractional release, which is partly determined by RyR2, was depressed in NOS1−/− and SMLT-treated myocytes (Fig. 1). It should be noted that the reduced SR Ca2+ load may also contribute to the decreased fractional release (Shannon et al. 2000). However, there are reported instances in the literature that demonstrate a change in fractional release without changes in SR Ca2+ load and Ca2+ current (duBell et al. 1996). Further Maier et al. (2003) showed that even with decreased SR Ca2+ load there was an increase in fractional release. Both studies concluded that the changes in fractional release were due to changes in RyR2 activity. We performed fractional release studies to obtain an indirect measure of RyR2 activity in contracting myocytes. Along with the decreased fractional release, we also observed a decrease in RyR2 S-nitrsoylation levels in NOS1−/− myocytes (Fig. 1). SNAP also increased SR Ca2+ fractional release in NOS1−/− and SMLT-treated myocytes, consistent with its effect on RyR2 activity (Fig. 6). Interestingly, our current results (Fig. 2C) and previous studies have found increased expression of RyR2 in NOS1−/− hearts (Sears et al. 2003; Gonzalez et al. 2007a). We suggest that this is a compensatory adaptation of NOS1−/− mice in view of the fact that we observed a decrease in RyR2 activity.

Effects of nitric oxide on RyR channels

RyR2 activity is regulated by many factors including Ca2+, calmodulin, phosphorylation, S-nitrosylation, etc. (Stamler & Meissner, 2001; Meissner, 2004). S-Nitrosylation is an important cGMP-independent signalling pathway of NO (Jaffrey et al. 2001; Hess et al. 2005; Ziolo, 2008). Studies have found that NO donors can increase RyR2 activity via S-nitrosylation (Stoyanovsky et al. 1997; Xu et al. 1998; Ziolo et al. 2001). In fact, Xu et al.'s data demonstrated that S-nitrosylation of up to 12 sites (3 per RyR2 subunit) led to progressive channel activation (Xu et al. 1998). While RyR2 can be endogenously S-nitrosylated (Xu et al. 1998; Gonzalez et al. 2007a), few studies have investigated the pathway(s) responsible. In the heart, NOS1 localizes to the SR and co-immunoprecipitates with RyR2 (Barouch et al. 2002) and we found that NOS1−/− hearts have deficient RyR2 S-nitrosylation levels (Fig. 1B). This is consistent with the reduced RyR2 activity (Figs 2–4). Hence, our data suggest that within cardiac myocytes NOS1 signalling leads to RyR2 S-nitrosylation.

Similar to RyR2, the skeletal muscle ryanodine receptor isoform, RyR1, can also be endogenously S-nitrosylated, which also leads to channel activation (Eu et al. 2003). We have found that S-nitrosylatation at a single cysteine (3635) of RyR1 is capable of activating the channel (Sun et al. 2001), although another study reported that there are multiple cysteines which can be S-nitrosylated (Aracena-Parks et al. 2006). For RyR1, this S-nitrosylation is also due to NOS1 signalling (Eu et al. 2003). In addition to physiological regulation, studies have found that hypernitrosylation of RyR1 can lead to abnormal channel activity during pathological states, such as impaired exercise capacity (Bellinger et al. 2008), dystrophic muscle (Bellinger et al. 2009), and environmental heat stroke and sudden death (Durham et al. 2008). It has also been hypothesized that S-nitrosylation may help protect RyR from oxidative stress (i.e. oxidation of RyR) (Eu et al. 2003; Sun et al. 2006). We have found that oxidation of RyR2 during heart failure also leads to abnormal channel activity and myocyte dysfunction (Terentyev et al. 2008). Thus, future studies are warranted to investigate the S-nitrosylation/oxidation status of RyR2 as a potential molecular basis for cardiac pathologies.

We further investigated RyR2 S-nitrosylation by using the NO donor SNAP. The greater cardiac contraction observed with the NO donor SNAP has been suggested to be due to, in part, enhanced S-nitrosylation and increased activity of RyR2 (Gonzalez et al. 2007b). Consistent with the lack of NO production via NOS1, SNAP had a much greater effect leading to normalized RyR2 S-nitrosylation and open probability in NOS1−/− and SMLT-treated myocytes compared to WT myocytes (Figs 5 and 6). SNAP also had a much larger effect in NOS1−/− and SMLT-treated myocytes, compared to WT myocytes, resulting in normalization of myocyte contractile amplitude (Fig. 6). Our data show that part of the contractile dysfunction with NOS1 knockout or inhibition is due to reduced RyR2 activity. We also infer that the mechanism of the SNAP-mediated normalization of NOS1−/− myocyte function is via increased RyR2 activity. Therefore, we suggest that NOS1 signalling leads to a positive inotropic effect, in part, via S-nitrosylation of RyR2.

A smaller SR Ca2+ leak should result in an increase in the SR Ca2+ load (Lukyanenko et al. 2001). However, NOS1 signalling also results in the phosphorylation of PLB, which will maintain SR Ca2+ load (Wang et al. 2008a). This NOS1-mediated effect on PLB may explain why NOS1−/− myocytes exhibited lower SR Ca2+ load despite a smaller SR Ca2+ leak (Fig. 4). Additionally, previous work has shown that altering RyR2 activity alone will not result in steady-state changes in myocyte contraction due to corresponding changes in SR Ca2+ load (Eisner et al. 2009). However, we hypothesize that NOS1 has multiple end targets which work in concert to increase myocyte contraction. That is, NOS1 signalling results in S-nitrosylation of RyR2 (which leads to increased activity) and phosphorylation of PLB (to maintain SR Ca2+ levels) (Wang et al. 2008a). Thus, NOS1 signalling via modulation of multiple end targets, including RyR2, is an important regulator of cardiac contraction resulting in positive inotropy.

Acknowledgments

This work was supported by the American Heart Association (Post-doctoral Fellowship 0725560B, H.W.; SDG 0435033N, S.V.-K.; Pre-doctoral Fellowship 0715159B, M.J.K.) and the National Institutes of Health (HL 055438, H.H.V.; HL 074045 and HL 063043, S.G.; HL 079283 and HL 094692, M.T.Z.). We thank Sean Little and Dr Jonathan Davis for their technical assistance.

Glossary

Abbreviations

NO

nitric oxide

NOS

nitric oxide synthase

SNAP

S-nitroso-N-acetylpenicillamine

SNO

S-nitrosothiol

SR

sarcoplasmic reticulum

Author contributions

H.H.V, E.M., S.G. and M.T.Z. designed the experiments. H.W., S.V.K., J.S., I.G., N.A.B., M.J.K. performed the experiments and analyzed the data. All authors approved the final version of the paper for publication.

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