Differential Effects of Fentanyl and Morphine on Intracellular Ca²⁺Transients and Contraction in Rat Ventricular Myocytes 

Isolated adult ventricular myocytes from rat hearts were obtained as previously described. [24]Briefly, the hearts were excised, cannulated via the aorta, attached to a modified Langendorff perfusion apparatus, and perfused with oxygenated (95% oxygen and 5% carbon dioxide) Krebs-Henseleit buffer (37 [degree sign]C) containing 118 mm NaCl, 4.8 mm KCl, 1.2 mm MgCl (²), 1.2 mm KH^2PO4, 1.2 mm CaCl², 37.5 mm NaHCO³, and 16.5 mm dextrose, with a pH of 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 (Worthington Biochemical, Freehold, NJ; lot M6C152; 347 units/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 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, with a pH of 7.35 and vigorously bubbled immediately before use with 100% oxygen. Typically, 6-8 x 10⁶cells/rat heart were obtained using this procedure. Viability, as assessed by the percentage of cells that retained a rod-shaped structure with no blebs or granulations, was routinely between 80% and 90%. Myocytes were suspended in HEPES-buffered saline (1 x 10 (⁶) cells/ml) and stored in an oxygen hood until they were used.

Simultaneously measurement of shortening and [Ca²⁺]ⁱwas performed as previously described. [25]Ventricular myocytes (0.5 x 10⁶cells/ml) were incubated in HEPES-buffered saline containing 2 [micro sign]m fura-2/AM (Texas Fluorescence Labs, Austin, TX) at 37 [degree sign]C for 20 min. Fura-2-loaded ventricular myocytes were placed in a temperature-regulated (28 [degree sign]C) chamber (Bioptechs, Butler, PA) mounted on the stage of an Olympus IX-70 (Olympus America, Lake Success, NY) inverted fluorescence microscope. The volume of the chamber was 1.5 ml. The cells were superfused continuously with HEPES-buffered saline at a flow rate of 2 ml/min and were field stimulated via bipolar platinum electrodes at a frequency of 0.3 Hz and a duration of 5 ms using a Grass SD9 stimulator (Grass Instruments, West Warwick, RI). Myocytes were chosen for study according to the following criteria:(1) a rod-shaped appearance with clear striations and no membrane blebs, (2) a negative staircase of twitch performance (typical for rats) when stimulated from rest, and (3) the absence of spontaneous contractions.

Fluorescence measurements were performed on single ventricular myocytes using a dual-wavelength spectrofluorometer (Deltascan RFK6002; Photon Technology International, South Brunswick, NJ) at excitation wavelengths of 340 and 380 nm and an emission wavelength of 510 nm. The cells also were illuminated with red light at a wavelength of more than 600 nm for simultaneous video edge detection. An additional postspecimen dichroic mirror deflects light at wavelengths of more than 600 nm into a charge-coupled device video camera (Phillips VC 62505T; Marshall Electronics, Culver City, CA) to measure myocyte shortening and relengthening. The fluorescence sampling frequency was 100 Hz, and data were collected using a software package (Felix) from Photon Technology International. The [Ca²⁺]ⁱwas estimated by comparing the cellular fluorescence ratio with fluorescence ratios acquired using fura-2 (free acid) in buffers containing known Ca²⁺concentrations.

Simultaneous measurement of cell shortening was monitored using a video edge detector (Crescent Electronics, Sandy, UT) with 16-ms temporal resolution. The video edge detector was calibrated using a stage micrometer so cell lengths during shortening and relengthening could be monitored. Myocytes typically contracted on one end with the other end lightly attached to the chamber. Thus contraction represented unloaded isotonic shortening. Myocyte length in response to field stimulation was measured (in micrometers) and is expressed as the change from resting cell length (twitch amplitude). Lab View (National Instruments, Austin, TX) was used for data acquisition of cell shortening using a sampling rate of 100 Hz.

Fluorescence data for the Ca²⁺transients were imported into Labview, and both the Ca²⁺transients and the myocyte contractile responses were analyzed synchronously and simultaneously. The following parameters were calculated for each contraction: diastolic and systolic [Ca (²⁺)]ⁱand cell length; change in [Ca²⁺]ⁱand twitch amplitude; time to speak (Tp) for [Ca²⁺]ⁱand shortening; and time to 50% recovery (Tr) for [Ca²⁺]ⁱand shortening. Parameters from 15 contractions were averaged to obtain mean values at baseline and in response to the various interventions. Averaging the parameters progressively minimizes beat-to-beat variation.

Changes in twitch amplitude in response to the interventions are expressed as a percentage of baseline shortening. Changes in timing were measured in milliseconds and were normalized to changes in amplitude. Changes in [Ca²⁺]ⁱwere measured as the change in the 340:380 ratio from baseline. Changes in the 340:380 ratio in response to the interventions were expressed as a percentage of the control response in the absence of any intervention.

Purification of Sarcoplasmic Reticulum Vesicles. Freshly isolated adult rat hearts were homogenized in five volumes of MOPS buffer (10 mm, pH 7.4, 4 [degree sign]C) containing 290 mm sucrose, 3 mm NaN³, 1 mm dithiothreitol, 1 [micro sign]m pepstatin A, 1 [micro sign]m leupeptin, and 0.8 mm phenylmethylsulfonyl fluoride using a Brinkmann Polytron homogenizer (Westbury, NY). The homogenate was centrifuged at 7,500g for 20 min. The supernatant was saved and centrifuged again at 40,000g for 60 min. The resultant pellet was suspended in three volumes of MOPS (10 mm, pH 6.8, 4 [degree sign]C) containing 600 mm KCl, 3 mm NaN³, 1 mm dithiothreitol, and protease inhibitors. The material was centrifuged at 140,000g for 40 min, and the final pellet was resuspended in a Ca²⁺-free sucrose buffer and stored at -80 [degree sign]C until it was used.

Measurement of Ca²⁺Uptake and Content in Sarcoplasmic Reticulum Vesicles. Double-distilled tap water was deionized using a Milli-Q reagent water system (Millipore, Bedford, MA) and further purified by dual ion-exchange chromatography and a Ca²⁺Sponge-S (Molecular Probes, Eugene, OR) to remove residual Ca²⁺. A buffering system representing intracellular conditions and capable of regenerating adenosine triphosphate was used to suspend the vesicles and contained 20 mm HEPES, 100 mm KCl, 5 mm NaCl, 5 mm MgCl², and 5 mm creatine phosphate (pH 7.2, 37 [degree sign]C) and creatine phosphokinase (0.4 units/ml). Oxalate (10 mm) was added to act as a Ca²⁺-precipitating anion inside the vesicles to minimize leakage of Ca²⁺and to maintain the Ca²⁺gradient across the vesicular membrane. [26]The solutions were prepared using an iterative solution-mixing program (Solwin v2.0, Philadelphia, PA). Binding constants for the ionic compounds were corrected for temperature and ionic strength. CaCl²was added back to the buffer to yield a free Ca²⁺concentrations of 1 [micro sign]m (pCa 6).

Measurements of Ca²⁺uptake and release were evaluated in real time using suspensions of SR vesicles and 2 [micro sign]m fura-2 free acid (Texas Fluorescence Labs) in the extravesicular compartment. Fluorescence experiments were performed using dual-wavelength fluorometry in a temperature-regulated (37 [degree sign]C) sample compartment. Microcuvettes (250 [micro sign]l) were washed in EGTA (2 mm) solution to remove all Ca²⁺and then thoroughly rinsed with Ca²⁺-free buffer and allowed to dry. The addition of adenosine triphosphate (1 mm) 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. Caffeine (20 mm) was used to release Ca²⁺from the vesicles to evaluate vesicular Ca²⁺content. Fluorescence data were collected using the Felix program at a sampling frequency of 20 Hz. The rate of Ca²⁺uptake was measured as the decrease in the fluorescence signal during a 45-s period in the presence or absence of opioid. The addition of 1,000 nm fentanyl or 100 [micro sign]m morphine did not alter the pH of the suspension buffer.

Protocols were designed so each cell could be used as its own control.

Protocol 1: Dose-dependent Effects of Opioids on [Ca²⁺]ⁱand Myocyte Shortening. Changes in myocyte shortening and [Ca²⁺]ⁱduring exposure to fentanyl or morphine were determined. Baseline measurements were collected from individual myocytes for 1.5 min in the absence of any intervention. Myocytes were exposed to sequential doses of the same opioid at four different concentrations (30, 100, 300, and 1,000 nm fentanyl; 3, 10, 30, and 100 [micro sign]m morphine). This was achieved by rapidly exchanging the buffer in the dish with new buffer containing the opioid at the desired concentration. Individual myocytes were exposed to only one opioid. Data were acquired for 1.5 min after a 5-min equilibration period in the presence of the opioid.

Protocol 2: Effects of Opioids on Sarcoplasmic Reticulum Ca²⁺Stores. To determine whether fentanyl or morphine alters Ca²⁺release from SR Ca²⁺stores, we measured caffeine-induced Ca²⁺release in the presence or absence of the opioid. Baseline [Ca²⁺]ⁱtransients were collected from individual myocytes for 1.5 min. Fentanyl (100, 1,000 nm) or morphine (10, 100 [micro sign]m) was then added to the superfusion buffer and allowed to equilibrate for 5 min. Field stimulation of the myocyte was discontinued and caffeine (20 nm) was applied to the cell 15 s later. The amplitude of the [Ca²⁺]ⁱtransient induced by caffeine was compared with the amplitude of the field-stimulated [Ca²⁺]ⁱtransient before the respective drugs were added and is reported as a percentage of the control amplitude.

Protocol 3: Effects of Opioids on Myofilament Ca²⁺Sensitivity. To determine whether fentanyl or morphine alters myofilament Ca (²⁺) sensitivity, we evaluated the dose-response curve to extracellular Ca (²⁺) in the presence or absence of the opioids. Baseline parameters were collected from individual myocytes for 1.5 min. Dose-response curves to extracellular Ca²⁺were performed by exchanging the buffer in the dish with a new buffer containing Ca²⁺at the desired concentration. Data were acquired for 1.5 min after a new steady state was established. Dose-response curves to extracellular Ca²⁺were then performed in the presence of either 100 nm fentanyl or 10 [micro sign]m morphine. Cells were allowed to stabilize for 5 min after each intervention. Changes in myocyte shortening and the [Ca²⁺]ⁱtransient were expressed as a percentage change from baseline in the control group. Similarly, changes in myocyte shortening and the [Ca²⁺]ⁱtransient in the presence of fentanyl or morphine were expressed as percentage changes from baseline 5 min after exposure to the opioids.

Fentanyl and morphine were obtained from the Cleveland Clinic Pharmacy. Caffeine was purchased from Sigma Chemical Company (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 all hearts were weighted equally. The dose-dependent effects of fentanyl or morphine on myocyte shortening and [Ca (²⁺)]ⁱwere assessed using one-way analysis of variance with repeated measures and the Bonferonni/Dunn post hoc test. Comparisons between groups were made by two-way analysis of variance. Results are expressed as the mean +/− SEM. Differences were considered significant at P < 0.05.

Baseline [Ca²⁺]ⁱand the diastolic cell length were 80 +/− 10 nm and 124 +/− 2 [micro sign]m, respectively. Peak [Ca²⁺]ⁱwas 360 +/− 30 nm. Twitch amplitude was 11%(14.0 +/− 0.7 [micro sign]m) of the baseline resting diastolic cell length. Time to peak [Ca²⁺]ⁱand shortening were 166 +/− 3 and 182 +/− 3 ms, respectively. The Tr for [Ca (²⁺)]ⁱand shortening was 307 +/− 4 and 326 +/− 5 ms, respectively. Baseline measurements were stable during the course of these experiments.

(Figure 1A) shows that the addition of fentanyl to an individual, field-stimulated ventricular myocyte results in dose-dependent inhibition of myocyte shortening and a concomitant decrease in peak [Ca²⁺]ⁱ. The myocardial depressant effects of fentanyl were completely restored after washout of fentanyl. Figure 1B shows individual contractions and [Ca²⁺]ⁱtransients. Figure 2displays the summarized data. Fentanyl caused dose-dependent decreases in myocyte shortening and peak [Ca (²⁺)]ⁱ. Fentanyl prolonged Tp and Tr for shortening and [Ca²⁺](ⁱ) at concentrations more than 30 nm (Figure 3).

(Figure 4A) shows that the addition of morphine to an individual, field-stimulated ventricular myocyte results in a decrease in myocyte shortening with no concomitant change in the amplitude of the [Ca²⁺]ⁱtransient. The inhibitory effect reached a plateau at approximately 10 [micro sign]m morphine, higher concentrations had no additional effect on shortening. At the highest concentration tested (100 [micro sign]m), morphine increased the amplitude of the [Ca²⁺]ⁱtransient. A decrease in diastolic [Ca²⁺]ⁱ(change in 340:380 ratio =-0.5 +/− 0.1) paralleled by an increase in diastolic cell length (3.0 +/− 0.2 [micro sign]m) was also observed (P < 0.05). The changes in myocyte shortening and [Ca²⁺]ⁱwere not immediately reversible after washout of morphine. Figure 4B shows individual myocyte twitches and [Ca²⁺]ⁱtransients. Figure 5displays summarized data. Morphine caused a decrease in myocyte shortening without a concomitant decrease in the amplitude of the [Ca²⁺]ⁱtransient, which was actually increased by 100 [micro sign]m morphine. Morphine prolonged Tp and Tr for myocyte shortening with no concomitant effect on Tp or Tr for [Ca²⁺]ⁱ(Figure 6).

Ca²⁺uptake by the vesicles was measured in real time as a decrease in the 340:380 ratio from the extravesicular compartment. Caffeine (20 mm) was used to release Ca²⁺from the vesicles. Figure 7A shows that fentanyl had no effect on the rate of Ca²⁺uptake at any concentration tested. Summarized data for fentanyl are shown in Figure 7B. Morphine only had an effect on the rate of Ca²⁺uptake into the SR vesicles at the highest concentration tested (Figure 8A). Summarized data for morphine are shown in Figure 8B. The total amount of Ca²⁺released from the vesicles in response to caffeine was unaltered be fentanyl (96 +/− 2% of control) or morphine (97 +/− 4% of control) compared with the vehicle control (Figure 7A and Figure 8A).

We also assessed the extent to which fentanyl or morphine altered the amount of Ca²⁺released from the SR in response to caffeine (20 mm) in intact cardiac myocytes. Fentanyl (100, 1,000 nm) did not alter the amplitude of the caffeine-induced [Ca²⁺]ⁱtransient compared with control (Figure 9). Only the highest dose of morphine (100 [micro sign]M) significantly decreased the caffeine-induced [Ca²⁺]ⁱtransient.

Myofilament responsiveness to Ca²⁺can be assessed by evaluating the relation between shortening and [Ca²⁺]ⁱ. To obtain a range of values for myocyte shortening and [Ca²⁺]ⁱ, we performed a dose-response curve to extracellular Ca²⁺([Ca²⁺]^o). Increasing [Ca²⁺]^ofrom 1 to 4 mm without opioids resulted in a dose-dependent increase in myocyte shortening and a concomitant increase in peak [Ca²⁺]ⁱ(Figure 10). Five minutes after 100 nm fentanyl, myocyte shortening and the [Ca²⁺]ⁱtransient decreased (P < 0.05) to 85 +/− 3% and 82 +/− 5% of preadministration values, respectively. Five minutes after 10 [micro sign]M morphine, myocyte shortening decreased (P < 0.05) to 93 +/− 2% of the preadministration value, whereas no change (100 +/− 3%) in the [Ca²⁺]ⁱtransient was observed. Fentanyl (100 nm) and morphine (10 [micro sign]M) caused a downward shift in the dose-response curve to [Ca²⁺]^ofor shortening with no concomitant effect on [Ca (²⁺)]ⁱ(Figure 10). Fentanyl and morphine caused a rightward shift in the relation between cell shortening and peak [Ca²⁺]ⁱ(Figure 11).

Previous studies that evaluated the effects of opioids on mammalian myocardial function yielded varying results, including evidence for positive [7,8]and negative [13,14,17]inotropic actions and no inotropic effect. [27]Those studies used intact animals, isolated perfused hearts, isometrically contracting papillary muscles, or left ventricular muscle strips. Thus, the experimental results probably reflect concomitant changes in preload, afterload, coronary blood flow and heart rate, and the effects of opioids to presynaptically modulate the release of norepinephrine and acetylcholine from nerve endings. To avoid these and other extrinsic factors, we used the individual, isolated ventricular myocyte preparation to evaluate the direct effects of fentanyl and morphine on cardiac excitation-contraction coupling at the cellular level.

In the current study, fentanyl had a direct negative inotropic action on isolated ventricular myocytes that was mediated, at least in part, by a decrease in peak [Ca²⁺]ⁱ. The cardiodepressant effect of fentanyl was reversible after washout of the opioid. Although most evidence suggests that fentanyl causes little change in myocardial contractility in vivo, several in vitro studies showed negative inotropic actions of fentanyl on cardiac contractility. In a canine blood-perfused papillary muscle preparation, fentanyl (95 [micro sign]M) reduced developed tension by 30%. [19]Taking into account binding of fentanyl to serum proteins, [28]the actual free plasma concentration in that study was approximately 20 [micro sign]M. In rat trabeculate carneae muscle, fentanyl (150 [micro sign]M) reduced peak developed tension by 30%. [15]Fentanyl (13 [micro sign]M) also depressed the velocity of isometric shortening of isolated cat papillary muscle by 30%. [16]These differences in concentrations for fentanyl-induced myocardial depression are probably because of species differences or the experimental preparation. Furthermore, it is difficult in multicellular preparations to exclude the possibility that fentanyl may alter the production of locally derived factors, which can regulate myocardial contractility.

In the current study, fentanyl (1 [micro sign]M) caused a 34% decrease in myocyte shortening and a concomitant 21% decrease in peak [Ca²⁺]ⁱ. Fentanyl (1 [micro sign]M) was recently shown to reduce the Ca²⁺current (I^Ca) in rabbit sinoatrial node by 20%. [29]Leucine enkephalin, an opioid receptor agonist, also has been shown to reduce the L-type Ca²⁺channel current in rat ventricular myocytes by 40%. [4]Therefore, the fentanyl-induced decrease in peak [Ca²⁺]ⁱin the current study may be a result of reduced Ca²⁺entry via L-type Ca²⁺channels.

Morphine causes bradycardia and hypotension via a direct effect on the central nervous system, which enhances parasympathetic nervous system outflow and inhibits sympathetic nervous system outflow. [30]However, the direct effects of morphine on intrinsic myocardial contractility have not been elucidated fully. Earlier in vivo studies reported that morphine induced a positive inotropic effect mediated by sympathoadrenal stimulation. [7,8]In contrast, other studies showed that morphine has a negative inotropic effect in isolated perfused heart and in atrial and ventricular muscle preparations. [13-17]In the current study, morphine decreased myocyte shortening, whereas [Ca²⁺]ⁱwas either unchanged or increased at the highest concentration of morphine. These results indicate that the cardiodepressant effect of morphine is not mediated by a reduction in Ca²⁺availability, but rather by a decrease in myofilament Ca²⁺sensitivity. The increase in [Ca²⁺]ⁱwith higher concentrations of morphine may act to counteract the reduction in shortening and is consistent with a previous observation in cultured cardiac myocytes. [18]In addition, morphine-induced myocardial depression exhibited a plateau effect that was difficult to wash out.

These findings suggest that morphine causes myocardial depression via a highly specific receptor-mediated pathway. A wide range of morphine doses was studied, because morphine is a mixed opioid receptor agonist and may activate multiple receptor subtypes with different affinities for the opioid. This could result in cross-talk between several signal transduction pathways, as previously described. [5]In preliminary studies, we observed that morphine-induced myocardial depression is completely prevented by naloxone, a mixed opioid receptor antagonist. [31] 

A prolongation in the time course for shortening by both opioids suggests changes in SR Ca²⁺dynamics. However, only fentanyl prolonged the timing of the [Ca²⁺]ⁱtransient. Thus, inhibition of SR Ca²⁺uptake could be one mechanism for fentanyl-induced myocardial depression but would not explain the effect of morphine. The fentanyl-induced prolongation of timing parameters could be caused by the concomitant decrease in peak [Ca²⁺]ⁱ, [32]or it could be a specific effect of fentanyl on SR Ca²⁺transport processes. To resolve this issue, we used isolated SR vesicles.

A decrease in the rate of Ca²⁺sequestered by the SR or a decrease in the amount of Ca²⁺available for release, or both, could be potential explanations for prolongation of the timing parameters for shortening by both opioids, as well as the depression in peak [Ca²⁺]ⁱobserved with fentanyl. Fentanyl had no effect on Ca²⁺uptake into isolated SR vesicles at any concentration tested, whereas morphine reduced Ca (²⁺) uptake only at the highest concentration tested. This is in contrast to our previous finding that thiopental directly alters the rate of Ca²⁺uptake into isolated SR vesicles in a dose-dependent manner. [33]Neither opioid had an effect on the amount of Ca²⁺released from SR vesicles in response to caffeine. These results indicate that fentanyl and morphine do not directly alter SR Ca²⁺dynamics, but this does not exclude a possible second-messenger-mediated effect of the opioids on SR Ca²⁺function in the intact cell. [34] 

Because second messengers, such as diacylglycerol, and activation of protein kinase C can be involved in altering SR Ca²⁺dynamics, [35,36]we wanted to determine whether the opioids altered the amount of Ca²⁺released from the SR of intact myocytes in response to caffeine. Caffeine-releasable Ca²⁺pools were unaltered by fentanyl pretreatment, indicating that the decreases in peak [Ca²⁺]ⁱand shortening were not caused by alterations in the amount of Ca²⁺released from the SR. These data are consistent with our findings in isolated SR vesicles. Therefore, the decrease in peak [Ca²⁺]ⁱinduced by fentanyl probably is related to inhibition of Ca²⁺influx across the sarcolemma. [4]Morphine decreased the amount of Ca²⁺released by caffeine at the highest concentration tested (100 [micro sign]m). Interestingly, morphine increased peak [Ca²⁺]ⁱin response to electric stimulation at the same concentration. Together, these results suggest that high-dose morphine may increase peak [Ca²⁺]ⁱby increasing Ca²⁺influx across the sarcolemma, [18]which could counteract its negative inotropic effect and refill depleted SR Ca²⁺stores.

In addition to decreased Ca²⁺availability, changes in myofilament Ca²⁺sensitivity also can alter cardiac contractile function. In the current study, morphine caused myocardial depression, independent of changes in peak [Ca²⁺]ⁱ. Furthermore, fentanyl and morphine both caused a downward shift in myocyte shortening without a concomitant change in peak [Ca²⁺]ⁱin response to elevated [Ca²⁺]^o. Therefore, opioid-induced myocardial depression may involve a decrease in the maximal response of the myofilament to Ca²⁺when [Ca²⁺]ⁱis increased. Ela et al. [18]observed that morphine decreased myofilament responsiveness to Ca²⁺in cultured neonatal rat cardiac myocytes and caused a downward shift in the cell motion-Ca²⁺transient relation induced by varying [Ca²⁺]^o. Our results in freshly dispersed adult myocytes are consistent with that study. [18]In addition, both opioids caused a rightward shift in the cell shortening versus [Ca²⁺]ⁱrelation, indicating a decrease in the affinity of the myofilament for Ca²⁺. Thus, a decrease in myofilament Ca²⁺sensitivity appears to be involved in opioid-induced myocardial depression.

Our results must be interpreted in the context of the experimental conditions (low temperature, 28 [degree sign]C; and low frequency of stimulation, 0.3 Hz). These conditions are necessary to maintain myocyte viability during these experiments. Peak plasma concentrations of 215 nm fentanyl have been reported after a 500-[micro sign]g intravenous injection in humans. [37]Assuming 80% protein binding, the peak concentration of non-protein-bound fentanyl would be less than 50 nm. Similarly, the total serum morphine concentration after intravenous bolus injection is approximately 10 [micro sign]m, resulting in a free plasma concentration of approximately 8 [micro sign]m because of 20% protein binding. [38]Thus, the concentrations of opioids in this study that caused significant cardiac depression are likely to encompass the concentrations encountered in the clinical setting. Serum protein levels, their capacity to bind opioids, or both may vary in certain pathologic conditions (e.g., hemodilution, liver disease, hypoproteinemia). Small changes in the amount or the binding capacity of proteins could result in an increase in the free plasma concentration of opioids. In addition, the myocardial depressant effect of fentanyl observed in this study may contribute to the profound hemodynamic depression observed during rapid infusion of high-dose fentanyl in the clinical setting.

The inhibitory effect of fentanyl on myocyte shortening appears to involve a decrease in the availability of intracellular free Ca²⁺and a decrease in myofilament Ca²⁺sensitivity. In contrast, the actions of morphine appear to be mediated primarily by a decrease in myofilament Ca²⁺sensitivity. Both opioids prolonged the timing for shortening, and fentanyl prolonged the timing for the Ca²⁺transient. Neither opioid had a direct effect on SR Ca²⁺uptake or content at clinically relevant concentrations. At high concentrations, morphine decreased the size of the caffeine-releasable Ca²⁺pool in intact cardiomyocytes.

The authors thank Cindy Shumaker for technical support and Ronnie Sanders and Cassandra Talerico for preparing the manuscript.