The contraction of mammalian cardiac myocytes is controlled by a transient increase in cytosolic Ca²⁺ concentration via sarcoplasmic reticulum (SR) Ca²⁺ release upon depolarization. Membrane depolarization induces Ca²⁺ influx through the L-type Ca²⁺ channels, which, in turn, triggers the release of large amounts of Ca²⁺ from the SR into the cytosol [1-4]. Confocal Ca²⁺ imaging of cardiac myocytes has revealed local Ca²⁺ releases through ryanodine receptors (RyRs) (“Ca²⁺ sparks”) triggered either spontaneously or by L-type Ca²⁺ current (I_Ca) [5-7]. Ca²⁺ sparks are thought to be elementary Ca²⁺ release events that underlie global Ca²⁺ release upon depolarization in cardiac myocytes [5-9].

Reactive oxygen species (ROS) oxidize macromolecules including RyRs and are extensively involved in physiological and pathological processes in cardiac muscle [10-13]. The NADPH oxidase (Nox) is one of the major sources for superoxide anion (O₂.^–) in cardiac myocytes under various external stimuli including mechanical stresses and plays a central role in Ca²⁺ mobilization via RyRs [14-19]. Diphenyleneiodonium (DPI) has been known as a representative Nox inhibitor [20] and often used for this purpose [16-18,21]. However, DPI has exerted diverse inhibitory effects in different mammalian cell types including mitochondrial Complex I [22-25], cholinesterase [26], nitrogen oxide synthase (NOS) [27,28], and xanthin oxidase [29]. These reports raise concerns about interpretation of data obtained using DPI and suggest an involvement of ubiquitous signaling molecule in the side effects by DPI.

We have previously found that DPI, at the concentrations used to eliminate the Nox activity (~3 µM), significantly reduces the occurrence of spontaneous Ca²⁺ sparks in rat ventricular myocytes under control conditions, although the specific inhibitor of Nox 2 (e.g., gp91-ds) did not alter the occurrence of Ca²⁺ sparks [18]. Therefore, we hypothesized that DPI may affect global Ca²⁺ signals and contraction independently of its action on Nox in cardiac myocytes. To test this hypothesis, we examined the effects of DPI on contraction, global Ca²⁺ signaling and I_Ca, and cellular mechanism for the effects in isolated rat ventricular myocytes using a video edge detection, confocal imaging, and whole-cell patch clamp technique. Here we describe inhibition of excitation-contraction coupling by DPI in ventricular myocytes.

Cell isolation

Cardiac myocytes were isolated from male Sprague-Dawley rats (200–300 g) as previously described [30]. The experiments were performed in accordance with the principles for the care and use of experimental animals published by the Korean Food and Drug Administration. The surgical method was approved by the Animal Care and Use Committees of the Chungnam National University (CNU-00992). Briefly, after rats were anesthetized with sodium pentobarbital (150 mg/kg, intraperitoneal injection), the hearts were taken out with thoracotomy. Then, aorta was tied onto the cannular of Langendorff apparatus for retrograde perfusion at 7 ml/min through the aorta (at 36.5°C). The heart was perfused first with Ca²⁺-free Tyrode's solution composed of (in mM) 137 NaCl, 5.4 KCl, 10 HEPES, 1 MgCl₂, and 10 glucose (pH 7.3) for 3 min, and then with Ca²⁺-free Tyrode's solution containing collagenase (1.4 mg/ml, type A; EC 3.4.24.3; Sigma-Aldrich) and protease (0.14 mg/ml, type XIV; EC 3.4.24.31; Sigma-Aldrich) for about 12 min. Then, the heart was perfused with Tyrode's solution containing 0.2 mM Ca²⁺ for further 5 min. The digested heart was then cut and chopped into pieces for further dissociation of myocytes.

Measurement of cell shortenings

Ventricular myocytes attached at the bottom of experimental chamber were continuously superfused with normal Tyrode’s solutions (pH = 7.4) containing 2 mM Ca²⁺ and field-stimulated at 1 Hz with two paralleled Pt wires connected to an electrical stimulator (D-7806; Hugo Sachs) at room temperature. Cell shortenings were detected with a video edge detector (Model VED-105; Crescent Electronics) connected to a CCD camera (LCL902C; Till Photonics) and video monitor (ViewFinder III, Polychrome V system; Till Photonics) as previously reported [30]. Changes of cell length were monitored using A/D converter (Digidata 1322A; Molecular Devices) connected to a PC software pClamp (9.0; Molecular Devices) and analyzed with Clampfit 9.0 (Molecular Devices) and Origin software (8.0; OriginLab Corporation).

Two-dimensional (2-D) confocal Ca²⁺ imaging and image analysis

To detect cytosolic Ca²⁺ ventricular myocytes were loaded with 3 µM fluo-4 acetoxymethyl (AM) ester (Thermo Fisher Scientific) for 30 min. The dye-loaded cells were continuously superfused with 2-mM Ca²⁺-containing normal Tyrode’s solutions (pH 7.4; see above). Dyes were excited at 488 nm using Ar ion laser, and fluorescence emission at > 510 nm was detected. Cytosolic Ca²⁺ fluorescence signals were recorded in 2-D images using a laser scanning confocal imaging system (A1; Nikon) attached to an inverted microscope (Eclipse Ti; Nikon) fitted with a ×60 oil immersion objective lens (Plan Apo, Numerical Aperture 1.4; Nikon) [30]. Acquisition and analysis of images were performed using a workstation software (NIS Elements AR, v3.2; Nikon).

To record global Ca²⁺ transients, images were captured at 120 Hz in the field-stimulated (1 Hz) cells with two paralleled Pt wires connected to an electrical stimulator (D-7806; Hugo Sachs). The average diastolic fluorescence intensity (F₀) was measured from several frames captured before the upstroke of Ca²⁺ transient. The time-courses of Ca²⁺ transients were evaluated as the average fluorescence of each area normalized relative to the F₀ (F/F₀) [30]. To measure Ca²⁺ spark frequency, the whole-cell images were recorded at 30 Hz. Recording of spontaneous sparks was preceded by a series of electrical pulses at 1 Hz to maintain the SR Ca²⁺ content. Ca²⁺ sparks were identified by a computerized algorithm in the “RealTimeMicroscopy” PC program (own written in C++) as previously described [18]. In order to calculate the frequency of sparks ([total number of sparks]/[10³ µm²·s]), whole-cell areas were measured using the NIS Elements AR software (v3.2; Nikon).

Measurements of I_Ca

I_Ca was recorded using the whole cell configuration of the patch-clamp technique using an EPC7 amplifier (HEKA) as previously described [31]. The patch pipettes were made of glass capillaries (Kimble Glass) to have resistance of 2–3 MΩ. Internal solution contained (in mM) 110 CsCl, 20 TEA-Cl, 20 HEPES, 5 MgATP, and 15 EGTA, with the pH adjusted to 7.2 (with CsOH). Outward K⁺ currents were suppressed by internal Cs⁺ and TEA⁺, and inward rectifier K⁺ current was suppressed by replacing external K⁺ with Cs⁺. Na⁺ current was inactivated by holding the membrane potential at –40 mV. Recording of I_Ca was carried out ~8 min after a rupture of the membrane with the patch pipette, when the rundown of Ca²⁺ channels were slowed and stabilized. Using pCLAMP (9.0; Molecular Devices) combined with an analog-to-digital converter (Digidata 1322; Molecular Devices) we applied voltage commands and acquired current data. The series resistance was ~2 times the electrode resistance and was compensated through the amplifier. The current signals were filtered with low pass filter at 1 kHz and digitized at 10 kHz. The current data were analyzed using Clampfit (9.0; Molecular Devices). The time constant (τ) of inactivation of I_Ca was obtained with single exponential curve fitting using the equation:

where A_i and A_f are the initial (t = 0) and final (t = infinity) values of the parameter, and τ is a time constant of exponential decay. Curve fitting was performed using OriginPro 8 SR0 software (OriginLab Corporation). The percent suppressions of I_Ca by various interventions were evaluated after a gradual decrease in I_Ca by rundown was subtracted from the raw current [31].

Measurements of mitochondrial ROS

To measure mitochondrial O₂.⁻, cells were loaded with MitoSOX Red (10 µM; Thermo Fisher Scientific) for 30 min. Fluorescence signal was imaged using the same confocal system at 5-s intervals [18]. MitoSOX Red was excited with light at 514 nm while measuring the emitted light collected at 570–620 nm. To prevent artefactual signal due to light exposure low imaging speed was used. The time course of the MitoSOX fluorescence was evaluated as the average fluorescence of each area normalized to control fluorescence detected prior to DPI exposure (F₀).

Solutions and reagents

DPI was purchased from Sigma-Aldrich. DPI was diluted in Tyrode’s solution for testing (dimethylsulfoxide [DMSO] ≤ 0.08% (v/v), e.g., 0.01% DMSO at 3-µM DPI solutions). Same concentrations of DMSO were added to external solutions without or with DPI. The drug solutions were applied to the cells by superfusion using custom-made solution switching apparatus except the experiments using caffeine, and zero Na⁺ and zero Ca²⁺ external solutions. To make zero Na⁺ and zero Ca²⁺ external solutions, 137 mM Na⁺ and 2 mM Ca²⁺ were removed with adding 1 mM EGTA and 137 mM LiCl. Zero Na⁺ and zero Ca²⁺ external solutions and 10 mM caffeine-containing external solutions were rapidly applied using own made puffing device. All the experiments were performed at room temperature (22°C–25°C).

Statistics

The numerical results are presented as means ± standard error of the mean. n indicates number of cells used. The Student's t-tests were used for statistical comparisons depending on the experiments. Differences were considered to be significant to a level of p < 0.05.

Effects of DPI on contraction in rat ventricular myocytes

Fig. 1 shows the effects of different concentrations of DPI on cell shortenings in rat ventricular myocytes stimulated at 1 Hz. Cell shortenings were reduced by the application of DPI in a concentration-dependent manner (Fig. 1): % inhibition: 7.65 ± 1.61, 22.8 ± 2.19, 29.7 ± 1.63, 42.0 ± 5.28, 64.5 ± 3.80, and 73.0 ± 5.30 at the concentrations of 0.01, 0.05, 0.1, 1, 10 and 100 µM, respectively. Curve fitting for the concentration-response plot using Hill equation showed 50%-inhibition in contraction by DPI at ≅0.17 µM with maximal inhibition of 69.5 ± 6.34% at 100 µM (Fig. 1B). DPI did not significantly alter the time-to-peak of contraction, the time-to-relaxation and the rates of contraction and relaxation (Table 1 and Fig. 1B, inset).

Table 1 . Effects of DPI on the kinetics of cell contraction and relaxation.

	Control	DPI (1 µM)
Time-to-peak (ms)	82.1 ± 6.24	89.3 ± 11.2
Time-to-relaxation (ms)	183 ± 10.3	206 ± 19.8
Rate of contraction (µm/s)	69.4 ± 8.61	66.8 ± 9.23
Rate of relaxation ([µm/s)	13.8 ± 2.52	12.6 ± 2.76

Data represents mean ± SEM. Stable six cells were analyzed. DPI, diphenyleneiodonium.

Figure 1. Negative inotropy induced by diphenyleneiodonium (DPI) in rat ventricular myocytes. (A) Representative contraction traces recorded immediately before (“Control”) and after the exposure to different concentrations of DPI in field-stimulated rat ventricular myocytes at 1 Hz. The traces were selected when a maximal decrease in cell shortening by DPI was observed. (B) Concentration-dependent decrease in cell shortenings (% suppression) by the extracellular application of DPI; 0.01 μM, n = 3, p > 0.05; 0.05 μM, n = 4, p < 0.01; 0.1 μM, n = 4, p < 0.01; 1 μM, n = 6, p < 0.001; 10 μM, n = 6, p < 0.0001; 10 μM, n = 4, p < 0.001. Paired t-test was used. The sigmoidal curve represents the fit of the Hill equation.

Attenuation of global Ca²⁺ signaling by DPI

To determine the cellular mechanism underlying the negative inotropy in the presence of DPI, we examined the effects of DPI on global Ca²⁺ signaling in these myocytes. Fig. 2 represents a series of Ca²⁺ transients measured from a field-stimulated ventricular myocyte, followed by a Ca²⁺ signal induced by the treatment of 10 mM caffeine in resting condition, before and after the application of DPI (3 µM). We tested DPI at the concentration of 3 µM, which showed submaximal effects on cell shortening and was used to examine the role of Nox in cardiac myocytes [16,18]. The results showed that the systolic Ca²⁺ levels and the magnitudes of Ca²⁺ transients were significantly decreased by DPI (% of control: systolic Ca²⁺, 93.1 ± 1.9, p < 0.01; magnitude of Ca²⁺ transient, 80.1 ± 2.7, p < 0.0001; n = 15; Fig. 2A, B). Diastolic Ca²⁺ levels were not significantly altered by DPI (% of control: 108 ± 2.4, p > 0.05, n = 15; Fig. 2A, B). The kinetics of release and decay of Ca²⁺ during depolarization, estimated as the time-to-90%-peak (T_P,90) and the half-decay time (D_1/2), was not significantly altered by DPI (Fig. 2C; % of control: T_P,90, 96.4 ± 6.4, p > 0.05; D_1/2, 103 ± 3.5, p > 0.05, n = 15).

Figure 2. Alterations of global Ca²⁺ signaling in the presence of diphenyleneiodonium (DPI). (A) Ca²⁺ transients measured in a representative ventricle cell followed by caffeine (10 mM)-induced Ca²⁺ signals in the absence and presence of 3 μM DPI. (B, C) Summary of mean values of the diastolic and systolic Ca²⁺ levels, and in the magnitude, time-to-90%-peak (T_P,90) and half decay time of Ca²⁺ transient in the absence (Con) and presence of DPI (3 μM) (n = 15). ****p < 0.0001, **p < 0.01 vs. Con (Paired t-test). (D) Comparison of magnitudes of the caffeine-induced Ca²⁺ transients and fractional release measured before (Control) and after application of DPI (3 μM) (n = 15). ****p < 0.0001 vs. Con (Paired t-test).

The SR Ca²⁺ loading status, evaluated by the magnitude of caffeine-induced Ca²⁺ transients (Fig. 1A), was significantly reduced by the exposure to DPI (Fig. 2D; % of control in 3 µM DPI: 82.6 ± 2.8%, p < 0.001, n = 15). Fractional release, the ratio of depolarization-induced Ca²⁺ release to caffeine-induced Ca²⁺ release, was not affected by DPI (Fig. 2D; % of control in 3 µM DPI: 96.4 ± 1.9, p > 0.05, n = 12). These results indicate that DPI significantly attenuates Ca²⁺ release from the SR on depolarization in ventricular myocytes, and that such reduction in Ca²⁺ release may be associated with the decrease in SR Ca²⁺ loading.

Suppression of I_Ca by DPI

The Ca²⁺ release in cardiac myocytes on depolarization is mainly controlled by Ca²⁺ influx through the L-type Ca²⁺ channels [1-4]. I_Ca also contributes to loading of Ca²⁺ into the SR. To understand the cellular mechanism for DPI-induced reductions in Ca²⁺ transient and SR Ca²⁺ content (Fig. 2), we next tested whether DPI alters I_Ca. The effects of different concentrations of DPI (0.3–200 µM) on I_Ca were tested using whole-cell patch clamp technique (see METHODS). The I_Ca was continuously measured at 0.1 Hz during a voltage step to 0 mV from a holding potential of –40 mV. There was no change in I_Ca in the presence of 0.3 µM (Fig. 3B). The application of DPI at 3 µM slightly but significantly reduced the peak I_Ca (Fig. 3; by 12 ± 3.3%, n = 6, p < 0.05). This effect by DPI was not mimicked by the application of the specific inhibitors of Nox 2 and Nox 4—the most abundantly expressed isoforms of Nox in adult cardiac myocytes [32] (Supplementary Fig. 1). Higher concentrations of DPI suppressed I_Ca more strongly in a concentration-dependent manner with IC₅₀ value of ≅40 µM (Fig. 3A–C; 30 µM: 41 ± 6.9%, n = 8, p < 0.01; 100 µM, 71 ± 4.7, n = 4, p < 0.05; 200 µM, 84 ± 4.1%, n = 3, p < 0.05). The time constant of inactivation of I_Ca, measured with curve fitting (see METHODS), was slightly increased by the application of 30 µM DPI (control, 24.4 ± 1.35 ms; 30 µM DPI, 27.8 ± 1.62 ms, p < 0.05, n = 9). We measured the current-voltage relationship of the I_Ca before and after application of submaximal concentrations of DPI (30 µM), and found there was no significant change in the current-voltage relationship by DPI (Fig. 3D, E). These results suggest that the attenuations of Ca²⁺ transients and SR Ca²⁺ loading in the presence of 3 µM DPI may be caused by I_Ca suppression. The concentration-dependent inhibition of contractility in the presence of DPI is also consistent with stronger suppression in I_Ca by the higher concentrations of DPI.

Figure 3. Suppression of Ca²⁺ current (I_Ca) by diphenyleneiodonium (DPI). (A) Superimposed I_Ca recorded at 0 mV (holding potential at –40 mV) before and after applications of 3 μM, 30 μM, 100 μM, and 200 μM DPI in the representative ventricular myocytes, showing concentration-dependent inhibition in I_Ca by DPI. Scale bars indicate 30 ms in x axis and 2 pA/pF in y axis. (B) Comparison of mean peak I_Ca measured under control conditions and after applications of different concentrations of DPI (0.3 μM, n = 3; 3 μM, n = 6; 30 μM, n = 8; 100 μM, n = 4; 200 μM, n = 3). **p < 0.01, *p < 0.05 vs. control (Con). Paired t-test. (C) Concentration-dependence curve for % inhibition in I_Ca versus concentrations of DPI. Plot was fit with Hill equation (Hill coefficient = 0.68). (D) Superimposed I_Ca measured at voltage steps ranging –40 to +70 mV (holding at –40 mV) with 10-mV-increment before (Control) and after application of 30 μM DPI in a representative ventricle cell. (E) Current-voltage relationships of averaged I_Ca at the step potentials in the control condition and after exposure to 30 μM DPI (n = 3). Cells were dialyzed with 15 mM EGTA containing internal solutions.

DPI-induced suppression of Ca²⁺ sparks independently of external Na⁺- and Ca²⁺-flux

We have previously observed that DPI at the concentrations of 3 µM decreased the frequency of spontaneous Ca²⁺ sparks in rat ventricular myocytes [18]. Since the Ca²⁺ sparks represent elementary Ca²⁺ releases through the RyR clusters composing global Ca²⁺ increase during I_Ca in cardiac myocytes [5-9], one of possible mechanisms for decrease in global Ca²⁺ releases on depolarization in the presence of DPI is such reduction in the activity of RyRs. We further examined whether external Ca²⁺- and/or Na⁺-mediated ionic flux through the cell membrane plays a role in suppression of Ca²⁺ releases in the cells treated with DPI, we tested the effects of removal of external Na⁺ and Ca²⁺ on DPI-induced Ca²⁺ spark suppression.

To quantify the occurrence of Ca²⁺ sparks, we performed 2 s-long 2-D confocal Ca²⁺ imaging at 30 Hz to monitor a major part of the ventricular cells. Conditioning electrical stimulations were applied to stabilize SR Ca²⁺ loading before measuring spontaneous Ca²⁺ sparks. Under this control conditions, Ca²⁺ sparks spontaneously occurred at a frequency of 1.82 ± 0.23 events/10³ µm²·s (n = 11). The treatment of 3 µM DPI suppressed the spark frequency to 1.17 ± 0.17 events/10³ µm²·s (n = 11) within 2–3-min of treatment (p < 0.01), consistent with the previous report [18]. When the external Na⁺ and Ca²⁺ were shortly removed (see METHODS), the spark frequency was not significantly changed (control, 1.83 ± 0.32 vs. zero Na⁺ and Ca²⁺, 1.81 ± 0.33, p > 0.05, n = 7). In the continued presence of external zero Na⁺ and zero Ca²⁺ solutions, application of DPI still reduced the spark frequency by ~25% (DPI in zero Na⁺ and Ca²⁺: 1.34 ± 0.30, p < 0.05, n = 7) (Fig. 4). This result suggests that other Ca²⁺ entry and Ca²⁺- and/or Na⁺-dependent membrane ionic flux may not contribute to suppression of spontaneous Ca²⁺ sparks in the presence of DPI.

Figure 4. Suppression of Ca²⁺ sparks by diphenyleneiodonium (DPI) in the absence of external Na⁺ and Ca²⁺. (A) A series of sequential confocal Ca²⁺ images recorded at 30 Hz from a representative resting rat ventricular myocyte in the control condition (“1”) and after brief exposure to zero Na⁺ and zero Ca²⁺ external solutions (0 Na, 0 Ca) for 10 s (“2”), followed by additional application of DPI (0 Na, 0 Ca, DPI) for 3 min (“3”). The images were selected from the periods marked with the boxes correspondingly numbered (“1”, “2”, and “3”) in the panel B. After 3-min DPI application, DPI was removed from the zero Na⁺ and zero Ca²⁺ solutions (2 min; (0 Na, 0 Ca, DPI)`), which was followed by the exchange of external solution with control solutions (5 min; “Wash”). Arrows indicate distinct Ca²⁺ sparks. (B) Plots of the total numbers of sparks occurred in each frame (33 frames/s) versus 2-s-long recording periods under each condition labeled above the plots. (C) Summary of mean spark frequency detected under each condition indicated underneath the graphs. Paired t-test was used.

Decrease in mitochondrial ROS level by DPI

A line of previous reports indicate that long-term injection of DPI can cause mitochondrial myopathy in animal models by impairing oxidative phosphorylation, particularly Complex 1 activity [22-25]. Complex I is essential for ATP synthesis and can generate ROS in the cardiac muscle mitochondria. Inhibition of Complex 1 by DPI has reduced mitochondrial O₂.^– in unstimulated monocyte/macrophage [33] and in isolated mitochondria from guinea-pig cardiac myocytes [34]. Oxidation of the thiol groups in RyR2 by ROS enhances its channel activity, whereas their reduction inhibits the channels [35-37]. Therefore, in the next series of experiments, we tested whether DPI also affects mitochondrial ROS level in rat ventricular myocytes under control conditions using confocal imaging with MitoSOX-red, the mitochondrial O₂.^– indicator. We found that application of DPI at 3–100 µM elicited significant decreases in the mitochondrial O₂.^– level in these myocytes (Fig. 5). These results support the notion that reduction in spontaneous spark frequency in the presence of 3 µM DPI with no external Ca²⁺ and Na⁺ (Fig. 4) may be due to decrease in mitochondrial ROS level.

Figure 5. Reduction of mitochondrial superoxide by diphenyleneiodonium (DPI). (A) Plots of averaged Mito-SOX fluorescence, normalized to the levels (F₀) just prior to the application of DPI (3 μM, n = 4; 30 μM, n = 5; 100 μM, n = 4), versus recording time, showing decrease of mitochondrial superoxide levels by DPI in a concentration-dependent manner. (B) Summary of Mito-SOX fluorescence ratio measured at 100-s after the onset of DPI applications, showing DPI-induced signal reductions at 3 μM, 30 μM, and 100 μM DPI. **p < 0.01, *p < 0.05 vs. control (Con) (paired t-test).

In this study, we demonstrate for the first time that DPI exerts negative inotropic effects on cardiac myocytes (Fig. 1), and that such negative inotropy can be mediated by the suppression of I_Ca and Ca²⁺-induced Ca²⁺ release upon depolarization (Figs. 2 and 3). The suppressive effect on I_Ca by DPI was not mimicked by other specific Nox inhibitors (Supplementary Fig. 1). We also showed that this chemical significantly reduces SR Ca²⁺ loading and the occurrence of Ca²⁺ sparks at the concentrations (3 µM) used to investigate the role of Nox in mammalian cells including cardiac myocytes (Figs. 2 and 4). The suppression of resting Ca²⁺ spark occurrence by DPI was not altered when the external Ca²⁺ and Na⁺ were removed (Fig. 4), suggesting that I_Ca-independent mechanism may be involved in spark suppression by DPI. Mitochondrial superoxide level was significantly reduced by DPI at the concentration of 3–100 µM. These results indicate that DPI suppresses Ca²⁺ channels and RyRs, thereby eliciting negative inotropy in cardiac myocytes, and suggest that the effects of DPI may involve its inhibitory action on mitochondrial metabolism independently of Nox.

Decreases in the I_Ca and SR Ca²⁺ loading can underlie decreases in the Ca²⁺ transient magnitudes in cells exposed to DPI (Figs. 2 and 3). However, the decrease in the Ca²⁺ transients may partly explain the negative inotropy of ventricular myocytes during the DPI exposure. At the concentrations of 3 µM, the magnitude of Ca²⁺ transients were reduced by approximately 20% (Fig. 2A, B), while cell shortenings were decreased by 50%–60% (Fig. 1). In addition, at the concentrations of 0.3 µM, contraction was significantly decreased by DPI (Fig. 1), although I_Ca was not altered (Fig. 3). These results suggest that other mechanisms may be involved in the negative inotropy by DPI. In this regard, it has been reported in skeletal muscle that DPI causes fatigue and force failure via irreversible impairment in oxidative phosphorylation, particularly Complex 1 activity [22-25]. Potent and strong inhibition in contraction by DPI may be caused by such reduction in ATP production by mitochondria.

Reduction in SR Ca²⁺ loading in the presence of DPI could be a result of decreased Ca²⁺ influx through the Ca²⁺ channels (Figs. 2D and 3). Because most of the Ca²⁺ removal during the decay phase of Ca²⁺ transient is thought to be mediated by SR Ca²⁺ pump in ventricular myocytes from rat heart [38], one can indirectly approximate the activity of SR Ca²⁺ pump with evaluating the speed of Ca²⁺ transient decay. No change in the half decay time of Ca²⁺ transient in the presence of 3 µM DPI (Fig. 2C) suggests that the decrease in SR Ca²⁺ content by DPI may not be caused by alteration in the activity of SR Ca²⁺ pump. Nevertheless, it should also be noted, in the microsome preparation from pig coronary artery, that DPI at the concentrations of > 10 µM has exerted mild inhibitory effects on SR Ca²⁺ pump activity [26].

Resting Ca²⁺ spark frequency was reduced by 3 µM DPI even in the absence of external Na⁺ and Ca²⁺ (Fig. 4), suggesting suppression of in situ activity of RyR2 clusters in cardiac myocytes exposed to this chemical regardless of I_Ca and/or Na⁺–Ca²⁺ exchanger. We have previously found that suppressive effect by DPI on the spark frequency is not mimicked by the application of inhibitor for either Nox 2 (gp91-ds) or NOS (L-NAME) in rat ventricular myocytes [18]. In addition, inhibition of xanthine oxidase, one of the ROS producing enzymes that is sensitive to DPI [29], has rather enhanced ventricular sparks in rats [39], which excludes its role in the effect of DPI on Ca²⁺ sparks. It is plausible to suggest that the suppressive effect by DPI on the resting spark frequency could be indirectly caused by decrease in SR Ca²⁺ content (Fig. 2), since SR luminal Ca²⁺ plays a role in sensitizing RyR to Ca²⁺ in cardiac myocytes [40,41]. Nevertheless, reversible and quick effects of DPI on the spark frequency (Fig. 4) may reflect other mechanism involved in modulation of RyRs by DPI, such as reduction of thiol groups on RyR by decrease of mitochondrial ROS level in the vicinity [11,35-37]. Since we observed mitochondrial ROS reduction during application of DPI at similar concentrations of DPI (Fig. 5), such notion may be possible.

Nox 2 and Nox 4 are the most abundant Nox isoforms in cardiac myocytes [32]. The former is a key signaling protein to generate ROS in the transverse-tubules of cardiac myocytes under external or mechanical stress [16,17] and the latter has been thought to play a role in mitochondrial ROS generation under pathological conditions, such as cardiomyopathy and hypertrophy [32,42]. Nox generates O₂.^– in a highly regulated manner with a stimulus dependent cell-signaling pathways unlike other ROS sources, such as xanthin oxidase and mitochondrial respiratory chain [14-19]. Although current experiments were performed in the control conditions with no stress, possibility that Nox inhibition contributes to DPI-induced mitochondrial ROS reduction in normal cardiac myocytes under control conditions may not be completely excluded. Inhibition of Complex I in the mitochondrial respiratory chain by DPI [22-25] has been shown to decrease cellular and mitochondrial ROS in macrophages and in isolated mitochondria from guinea-pig cardiac myocytes [33,34]. These reports seem to be consistent with our observation on DPI-induced ROS reduction in the mitochondria of rat ventricular myocytes (Fig. 5). It should be noted, however, that increase in ROS level by DPI has also been reported in other cell type and/or experimental conditions [43].

Slight but significant suppression in I_Ca in the presence of 3 µM DPI was not mimicked by other specific Nox inhibitors, such as the Nox-2-blocking peptide gp91ds-tat and the Nox-4-specific inhibitor setanaxib (Supplementary Fig. 1). In addition, inhibition of NOS using L-NAME did not significantly alter I_Ca in these myocytes (Supplementary Fig. 2). These support that Nox and NOS are not involved in the suppressive effect by DPI on I_Ca in cardiac myocytes under control conditions. The effect of DPI on membrane ion channels including Ca²⁺ channels have been recognized previously at the similar concentrations in other cell types, for examples, neuron from carotid body and smooth muscle cells of pulmonary vasculature [44,45]. The mechanism for DPI-induced I_Ca suppression, however, has not been fully understood. In this regard, it has been shown that an addition of H₂O₂ in the presence of DPI has not reversed the inhibitory effect by DPI on I_Ca in pulmonary smooth muscle cells [44]. Molecular mechanism for the I_Ca suppression by DPI via the mitochondrial inhibition (e.g., Complex I) remains uncertain. Further studies will be required to determine whether the mechanisms including 1) attenuated mitochondrial Ca²⁺ uptake to inactivate I_Ca [46], 2) a drop of cytosolic pH [47] resulting from suppression of oxidative phosphorylation (increased anaerobic glycolysis), or 3) loss of ATP and/or Ca²⁺-dependent channel phosphorylation by kinases [48] such as CaMKII or PKC, play a role in the inhibition of I_Ca by DPI.

Strong suppressions in I_Ca and contraction at high concentrations of DPI appear to elicit cardiac toxicity. Such DPI effects appear to be similar to the H₂S- or rotenone-mediated toxicity in cardiac myocytes in terms of the modulations of contraction and mitochondrial metabolism [49,50]. Interestingly, H₂S- or rotenone-mediated cardiotoxicity was reversed by the application of methylene blue [50]. It would be worth investigating the effects of DPI in combination with methylene blue, H₂S and/or rotenone to further delineate the mode of action of DPI in the modulation of cardiac excitation-contraction coupling.

DPI induces negative inotropy in cardiac myocytes and attenuate Ca²⁺-induced Ca²⁺ releases via suppression of L-type Ca²⁺ channel, SR Ca²⁺ loading and RyRs in the presence of popular concentrations of DPI used to inhibit Nox. In addition, the results suggest that ventricular contractile machinery is more sensitive to DPI compared to Ca²⁺ channels. Its effects on I_Ca and RyRs appear to be independent of inhibition of Nox or NOS. Since DPI significantly reduced ROS in the mitochondria the suppression of RyRs by such ROS decrease is likely. Since Ca²⁺ is a ubiquitous signaling molecule, DPI-induced secondary effects on other proteins are expected. This raises needs for a caution to interpret experimental results collected with the use of DPI. In this regard, our data obtained in cardiac Ca²⁺ signaling and contraction may help dissecting a role of Nox itself in cardiac functional regulation under various environments.

Supplementary data including two figures can be found with this article online at https://doi.org/10.4196/kjpp.2024.28.4.335

None.

The work was supported by National Research Foundation of Korea (NRF) grants funded by the Korea government (2022R1A2C2091583 and 2022R1A5A7085156).

The authors declare no conflicts of interest.