Droperidol Inhibits Intracellular Ca²⁺, Myofilament Ca²⁺Sensitivity, and Contraction in Rat Ventricular Myocytes

All experimental procedures and protocols were approved by the Cleveland Clinic Institutional Animal Care and Use Committee (Cleveland, Ohio) and conformed to the guidelines for the care and use of laboratory animals.

Ventricular myocytes were freshly isolated from adult male Sprague-Dawley rat hearts, as previously described.7Immediately after the animals were killed, the hearts were rapidly removed and perfused in a retrograde manner at a constant flow rate (8 ml/min) with oxygenated (95% O²–5% CO²) Krebs-Henseleit buffer (KHB; 37°C) containing the following: 118 mm NaCl, 4.8 mm KCl, 1.2 mm MgCl², 1.2 mm KH²PO⁴, 1.2 mm CaCl², 37.5 mm NaHCO³, and 16.5 mm dextrose, at a pH of 7.35. After a 5-min equilibration period, the perfusion buffer was changed to a Ca²⁺-free KHB buffer containing collagenase type II (309 U/ml; Worthington Biochemical Corp., Freehold, NJ). After digestion with collagenase (20 min), the ventricles were minced and shaken in KHB, and the resulting cellular digest was washed, filtered, and resuspended in phosphate-free HEPES-buffered saline (HBS; 23°C) containing the following: 118 mm NaCl, 4.8 mm KCl, 1.2 mm MgCl², 1.25 mm CaCl², 11.0 mm dextrose, 25.0 mm HEPES, and 5.0 mm pyruvate, at a pH of 7.35.

Simultaneous measurement of [Ca²⁺]ⁱand cell shortening was performed, as previously described by our laboratory.7Ventricular myocytes exhibiting a rod-shaped appearance with clear striations were chosen for study. Myocytes (0.5 × 10⁶cells/ml) were incubated in HBS containing 2 μm fura-2/acetoxy methylester at room temperature for 15 min. Fura-2–loaded ventricular myocytes were placed in a temperature-regulated (30°C) chamber (Bioptechs, Inc., Butler, PA) mounted on the stage of an Olympus IX-70 inverted fluorescence microscope (Olympus America, Lake Success, NY). The cells were superfused continuously with HBS at a flow rate of 2 ml/min and field-stimulated via  bipolar platinum electrodes at a frequency of 0.3 Hz with a 5-ms pulse using a Grass SD9 stimulator (Grass Instruments, West Warwick, RI).

Fluorescence measurements were performed on individual myocytes using a dual-wavelength spectrofluorometer (Deltascan RFK6002; Photon Technology International, Lawrenceville, NJ) at excitation wavelengths of 340 and 380 nm and an emission wavelength of 510 nm. Because calibration procedures rely on a number of assumptions, the ratio of the light intensities at the two wavelengths was used to measure qualitative changes in [Ca²⁺]ⁱ. Just before data acquisition, background fluorescence was measured and automatically subtracted from the subsequent experimental measurement. The fluorescence sampling frequency was 100 Hz, and data were collected using software from Photon Technology International.

To simultaneously monitor cell shortening, the cells were also illuminated with red light. A dichroic mirror (600-nm cutoff) in the emission path deflected the cell image through a charge-coupled device video camera (Phillips VC 62505T; Marshall Electronics, Culver City, CA) into a video-edge detector (Crescent Electronics, Sandy, UT) with 16-ms resolution. The video-edge detector was calibrated using a stage micrometer so that cell lengths during shortening and relengthening could be measured.

Measurement of pHⁱwas performed, as previously described.8Ventricular myocytes were incubated in HBS containing 2 μm of the acetoxy methylester form of 2′,7′-bis(2-carboxy-ethyl)-5,6-carboxyfluorescein (BCECF/AM) at 23°C for 20 min. Similar to the procedure described for fura-2–loaded myocytes, BCECF-loaded ventricular myocytes were placed in a temperature-regulated (30°C) chamber mounted on the stage of an Olympus IX-70 inverted fluorescence microscope. The volume of the chamber was 1.5 ml. The cells were superfused continuously with HBS at a flow rate of 2 ml/min (30°C). Fluorescence measurements were performed on single ventricular myocytes as described above for [Ca²⁺]ⁱ; however, excitation wavelengths of 440 and 500 nm and an emission wavelength of 530 nm were used, as previously described.8The fluorescence sampling frequency was 10 Hz, and background fluorescence was determined as described above. To estimate the pHⁱvalue from the ratio of 500/440 nm fluorescence, we used an in situ  calibration procedure.8–10At the end of each experiment, the fluorescence ratio from each cell was calibrated in situ  by exposing the cell to solutions of varying pH. Each solution contained 140 mm KCl, 1.0 mm MgCl², 4.0 mm HEPES, 2.0 mm EGTA, 30 mm 2,3-butanedione monoxime, 50 μm BAPTA-AM, and 14 μm nigericin and was titrated to varying pH values (6.6, 6.8, 7.0, 7.2, 7.4, 7.6, 7.8) using KOH (1.0 N). The pHⁱfor each cell was then determined from a linear regression of the fluorescence ratio versus  the pH value of the calibration buffer. We previously determined that a linear relation exists between the 500/440-nm ratio and pHⁱin the physiologic range (pH 6.6–7.8).8 

The following variables were calculated for each individual contraction: resting [Ca²⁺]ⁱand cell length, peak [Ca²⁺]ⁱand cell length, change in [Ca²⁺]ⁱ(peak [Ca²⁺]ⁱminus resting [Ca²⁺]ⁱ) and twitch amplitude, time to peak (Tp) for [Ca²⁺]ⁱand shortening, and time to 50% (Tr) resting [Ca²⁺]ⁱand relengthening. Variables from 10 contractions were averaged to obtain mean values at baseline and in response to the various interventions. Averaging the variables over time minimizes beat-to-beat variation.

Only rod-shaped, quiescent cells with smooth striations were selected. The whole cell configuration was achieved with fire-polished and Sylgard-coated tipped glass pipettes (Corning G86165T-4; World Precision Instruments, Sarasota, FL) and a resistance of 2–3 mV when filled with pipette solution containing 135 mm KCl, 1 mm MgCl², 10 mm EGTA, 10 mm HEPES, and 5 mm glucose, at a pH of 7.3. The bathing solution contained the following: 140 mm NaCl, 4 mm KCl, 2 mm CaCl², 2 mm MgCl², 10 mm HEPES, and 10 mm glucose, at a pH of 7.3. Using the Axopatch 1C (Axon Instruments, Foster City, CA), series resistance was 4–8 mV and was 30–80% electronically compensated. Action potentials were evoked by repetitive square pulses of 3-ms duration at 1.5× stimulus threshold. Experiments were performed under constant flow conditions at 30°C using a temperature-controlled experimental chamber (Delta T Culture Dish; Bioptechs, Butler, PA). Myocytes were paced under current clamp conditions at a cycle duration of 1,000 ms. Runs of 12 steady state action potentials were recorded with filtering at 2 kHz and sampled at 10 kHz. Data were acquired using a Pentium computer that controlled data acquisition hardware and software (pClamp 6.03; Axon Instruments). Analysis of action potentials was performed by taking the average of each sweep of 12 stable and steady state records.

Nitric oxide production was assessed by measurement of nitrate/nitrite production using a colorimetric kit from Cayman Chemical (Ann Arbor, MI). Suspensions of cardiomyocytes were placed in wells and incubated in the presence or absence of droperidol (0.1, 0.3, 1 μm) at 30°C for 15 min. Total nitrite levels (after conversion of nitrate to nitrite) were determined with Griess reagent using a microplate reader (absorbance, 540 nm). Aliquots (150 μl) from each well (before and after addition of droperidol) were collected and mixed with an equal volume of Griess reagent (1% sulfanilic acid and 0.1% N -(1-napthyl) ethylenediamine dihydrochloride in 2% phosphoric acid). The mixture was incubated at 20°C for 10 min. Nitrite concentrations in the samples were determined based on standard calibration curves by using an aqueous solution of sodium nitrite. The background value from buffer alone was subtracted from the experimental value.

Male Sprague-Dawley rats weighing 250–300 g, were given an intraperitoneal injection of heparin (200 U). After the animals were killed, hearts were excised rapidly and placed in ice-cold KHB before being mounted on a Langendorff apparatus for perfusion at 37°C with KHB at 330 beats/min and a constant pressure of 65– 70 mmHg. The buffer was equilibrated with 95% O²and 5% CO²and had the following composition: 118 mm NaCl, 4.7 mm KCl, 1.25 mm CaCl², 1.2 mm KH²PO⁴, 1.2 mm MgSO⁴, 25 mm NaHCO³, and 11 mm dextrose. A balloon-tipped catheter was inserted through the left atrium into the left ventricle, and the left ventricular end-diastolic pressure in all hearts was adjusted to between 8 and 12 mmHg. Left ventricular developed pressure was monitored continuously throughout the experiment. Coronary blood flow was measured by timed collection of the effluent into a graduated cylinder.

All experimental protocols were performed at 30°C, with the exception of the perfused Langendorff heart experiments performed at 37°C.

A stock solution of droperidol was obtained by dissolving the drug in dimethyl sulfoxide. Baseline measurements were collected from individual myocytes for 1.5 min in the absence of any intervention. Myocytes were exposed to four concentrations of droperidol (0.03, 0.1, 0.3, and 1 μm) by exchanging the buffer in the dish with new buffer containing droperidol at the desired concentration. Data were acquired for 1.5 min after a 5-min equilibration period in the presence of droperidol. Summarized results for the concentration–response curves are expressed as percent of the control value. Dimethyl sulfoxide (0.05% vol/vol) alone has no effect on [Ca²⁺]ⁱor shortening.7We verified that droperidol had no effect on fura-2 fluorescence at the concentrations tested. This was confirmed in separate cell-free experiments using fura-2 (pentapotassium salt) in buffers ranging from pCa (−log Ca²⁺concentration) 9 to pCa 5 in the presence or absence of droperidol (data not shown).

Droperidol was added to the superfusion medium for 15 min, and then KCl (35 mm) was applied to the cell. This concentration of KCl was chosen because it stimulates an increase in [Ca²⁺]ⁱapproximately 50% of the maximum response. Peak [Ca²⁺]ⁱinduced by KCl was compared with peak [Ca²⁺]ⁱbefore addition of droperidol (1 μm) and is reported as percent change from control.

Action potentials were recorded before and after addition of droperidol (1 μm) in individual cardiomyocytes. Changes in action potential duration at 90% repolarization were determined.

Baseline values for [Ca²⁺]ⁱwere measured in individual, field-stimulated myocytes for 1.5 min. Droperidol (0.1 and 1 μm) was then added to the superfusion buffer and allowed to perfuse the cell for 5 min. Field stimulation of the myocyte was then discontinued, and caffeine (20 mm) was applied to the cell 15 s later in the continued presence of droperidol. Peak [Ca²⁺]ⁱinduced by caffeine was compared with peak [Ca²⁺]ⁱbefore addition of droperidol and as a percent increase above the peak [Ca²⁺]ⁱachieved during field stimulation (fractional release).

Changes in the extracellular Ca²⁺concentration ([Ca²⁺]^o)–shortening relation (indirect measurement of myofilament Ca²⁺sensitivity), were examined as previously described.10,11Baseline variables were collected from individual myocytes for 1.5 min. Concentration–response curves for [Ca²⁺]^owere performed by exchanging the buffer in the dish with a new buffer containing the desired [Ca²⁺]^o. Data were acquired for 1.5 min after establishment of a new steady state. Concentration–response curves for [Ca²⁺]^owere then performed in the presence of droperidol (1 μm). Cells were allowed to stabilize for 5 min after addition of droperidol.

Baseline pHⁱwas collected from individual myocytes for 1 min. Droperidol (0.03, 0.1, 0.3, and 1 μm) was added by exchanging the superfusion buffer in the dish with new buffer containing droperidol at the desired concentration. Each myocyte was exposed to only one concentration of droperidol. Results are expressed as the change in pHⁱover time with each concentration of droperidol.

Suspensions of ventricular myocyte were exposed to droperidol (0.1, 0.3, and 1 μm) for 15 min at 37°C in HBS buffer. Total nitrite/nitrate production was measured as an indicator of nitric oxide production. Results are expressed as percent change in total nitrate/nitrite from baseline.

Left ventricular pressure was continuously monitored in perfused Langendorff hearts paced at 330 beats/min at 37°C before (20 min) and during administration of droperidol (1 μm, 20 min). Results are expressed as percent change in pulse pressure (left ventricular end systolic pressure minus end diastolic pressure) from baseline.

Each experimental protocol was performed on multiple myocytes from the same heart and repeated in five hearts. Results obtained from myocytes in each heart were averaged so that all hearts were weighted equally. Comparison of several means was performed using repeated measures and two-way analysis of variance.12The Bonferroni post hoc  test was used when significant differences among groups were detected. Differences were considered statistically significant at P < 0.05. All results are expressed as mean ± SD.

Droperidol, caffeine, and dimethyl sulfoxide were purchased from Sigma Chemical Co. (St. Louis, MO). Collagenase was purchased from Worthington Biochemical Corp. (Lakewood, NJ). BCECF/AM and fura-2/AM were purchased from Texas Fluorescence Labs (Austin, TX). The nitrate/nitrite colorimetric assay kit was obtained from Cayman Chemical.

Resting cell length was 125 ± 5 μm, and the baseline 340/380 ratio was 0.8 ± 0.1. Twitch height was 12 ± 1.5 μm (10.4 ± 1.6% of the resting cell length). The change in 340/380 ratio from baseline with shortening was 0.5 ± 0.1. Tp [Ca²⁺]ⁱand shortening were 151 ± 22 and 177 ± 16 ms, respectively. Times to 50% recovery (Tr) for [Ca²⁺]ⁱand shortening were 192 ± 19 and 228 ± 21 ms, respectively.

A representative trace depicting the concentration-dependent effects of droperidol on [Ca²⁺]ⁱand shortening in a single, field-stimulated ventricular myocyte is shown in figure 1A. Droperidol (1 μm) reduced peak [Ca²⁺]ⁱand shortening by 22 ± 4 and 43 ± 5%, respectively. An increase in resting cell length of 2 ± 0.6 μm with no change in resting [Ca²⁺]ⁱwas observed in most cells. The myocardial depressant effects of droperidol were reversible after washout. Summarized data for the concentration-dependent effects of droperidol on [Ca²⁺]ⁱand shortening are also shown in figure 1B. Droperidol caused concentration-dependent decreases in [Ca²⁺]ⁱand shortening. Figure 2represents an exploded overlay view of the individual [Ca²⁺] transient and shortening event before (A and B) and after being normalized to peak height (C and D) in the presence or absence of droperidol (1 μm) to illustrate changes in timing. Droperidol (1 μm) had no effect on Tp [Ca²⁺]ⁱ(98 ± 5% of control), Tp shortening (94 ± 6% of control), Tr [Ca²⁺]ⁱ(96 ± 8% of control), or Tr shortening (102 ± 3% of control). The continuous [Ca²⁺]ⁱ–shortening relations are also depicted as hysteresis loops in figure 2E. Droperidol caused a marked concentration-dependent downward shift in the continuous [Ca²⁺]ⁱ–shortening relation.

Addition of KCl (35 mm) to quiescent myocytes resulted in a sustained increase in [Ca²⁺]ⁱ(fig. 3A). Pretreatment with droperidol (0.1 and 1 μm) attenuated the KCl induced-increase in [Ca²⁺]ⁱby 24 ± 5 and 27 ± 5%, respectively (fig. 3B).

Figure 4Ashows that exposure of the myocyte to droperidol (1 μm) had no effect on action potential duration compared with control myocytes not exposed to droperidol. Summarized data depicting the effect of droperidol on action potential duration at 90% repolarization are also shown in figure 4B.

Although the peak [Ca²⁺]ⁱachieved with caffeine was reduced by 22 ± 5% in the presence of droperidol (0.1 μm) compared with control, the fractional release of Ca²⁺from the SR was not different from that observed in the absence of droperidol (fig. 5A). Summarized data for the effects of droperidol on SR Ca²⁺content and fractional release are depicted in figure 5B. The decrease in SR Ca²⁺content is likely due to the inhibitory effect of droperidol on the L-type Ca²⁺channel, which reduces the driving force for refilling the SR.

Figure 6demonstrates that increasing [Ca²⁺]^ofrom 1 to 4 mm (control, without droperidol) resulted in a concentration-dependent increase in shortening (A) and a concomitant increase in peak [Ca²⁺]ⁱ(B). Droperidol (0.3 μm) caused a significant downward shift in the concentration–response curve to increasing [Ca²⁺]^ofor shortening, with no concomitant effect on peak [Ca²⁺]ⁱ.

Baseline pHⁱwas 7.11 ± 0.04. Summarized data depicting the effect of droperidol on pHⁱare shown in figure 7. Droperidol at 0.1, 0.3, and 1 μm caused concentration- and time-dependent decreases in pHⁱ. The effects of droperidol on pHⁱwere reversible after washout with HBS (pHⁱ= 7.08 ± 0.04). There was no significant change in extracellular pH in the presence of 1 μm droperidol (control: 7.35 ± 0.13, droperidol: 7.34 ± 0.11; not significant).

Droperidol at concentrations of 0.1, 0.3, and 1 μm increased nitric oxide production by 22 ± 6, 53 ± 7, and 74 ± 10% compared with control. The nitric oxide donor, S -nitroso-N -acetylpenicillamine (10 μm), increased nitric oxide by 128 ± 12% compared with control.

Addition of droperidol (0.1, 1 μm) to the perfusate resulted in concentration-dependent decreases in pulse pressure of 29 ± 6 and 43 ± 8%, respectively. There were no significant changes in coronary blood flow after perfusion with 1 μm droperidol (18 ± 1.4 ml/min before droperidol vs.  16 ± 1.8 ml/min after droperidol).

The major findings of this study are that droperidol causes a decrease in cardiomyocyte contractility via  a decrease in [Ca²⁺]ⁱand a decrease in myofilament Ca²⁺sensitivity. The decrease in [Ca²⁺]ⁱis due to a decrease in sarcolemmal Ca²⁺influx, whereas the decrease in myofilament Ca²⁺sensitivity is likely mediated by a decrease in cardiomyocyte pHⁱ, an increase in nitric oxide production, or both.

There are no previous studies that have evaluated the effects of droperidol on ventricular cardiomyocyte [Ca²⁺]ⁱand contractility. However, droperidol has been shown to slow pacemaker activity and depress maximum velocity of contraction in guinea pig ventricular muscle, although the cellular mechanisms for these effects were not investigated.13,14In the current study, we observed that clinically relevant concentrations of droperidol inhibit the peak [Ca²⁺]ⁱachieved in response to field stimulation in a concentration-dependent manner. In addition, droperidol caused an increase in resting cell length in the absence of any change in diastolic [Ca²⁺]ⁱ, suggesting a decrease in myofilament Ca²⁺sensitivity. Taken together, these data suggest that droperidol exerts its effects on cellular mechanisms that regulate [Ca²⁺]ⁱand myofilament Ca²⁺sensitivity. Because there were no changes in the timing variables of [Ca²⁺]ⁱ, which would have been reflected in the shortening and relengthening of the myocyte, it is unlikely that droperidol has any effect on the Na⁺–Ca²⁺exchanger or the SR Ca²⁺pump. Therefore, we investigated the effects of droperidol on Ca²⁺influx via  voltage-gated Ca²⁺channels.

The increase in cardiomyocyte [Ca²⁺]ⁱafter addition of KCl is known to result from a depolarization-induced activation of the L-type Ca²⁺channel. We observed a decrease in the KCl-induced increase in [Ca²⁺]ⁱin myocytes pretreated with droperidol. These findings suggest that droperidol may have a direct inhibitory effect on L-type Ca²⁺channels. Alternatively, a droperidol-induced activation of the transient outward K⁺current or the delayed rectifier K⁺current could abbreviate action potential duration and thereby reduce the time in which Ca²⁺enters the cell via  L-type Ca²⁺channels. Therefore, we examined the effect of droperidol on action potential duration.

Previous studies have demonstrated a prolongation in action potential duration and the presence of early afterdepolarizations with low concentrations of droperidol (10–300 nm), whereas a shortening of action potential duration was observed with high concentrations (10– 30 μm).13,15The prolongation in action potential duration was due to a droperidol-induced inhibition of the delayed rectifier K⁺current.15Droperidol has been reported to cause a concentration-dependent prolongation of the QT interval in humans1and subsequent torsade de pointes, which can progress to ventricular fibrillation and sudden cardiac death.2In the current study using rat ventricular myocytes, we did not observe a prolongation in action potential duration with any concentration of droperidol. This may be because rat ventricular myocytes lack a prominent delayed rectifier K⁺current and primarily rely on the transient outward K⁺current for repolarization. Therefore, we conclude that the inhibitory effect of droperidol on the increase in [Ca²⁺]ⁱinduced by electrical field stimulation or KCl is not due to an indirect effect of droperidol on action potential duration.

Ca²⁺-induced Ca²⁺release is the process by which influx of Ca²⁺through sarcolemmal L-type Ca²⁺channels triggers Ca²⁺release from the SR through an activation of the ryanodine receptor (Ca²⁺release channel) in cardiomyocytes.16Rapid application of caffeine to quiescent myocytes results in direct activation of the ryanodine receptor on the SR, triggering the release of Ca²⁺from the SR. The difference between the peak [Ca²⁺]ⁱinduced by electrical stimulation and the caffeine-induced increase in [Ca²⁺]ⁱrepresents the fractional release of Ca²⁺from the SR and is used to assess whether interventions have a direct effect on SR Ca²⁺content or an indirect effect due to decreased sarcolemmal Ca²⁺influx. Our results indicate that although the peak increase in [Ca²⁺]ⁱin response to caffeine is attenuated by droperidol, the fractional release of Ca²⁺is unaltered. This is because droperidol inhibits sarcolemmal Ca²⁺influx through the L-type Ca²⁺channels, resulting in an overall decrease in the size of the releasable pool of Ca²⁺in the SR. Therefore, it seems that droperidol does not exert a direct inhibitory effect on SR Ca²⁺release but does have an indirect effect on the size of the releasable pool of Ca²⁺.

We hypothesized that a decrease in myofilament Ca²⁺sensitivity may also play a role in the inhibitory effects of droperidol on contractility, based on our observations of a droperidol-induced increase in resting cell length and a downward shift in the [Ca²⁺]ⁱ–shortening relation. Although changes in sensitivity can also be reflected in the timing variables of contraction, we did not observe a change in the timing variables. This indicates that droperidol likely exerts its effects at multiple sites of regulation for myofilament Ca²⁺sensitivity, which may offset or mask the effect of one another. Because alterations in myofilament Ca²⁺sensitivity can alter contractility,17we examined whether droperidol altered the concentration–response relation to [Ca²⁺]^o. This protocol is an indirect assessment of myofilament Ca²⁺sensitivity that allows for a paired comparison of [Ca²⁺]ⁱand contractile amplitude in the same cell in the presence or absence of droperidol over a range of values for [Ca²⁺]^o. Droperidol caused a downward shift in the concentration–response curve to [Ca²⁺]^ofor shortening, with no concomitant effect on [Ca²⁺]ⁱ. These data suggest that droperidol decreases the maximal response of the myofilament to Ca²⁺as [Ca²⁺]ⁱincreases. Therefore, it seems that in addition to a decrease in sarcolemmal Ca²⁺influx, a droperidol-induced decrease in myofilament Ca²⁺sensitivity contributes to the inhibitory effect of droperidol on cardiomyocyte contractility. We next assessed potential cellular mechanisms that may be responsible for the decrease in myofilament Ca²⁺sensitivity.

One possible mechanism for a decrease in myofilament Ca²⁺sensitivity is intracellular acidification. It is well known that intracellular acidosis decreases the contractility of cardiac muscle,17although the mechanisms responsible for the decrease are complicated. Acidosis affects every step of the excitation–contraction coupling pathway, including the availability and delivery of Ca²⁺to the myofilaments, as well as the response of the myofilaments to Ca²⁺.18–21Droperidol decreased pHⁱin a time- and concentration-dependent manner in cardiomyocytes. These data suggest that droperidol may have an inhibitory effect on the Na⁺–  H⁺exchanger, which would promote accumulation of H⁺in the cytoplasm, resulting in intracellular acidosis. Because these studies were conducted in a HBS buffer in the absence of carbon dioxide and bicarbonate, a possible interaction between droperidol and Na⁺–  HCO³⁻symport or Cl⁻–HCO³⁻exchange is unlikely, because these transport systems are inactive in myocytes bathed in HCO³⁻-free solution.22,23The magnitude of the change in pHⁱin response to droperidol (0.1 μm) was similar in magnitude, although opposite in direction, to that previously reported by our laboratory for thiopental- or propofol-induced changes in pHⁱand myofilament Ca²⁺sensitivity.8,10,24Moreover, other studies have documented similar changes in pHⁱfor phenylephrine- and endothelin-induced intracellular alkalosis, respectively, resulting in a positive inotropic response.25,26It is also possible that a droperidol-induced decrease in pHⁱcontributes to the inhibitory effect of droperidol on the KCl-induced increase in [Ca²⁺]ⁱ. Further studies are required to confirm this possibility.

An increase in cardiomyocyte nitric oxide has been shown to decrease myofilament Ca²⁺sensitivity, resulting from an alteration in troponin I phosphorylation27and/or an alteration in actin–myosin cross-bridge cycling by modulating critical thiols on the myosin head.28In addition, nitric oxide is known to activate the cyclic guanosine monophosphate signaling pathway, which has been shown to reduce the myofilament response to Ca²⁺in cardiac myocytes29but augment release of Ca²⁺from the SR by caffeine.30Droperidol caused a concentration-dependent increase in nitric oxide production in cardiomyocytes. The increase in nitric oxide production, as well as the decrease in pHⁱ, are likely involved in the droperidol-induced decrease in myofilament Ca²⁺sensitivity. However, direct confirmation of this hypothesis and the relative roles of each will require additional studies.

To identify whether the changes in cardiomyocyte function could be extrapolated to the working heart, we assessed the effect of droperidol on overall cardiac function in buffer-perfused Langendorff hearts. Only one study assessing the effects of droperidol on isolated hearts has been performed, although the focus was on action potential duration and cardiac repolarization with no assessment of inotropic status.15An in vivo  study in humans demonstrated a droperidol-induced decrease in left ventricular performance,4whereas another in vivo  study focused on prolongation of the QT interval.1In the current study, droperidol caused decreases in left ventricular developed pressure that correlated with changes seen at the level of the individual cardiomyocyte. These data indicate that droperidol, at clinically relevant concentrations, exerts a negative inotropic effect in isolated perfused hearts. However, the magnitude of the negative inotropic effect of droperidol (1 μm) observed in cardiomyocytes, where the experiments were performed at 30°C, was greater than that observed in the perfused Langendorff hearts, where the experiments were performed at 37°C. The negative inotropic effect of droperidol in the cardiomyocytes is likely greater because of a reduced diffusion gradient for drug interaction with the cardiomyocytes.

All anesthetic agents bind to plasma proteins, reducing the concentration available to bind to tissues. In the clinical setting, the use of droperidol (0.125 mg/kg) is standard for the prevention of postoperative nausea and vomiting. The peak plasma concentration with this dose has been estimated at 2 μm.31Taking into account that 90% of droperidol is bound to protein, the free concentration of droperidol is approximately 0.2 μm.32However, the microkinetic behavior of droperidol within the vascular space has not been defined. In addition, small changes in the amount or binding capacity of proteins could result in significant increases in the free plasma concentrations of droperidol. Not only is there uncertainty in calculating the in vivo  concentration of droperidol during normal circumstances, but the concentration in free plasma would certainly be higher when the concentration of protein serum is reduced (e.g. , hemodilution, liver disease, hypoproteinemia). Another potential limitation is the use of rat cardiomyocytes as a model for human cardiomyocytes, because species differences may exist. Action potentials in rodent cardiomyocytes are abbreviated compared with those recorded in human myocytes and are comprised primarily of the transient outward K⁺current, whereas in humans the delayed rectifier K⁺current predominates. This may explain our inability to observe a droperidol-induced prolongation in action potential duration. Moreover, this may also contribute to differences in the response of human and rat myocardium to anesthetic agents. It should be noted that this study deals only with intrinsic properties of the heart and that cardiac function also depends on preload, afterload, venous return, and heart rate, which are not factors in isolated cardiomyocytes. However, we have tried to overcome this limitation with the perfused Langendorff heart preparation. Despite these limitations, our results demonstrate that clinically relevant concentrations of droperidol decrease [Ca²⁺]ⁱand myofilament Ca²⁺sensitivity in cardiomyocytes, resulting in a negative inotropic effect. Moreover, in addition to the α-adrenergic blocking effect of droperidol in the vasculature,33,34a decrease in the inotropic state of the heart will likely exacerbate the hypotensive effect of droperidol observed in the clinical setting.

Our results provide the first direct evidence that droperidol causes a negative inotropic effect in individual cardiomyocytes. This effect is mediated by both a decrease in [Ca²⁺]ⁱand a decrease in myofilament Ca²⁺sensitivity. The decrease in [Ca²⁺]ⁱis not due to an effect on action potential duration, but rather due to an effect on the L-type Ca²⁺channel to limit sarcolemmal Ca²⁺influx. The decrease in myofilament Ca²⁺sensitivity is likely mediated by a decrease in pHⁱand an increase in nitric oxide production.