Propofol Increases Myofilament Ca²⁺Sensitivity and Intracellular pH via  Activation of Na⁺–H⁺Exchange in Rat Ventricular Myocytes

All experimental procedures were approved by the Cleveland Clinic Institutional Animal Care and Use Committee (Cleveland, OH).

Adult ventricular myocytes from rat hearts were isolated as previously described. 5In brief, the hearts were excised, cannulated via  the aorta, attached to a modified Langendorff perfusion apparatus, and perfused with oxygenated (95% O², 5% CO²) Krebs-Henseleit buffer (37°C) containing 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, pH 7.35. After a 5-min equilibration period, the perfusion buffer was changed to a Ca²⁺-free Krebs-Henseleit buffer containing 30 mg collagenase type II (347 U/ml). After collagenase digestion (20 min), the ventricles were minced and shaken in Krebs-Henseleit buffer, and the resulting cellular digest was washed, filtered, and resuspended in phosphate-free HEPES-buffered saline (HBS) containing 118 mm NaCl, 4.8 mm KCl, 1.2 mm MgCl², 1.25 mm CaCl², 11 mm dextrose, 25 mm HEPES, and 5 mm pyruvate, pH 7.35, and vigorously bubbled immediately before use with 100% O². Typically, 6–8 × 10⁶cells per rat heart were obtained using this procedure. Viability, as assessed by the percentage of cells retaining a rod-shaped morphology, was routinely between 80 and 90%. Myocytes were suspended in HBS (1 × 10⁶cells/ml) and stored in an oxygen hood until used.

Two-milliliter aliquots of the freshly isolated myocyte suspension were incubated in the presence or absence of propofol (10, 30, 100 μm) for 10 min at 37°C with gentle agitation. The myocytes were immediately washed twice in ice-cold HBS containing protease and phosphatase inhibitors and pelleted at 400 g  for 3 min, after which an equivalent volume of extraction buffer, 50 mm Tris (pH 7.5) containing Triton-X 100 (0.1%), NaF (20 mm), dithiothreitol (0.5 mm), MgCl²(0.5 mm), EDTA (0.125 mm), antipain (5 μg/ml), leupeptin (10 μg/ml), pepstatin A (5 μg/ml), and paramethylsulfonic acid (43 μg/ml) was added to the suspension. The cells were homogenized and kept on ice for 30 min. The triton-extracted myofibrils were pelleted at 10,000 g  (5 min, 4°C). The detergent solubilized supernatant was set aside and the pellet was resuspended in an equivalent volume of extraction buffer and washed twice again. The resultant myofibrillar fraction was resuspended in Ca²⁺-free extraction buffer and stored at −20°C. Examination of the pellet under the microscope indicated that it was enriched in myofibrils.

The Ca²⁺-stimulated actomyosin ATPase activity of the myofibrillar fraction was measured from the rate of decrease of nicotinamide adenine dinucleotide (NADH) absorbance (340-nm excitation wavelength) in a reaction coupled to the pyruvate kinase and lactate dehydrogenase reactions. 16Triton-X 100–extracted myofibrils were solubilized in a buffer containing 25 mm BES (N,N -bis-(2-hydroxyethyl)-2-aminoethanesulfonic acid, pH 7.4), 85 mm potassium methanol sulfonic acid, 3 mm MgCl², 2 mm EGTA, 10 mm NaF, 0.5 mm dithiothreitol, and 0.5 mm leupeptin. The protein concentration of the extracted myofibrils was determined using the Lowry protein assay. The reaction mixture consisted of 25 mm BES (pH 7.0), 2.7 mm MgCl², 2 mm EGTA, 10 mm NaF, and 126 mm potassium methanol sulfonic acid and varying free CaCl²concentrations, giving free Ca²⁺concentrations from pCa 9 to 4, according to an iterative computer program provided by Dr. Frank Brozovich (Case Western Reserve University School of Medicine, Cleveland, OH). The reaction mixture also contained 200 mm phosphoenolpyruvate, 10 mm NADH, 0.5 mg/ml lactate dehydrogenase, 12.5 mg/ml pyruvate kinase, and 1 ml of one of the calcium buffers (pCa 4–9) containing myofibrillar fractions. The reaction was initiated by addition of 2 mm ATP and allowed to continue for up to 10 min, although the reaction was usually complete within 5 min. Ca²⁺-stimulated actomyosin ATPase activity was monitored by the formation of adenosine diphosphate, coupled to the oxidation of NADH, and recorded by the change in absorption at 340 nm. The enzyme activity was determined from the rate of ATP hydrolysis and expressed as the percent of maximal actomyosin ATPase activity per milligram of protein. Because crude myofibrillar fractions were used for these assays, we determined whether there was a contribution to the activity measured from other ATPases present in the sample. We found no significant difference in ATPase activity measured in the presence or absence of ATPase inhibitors, including 2 mm thapsigargin, 200 mm ouabain, 2 mm rotenone, and 2 mm oligomycin.

For simultaneous measurement of shortening and pHⁱ, ventricular myocytes (0.5 × 10⁶cells/ml) were incubated in HBS containing 2 μm 2′,7′-bis-(2-carboxyethyl)-5 (6′)-carboxyfluorescein–acetoxymethyl ester (BCECF–AM) at 37°C for 20 min. BCECF-loaded ventricular myocytes were placed in a temperature-regulated (30°C) chamber mounted on the stage of an 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 and field-stimulated via  bipolar platinum electrodes at a frequency of 0.3 Hz and a duration of 5 ms using a stimulator.

Fluorescence measurements were performed on single ventricular myocytes using a dual-wavelength spectrofluorometer at excitation wavelengths of 440 and 500 nm and an emission wavelength of 530 nm. 17The cells were also illuminated with red light at a wavelength greater than 600 nm for simultaneous video edge detection. An additional postspecimen dichroic mirror deflects light at wavelengths greater than 600 nm into a charge coupled device video camera for measurement of myocyte shortening. The fluorescence sampling frequency was 10 Hz, and data were collected using a software package. Background fluorescence was determined from the blank dish and subtracted from fluorescence at each wavelength. To estimate the pHⁱvalue from the ratio of 500/440 nm fluorescence, we used an in situ  calibration procedure. 17,18At 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 K⁺, M1.0 mm gCl², 4.0 mm HEPES, 2.0 mm EGTA, 30 mm 2,3-butanedione monoxime, 50 μm 1,2-bis(o-Aminophenoxy)ethane-N,N,N′,N′- tetraacetic acid, 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 1.0 N KOH. The pHⁱfor each cell was then determined from a linear regression of 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). 17 

Simultaneous measurement of cell shortening was monitored using a video edge detector with 16-ms temporal resolution. The video edge detector was calibrated using a stage micrometer so that cell lengths during shortening could be monitored. LabVIEW (National Instruments, Austin, TX) was used for data acquisition of cell shortening using a sampling rate of 100 Hz.

Fluorescence data for the pHⁱmeasurements were imported into LabVIEW, which allowed the simultaneous analysis of pHⁱand shortening. Myocyte length in response to field stimulation was measured (micrometers) and expressed as the change from resting cell length (twitch amplitude). Changes in twitch amplitude in response to the interventions are expressed as a percent of baseline shortening. Contractile parameters from 15 contractions were averaged to obtain mean values at baseline and in response to the various interventions. Averaging the parameters over time minimizes beat-to-beat variation. Changes in pHⁱwere measured as the change in the 500/440 ratio from baseline.

Where applicable, protocols were designed so that each cell could serve as its own control.

To determine whether propofol alters myofilament Ca²⁺sensitivity, changes in actomyosin ATPase activity were measured in myofibrils isolated from control and propofol-treated cardiomyocytes. The myocyte suspension was divided into separate aliquots, and each aliquot was treated with one concentration of propofol (10, 30, 100 μm) or intralipid equivalent for 10 min at 37°C with gentle agitation. Propofol had no effect on extracellular pH at the concentrations used in this study and did not appear to alter the assay conditions. Activity is expressed as a percentage of the maximum rate per milligram of protein.

To identify the effects of changing pHⁱon myocyte shortening, changes in myocyte shortening and pHⁱin response to 10 mm NH⁴Cl (2 min) followed by washout were determined. Baseline measurements were collected from individual myocytes for 1 min in the absence of any intervention.

To determine whether propofol alters pHⁱ, we measured pHⁱfor 15 min after addition of propofol. Baseline pHⁱwas collected from individual myocytes for 1 min. Propofol (10, 30, 100 μm) or intralipid equivalent was added by exchanging the superfusion buffer in the dish with new buffer containing propofol at the desired concentration. Each myocyte was exposed to only one concentration for propofol.

To examine the role of the Na⁺–H⁺exchange and PKC in modulating propofol-induced changes in pHⁱand shortening, we assessed propofol-induced intracellular alkalinization and shortening in myocytes pretreated (5 min) with a Na⁺–H⁺exchange inhibitor, 5-(N -ethyl-N -isopropyl)-amiloride (EIPA, 1 μm), or a PKC inhibitor, bisindolylmaleimide I (Bis, 1 μm). We have verified that propofol, EIPA, and Bis have no effect on the BCECF signal at the concentrations used in these studies.

To determine whether propofol alters NH⁴Cl-induced changes in pHⁱand contractility, we simultaneously measured changes in pHⁱand myocyte shortening in the presence or absence of propofol. Baseline pHⁱand shortening were measured in individual myocytes for 1 min. In the propofol group, the cells were superfused with buffer containing propofol (30 μm) 5 min before exposure to NH⁴Cl. Washout was performed with HBS containing propofol.

To determine whether propofol alters pHⁱrecovery from acid load, we measured the changes in pHⁱfrom intracellular acidosis after washout with 10 mm NH⁴Cl in the presence or absence of propofol. Baseline pHⁱwas measured in individual myocytes for 1 min. At the time of NH⁴Cl washout, each myocyte was superfused with buffer either in the presence or absence of propofol (10, 30, 100 μm). Rate of recovery of pHⁱfrom NH⁴Cl-induced acidosis was assessed by calculating the derivative of the slope of the pHⁱtime course trace for 10-s intervals through the first 120 s of recovery.

We previously reported that propofol caused a leftward shift in the extracellular Ca²⁺concentration ([Ca²⁺]^o)–shortening relation, with no concomitant effect on the intracellular Ca²⁺transient. 5Baseline parameters were collected from individual myocytes for 1.5 min. Dose–response curves to [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. Dose–response curves to [Ca²⁺]^owere then performed in the presence of propofol (30 μm). Cells were allowed to stabilize for 5 min after each intervention. The relative contribution of the Na⁺–H⁺exchanger in mediating changes in the [Ca²⁺]^o–shortening relation was assessed by pretreating the cells (5 min) with 1 μm EIPA.

Collagenase type II was obtained from Worthington Biochemical (Freehold, NJ). Propofol and intralipid were obtained from The Cleveland Clinic Pharmacy. BCECF–AM was obtained from Texas Fluorescence Labs (Austin, TX). Nigericin, EIPA, and Bis were purchased from Sigma Chemical Co. (St. Louis, MO).

Each experimental protocol was performed on multiple myocytes from the same heart and repeated in at least four hearts. Results obtained from myocytes in each heart were averaged so that all hearts were weighted equally. The effects of each concentration of propofol on myocyte shortening and pHⁱwere assessed using one-way analysis of variance with repeated measures and the Bonferonni correction for multiple comparisons. Comparisons between groups were made by two-way analysis of variance and unpaired t  test. Results are expressed as mean ± SD. P < 0.05 was considered significant.

To directly test the hypothesis that propofol increases myofilament Ca²⁺sensitivity, we assessed the effects of propofol on myofibrillar actomyosin ATPase activity in myocytes pretreated with propofol (10–100 μm). Propofol caused a concentration-dependent leftward shift in the actomyosin ATPase activation curve (fig. 1A). Propofol increased the EC⁵⁰(pCa) value (i.e. , decreased the Ca²⁺requirement) for Ca²⁺-activated actomyosin ATPase (fig. 1B). Propofol (30 and 100 μm) reduced (P < 0.05) the maximal activation of actomyosin ATPase (V^max) by 11 ± 4% and 17 ± 5%, respectively, from a control value of 175 ± 8 nmol · min⁻¹· mg⁻¹. The intralipid vehicle had no effect on actomyosin ATPase activity (EC⁵⁰= 5.6 ± 0.1; V^max= 169 ± 9 nmol · min⁻¹· mg⁻¹) at a concentration equivalent to 100 μm propofol.

Baseline pHⁱand diastolic cell length were 7.05 ± 0.03 and 130 ± 15 μm, respectively. Twitch amplitude was 9 ± 1% (11.7 ± 2.8 μm) of the resting cell length. To test the hypothesis that changes in pHⁱalter the contractile properties of ventricular myocytes, we measured pHⁱand shortening simultaneously in response to NH⁴Cl followed by washout to cause intracellular alkalinization and acidification, respectively. Figure 2illustrates the effects of NH⁴Cl (10 mm) on shortening and pHⁱin an individual cardiomyocyte. Field stimulation alone had no effect on baseline pHⁱ. NH⁴Cl initially caused intracellular alkalinization (7.39 ± 0.06), a 4.4 ± 1.7% decrease (P < 0.05) in diastolic cell length, and an increase (P < 0.05) in myocyte shortening (135 ± 10% of baseline). Washout of NH⁴Cl caused intracellular acidification (6.92 ± 0.02), a return of diastolic cell length to baseline (1.0 ± 0.1%), and a decrease (P < 0.05) in myocyte shortening (48 ± 8% of baseline). Myocyte shortening gradually increased after washout of NH⁴Cl and reached 95 ± 8% of the baseline value after 10 min.

To test the hypothesis that the propofol-induced increase in myofilament Ca²⁺sensitivity was caused by intracellular alkalinization, we assessed the extent to which propofol altered pHⁱ. Propofol at 10 μm had no significant effect on pHⁱor shortening. Propofol at 30 and 100 μm caused a concentration- and time-dependent increase in pHⁱof 0.03 ± 0.01 and 0.07 ± 0.02, respectively, after a 15-min exposure (fig. 3). Despite the increase in pHⁱ, propofol (30 and 100 μm) reduced (P < 0.05) shortening by 10 ± 3% and 33 ± 7%, respectively. The intralipid vehicle had no effect on pHⁱ(7.03 ± 0.03) or shortening at a concentration equivalent to 100 μm propofol. In some cells (6 of 16), 30 μm propofol decreased resting cell length from 127 ± 10 μm to 124 ± 7 μm. Similarly, 100 μm propofol decreased resting cell length to 122 ± 9 μm in 10 of 16 cells studied.

To test the hypothesis that the propofol-induced increase in pHⁱwas mediated by an increase in Na⁺–H⁺exchange, we assessed the effects of propofol in cells pretreated with EIPA, a Na⁺–H⁺exchange inhibitor. Pretreatment with EIPA had no effect on baseline pHⁱor shortening. Figure 4summarizes changes in pHⁱafter 15-min exposure to 100 μm propofol. The propofol-induced intracellular alkalinization was inhibited by pretreatment with EIPA. PKC inhibition with Bis also inhibited the propofol-induced intracellular alkalinization (fig. 4). In the presence of EIPA, 30 μm propofol decreased (P < 0.05) cell shortening by an additional 17 ± 3% compared with propofol alone. In the presence of Bis, propofol (30 μm) decreased (P < 0.05) cell shortening by an additional 24 ± 4% compared with propofol alone.

Administration of NH⁴Cl results in activation of Na⁺–H⁺exchange without the involvement of second messengers. We used NH⁴Cl to test the hypothesis that propofol-induced changes in pHⁱare mediated by changes in Na⁺–H⁺exchange activity. Baseline pHⁱand myocyte shortening were 7.03 ± 0.02 and 8 ± 1%, respectively. Pretreatment with propofol (30 μm, 10 min) increased (P < 0.05) pHⁱ(+0.03 ± 0.01), whereas shortening decreased (P < 0.05) to 92 ± 4% of control. As expected, pretreatment with propofol had no effect on NH⁴Cl-induced increases in pHⁱand cell shortening (fig. 5), because these changes are not mediated by Na⁺–H⁺exchange activation. However, the acidification-induced decreases in pHⁱ(mediated by an increase in Na⁺–H⁺exchange) and myocyte shortening in response to NH⁴Cl were attenuated by propofol (fig. 5).

To further test the hypothesis that propofol enhances the activity of the Na⁺–H⁺exchanger, we assessed the extent to which propofol altered the rate of recovery after acidosis (dpHⁱ/dt) induced by a 2–3-min exposure to 10 mm NH⁴Cl. NH⁴Cl produced an intracellular alkalosis, whereas subsequent washout produced a transient intracellular acidosis (fig. 6A). EIPA abolished the recovery from acidosis induced by NH⁴CI (fig. 6A). The time course of pHⁱrecovery from acidosis was compared in the presence or absence of propofol. Propofol at 30 μm accelerated the rate of recovery from NH⁴Cl-induced intracellular acidosis (fig. 6B). This effect of propofol on the rate of recovery is even more apparent when the data are normalized to peak acidosis (fig. 6C). As summarized in figure 7, propofol increased the recovery rate from acidosis in a concentration-dependent manner.

Confirming our previous results, 5the propofol-induced increase in myofilament Ca²⁺sensitivity is manifested as a leftward shift in the [Ca²⁺]^o–shortening relation (fig. 8). To test the hypothesis that this effect is caused by an increase in Na⁺–H⁺exchange, we assessed the effect of 1 μm EIPA on the propofol-induced change in the [Ca²⁺]^o–shortening relation. EIPA attenuated, but did not abolish, the propofol-induced leftward shift in the [Ca²⁺]^o–shortening relation (fig. 8).

This is the first study to directly assess the effects of propofol on myofilament Ca²⁺sensitivity, Na⁺–H⁺exchange, and pHⁱin isolated cardiac muscle cells. A previous study from our laboratory using isolated rat ventricular myocytes, as well as a recent study using intact beating guinea pig hearts, provided indirect evidence that propofol may increase the sensitivity of the contractile machinery to Ca²⁺. 5,13In contrast, a recent study using phase-loop diagrams depicting the continuous relation between cell length and the fura-2 fluorescence ratio in rat ventricular myocytes has suggested that propofol may reduce myofilament Ca²⁺sensitivity. 19In addition, several previous studies have indicated that propofol does not alter myofilament Ca²⁺sensitivity. 2,6In this study, we measured changes in Ca²⁺-activated myofibrillar actomyosin ATPase activity in response to propofol to directly assess changes in myofilament Ca²⁺sensitivity. The key finding of this study is that propofol markedly reduced the Ca²⁺requirement for activation of myofibrillar actomyosin ATPase. Moreover, the mechanism responsible for this effect involves, at least in part, a propofol-induced increase in pHⁱmediated via  PKC-dependent activation of the Na⁺–H⁺exchanger.

Cardiac myofilaments contain an intrinsic actomyosin ATPase that represents the molecular basis for contraction. Sensitivity of the actomyosin ATPase to Ca²⁺is mediated by the troponin complex in the thin filament. This multiprotein complex is comprised of the Ca²⁺binding TnC subunit, the ATPase inhibiting TnI subunit, and the tropomyosin binding TnT subunit. It is well established that changes in pHⁱand/or direct phosphorylation of the contractile proteins, TnI, TnT, and myosin light chain 2, can alter the sensitivity of the contractile machinery to Ca²⁺. Although propofol had no direct effect on actomyosin ATPase activity in isolated myofibrils, it caused a leftward shift in the Ca²⁺-activated actomyosin ATPase activity curve (i.e. , a decrease in the amount of Ca²⁺required for activation) with a small decrease in maximal activity (V^max) in myofibrils isolated from propofol-treated cardiomyocytes. These results imply the requirement of a cytosolic mediator for the actions of propofol on actomyosin ATPase activity. Taken together, it appears that propofol may alter multiple cellular mechanisms that regulate calcium sensitivity. PKC activators have similar effects on actomyosin ATPase activity. 20–22 

Control of pHⁱwithin the physiologic range is essential for the optimal function of enzymes, cell growth, and protein synthesis. 23,24In cardiac muscle, pHⁱis a key factor regulating myocardial contractility, because changes in pHⁱalter the sensitivity of the contractile apparatus to Ca²⁺. 15The Na⁺–H⁺exchanger represents the most important mechanism for regulation of cardiomyocyte pHⁱ. Baseline pHⁱin this study was 7.05 ± 0.03, which is similar to that in other reports. 18,25 

Both the negative and positive inotropic effects of acidosis and alkalosis, respectively, have been documented. 26These effects appear to result almost entirely from changes in pHⁱand not from changes in extracellular pH. 27However, the precise mechanisms underlying the changes in myocardial contractility in response to changes in pHⁱare not known, because the development of force on Ca²⁺activation involves many pH-dependent biochemical processes. To date, several mechanisms have been postulated to contribute to pHⁱ-mediated changes in myocardial contractility, including changes in the affinity of myofibrillar troponin C for Ca²⁺, 28pH dependence of the myofibrillar actomyosin ATPase activity, 29and changes in sarcoplasmic reticulum Ca²⁺handling. 15 

Propofol caused a time- and dose-dependent increase in pHⁱ. The magnitude of the increase in pHⁱwas comparable to that observed in response to α¹-adrenergic agonists, endothelin, and angiotensin II. 25,30,31Despite the increase in pHⁱ, propofol did not exert a positive inotropic effect. However, propofol caused a slight decrease in resting cell length in approximately half of the cells studied. Propofol likely alters multiple cellular mechanisms that negatively regulate contractility (e.g. , sarcoplasmic reticulum Ca²⁺handling, L-type Ca²⁺channel activity), which could offset the effects of increasing myofilament Ca²⁺sensitivity. In fact, a propofol-induced depression of the intracellular Ca²⁺transient has been observed in beating guinea pig hearts with no concomitant decrease in contractility, implying enhanced myofilament Ca²⁺sensitivity. 13Moreover, intracellular alkalosis can cause a decrease in peak intracellular Ca²⁺concentration, 14which could partially offset the effect of increased pHⁱon myofilament Ca²⁺sensitivity. Propofol may also alter myofilament Ca²⁺sensitivity via  PKC-dependent phosphorylation of contractile proteins. 32 

Protein kinase C activation is the primary regulator of Na⁺–H⁺exchange in cardiomyocytes. We hypothesized that inhibition of Na⁺–H⁺exchange and/or PKC should attenuate the ability of propofol to induce intracellular alkalinization. Consistent with this, the propofol-induced increase in pHⁱwas markedly attenuated by EIPA and Bis, indicating that the effects are mediated by a PKC-dependent activation of Na⁺–H⁺exchange. Pretreatment with EIPA or Bis also resulted in an even greater propofol-induced reduction in shortening. Moreover, the propofol-induced decrease in shortening was greater in Bis-treated myocytes compared with EIPA-treated myocytes. Taken together, these results suggest that PKC-dependent intracellular alkalinization offsets the negative inotropic effect of propofol (likely because of a decrease in intracellular Ca²⁺availability) 13 via  an increase in myofilament Ca²⁺sensitivity.

We used NH⁴Cl as a tool to assess the effects of changing pHⁱon myocardial contractility in our experimental model. NH⁴Cl initially causes intracellular alkalosis because of the highly permeant NH³crossing the membrane more quickly than NH⁴⁺. Once inside the myocyte, NH³combines with H⁺to form NH⁴⁺, which increases pHⁱand shortening. With continued exposure to NH⁴Cl, pHⁱgradually decreases as NH⁴⁺enters the cell. The subsequent abrupt removal of external NH⁴Cl produces an intracellular acidosis. This occurs because NH⁴⁺ions that had accumulated in the cytosol dissociate to form NH³, which rapidly leaves the cell, resulting in intracellular retention of H⁺and a decrease in shortening. We hypothesized that NH⁴Cl-induced alkalinization (independent of Na⁺–H⁺exchange) would not be affected by propofol, whereas the acidification (dependent on Na⁺–H⁺exchange) would be attenuated if propofol caused intracellular alkalinization via  activation of Na⁺–H⁺exchange. We found that propofol had no effect on NH⁴Cl-induced intracellular alkalinization but attenuated the acidification. In addition, propofol attenuated the decrease in shortening induced by acidification. These data give further support to the idea that propofol activates the Na⁺–H⁺exchange mechanism in ventricular myocytes.

Because these experiments were performed in the absence of HCO³⁻and carbon dioxide, the acid-load recovery is likely mediated by Na⁺–H⁺exchange. 33We hypothesized that if propofol activated the Na⁺–H⁺exchanger as a mechanism to increase pHⁱ, propofol should stimulate a faster rate of recovery from an acid load in response to NH⁴Cl washout. At concentrations greater than 10 μm, propofol enhanced the rate of recovery from an acid load. Because the recovery from an acid load is dependent on Na⁺–H⁺exchange activity, these data give further support to the hypothesis that propofol enhances the activity of the Na⁺–H⁺exchanger during acid-load conditions.

If intracellular alkalinization mediates the propofol-induced leftward shift in the [Ca²⁺]^o–shortening relation, then this effect should be blocked by inhibiting Na⁺–H⁺exchange. Our results indicate that inhibition of Na⁺–H⁺exchange with EIPA attenuates the propofol-induced increase in myofilament Ca²⁺sensitivity by approximately 50%. It is possible that phosphorylation of contractile proteins by propofol also plays a role in mediating the increase in myofilament Ca²⁺sensitivity. 32Inhibition of Na⁺–H⁺exchange with amiloride has been reported to attenuate the increase in myofilament Ca²⁺sensitivity in response to endothelin 31and phenylephrine 14in rat ventricular myocytes. In contrast to the present study, EIPA abolished the leftward shift in the [Ca²⁺]^o–shortening relation induced by thiopental. 17 

Because propofol partitions in vivo  between serum proteins, lipid microsomes, and into tissue, it is difficult to know the precise concentration of free and active propofol at the tissue level. The concentrations of propofol used in these in vitro  studies are supraclinical. Because we used isolated cells rather than intact tissue, the effects of propofol may be enhanced because of optimal solute diffusion distances between the cytosol and extracellular medium. In addition, this in vitro  study only deals with intrinsic myocardial contractility, whereas changes in cardiac function in vivo  after propofol administration also depend on venous return, afterload, and compensatory mechanisms. We also acknowledge that the experimental conditions (temperature, stimulation frequency, unloaded cells) do not perfectly simulate the in vivo  heart. However, the strength of this model is that we can directly assess the effects of propofol on the fundamental contractile unit, the individual cardiomyocyte. Finally, we now have direct evidence that propofol increases the Ca²⁺sensitivity of the actomyosin ATPase, causes intracellular alkalosis, and stimulates both PKC-dependent phosphorylation of contractile proteins 32and Na⁺–H⁺exchange activity in rat cardiomyocytes. However, we have not directly demonstrated that intracellular alkalosis increases the Ca²⁺sensitivity of actomyosin ATPase activity.

In summary, our results provide the first direct evidence that propofol increases the sensitivity of myofibrillar actomyosin ATPase to Ca²⁺(i.e. , increases myofilament Ca²⁺sensitivity), at least in part by increasing pHⁱvia  PKC-dependent activation of Na⁺–H⁺exchange.