Propofol is used with increasing frequency for sedation, sometimes in high-risk patients with compromised myocardial function. A typical anesthetic induction dose of propofol (2 mg/kg) results in an approximate 30% reduction in systolic blood pressure (1). The majority of literature recognizes that this hypotension is mainly attributable to a decrease in sympathetic activity (2) and/or direct vasodilation (3) and less is attributable to direct myocardial depression causing a decrease in cardiac output (4). In patients with good ventricular function undergoing coronary artery bypass graft surgery, propofol causes no significant changes in cardiac output (5). Pagel et al. (6) studied cardiovascular effects of propofol in dogs with dilated pacing-induced cardiomyopathy and found significant reduction of left ventricular preload, afterload, and relatively minor effects on contractility. Indeed, findings from clinical studies suggest what several in vitro studies (7–9) have already shown, i.e., that depression of myocardial contractility may not be the primary cause of propofol-induced hypotension. The effects of propofol on myocardial contractility in the failing human myocardium have not been studied. Because it is difficult to study anesthetic-induced direct negative myocardial inotropy in vivo (as a result of simultaneous changes in left ventricular loading conditions (6) and changes in sympathetic nervous system activation), we have assessed the effects of propofol on myocardial contractility in an in vitro preparation of human cardiac muscles.
Our first goal was to quantify the effects of propofol on inotropy in failing cardiac muscles. Our second goal was to test the hypothesis that inotropic responsiveness of failing human myocardium is attenuated in the presence of propofol. Even in the absence of anesthetics, failing human myocardium may exhibit decreased responsiveness to β-adrenergic stimulation (10) because of chronic hyperstimulation of the adrenergic receptors in patients with heart failure. This adrenergic hyperstimulation causes a reduction in myocardial β1 receptors, uncouples myocardial β2 receptors, and decreases the catalytic activity of adenyl-cyclase (11). We have already shown that etomidate can reduce (12) and ketamine can abolish (13) responsiveness to β-adrenergic stimulation; however, the effects of propofol, especially on failing myocardium, have never been studied. Finally, to elucidate the cellular mechanism(s) of negative inotropy which occurred at larger propofol concentrations, we examined the effects of propofol on Ca2+- stimulated actomyosin adenosine triphosphatase (ATPase) activity of the myofibrils and on sarcoplasmic reticulum (SR) Ca2+ uptake by SR vesicles from cardiac muscle.
Materials and Methods
The IRB at the Cleveland Clinic Foundation approved procurement of tissue for this study. All muscles were obtained from patients undergoing either cardiac transplantation or myocardial revascularization. In the first group, 14 trabecular muscles (8 exposed to propofol and 6 to intralipid) removed from the right atrial appendages of 6 cardiac transplant patients and 27 muscles (16 exposed to propofol and 11 to intralipid) removed from the left ventricle of 9 cardiac transplant patients were used. During the average 10-min transport to the laboratory, the heart was maintained in ice-cold, oxygenated cardioplegic solution as described previously (12,13). The second group of human myocardial trabeculae (n = 17; 8 exposed to propofol and 9 to intralipid) was obtained from 7 patients with nonfailing hearts undergoing routine coronary artery bypass graft surgery. Trabecular muscles were dissected from the right atrial appendage, which was removed during surgery for the purpose of atrial cannulation and from the failing right atrial appendage or the left ventricle before the heart was explanted. The anesthetic regimen used in this study consisted of reduced doses of propofol or sodium thiopental for induction, pancuronium for muscle relaxation, midazolam and large-dose fentanyl, and isoflurane for maintenance. The decision about anesthetic regimen was left to the discretion of the primary anesthesiologist. All tissues were procured at least 1 h after anesthetic induction; therefore, the possibility of any residual propofol or sodium thiopental effect at the time of in vitro experimentation was remote.
Measurement of Isometric Contraction
Isometric contraction variables were recorded using the previously described protocol (12,13). In short, the muscle was mounted in a water-jacketed tissue bath containing Krebs-Henseleit buffer. The bath temperature was maintained at 37°C and buffer was oxygenated with 95% O2/5% CO2. After a 40-min stabilization period, stimulation (frequency 1 Hz, duration 5 ms, voltage 20% above threshold) was initiated through two parallel platinum electrodes. The muscle length was gradually increased until maximal developed tension (Lmax) was reached. Contractile variables recorded consisted of the resting tension, maximal developed tension (DT), time-to-peak tension, time-to-half relaxation, and maximal rate of contraction and relaxation (+dT/dt and −dT/dt, respectively). Once the response was stable at Lmax, propofol was added. In the present study, propofol was used in the commercial 10% intralipid solvent used clinically (Diprivan®; Zeneca Pharmaceuticals, Wilmington, DE) (concentration 10 mg/mL). Propofol was added cumulatively to the bath in concentrations between 0.056 and 560 μM to encompass a wide range of clinical and supratherapeutic concentrations (see Discussion). Muscle contractility in the presence of 10% intralipid solution [100 mg/mL soybean oil, 22.5 mg/mL glycerol, and 12 mg/mL egg lecithin in water] was compared with that after propofol. After the concentration-response curves to propofol or intralipid were completed, a single dose of 1 μM isoproterenol, a β-adrenergic agonist, was added to the bath to determine the inotropic response of cardiac muscle in the presence and absence of propofol. The muscle was not washed between the exposure to propofol or intralipid and isoproterenol addition. At the end of each experiment, the muscle length was measured (using Vernier calipers) and weighed, cross-sectional area was calculated, and this calculation was used to normalize the contractility variables.
Measurements of Actomyosin ATPase Activity
Measurements of the Ca2+-stimulated actomyosin ATPase activity were completed in eight muscles (four propofol, and four intralipid) from six nonfailing atrial hearts. In brief, muscles were mounted in a muscle bath and stimulated to contract at Lmax. Muscles were treated with either 56 μM propofol in intralipid, an equivalent volume of intralipid alone, or no treatment (control). The dose of 56 μM propofol was chosen as the dose at which the negative inotropic effect starts to be significant in vitro. Muscles were homogenized in 1.0 mL of ice-cold “inhibiting buffer” containing (in mM): 50 KH2PO4, 70 NaF, and 5 EDTA, with protease inhibitors (5 μg/mL antipain, 10 μg/mL leupeptin, 5 μg/mL pepstatin A, 43 μg/mL phenylmethylsulfonyl fluoride, 5 mM EGTA, and phosphatase inhibitor (2 nM calyculin A) and pelleted at 100 g for 3 min at 4°C. The myofibrillar proteins were then extracted in 1 mL of ice-cold “inhibiting buffer” plus 1% Triton X-100 and the protease and phosphatase inhibitors described above, and kept on ice for 30 min. The Triton X-100 extracted myofibrils were then pelleted at 5000 g for 5 min, using an Eppendorf microcentrifuge. The myofibrils were solubilized in an N, N-bis-(2-hydrozyethyl)-2-aminoethanesulfonic acid (BES)-buffer containing the following (in mM): 85 potassium methanol sulfonic acid (KMS), 3 MgCl2, 2 EGTA, 10 NaF, 0.5 dithiothreitol (DTT), 0.5 leupeptin, and 30 BES, pH 7.00, plus 3% Tween 20. The protein concentration of the extracted myofibrils was determined by using the Lowry method (14).
The Ca2+-stimulated actomyosin ATPase activity was then measured in a reaction mixture consisting of (in mM): 25 BES (pH 7.0), 2.7 MgCl2, 2 EGTA, 10 NaF, 126 KMS, and 2.26 CaCl2 (the free Ca2+ concentration was calculated to be 100 μM, pCa 4). Ca2+ buffers were prepared according to an iterative computer program. Binding constants were corrected for temperature and ionic strength. KMS supplied the major anion and the ionic strength was set at 200 mM. Solutions were prepared for pCa 4 (activating solution) and pCa 9 (relaxing solution). The absolute free Ca2+ concentrations from pCa 9 to pCa 4 (0.001 to 100 μM) were verified by recording the fura-2-fluorescence (340 nm excitation, 530 nm emission) from the reaction mixtures, using a fluorimeter. The reactions were prepared by adding 200 mM phosphoenolpyruvate, 10 mM nicotinamide dinucleotide, 0.5 mg/mL lactate dehydrogenase, 12.5 mg/mL pyruvate kinase to 1 mL of the various Ca2+ solutions (containing myofibrillar fractions), and initiated by the addition of 2 mM adenosine 5′-triphosphate (ATP). The Ca2+-stimulated actomyosin ATPase activity was monitored by the formation of adenosine 5′-diphosphate, coupled to the oxidation of nicotinamide dinucleotide, and recorded by the change in absorption at 340 nm for a duration of 5 to 10 min. The enzyme activity was determined from the rate of ATP hydrolysis and subsequently expressed as the percent of maximal Ca2+-stimulated actomyosin ATPase activity per milligram of protein. The kinetic data for the Ca2+-stimulated actomyosin ATPase activity (as a function of Ca2+ concentration) was calculated by nonlinear regression of a sigmoidal concentration-response curve with a fixed Hill coefficient.
In addition, because the myofibrillar fractions used for these assays were not purified myofibrillar proteins, there was a potential contribution of other ATPases present in the sample. However, we expected that the activity of all membrane-associated ATPases would be eliminated during extraction of the myofibrils in 1% Triton X-100. Therefore, in control experiments, we verified that there was no significant difference in ATPase activity measured in the presence of ATPase inhibitors including (in mM): 2 thapsigargin, 200 ouabain, 2 rotenone, and 2 oligomycin. Additionally, we determined whether the level of troponin I (Tn I) phosphorylation was preserved during the actomyosin ATPase activity assay. Because protein kinase A-dependent Tn I phosphorylation alters myofilament Ca2+ sensitivity by altering troponin C affinity for Ca2+, different concentrations of free [Ca2+] would be required to achieve the same degree of actomyosin ATPase activity or force development in response to phosphorylation. Therefore, in control experiments, we measured the extent of Tn I phosphorylation in myofibrillar fractions for the duration of the assay. We found no kinase- or phosphatase-dependent change in the 32Pi incorporation into Tn I for the duration of the 5-min assay (data not shown).
Preparation of SR Vesicles
Human left ventricular tissue from five failing hearts was homogenized in five volumes of 3-(N-morpholino) propanesulfonic acid (MOPS) buffer (10 mM, pH 7.4, 4°C) containing 290.0 mM sucrose, 3.0 mM NaN3, 1.0 mM DTT, 1.0 μM pepstatin A, 1.0 μM leupeptin, and 0.8 mM phenylmethylsulfonyl fluoride, using a Brinkmann (Westbury, NY) Polytron homogenizer. The homogenate was centrifuged at 7500 g for 20 min. The supernatant was separated and centrifuged again at 40,000 g for 60 min. The resultant pellet was suspended in three volumes of 3-(N-morpholino) propanesulfonic acid (10 mM, pH 6.8, 4°C) containing 600.0 mM KCl, 3.0 mM NaN3, 1.0 mM DTT, and protease inhibitors, and was centrifuged at 140,000 g for 40 min. The final pellet was resuspended in a Ca2+-free sucrose buffer and stored at −80°C until used for the experiments.
Measurement of SR Ca2+ Uptake
Double-distilled tap water was deionized by using a Milli-Q reagent water system (Millipore Corp., Bedford, MA) and was further purified by dual ion-exchange chromatography and the use of Ca2+ Sponge-S (Molecular Probes, Eugene, OR) to remove residual Ca2+. A buffering system, which contained ions in similar concentrations to the intracellular environment, and which was also capable of regenerating ATP, was used for suspending the vesicles. The buffer contained the following (in mM): 20.0 HEPES, 100.0 KCl, 5.0 NaCl, 5.0 MgCl2, 5.0 creatine phosphate, pH 7.2, 37°C, and creatine phosphokinase (0.4 U/mL). Oxalate (10.0 mM) was used as a Ca2+ precipitating anion inside the vesicles. The use of oxalate minimized leakage of Ca2+ and maintained the Ca2+ gradient across the vesicular membrane. All solutions were prepared by using an iterative solution mixing program (Solwin version 2.0, Philadelphia, PA). Binding constants for the ionic compounds were corrected for both temperature and ionic strength. CaCl2 was added back to the buffer to yield a free Ca2+ concentration of 1 μM (pCa 6).
Measurements of Ca2+ uptake and release were completed in real time, using suspensions of SR vesicles and 2 μM fura-2 free acid in the extravesicular compartment. Fluorescence experiments were performed by using dual wavelength fluorometry in a temperature-regulated sample compartment (37°C). Microcuvettes (250 μL) were washed in 2.0 mM EGTA to remove all Ca2+, and then thoroughly rinsed with Ca2+-free buffer and allowed to dry. For Ca2+ uptake studies, the addition of 1.0 mM ATP to the vesicular suspension triggered the uptake of Ca2+ into the vesicles, which was measured as a decrease in the fluorescence signal (340:380 ratio) from the extravesicular compartment. Fluorescence data were collected using the computer program at a sampling frequency of 20 Hz. The rate of Ca2+ uptake was measured as the decrease in the fluorescence signal over a 60-s period in the presence or absence of propofol (5.6–560 μM). The addition of propofol did not alter the pH of the suspension buffer. An equal volume of intralipid was added to a different suspension of SR vesicles from the same heart, and effects on Ca2+ uptake were monitored.
Contractile variables for atrial muscles and ventricular muscles taken from the same heart were averaged separately, such that each heart was used as a single observation and contributed only once to each overall data set. Responses to propofol and intralipid were compared by using a repeated-measures analysis of variance with Student-Newman-Keuls testing for individual measurements. All values in tables and figures were expressed as mean ± sd. The accepted level of significance was P < 0.05. For the actomyosin ATPase assay, the 50% effective concentration (EC50) values for propofol, intralipid, and control muscles were averaged per heart and then per group. An unpaired t-test was used to determine significant differences between propofol- and intralipid-treated groups and between propofol- and control-treated groups. For the SR Ca2+ uptake experiments, the rate of Ca2+ uptake at each dose of propofol was averaged per heart, and a repeated-measures analysis of variance was used to compare propofol- versus intralipid-treated groups.
Table 1 shows the patients’ demographic data, preoperative medications, and preoperative left ventricular ejection fractions. Table 2 summarizes the muscle baseline contractile variables during control conditions at Lmax from all human atrial and ventricular muscles used. None of the baseline contractile variables were significantly different between nonfailing and failing atrial muscles.
Effects of Propofol and Intralipid on Maximal DT in Nonfailing and Failing Hearts
Over the wide range of propofol concentrations, myocardial contractility decreased in a concentration-dependent manner in all three tissue types; however, this decrease was not significant at typical clinical propofol concentrations. Maximal decreases in DT after exposure to 560 μM propofol were similar for failing and nonfailing muscles, between 40% and 50% (Fig. 1). In nonfailing atrial and failing ventricular muscles, contractility started to decrease significantly at 56 μM (P < 0.05) which may be considered a large clinical concentration range. In failing atrial muscle, contractility did not significantly decrease until the muscle was exposed to a supratherapeutic concentration of 560 μM (Fig. 1, P < 0.05). None of the muscles exposed to intralipid demonstrated a change in myocardial contractility (P > 0.05 compared with baseline). In nonfailing and failing atrial muscles exposed to propofol, 1 μM isoproterenol increased average maximal DT to 44% and 36% above baseline, respectively (P = nonsignificant compared with baseline, Fig. 2). At the same time, the increase in myocardial contractility in failing ventricular muscles was somewhat lower (4% below the baseline), and this represented a diminished response compared with failing ventricular muscles exposed to intralipid (P < 0.05). The responsiveness of intralipid-exposed muscles to isoproterenol was uniformly preserved: DT increased to 65% above baseline in nonfailing atrium and 62% and 73% above baseline in both failing atrium and ventricle, respectively (Fig. 2).
Effects of Propofol on Other Contractility Variables
Figures 3 and 4 show changes in +dT/dt and −dT/dt in the nonfailing atrial and failing atrial and ventricular muscles after treatment with propofol; these changes are compared with intralipid only. The relative changes in maximal rate of contraction and relaxation paralleled changes in DT. There were no significant effects of propofol on resting tension, time-to-peak tension, or time-to-half relaxation in any group of muscles exposed to either propofol or intralipid alone (P > 0.05 for all groups).
Effects of Propofol and Intralipid on Actomyosin ATPase Activity
In the presence of 56 μM propofol, the Ca2+-activated actomyosin ATPase activity of human nonfailing atrial trabecular muscles was shifted significantly leftward compared with control muscles (Fig. 5), demonstrating an increase in the myofilament sensitivity to Ca2+ after propofol. The same volume of intralipid alone had no effect on the Ca2+ sensitivity of actomyosin ATPase activity (Fig. 5). In control human atrial trabecular muscles, the EC50 for Ca2+ in pCa units was 6.14. The EC50 for Ca2+ was 6.24 pCa units in muscles treated with intralipid alone, but moved to 6.70 pCa units in muscles treated with propofol in intralipid, which was significantly different from control muscles (P < 0.001) and from muscles treated with intralipid alone (P < 0.01).
Effect of Propofol on Uptake of Ca2+ into SR Vesicles
In control vesicles, in response to the addition of ATP, there was a rapid linear uptake of Ca2+ into SR vesicles, measured as a decrease in the fura-2 fluorescence signal (Fig. 6, Control). In response to increasing doses of propofol (5.6–560 μM), there was a significant, concentration-dependent decrease in the uptake of Ca2+ into SR vesicles (P < 0.05, Fig. 6). In the presence of equal volumes of intralipid, Ca2+ uptake into SR vesicles was unaffected (not shown).
Typical clinical peak serum propofol concentrations may be as large as 44 μM after bolus injection and 10 to 20 μM during maintenance infusion (15). At these concentrations, propofol induces hypotension which is primarily attributed to a decrease in sympathetic activity (2) and vasodilation (3). Earlier studies by Pagel et al. (6,16) suggested several mechanisms of propofol action, namely a direct negative inotropy, as well as direct effects on arterial and venous tone. Patrick et al. (17) suggested that the primary mechanism of propofol-induced hypotension is a decrease in peripheral vascular resistance, with little or no change in cardiac output. Several subsequent studies demonstrated that propofol has no significant negative inotropic action on the myocardium of the rat (7), dog (8), and rabbit (9). Gelissen et al. (18) studied the inotropic effects of propofol on human atrial muscles and demonstrated that a concentration-dependent direct negative inotropic effect in vitro manifests at concentrations far larger than those clinically used. Mouren et al. (9) showed that, even at supratherapeutic propofol concentrations (up to 1000 μM), propofol did not induce direct cardiac depressant effects in an isolated blood-perfused rabbit heart. Ismail et al. (8) demonstrated that propofol at clinically relevant blood concentrations has no direct negative inotropic effect in the in situ canine heart, although at supratherapeutic concentrations, it caused cardiac depression. These data are in agreement with the findings of our study. All of these observations suggest that the risk of direct myocardial depression after anesthetic induction with propofol is minimal; however, these studies do not exclude other possible mechanisms of hemodynamic compromise by propofol that may be present in vivo.
The effects of propofol have been studied in failing animal hearts (6,19,20), but never on the failing human myocardium. Hebbar et al. (19) studied the effects of 1–6 μg/mL propofol (5.6–33.6 μM) on failing pig hearts and found that propofol in vitro has a direct negative effect on basal myocyte contractile properties in the setting of pacing-induced congestive heart failure. Another study on cardiomyopathic hamster muscles at physiologic propofol concentrations (between 5.6 and 56 μM) but at a nonphysiologic temperature (29°C) showed that propofol exerted only a slight negative effect on the intrinsic mechanical properties of failing myocardium (20). Indirect cardiac effects such as modifications in venous return, afterload, compensatory mechanisms, and sympathetic activity may be more important than direct effects, even when cardiac function is impaired (6,20). Our study is the first to test the effects of propofol at physiologic temperature and rate of stimulation on the intrinsic contractility of failing human heart muscles. We found a concentration-dependent depression of contractility with propofol, which became statistically significant at concentrations much larger than achieved with typical clinical doses. At the same time, we found no significant difference between contractility of nonfailing atrial and failing atrial and ventricular muscles in response to propofol.
Although increased sympathetic tone and preserved adrenergic myocardial responsiveness can minimize anesthetic-induced negative inotropy in the normal myocardium, it is not known whether the same effects are present in failing human myocardium. Patients with end-stage heart failure have a reduced number of β1-adrenergic receptors (10), and in isolated cardiac preparations from failing human hearts, a reduced responsiveness to drugs acting on β-adrenoreceptors has been reported (21). Hebbar et al. (19), described decreased β-adrenergic responsiveness of pig myocytes exposed to 33.6 μM of propofol. In the present study, failing and nonfailing muscles have a preserved response to isoproterenol: 1 μM of isoproterenol increased the contractility of failing heart muscles exposed to intralipid to between 60% and 73% above baseline. At the same time, myocardium exposed to propofol exhibited reduced inotropic responsiveness: after 1 μM of isoproterenol, failing ventricular muscles recovered contractility to around baseline whereas the contractility of failing and nonfailing atrial muscles increased to between 36% and 44% above baseline. This recovery of contractility after propofol is similar to that found after etomidate in our previous study (12), and much higher than that measured after exposure to ketamine (13). At the same time, we did not find any significant effect of intralipid on human myocardial contractility, and the effects of this solvent were similar to those of buffer solutions as reported previously (12,13).
There are still doubts about the cellular mechanism of the negative inotropic effects of propofol at large concentrations. In the ferret myocardium, the negative inotropic effect was attributed to decreased calcium availability within the myocardium without a change in myofilament calcium sensitivity (22). Propofol does not have a major effect on sarcoplasmic reticulum function (23); therefore, the negative inotropic effects of propofol may involve interfering with transsarcolemmal calcium flux (22,24). Other investigators, who demonstrated a negative inotropic effect of propofol, have suggested that the effect is mediated primarily through its action on the inward calcium current (25,26) : at 5 and 30 μM of propofol, peak amplitude of inward calcium current was decreased by 10% and 21%, respectively. The results of Zhou et al. (27) suggest that propofol may inhibit cardiac L-type calcium current by interacting with the dihydropyridine binding site. The recent study by Nakae et al. (28) showed that propofol decreases available intracellular calcium without decreasing cardiac contraction, and enhances myofilament Ca2+ sensitivity even at clinical concentrations. It is also postulated that inhibition of available intracellular Ca2+ by propofol may be mainly mediated by an impairment of SR Ca2+ handling rather than the sarcolemmal L-type Ca2+ current. The finding of our study that propofol increases myofilament sensitivity to Ca2+ is somewhat surprising, because this mechanism of action should result in a net positive inotropic effect. Similar results have been reported for the effects of propofol in rat cardiac myocytes by Kanaya et al. (29) who showed that propofol caused a leftward shift in the concentration-response curve to extracellular Ca2+ with no effect on peak Ca2+ levels. Several investigators have demonstrated that propofol decreases the entry of Ca2+ into the cardiac myocyte during the action potential (29,30), and have speculated that this is the mechanism for the negative inotropic effect of propofol. Our data from the SR vesicle experiments demonstrate a direct effect of propofol on the uptake of Ca2+ by the SR. This intracellular mechanism of action could cause the negative inotropic effects of propofol on cardiac muscle, inasmuch as the decreased uptake of Ca2+ into the SR would make less Ca2+ available for release with the next action potential. However, because propofol simultaneously sensitizes the myofilaments to Ca2+, this would result in the same amplitude of contraction for a smaller amount of Ca2+. It is possible that the increased myofilament Ca2+ sensitivity, which others and we have shown with propofol, is actually a compensatory mechanism, which effectively offsets the decreased amounts of activator Ca2+. These findings are in agreement with those of previous investigators who studied the mechanism of action of propofol in the isolated rat cardiac myocyte (29) and the isolated guinea pig heart (28).
Most of our patients undergoing heart transplantation or myocardial revascularization were preoperatively treated with β-adrenergic agonists or β-adrenergic blockers until the morning of surgery. Although the muscle washout period in our experiments was substantial, the possibility exists that the long-term use of the above cardiac medications caused myocardial receptor alteration which may have changed the inotropic responsiveness of tissues. Despite this possibility, our results have significance because our experiments included a clinically relevant tissue source, i.e., all these interactions between diseased myocardium, preoperative cardiac medications, and propofol would normally be present in the actual clinical situation. Furthermore, we did not directly measure propofol concentrations in the tissue bath. Because a large fraction (98%) of propofol is bound to plasma proteins during clinical use, the concentration of free propofol in protein-free medium (tissue bath) may be larger than the concentrations of unbound (free) drug in blood.
In summary, our study revealed that propofol produces a concentration-dependent decrease in myocardial contractility at supratherapeutic concentrations in vitro. However, in concentrations equivalent to those used clinically, this negative inotropic effect was not significant even in failing myocardium. The mechanism of propofol-induced negative inotropy seems to be inhibition of Ca2+ uptake into the SR. At the same time, propofol increases myofilament sensitivity to Ca2+ at clinical concentrations and this mechanism may help to counteract the net negative inotropic effect of propofol.
The authors gratefully acknowledge the assistance of the cardiac transplant teams (Department of Cardiac Surgery) at the Cleveland Clinic Foundation and Life Banc of Northeast Ohio for aid in tissue procurement, as well as Wendy Sweet, Center for Anesthesiology Research, for help in preparing the figures. CSM is an Established Investigator of the American Heart Association.
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