METHYLENE BLUE COUNTERACTS H₂S-INDUCED CARDIAC ION CHANNEL DYSFUNCTION AND ATP REDUCTION

Abstract

We have previously demonstrated that methylene blue (MB) counteracts the effects of hydrogen sulfide (H₂S) cardiotoxicity by improving cardiomyocyte contractility and intracellular Ca²⁺ homeostasis disrupted by H₂S poisoning. In vivo, MB restores cardiac contractility severely depressed by sulfide and protects against arrhythmias, ranging from bundle branch block to ventricular tachycardia or fibrillation. To dissect the cellular mechanisms by which MB reduces arrhythmogenesis and improves bioenergetics in myocytes intoxicated with H₂S, we evaluated the effects of H₂S on resting membrane potential (E_m), action potential (AP), Na⁺/Ca²⁺ exchange current (I_NaCa), depolarization-activated K⁺ currents and ATP levels in adult mouse cardiac myocytes and determined if MB could counteract the toxic effects of H₂S on myocyte electrophysiology and ATP. Exposure to toxic concentrations of H₂S (100 μM) significantly depolarized E_m, reduced AP amplitude, prolonged AP duration at 90% repolarization (APD₉₀), suppressed I_NaCa and depolarization-activated K⁺ currents, and reduced ATP levels in adult mouse cardiac myocytes. Treating cardiomyocytes with MB (20 μg/ml) 3 min after H₂S exposure restored E_m, APD₉₀, I_NaCa, depolarization-activated K⁺ currents, and ATP levels towards normal. MB improved mitochondrial membrane potential (Δψ_m) and oxygen consumption rate (OCR) in myocytes in which Complex I was blocked by rotenone. We conclude that MB ameliorated H₂S-induced cardiomyocyte toxicity at multiple levels: (i) reversing excitation-contraction coupling defects (Ca²⁺ homeostasis and L-type Ca²⁺ channels); (ii) reducing risks of arrhythmias (E_m, APD, I_NaCa and depolarization-activated K⁺ currents); and (iii) improving cellular bioenergetics (ATP, Δψ_m).

Introduction

The justification of the present study is the persistent risk of accidental, environmental and industrial H₂S exposure (3, 21), and the use of sulfide as a method of suicide (56). One of the major toxic effects of H₂S resides in its ability to dramatically depress ventricular contractility that can lead to pulseless electrical activity (PEA) within a few minutes (37, 43, 73). In addition, we have recently demonstrated in rats that H₂S infusion led to frequent premature ventricular complexes, severe bradycardia, and in some instances, ventricular tachycardia or fibrillation (43). Finally, when a patient or an animal recovers from H₂S-induced coma, the continued presence of cardiogenic shock has been shown to contribute to the development of diffuse cortical and subcortical neuronal necrosis. This effect is mediated by a reduction in cerebral blood flow often associated with a decrease in arterial partial pressure of O₂ due to H₂S-induced ventilatory depression (6), which potentiates the direct toxicity of sulfide on neurons.

We have recently reported that methylene blue (MB) exerts a significant salutatory effect on the immediate outcome of sulfide intoxication (70) ; largely accounted for by the very rapid counteraction by MB of H₂S-induced depression in cardiac contractility in sedated or un-sedated rats (70) and in isolated cardiomyocytes (43).

MB is a redox molecule, which is reduced to leucomethylene blue (LMB) in the blood and in cells by NADH, NADPH (59) or reduced glutathione (44). LMB can provide electrons to a variety of oxidizing molecules (including O₂ and metallo-compounds) and is re-oxidized back to MB in the process, allowing for a new cycle of reduction-oxidation to occur. The theoretical mechanisms by which MB/LMB counteract H₂S toxicity include: (i) MB/LMB oxidation of ferrous iron into ferric iron present in various metallo-compounds. This occurs when high concentrations of MB (87) trap H₂S and catalyzes its oxidation in a cyclic manner. Of note, LMB acts as a reducing agent on ferric iron, an effect typically observed with low concentrations of MB, a property used for the treatment of methemoglobinemia (91); (ii) direct oxidation of H₂S by MB, a condition where H₂S could become a reducing agent allowing MB to form LMB (58); (iii) restoration by LMB of the redox environment of cardiac ion channels altered directly or indirectly, i.e., though ROS production (99) or protein sulfhydration (93); and (iv) a direct effect of MB/LMB on the mitochondrial electron chain transport complexes (89, 94).

The purpose of the present study was to determine the cellular mechanisms by which MB reduces the risks of arrhythmogenesis in animals exposed to H₂S; and to evaluate if MB confers beneficial effects on cellular bioenergetics in myocytes. Using adult mouse cardiomyocytes, we determined the effects of H₂S on resting membrane potential (E_m), action potential (AP), Na⁺/Ca⁺ exchange current (I_NaCa) and depolarization-activated K⁺ currents, and whether H₂S-induced alterations in electrophysiological parameters can be ameliorated by MB, thereby accounting for the beneficial anti-arrhythmic effects of MB observed in vivo (43). To simulate a clinically relevant scenario, MB was administered after H₂S exposure to isolated cardiomyocytes, at a time when signs of toxicity were already manifest. The effects of H₂S, with and without MB, on myocyte ATP levels were then measured. Finally, as a first step to dissect the mechanisms by which MB improves cellular bioenergetics, we measured mitochondrial membrane potential (Δψ_m) and oxygen consumption rate (OCR) in adult myocytes in which the electron chain transport activity was inhibited and tested whether MB could restore this function.

Methods

Isolation of adult murine cardiac myocytes

Cardiac myocytes were isolated from the LV free wall and septum of mice according to the protocol of Zhou et al. (98) and modified by us (68, 69, 79, 82–84). Myocytes were used within 2-8 h of isolation.

Myocyte shortening measurements

Myocytes adherent to coverslips were bathed in 0.7 ml of air- and temperature-equilibrated (37°C), HEPES-buffered (20 mM, pH 7.4) medium 199 containing 1.8 [Ca²⁺]_o. Myocytes were field stimulated to contract (2 Hz) between platinum wire electrodes spaced 2 mm apart. Images of myocytes viewed through an Olympus DApoUV x40/1.30 numerical aperture (NA) oil objective situated in a Zeiss IM35 inverted microscope were captured by a charge-coupled device video camera (Myocam; Ionoptix, Milton, MA). Edge detection algorithm was used to measure myocyte motion, and data were analyzed off-line by Ionoptix software as previously described (68, 69, 79, 82–84).

Electrophysiological measurements

Na⁺/Ca²⁺ exchanger current (I_NaCa)(68, 69, 82, 83, 95) and action potential (1 Hz)(68, 79, 82, 83) were measured in isolated LV myocytes (30°C) with whole cell patch-clamp. Fire-polished pipettes (tip diameter 4-6 μm) with resistances of 0.8-1.4 MΩ when filled with pipette solutions were used. Compositions of solutions and voltage protocols are given in Figure Legends.

For measurement of depolarization-activated K⁺ currents, only myocytes isolated from the LV free wall were used since the slow component of transient outward current (I_to,s) is absent in this subset of mouse myocytes (92). Pipette solution contained (in mM): 135 KCl, 1 CaCl₂, 14 EGTA, 10 HEPES and 5 MgATP, pH 7.1. ATP was included to block ATP-sensitive outward K⁺ current. External solution contained (in mM): 132 NaCl, 5.4 KCl, 1.8 CaCl₂, 1.8 MgCl₂, 0.6 NaH₂PO₄, 10 HEPES, 0.5 CdCl₂ and 10 glucose, pH 7.4. CdCl₂ was added to block L-type Ca²⁺ current and I_NaCa. Holding potential was at −70 mV. Voltage-gated K⁺ currents (30°C) were evoked during 5 s depolarizing voltage steps (from −40 to +60 mV; 10 mV increments). Myocytes were returned to −70 mV for 200 ms prior to next voltage step (68). After identifying peak currents, the decaying phases of the currents were fitted with 2 exponentials of the form: A₁e^−t/τ1 + A₂e^−t/τ2 + A_ss; where A₁, A₂ and A_ss are the amplitudes of fast component of the rapidly inactivating transient outward current (I_to,f), slowly inactivating K⁺ current (I_K,slow), and non-inactivating steady-state current (I_ss), respectively; and τ1 and τ2 are the time constants of decay of I_to,f and I_K,slow, respectively (92). Only values derived from curve fits with correlation coefficients of ≥ 0.97 were reported.

In vitro study protocol

For contraction experiments, at time zero, myocytes were exposed to either MB (0 to 500 μg/ml) or MB (0 to 500 μg/ml) + H₂S (100 μM; prepared from NaHS) and contraction from 3-5 myocytes per glass coverslip was measured at 10 min. For electrophysiological experiments, either saline or NaHS (100 μM) was added at time zero. At 3 min., MB (20 μg/ml) or saline was added and I_NaCa, depolarization-activated K⁺ currents and AP were measured at 7 min.

Measurement of mitochondrial membrane potential (ΔΨ_m)

LV cardiac myocytes were suspended in an intracellular-like medium (ICM) containing (in mM): KCl 120, NaCl 10, KH₂PO₄ 1, HEPES–Tris 20, thapsigargin (2 μg/ml), digitonin (80 μg/ml), pH 7.2; and protease inhibitors (EDTA-free complete tablets, Roche Applied Science)(40, 52). Permeabilized myocytes were supplemented with succinate (10 mM) and gently stirred. JC-1 (800 nM; Molecular Probes) was added to measure Δψ_m. Fluorescence signals were monitored in a temperature-controlled (37 °C) multiwavelength-excitation and dual wavelength-emission spectrofluorometer (Delta RAM, Photon Technology International), using 490-nm ex/535-nm em for the monomer and 570-nm ex/595-nm em for the J-aggregate of JC-1. Δψ_m was calculated as the ratio of the fluorescence of the JC-1 oligomeric to monomeric forms.

Measurement of mitochondrial O₂ consumption and ATP levels

The oxygen consumption rate in intact adult LV myocytes was measured at 37°C in an XF96 extracellular flux analyzer (Seahorse Bioscience). Myocytes were seeded in laminin-coated wells and sequentially exposed to oligomycin, FCCP, and rotenone plus antimycin A, using the XF Cell Mito Stress Kit (Seahorse Bioscience) according to the manufacturer’s instructions (52). Preliminary experiments were performed to select optimal seeding density (10⁴ cells/well) and compound concentrations, according to manufacturer’s instructions.

To measure ATP levels, isolated myocytes were lysed and ATP (luminescence) levels were measured using the CellTiter-Glo luminescent cell viability assay kit as described previously (41).

Statistics

All results are expressed as means ± SE. For analysis of I_NaCa and depolarization-activated K⁺ currents as a function of group (control, MB, H₂S, and MB + H₂S) and voltage, 2-way ANOVA was used. For analysis of action potential parameters, contraction amplitudes, ATP levels, ΔΨ_m and OCR, 1-way ANOVA was used. A commercially available software package (JMP version 12, SAS Institute, Cary, NC) was used. In all analyses, P<0.05 was taken to be statistically significant.

Ethics Statement

All protocols and procedures applied to the mice in this study were approved by the Institutional Animal Care and Use Committees of Temple University and The Pennsylvania State University.

Results

Rescue of H₂S-induced myocyte contractile dysfunction by MB: dose response

We have previously demonstrated that MB (20 μg/ml) ameliorated myocyte contractile dysfunction after exposure to H₂S (100 μM)(43). To determine the effective dose and to evaluate potential toxicity of MB, a dose response curve for MB was performed. MB alone at concentrations as high as 500 μg/ml had no negative inotropic effects after 10 min of exposure (Fig. 1). By contrast, myocyte contraction amplitudes decreased by ~50% (p<0.0001, control vs. H₂S) 10 min after addition of H₂S (100 μM)(Fig. 1), in agreement with our previous report (43). MB at 5 (p=0.0004, H₂S vs. H₂S + MB) but not at 1 μg/ml (p=0.2068, H₂S vs. H₂S + MB) significantly improved contractile dysfunction measured in H₂S-treated myocytes, with full therapeutic benefit (p<0.11; control vs. H₂S + MB) achieved at 10 μg/ml of MB (Fig, 1, inset).

Figure 1.

MB rescues contractile dysfunction of cardiomyocytes exposed to H₂S: MB dose response. Freshly isolated myocytes from mouse LV and septum were plated on laminin-coated coverslips, bathed in medium 199 ([Ca²⁺]_o 1.8 mM) and paced (2 Hz) to contract (37°C)(Methods). MB (0 to 500 μg/ml) or NaHS (100 μM) + MB (0 to 500 μg/ml) were added at time 0, and contractions were measured at 10 min. Maximal contraction amplitudes (% of resting cell length, %RCL) are shown for MB (□) and NaHS + MB (●) myocytes. For MB alone, there were 20, 7, 10, 6, 10, 8 and 6 myocytes at 0, 1, 5, 10, 20, 100 and 500 μg/ml, respectively. For NaHS + MB, there were 25, 8, 9, 8, 9, 7, and 8 myocytes at 0, 1, 5, 10, 20, 100 and 500 μg/ml of MB, respectively. Inset: expanded view of contractile response at MB doses from 0 to 20 μg/ml.

Effects of H₂S on action potential: rescue by MB

Various abnormal cardiac rhythms were observed in rat (45) and sheep within minutes of exposure to H₂S (72). These include frequent premature ventricular contractions, bradycardias, and less frequently, ventricular tachycardia and ventricular fibrillation. Since alterations in action potential morphology underlie the cellular basis of increased arrhythmogenesis, AP morphology and parameters in ventricular myocytes exposed to H₂S ± MB were measured. Compared to control myocytes, exposure to H₂S (100 μM) for 7 min resulted in E_m depolarization from −76.1 ± 1.5 to −67.2 ± 1.1 mV (p<0.0015), reduction in AP amplitude from 130.9 ± 3.3 to 110.1 ± 2.5 mV (p<0.0015), and prolongation of APD₉₀ from 43.6 ±3.8 to 81.3 ± 9.1 ms (p=0.005)(Fig. 2). APD₅₀ tended to be prolonged by H₂S although the differences did not reach statistical significance. Addition of MB (20 μg/ml) at 3 min had no significant effect on E_m, AP amplitude, APD₅₀ and APD₉₀ compared to myocytes treated with saline (Fig. 2). Compared to myocytes exposed to H₂S only, MB added 3 min after H₂S addition significantly restored E_m (p<0.01) and APD₉₀ (p<0.02) but not AP amplitude (p<0.37) toward values observed in control myocytes (Fig. 2).

Figure 2.

H₂S depolarizes resting membrane potential (E_m) and prolongs action potential duration (APD): rescue by MB. E_m and APD were measured in myocytes isolated from mouse LV and septum with whole cell patch-clamp. Myocytes were paced at 1 Hz. Pipette solution consisted of (in mM) 125 KCl, 4 MgCl₂, 0.06 CaCl₂, 10 HEPES, 5 K⁺-EGTA, 3 Na₂ATP, and 5 Na₂-creatine phosphate (pH 7.2). External solution consisted of (in mM) 132 NaCl, 5.4 KCl, 1.8 CaCl₂, 1.8 MgCl₂, 0.6 NaH₂PO₄, 7.5 HEPES, 7.5 Na⁺-HEPES, and 5 glucose, pH 7.4. At time 0, either saline or NaHS (100 μM) was added followed by MB (20 μg/ml) or saline at 3 min before action potential was measured at 7 min. Top. Representative action potentials from myocytes treated with saline (control), MB alone, NaHS alone, and NaHS + MB recorded using current-clamp configuration at 1.5x threshold stimulus, 4-ms duration and at 30°C (75, 79, 96, 97). Bottom: Means ± SE of resting E_m, action potential amplitude, action potential duration at 50% (APD₅₀) and at 90% repolarization (APD₉₀) from 5 control, 4 MB, 5 NaHS and 4 NaHS + MB myocytes are shown. * P<0.045, control vs. NaHS or NaHS + MB; ⁺ P<0.02, NaHS vs. NaHS + MB.

Effects of H₂S on I_NaCa and depolarization-activated K⁺ currents: rescue by MB

Action potential duration is predominantly determined by L-type Ca²⁺ current (I_Ca), I_NaCa and depolarization-activated K⁺ currents. We have previously demonstrated that H₂S significantly depressed I_Ca and that MB added 3 min after H₂S exposure was effective in restoring I_Ca towards normal (43). Exposure to H₂S (100 μM) for 7 min significantly (p<0.0001; group × voltage interaction effect) suppressed I_NaCa when compared to control myocytes treated with saline (Fig. 3). Addition of MB 3 min after H₂S exposure significantly (p<0.0001) ameliorated the toxic effects of H₂S on I_NaCa (H₂S vs. H₂S + MB; Fig. 3) although not to the normal control values (p<0.0012; control vs. H₂S + MB).

Figure 3.

H₂S inhibits Na⁺/Ca²⁺ exchanger current (I_NaCa): reversal by MB. Pipette solution contained (in mM) 100 Cs⁺ glutamate, 7.25 NaCl, 1 MgCl₂, 20 HEPES, 2.5 Na₂ATP, 10 EGTA and 6 CaCl₂, pH 7.2. Free Ca²⁺ in the pipette solution was 205 nM, measured fluorimetrically with fura-2. External solution contained (in mM) 130 NaCl, 5 CsCl, 1.2 MgSO₄, 1.2 NaH₂PO₄, 5 CaCl₂, 10 HEPES, 10 Na⁺ HEPES and 10 glucose, pH 7.4. Verapamil (1 μM) was used to block I_Ca,L. Our measurement conditions were biased towards measuring outward (3 Na⁺ out: 1 Ca²⁺ in) I_NaCa. (A). After holding the myocyte at the calculated reversal potential (−73 mV) of I_NaCa for 5 min (to minimize fluxes through Na⁺/Ca²⁺ exchanger and thus allowed [Na⁺]_i and [Ca²⁺]_i to equilibrate with those in pipette solution), I_NaCa (30°C) was measured in myocytes using a descending (from +100 to −120 mV; 500 mV/s) – ascending (from −120 to +100 mV; 500 mV/s) voltage ramp, first in the absence and then in the presence of 1 mM NiCl₂. (B). Raw currents measured in a WT myocyte. I_NaCa was defined as the difference current measured in the absence and presence of Ni⁺ during the descending voltage ramp. Note that with the exception of small contamination of the ascending ramp by the cardiac Na⁺ current, there were little to no differences in I_NaCa measured between the descending and ascending voltage ramps. This suggests that [Ca²⁺]_i and [Na⁺]_i sensed by Na⁺/Ca²⁺ exchanger did not appreciably change by ion fluxes during the brief (880 ms) voltage ramp. I_NaCa was divided by C_m prior to comparisons. (C). At time 0, either saline or NaHS (100 μM) was added followed by MB (20 μg/ml) or saline at 3 min before I_NaCa was measured at 7 min. Current-voltage relationships of I_NaCa (means ± SE) from control (▲; n=6), MB (●; n=3), NaHS (■; n=5) and NaHS + MB (◆; n=5) myocytes are shown. The reversal potential of I_NaCa was ~ −60 mV, close to the theoretical reversal potential of −73 mV. Error bars are not shown if they fall within the boundaries of the symbol.

Peak amplitudes of depolarization-activated K⁺ currents (I_peak) were significantly (p<0.0001; H₂S vs. control, group × voltage interaction effect) depressed by H₂S but were rescued by addition of MB 3 min post H₂S exposure (p<0.0003; H₂S vs. H₂S + MB) (Fig. 4). When depolarization-activated K⁺ currents were segregated into their respective components (68, 92), H₂S consistently decreased I_to,f, I_{K slow}, and I_ss (Fig. 4; p<0.025, control vs. H₂S). MB added 3 min after H₂S exposure restored I_peak, I_to,f, I_K,slow and I_ss towards values measured in control myocytes exposed to saline (Fig. 4; p<0.05, H₂S vs. H₂S + MB). MB alone had no effects on I_peak, I_to,f, I_K,slow and I_ss (Fig. 4; p<0.85; control vs. MB). The time constants of decay (at +40 mV) for I_to,f (p<0.45) and I_K,slow (p<0.40) were not affected by either H₂S or MB.

Figure 4.

H₂S decreases depolarization-activated K⁺ currents: rescue by MB. Depolarization-activated K⁺ currents (30°C, 135 mM [K⁺]_i) were measured in control (□; n=3), MB (▲; n=4), NaHS (◆; n=4) and NaHS + MB (○; n=3) myocytes isolated from the LV free wall (Methods). Top. Raw tracings of depolarization-activated K⁺ currents from control, MB, NaHS and NaHS + MB myocytes. K⁺ currents were separated into 3 components (Methods). Bottom. Current-voltage relationships of peak currents, fast component of transient outward currents (I_to,f), slowly inactivating K⁺ currents (I_K,slow) and steady-state non-inactivating K⁺ currents (I_ss) are shown. Values are means ± SE. Error bars are not shown if they fall within the boundaries of a symbol. Data for K⁺ currents are fitted by linear regression.

Effects of MB on cellular ATP and cardiac mitochondrial respiration

Improved cellular bioenergetics is a potential mechanism by which MB ameliorated contractile dysfunction in myocytes exposed to H₂S (Fig. 1). Indeed, ATP levels were depressed after 10 min of exposure to H₂S but rescued by MB (Fig. 5A). Since ATP is primarily synthesized via oxidative phosphorylation in cardiac myocytes, the effects of MB on mitochondrial function were measured. As shown in Fig. 5B, collapse of mitochondrial membrane potential (Δψ_m) by the Complex I inhibitor rotenone (10 nM) was partially restored by MB (20 μg/ml) although addition of MB alone had no effect on Δψ_m. MB prevented the decrease in mitochondrial oxygen consumption rate (OCR) induced by rotenone and the Complex III inhibitor antimycin A (1 μM)(Fig. 5C). One interesting observation is that OCR was higher in myocytes exposed to MB, regardless of presence or absence of inhibitors of mitochondrial electron transport (Fig. 5C). In the absence of cardiomyocytes, neither MB alone nor MB + NaHS had any significant OCR (data not shown).

Figure 5.

Effects on MB on cardiomyocyte bioenergetics: ATP levels, mitochondrial membrane potential (Δψ_m), and O₂ consumption rate (OCR). (A). ATP (luminescence, relative light units) levels from LV myocytes treated with saline (CNTL, n=8), MB (n=8), NaHS (n=7) and MB + NaHS (n=7) for 10 min were determined with CellTiter-Glo luminescent cell viability kit (Methods). *p<0.0005, CNTL vs. NaHS; #p<0.03; CNTL vs. MB. There are no differences in ATP levels between CNTL and NaHS + MB myocytes (p=0.53). (B). LV myocytes were permeabilized with digitonin and supplemented with succinate. Left: the ratiometric indicator JC-1 was added at 20s and used to monitor Δψ_m. Arrows indicate addition of JC-1 and the mitochondrial uncoupler CCCP (2 μM), respectively. Rotenone (10 nM) and/or MB (20 μg/ml) were added at time 0. Right: Summary of Δψ_m before CCCP addition (n=3 each). **p<0.01; ns, not significant. (C). OCR was measured in intact myocytes (Methods). After basal OCR was obtained, either saline control (●), MB alone (20 μg/ml; ○), rotenone (1 μM; □), or MB (20 μg/ml) + rotenone (■) were added (indicated by “Compound”). At times indicated, oligomycin (1 μM) was added to inhibit F₀F₁ATPase (Complex V) followed by addition of the uncoupler FCCP (1 μM) to measure maximal OCR. Finally, antimycin A + rotenone (1 μM each) were added to inhibit cytochrome bc1 complex (Complex III) and NADH dehydrogenase (Complex I), respectively. Each point in the traces represents the average of 8 different wells. This experiment was repeated 3 times with similar results.

Discussion

The major finding of the present study is that H₂S greatly affected the electrophysiology of isolated cardiomyocytes beyond the inhibition of L-type Ca²⁺ currents (I_Ca,L) that we and others have previously reported (43, 74, 93). A significant alteration of Na⁺/Ca²⁺ exchange and depolarization-activated K⁺ currents was effected by toxic level of H₂S, i.e., at concentrations that in vivo lead to lethal cardiogenic shock and several types of arrhythmias (73). MB was capable of restoring, for most part, normal cardiomyocyte contractile and electrophysiological activity.

On the use of high μM H₂S in solution to study toxicity in isolated cardiac myocytes

Typically, whenever solutions of H₂S at concentrations ≥ 50 μM were used in isolated hearts or isolated cells, the effect was a measurable depression in cardiac contractility (43, 74). This depression is an obvious pathological and toxic change in the function at the level of both individual myocytes and the intact heart. In vitro, the activity of the mitochondrial cytochrome c oxidase is abolished by a solution of H₂S at concentrations of H₂S/HS⁻ ranging from 10 to 30 μM (17, 49). In vivo, severe depression in cardiac contractility can be produced in rodents and in large mammals by infusing or inhaling H₂S at levels yielding blood concentrations of gaseous H₂S between 2–5 μM (38, 46, 71, 73), corresponding to level of total dissolved/free sulfide of < 20 μM. In a recent study, we found that the ejection fraction of the left ventricle started to decrease for an average level of gaseous H₂S of 3 μM, while blood pressure was decreased ~ 5 μM. This corresponds to a total level dissolved H₂S/HS⁻ of about 15–20 μM (assuming that 2/3 of total dissolved sulfide present is in the form of HS⁻ at physiological pH). Therefore, the H₂S concentration used in our present study was consistent with previous studies to produce toxic effects on the cardiomyocyte.

Early studies reported H₂S concentrations in the blood or in the tissues in the high μM range and was assumed to be due to endogenous H₂S production only (24). Subsequent work (50, 88) demonstrated that the high μM concentrations of free/soluble H₂S are unrealistically high by several orders of magnitude. Two main hypotheses have been advanced to explain why high μM levels of “endogenous” H₂S were initially (and are still) reported in the blood and tissues. The first reason relates to the nature of the pools of sulfide present in the blood and in tissues found in post-mortem conditions. No relevant (or trivial) levels of free/dissolved H₂S, which should smell like rotten eggs, can be found in the blood, the brain (24, 50) or heart homogenates (50). However, significant amount of H₂S can be mobilized after exposing these tissues to a strong acid (lowering the pH of the tissues to <2) and can be measured after evaporating from the brain or the heart (50, 55, 80). This pool of H₂S represents several micromoles of sulfide per kg of tissue (26, 42, 50, 55, 80), but is “trapped”, literally fossilized, in the form of metallo-sulfides (including Fe²⁺, Zn²⁺, Co²⁺, etc.). Similarly, strong reducing agents can release H₂S from a solution of proteins or from various tissues, where H₂S is present under the form protein-bound thiols (55, 85). The presence of H₂S evaporating after such reactions should not be confused with the presence of dissolved H₂S, able to freely diffuse and to potentially act as a gaseous-transmitter. We have discussed the complex issues regarding solutions containing H₂S (prepared from NaHS) and H₂S in tissues in a recent review (33).

H₂S poisoning induced cardiac toxicity

H₂S is commonly referred to as a mitochondrial Complex IV inhibitor, mimicking the effects of cyanide or sodium azide, by impeding the activity of the mitochondrial cytochrome C oxidase (17, 19, 45). Our results that H₂S inhibited various cardiac ionic currents (I_Ca,L (43), I_NaCa and depolarization-activated K⁺ currents) suggest that blockade of the mitochondrial cytochrome C oxidase may not be the sole mechanism of sulfide toxicity. H₂S depressed the rapidly activating and inactivating transient outward current (I_to,f), the rapidly activating but slowly inactivating delayed rectifier K⁺ current (I_K,_slow), and the fast activating and non-inactivating steady-state K⁺ current (I_ss). In addition, the depolarized resting membrane potential and reduced action potential amplitude in ventricular cells exposed to H₂S are consistent with suppression of the inwardly rectifying Kir currents (I_K1) and fast Na⁺ current I_Na, respectively. The effects of H₂S on APD₅₀ are complicated since it suppresses both I_Ca,L (shortens APD₅₀) and I_to,f (prolongs APD₅₀). Our observation that H₂S tended to prolong APD₅₀ suggests that its effects on I_to,f are dominant. Similar considerations apply to the toxic effects of H₂S on I_NaCa (shortens APD₉₀) and I_K,slow (prolongs APD₉₀). Our observation that APD₉₀ was significantly prolonged supports the view that the effects of H₂S on depolarization-activated K⁺ currents on APD₉₀ are dominant. Wei and al. (86), using cardiomyocytes of ventricular, atrial and nodal subtypes differentiated from H9 embryonic stem cells and human induced pluripotent stem cells, have reported similar effects of high concentrations of H₂S (100 to 300 μM) on APD, I_Ca,L, I_K,slow, the rapidly activating and inactivating delayed rectifier K⁺ current (I_K,rapid), and the hyperpolarization-activated inward current (I_f) found in sinoatrial nodal cells.

Since one of the primary functions of Na⁺/Ca²⁺ exchanger in adult cardiomyocytes is to extrude during diastole the amount of Ca²⁺ that has entered during systole via the L-type Ca²⁺ channel, thereby maintaining steady-state beat-to-beat Ca²⁺ balance (9), inhibition of Na⁺/Ca²⁺ exchanger by H₂S would be expected to result in Ca²⁺ overload and progressive elevation of diastolic [Ca²⁺]_i. However, diastolic [Ca²⁺]_i was similar between control and H₂S-toxic myocytes (43). Absence of overt Ca²⁺ overload in H₂S intoxicated cardiomyocytes is likely due to the simultaneous inhibition of L-type Ca²⁺ channels by H₂S, thereby limiting Ca²⁺ influx during systole and maintaining steady-state Ca²⁺ balance. This parallels the situation in adult cardiac-specific Na⁺/Ca²⁺ exchanger knockout ventricular myocytes in which L-type Ca²⁺ channel activity was substantially reduced when compared to wild-type myocytes (39).

The mechanisms by which most channels are affected at the same time remain undetermined. As recently proposed for Ca²⁺ channels, H₂S could react with cysteine residues not only in L-type Ca²⁺ channels (74, 93), but also the sarcoplasmic reticulum ryanodine receptors (RyR), thereby altering the 3D configurations and function of the channels. Whether other cardiac channels can be affected through a similar mechanism remains unknown. H₂S induced inhibition of the mitochondrial electron transport chain has been shown to result in an increase in ROS production (12) which could also contribute to this effect (99) through alterations in the redox environment of these ion channels.

MB as a treatment of H₂S intoxication

H₂S is the second most common cause of death by gas exposure in the workplace after carbon monoxide (8, 23, 28, 57). H₂S is a primary chemical hazard in oil and gas production, well drilling and gas refining industries (3, 21), and a significant occupational hazard during various farming activities (14). H₂S can be “weaponized”, as demonstrated by British troops during World War I (22). It is also used as a method of suicide (11). This form of suicide has increased in an alarming manner in the US (30) and is accomplished by mixing a source of sulfide and various types of acidic solutions readily available in most household chemicals. This has created major challenges for first responders (56, 78).

As soon as it enters the blood, only a very small portion of sulfide remains in a “free/soluble” or diffusible form, comprising the gaseous form H₂S (7, 13, 18, 20) and the sulfhydryl anion HS⁻ (2, 53). A larger pool of sulfide will combine with metallo-proteins such as the ferrous iron in hemoglobin or will react with cysteine residues present in proteins (29, 63, 66, 90), creating a large sink for H₂S. While free/soluble H₂S is the only form able to diffuse into the cells, bound H₂S represents the toxic pool (38, 46). The most remarkable feature of H₂S metabolism is that H₂S disappears from the blood and the tissues (29, 46, 76) at a very rapid rate. Indeed H₂S is almost immediately oxidized in the mitochondria, a reaction catalyzed by various enzymes such as the sulfide quinone oxido-reductase, sulfur dioxigenase and the sulfur transferase enzyme rhodenase (12, 48, 49, 54). We found that this oxidation can account for the disappearance of soluble H₂S following sulfide poisoning, within one minute in large and small mammals (38, 46), while the form “hidden” in disulfide bonds persists for a much longer period of time (46) in the tissues (85). As a consequence, the toxic effects of H₂S persist beyond the phase of exposure without being accessible to antidotes, which primarily act on the soluble form.

The challenge is therefore to find compounds antagonizing the effects of H₂S toxicity while the pool of exchangeable sulfide is already gone. The treatment of H₂S poisoning has been traditionally aimed at trapping free H₂S using metallo-compounds, e.g. ferric iron contained in methemoglobin (15, 36, 81) or cobalt in hydroxycobalamin (HyCo) (35, 62, 64, 77, 81). Other antidotes are based on empirical observations, such as sodium bicarbonate (1, 27) and hyperoxia (10, 34, 60, 67) with no proven efficacy. Nitrite-induced methemoglobinemia (47, 61–63, 66, 81) however can only trap H₂S outside the cells and has little or no effects on the combined forms after exposure (38). Sodium nitrite further decreases arterial blood pressure in subjects already in shock (31, 32) and affects oxygen transport (4, 65, 66). Cobalt contained in HyCo (81) has several theoretical advantages over methemoglobinemia (51, 77). Whether this very large molecule can penetrate cells at sufficient concentrations and rapidly enough (5) to neutralize the effects of H₂S remains to be demonstrated.

Although methylene blue (MB) is an old compound (16, 25), its use in H₂S intoxication represents a novel and very promising paradigm, as it counteracts the consequences of H₂S intoxication. Our data demonstrate that MB is effective even when given after H₂S exposure provide support for its use as an emergency treatment in the most severe forms of H₂S-induced coma with shock but also in non-life threatening conditions.

CONCLUSIONS

MB ameliorated changes in E_m, APD₉₀, I_NaCa and depolarization-activated K⁺ currents in cardiac myocytes exposed to H₂S. In addition, MB restored depressed ATP levels after H₂S exposure. Our data support the hypothesis that MB exerts a potent antidotal effect during H₂S intoxication through restoration of cardiac function depressed by sulfide. The redox properties of the couple MB/LMB may exert widespread effects ranging from modification of the activity of metallo-proteins or sulfhydrated proteins, to the alteration of essential metabolic pathways.