Mechanisms of Direct Inhibitory Action of Propofol on Uterine Smooth Muscle Contraction in Pregnant Rats

The experimental protocol used in this study was approved by the Sapporo Medical University Animal Care and Use Committee (Sapporo, Japan). One hundred thirty rats (Sprague Dawley, weighing 200–250 g) in the late stage of gestation (19–21 days after fertilization) were anesthetized with sevoflurane (3–4%). After a surgical level of anesthesia had been attained, the uteri were excised quickly and placed in a modified Krebs solution equilibrated with 95% O²–5% CO²at room temperature (22–24°C) (composition: 118 mm NaCl, 4.7 mm KCl, 21 mm NaHCO³, 1.2 mm MgSO⁴, 1.2 mm KH²PO⁴, 10 mm glucose, and 2.5 mm CaCl²; pH, approximately 7.4). The longitudinal smooth muscle layer of the uterus was carefully isolated by removing the endometrium and circular smooth muscle layer under a dissecting microscope.

In a preliminary study, the effect of a single dose of a high concentration of propofol (10⁻⁴m) on the oxytocin-receptor binding affinity was confirmed according to previously described methods. 25,26Briefly, smooth muscle tissue from a rat was minced in ice-cold 0.25 m sucrose buffer (10 times volume) with 10 mm Tris-HCl (pH 7.4) and homogenized twice with a homogenizer. The homogenate was centrifuged at 1,500 g  for 10 min at 4°C, and the supernatant was filtered through 120-μm nylon mesh and then centrifuged at 100,000 g  for 30 min at 4°C. The resting pellet was resuspended in 3 ml of 5 mm HEPES buffer with 1.0 mm MgSO⁴(pH 7.4), yielding a protein concentration of 500-100 μg/ml as measured by the method described by Lowry et al.  27 

Oxytocin receptor binding was determined using [³H]oxytocin (30–60 Ci/mmol). Nonspecific binding was determined in the presence of oxytocin (10⁻⁴m) and was subtracted from total binding to give specific binding. Labeled and unlabeled drugs were added as 20-μl aliquots to give a final assay volume of 540 μl. After incubation at room temperature with 10⁻⁴m propofol, the plate was placed on ice, and 50-μl aliquots of buffer were removed for determination of free (equilibrium) concentration of [³H]oxytocin. The remaining buffer was pipetted out, and the wells were washed twice for 5 min. The wells were finally drained, and punches were removed by touching with a small square of glass-fiber filter paper. Both were placed in a vial containing 1.8 ml scintillation fluid and counted. The density of receptors (B^max) and the dissociation constant for the ligand (K^d) were determined using linear regression and Scatchard transformation (n = 5 each).

The tissue was cut into small strips (2 mm wide and 15 mm long). The muscle strips were loaded with 5 μm acetoxymethyl ester of fura-2, an indicator of Ca²⁺, in a physiologic salt solution (PSS) containing 0.02% (vol/vol) cremophor EL for 3–4 h at room temperature. The PSS contained: 136.9 mm NaCl, 5.4 mm KCl, 1.5 mm CaCl², 1.0 mm MgCl², 23.9 mm NaHCO³, 5.5 mm glucose, and 0.01 mm EDTA. This solution was saturated with a gas mixture of 95% O²–5% CO²(pH, approximately 7.4). Each fura-2–loaded muscle strip was held in a temperature-controlled (37°C) organ bath, and one end of the muscle strip was connected to a strain gauge transducer (LVS-20GA; Kyowa, Tokyo, Japan). Experiments were performed using a fluorescence spectrometer (CAF-100; Japan Spectroscopic, Tokyo, Japan). Excitation light was passed through a rotating filter wheel (48 Hz) that contained 340- and 380-nm filters. The light emitted from the muscle strip at 500 nm was measured using a photomultiplier. The ratio of the fluorescence resulting from excitation at 340 nm to that at 380 nm (R^340/380) was calculated and used as an indicator of [Ca²⁺]ⁱ. 20,28 

Physiologic salt solution aerated with 95% O²–5% CO²was used for the control bath solution, and the uterine smooth muscle strips were allowed to equilibrate for 30 min after being mounted in the bath. To establish an optimal length, the resting tension was adjusted to 0.5 g. As previously reported, 29this value was selected as the optimal value for maximal active force generation determined in preliminary experiments using repeated oxytocin contractions and various baseline tensions. The tissue was contracted with half-maximal effect (ED⁵⁰) concentration of oxytocin (20 nm), a potent contractile agonist, released reflexively from the pituitary gland. After the force, frequency, and duration of contractions had reached a steady state, a single concentration of propofol (10⁻⁷–10⁻⁴m) was introduced into the tissue bath in the presence of oxytocin. To compare the quantitative effect of propofol with those of other intravenous anesthetics, ketamine (10⁻⁷–10⁻⁴m) or pentobarbital (10⁻⁷–10⁻⁴m) was also introduced into the tissue bath. Similar to this experiment, the tissue strips were exposed to 1 μm nifedipine, a dihydoropyridine-sensitive VDCC antagonist, or to Ca²⁺-free PSS with 5 μm EGTA during oxytocin-induced contraction.

To express the quantitative changes in muscle contractility and in [Ca²⁺]ⁱ, we measured the areas under the contraction and [Ca²⁺]ⁱcurves. 29,30Briefly, all data were digitized (2,000 samples/s) with an ITC-16 computer interface (Instrutech, Greatneck, NY) and stored on a hard disk. Subsequently, the digitalized data were analyzed using a 8100/100AV Power Macintosh computer (Apple, Cupertino, CA) to determine the areas under the contraction and [Ca²⁺]ⁱcurves for 5-min intervals (beginning 3 min after addition of each reagent to the organ bath) using Sigma Plot (Jandel Scientific, Corte Madera, CA).

The longitudinal smooth muscle tissues were cut into large pieces (15 mm wide and 20 mm long) and used for measurement of [IP³]ⁱ. Two or three pieces of the muscle tissues were incubated at 37°C in PSS containing 20 nm oxytocin for 20 min, in PSS containing 20 nm oxytocin for 10 min and subsequently in PSS containing 20 nm oxytocin and propofol (10⁻⁷–10⁻⁴m) for 10 min, or in PSS alone for 20 min. The muscle tissues were then frozen quickly in a liquid nitrogen and stored at −70°C until used for the assay.

The technique of Bredt et al.  21was used to measure [IP³]ⁱ. The frozen tissue sample was homogenized with 10% (wt/vol) ice-cold HClO⁴. The homogenates were centrifuged at 2,000 g  for 15 min at 4°C to remove insoluble materials. The pH of the supernatant was adjusted precisely to 7.5 with 1.5 m KOH containing 60 mm HEPES buffer. Insoluble precipitates (primarily KClO⁴) were removed by centrifugation at 2,000 g  for 15 min at 4°C. A 200-μl aliquot of the resultant supernatant was used to measure concentrations of protein. 27The extraction of IP³from the remaining resultant supernatant was conducted by solid-phase extraction procedures using Amprep SAX minicolumns (Amersham Pharmacia Biotech, Buckinghamshire, United Kingdom). [IP³]ⁱwas measured using a Biotrak D-myo -IP³[³H]assay system (code TRK 1,000; Amersham Pharmacia Biotech). This assay is based on competition between unlabeled IP³in the samples and a fixed quantity of [³H]IP³for a limited number of high-affinity binding sites on a specific IP³binding protein. 22The determinations were made in duplicate, and the results are expressed as picomoles per milligram of protein.

Ba²⁺was used, instead of Ca²⁺, to prevent an indirect inhibitory effect of oxytocin on VDCC activity, 31and conventional whole cell patch clamp techniques 23,24were used to observe inward Ba²⁺currents (I^Bas) through VDCCs. Uterine smooth muscle tissues were minced and digested for 20 min at 37°C in Ca²⁺-free modified Tyrode solution, to which 0.05% (wt/vol) collagenase (Lot#: 034-10533) was added. Cells were then dispersed by repeated aspiration into a plastic pipette, and debris was removed by filtration through 120-μm nylon mesh. The cell suspension was centrifuged at 200 g  for 3 min, and the pellet was resuspended in modified Kraftbrühe solution 32and stored at 4°C for up to 5 h before use. The modified Kraftbrühe solution contained: 85 mm KCl, 30 mm K²HPO⁴, 5.0 mm MgSO⁴, 5.0 mm Na²ATP, 5.0 mm pyruvic acid, 5.0 mm creatine, 20 mm taurine, 5.0 mm β-hydroxybutyrate, and 0.1% (wt/vol) fatty acid-free bovine serum albumin, with the pH adjusted to 7.25 with 0.5 m Tris.

The experiments were performed at room temperature (22–24°C). Micropipettes were pulled from soda lime tubing (GC-1.5; Narishige, Tokyo, Japan) using a brown-flaming horizontal puller (model P-97; Sutter Instrument, Novato, CA). These had resistances of 3–5 MΩ when filled with solution. An aliquot (approximately 0.5 ml) of the cell suspension was placed in a perfusion chamber on the stage of an inverted microscope (IX-70; Olympus, Tokyo, Japan). At ×600 magnification, a three-dimensional oil-driven micromanipulator (ONM-1; Narishige) was used to position the patch pipettes against the membrane of the uterine smooth muscle cells. After obtaining a high-resistance seal (5–20 GΩ) with slight suction (10–20 cm H²O), the patch membrane was disrupted by strong negative pressure, which allowed the voltage of the entire cell membrane to be controlled 23and permitted the pipette solution to diffuse into the cytoplasm. Membrane currents were monitored using a CEZ-2400 patch clamp amplifier (Nihon Kohden, Tokyo, Japan), and the amplifier output was low-pass filtered at 2,000 Hz. Leak currents, estimated by appropriate scaling of currents during 20-mV hyperpolarizing pulses, were subtracted from each of these records. Membrane capacitance and series resistance were compensated for by using the internal circuitry of the patch clamp amplifier. All data were digitized (10,000 samples per second), stored on a hard disk, and analyzed using a Power Macintosh computer (Apple) with Pulse+PulseFit 8.02 and Igor Pro 2.04 analysis software programs (Heka, Wiesenstrasse, Lambrecht, Germany).

To measure I^Bathrough VDCCs, recording solutions were chosen to inhibit K⁺currents and enhance Ba²⁺currents. The pipette solution contained: 20 mm CsCl, 110 mm CsOH, 5.2 mm MgCl², 112 mm l-glutamate, 10 mm EGTA, 5.0 mm Na²ATP, and 10 mm HEPES, with the pH adjusted to 7.2 with Tris. The bath solution contained: 130 mm tetraethylammonium chloride, 1.0 mm MgCl², 5.0 mm BaCl², 10 mm glucose, and 10 mm HEPES, with the pH adjusted to 7.35 with Tris. Whole cell I^Bas were elicited at 5-s intervals by 150-ms depolarizing pulses (−50 to +40 mV in 10-mV increments) from a holding potential of −60 mV. To identify the characteristics of the I^Baobserved in this study, the effect of an L-type VDCC antagonist, nifedipine (10⁻⁶m), on I^Bawas evaluated. Inactivation curves were determined using a double-pulse protocol that consisted of a 3-s prepulse to a potential of −60 to +10 mV, followed by a 150-ms depolarization to +10 mV. The peak change in the current during the test pulse was expressed as a fraction of that obtained with the −60-mV prepulse, and this quantity was fitted to a Boltzmann expression 33,34using least-squares analysis to estimate the potential of half-maximal inactivation (V1/2) and the slope factor (k). Data from the cells that showed unstable current amplitudes, less than 100 pA of peak I^Ba, or a greater than 10% reduction in amplitude during the control recording period were discarded. The GΩ seal was maintained for a period sufficient to evaluate the reversibility of the effects of propofol in 144 of 192 experiments (75%).

To measure outward K⁺currents (I^Ks), recording solutions were chosen to enhance the K⁺currents. The pipette solution contained: 70 mm KCl, 60 mm K⁺-glutamate, 5.0 mm K²ATP, 1.0 mm MgCl², 2.5 mm EGTA, 1.8 mm CaCl², and 10 mm HEPES, with pH adjusted to 7.2 with Tris. The computer-calculated [Ca²⁺]ⁱwas approximately 10⁻⁶m. The bath solution contained: 135 mm NaCl, 5.2 mm KCl, 1.8 mm CaCl², 1.0 mm MgCl², 10 mm HEPES, and 10 mm glucose, with pH adjusted to 7.35 with Tris. Whole cell I^Ks were elicited at 5-s intervals by 150-ms depolarizing pulses (−40 to +60 mV) from a holding potential of −60 mV.

Voltage-pulse protocols were performed in control solutions for more than 5 min to obtain a stable baseline. Cells were exposed to a single concentration of propofol (10⁻⁷–10⁻⁴m) with or without 20 nm oxytocin by changing the inflow perfusate of the chamber. The perfusion chamber consisted of a glass coverslip bottom, with needles placed for rapid solution changes. The chamber volume was approximately 1 ml, and complete solution changes in the chamber could be obtained within 1 min using a peristaltic pump (CTP-3; Iuchi, Tokyo, Japan) attached to the input and output ports. After 6-min exposure, the perfusate was switched again to the control solution.

The following drugs and chemicals were used: acetoxymethyl ester of fura-2, EDTA (Dojindo, Kumamoto, Japan), fatty acid-free bovine serum albumin, pyruvic acid, creatine, Na²ATP, taurine, β-hydroxybutyrate, pentobarbital, ketamine hydrochloride, nifedipine, EGTA, tetraethylammonium chloride, cremophor EL, oxytocin (Sigma Chemical, St. Louis, MO), collagenase (Wako Pure Chemical, Osaka, Japan), sevoflurane (Maruishi Pharmaceutical, Osaka, Japan), [³H]oxytocin (New England Nuclear, Boston, MA) and propofol (Diprivan^®; Astrazeneca, London, United Kingdom). Nifedipine and pentobarbital were dissolved in ethanol and in dimethyl sulfoxide, respectively (0.1% final concentration). We used commercially available propofol, which included 10% Intralipid^®(Pharmacia & Upjohn, Stockholm, Sweden) as the solvent (10%[vol/vol] soybean oil, 2.5% glycerol, and 1.2% purified egg lecithin). The effects of Intralipid per se  were therefore tested in each experiment, and the concentration of the solvent used alone corresponded to each concentration of propofol.

Data are expressed as mean ± SD. For the measurement of [Ca²⁺]ⁱand muscle tension, oxytocin-induced changes in [Ca²⁺]ⁱ(indicated by R^340/380) and muscle tension were used as references. 20,25Changes in peak I^Baor in the inactivation parameters V1/2 and k with exposure to propofol were compared at each applied potential by the paired, two-tailed t  test. Other data were analyzed using one-way analysis of variance for repeated measurements, and the Fisher exact test was used as a post hoc  test. In all comparisons, a P  value < 0.05 was considered significant.

Oxytocin binding experiments using [³H]oxytocin revealed saturable and high-affinity binding. Scatchard transformation of the data yielded linear plots from which B^maxand K^dcould be determined. Treatment of uterine cell membranes with a single dose of a high concentration of propofol (10⁻⁴m) had no effect on oxytocin receptor density or agonist affinity (n = 5 each, table 1). This result indicates that propofol does not interfere with the binding characteristics of the oxytocin receptors.

Even in the resting condition with PSS alone, oscillatory phasic contractions of the uterine smooth muscle strips were observed with a concomitant oscillatory increase in [Ca²⁺]ⁱ(fig. 2A). Oscillatory rates were constant (3.2 ± 0.8 cycles/min). Treatment with 20 nm oxytocin significantly increased oscillatory rates (4.5 ± 1.2 cycles/min) by approximately 41% and elevated the peaks of the contraction by approximately 35% and of the [Ca²⁺]ⁱby approximately 21%. When calculating the areas under the contraction and [Ca²⁺]ⁱcurves to express the quantitative changes in muscle contractility and in [Ca²⁺]ⁱ, 29,30introduction of 20 nm oxytocin increased muscle contraction by 123.5 ± 20.6% and [Ca²⁺]ⁱby 85.4 ± 12.6%. Nifedipine (1 μm), both in the resting condition or in the oxytocin-activated condition, completely blocked the oscillatory changes both in the muscle contraction and in [Ca²⁺]ⁱ(fig. 2A). Deletion of free Ca²⁺from the bath solution (with 5 μm EGTA) also blocked these oscillations (n = 3, data not shown). These results indicate that Ca²⁺influx through VDCCs is important for phasic contraction and increase in [Ca²⁺]ⁱ.

As shown in figure 2B, 10⁻⁴m propofol inhibited both the muscle contraction and increase in [Ca²⁺]ⁱinduced by 20 nm oxytocin. Propofol significantly decreased oscillatory rates by approximately 21% and suppressed the oxytocin-induced peaks of muscle contraction by approximately 34% and of [Ca²⁺]ⁱby approximately 41%. We determined the dose dependence of the inhibition of oxytocin-induced muscle contraction and increase in [Ca²⁺]ⁱby propofol, pentobarbital, and ketamine. Figure 3shows the relations of percent responses of oxytocin (20 nm)-induced contraction (fig. 3A) and increase in [Ca²⁺]ⁱ(fig. 3B) with concentrations of the anesthetics. Each intravenous anesthetic significantly decreased muscle contraction and [Ca²⁺]ⁱin a dose-dependent manner (n = 6 each). Based on total concentration in the solution, propofol and pentobarbital had similar potencies, whereas ketamine required somewhat greater concentrations to achieve the same inhibitory effect. These results indicate that propofol as well as pentobarbital and ketamine can inhibit oxytocin-induced uterine muscle contraction, at least in part, by decreasing [Ca²⁺]ⁱ, a main regulator of the contraction. 14,18 

The [IP³]ⁱin the resting condition was 4.2 ± 0.6 nmol/mg protein (n = 6). Because [IP³]ⁱis believed to oscillate phasically, as do muscle tension and [Ca²⁺]ⁱ, 18we did not measure the time course of [IP³]ⁱactivated by oxytocin. Instead, [IP³]ⁱat 20 min after the oxytocin activation was measured and found to be significantly elevated to 8.4 ± 1.2 nmol/mg protein. Figure 4summarizes the effects of various concentrations of propofol on [IP³]ⁱat 10 min after the propofol introduction (10⁻⁷–10⁻⁴m) during the condition of 20-min oxytocin incubation. Propofol significantly inhibited the increase in [IP³]ⁱinduced by oxytocin in a dose-dependent manner.

The inward Ba²⁺current (I^Ba) observed in enzymatically dispersed rat uterine smooth muscle cells during depolarizations from −60 mV peaked at approximately 10–15 ms and was inactivated slowly with a time constant of approximately 300–500 ms (fig. 5A: control). In baseline conditions, threshold activation of I^Baoccurred at approximately −40 mV, and maximum peak current amplitude was obtained at approximately +10 mV (fig. 5B). In 72 cells, the maximum peak I^Bawas −421 ± 46 pA (range, −259 to −514 pA). The inactivation parameters obtained in 24 cells in control conditions were V1/2 =−28.6 ± 3.3 mV and k = 5.1 ± 0.9 mV. The addition of 10⁻⁶m nifedipine virtually eliminated the I^Baof rat uterine smooth muscle cells by 91 ± 3% (data not shown, n = 4).

As shown in a representative trace for depolarization from −60 to +10 mV (fig. 5A: propofol), propofol (10⁻⁴m) inhibited the magnitude of I^Babut did not obviously alter the time course of the current. Figure 5Bshows the relation between peak I^Baagainst applied potential before and after exposure to 10⁻⁴m propofol. Propofol significantly inhibited I^Baat step potentials ranging from −20 to +40 mV and decreased peak I^Baat +10 mV by approximately 68%. There was no apparent shift in the voltage dependence of I^Bawith propofol. We determined the dose dependence of the inhibition of peak I^Baby propofol. Figure 5Cshows the relation between the percentage of control peak I^Baat +10 mV and the molar concentration of propofol in the bath solution. Propofol significantly inhibited peak I^Bain a dose-dependent manner (IC⁵⁰= approximately 3 × 10⁻⁶m). Introduction of 20 nm oxytocin had no effect either on the magnitude of I^Baor on the inhibitory effect of propofol (n = 4, data not shown). Figure 6shows the effect of 10⁻⁴m propofol on the inactivation curve of I^Ba. Propofol significantly shifted the inactivation curve to a more negative potential (V1/2 =−32.2 ± 2.3 mV) without changing the shape of the curve (k = 5.2 ± 1.1 mV).

Figure 7Ashows a macroscopic outward K⁺current (I^K) obtained from a freshly dispersed rat uterine smooth muscle cell dialyzed with a pipette solution containing a [Ca²⁺]ⁱof approximately 10⁻⁶m to enhance I^Kthrough Ca²⁺-activated K⁺channels. The I^Kwas activated progressively by 150-ms depolarizing pulses from a holding potential of −60 mV to consecutively more positive membrane potentials. Stepwise depolarization from a holding potential of −60 mV to more than −30 mV elicited an outward I^Kwith a mean peak amplitude of 2.11 ± 0.18 nA at +60 mV (fig. 7B, n = 6). The effect of propofol on I^Kwas examined in 24 cells. Only the highest concentration of propofol (10⁻⁴m) had a slight inhibitory effect on I^K(by approximately 8%) without any apparent effect on the time course of the current (fig. 7C).

We used Intralipid as the vehicle for propofol. Intralipid, the concentration of which corresponded to that used as the vehicle for propofol (10⁻⁴–10⁻⁷m), did not affect any parameters tested in this study (n = 4 each, data not shown). Neither ethanol nor dimethyl sulfoxide, the final concentration of which was less than 0.1%, affected any parameters tested in this study (n = 4 each, data not shown).

One of the major findings of our study is that propofol had a significant inhibitory effect on oxytocin-induced uterine smooth muscle contraction and increased [Ca²⁺]ⁱin pregnant rats without affecting the agonist-receptor binding affinity (table 1and figs. 2 and 3). For the agonist-receptor binding interactions, this result is in agreement with the general observations that volatile anesthetics 35–37or propofol 38,39have little effect on the binding of some other agonists. The result for muscle contraction and [Ca²⁺]ⁱis consistent with that obtained by Shin et al.  13using human pregnant uteri. Similar inhibitory effects were observed in this study with other intravenous anesthetics, ketamine and pentobarbital. The inhibitory effect of propofol on muscle tension was parallel to its inhibitory effect on [Ca²⁺]ⁱ. Because [Ca²⁺]ⁱplays a central role in the regulation of uterine smooth muscle tone, 14,15it is thought that propofol inhibits uterine smooth muscle contraction, at least in part, by decreasing [Ca²⁺]ⁱ.

Oxytocin receptor occupancy, coupled through G proteins, results in stimulation of the phosphatidylinositol-signaling pathway. 40Phillippe et al.  18,29,41demonstrated that activation of the phosphatidylinositol-signaling pathway results in the development of intracellular Ca²⁺oscillation-like phenomena. Similar intracellular Ca²⁺transients have been reported by other investigators in myometrium tissue and cultured uterine myocytes. 42–44When these phenomena occur, increased [Ca²⁺]ⁱis produced by both the release of stored intracellular Ca²⁺and the influx of extracellular Ca²⁺. Both events appear to be essential for the maintenance of repetitive myometrial contractions, 14,29,30because the introduction of Ca²⁺-free solution or the blockade of Ca²⁺influx through VDCCs by nifedipine completely blocked the oscillatory contractions in this study. Therefore, the mechanisms by which propofol decreased [Ca²⁺]ⁱare thought to be inhibition of the phosphatidylinositol-signaling pathway or inhibition of VDCC activity. To investigate the mechanisms, [IP³]ⁱand the changes in inward Ba²⁺currents (I^Ba) through VDCCs in uterine smooth muscles obtained from pregnant rats were examined using radioimmunoassay and whole cell patch clamp techniques, respectively.

The results of the current experiment showed that propofol inhibited the increase in [IP³]ⁱin a dose-dependent manner (fig. 4). The decrease in [IP³]ⁱcaused by propofol lead to a decrease in [Ca²⁺]ⁱ, resulting in inhibition of the uterine contraction. This result is in agreement with that in a study on canine smooth muscle 39showing that treatment with propofol is associated with inhibition of the increase in [Ca²⁺]ⁱmediated by IP³. Hirakata et al. , 45however, showed that high concentrations of propofol (3 × 10⁻⁵–10⁻⁴m) induced formation of IP³in human platelets. This discrepancy may result from the differences in cell types and species or in the selective effects of propofol on certain receptors, G proteins, or phospholipase C isozymes. 46 

As shown in figure 1, stimulation of the oxytocin receptor activates the G protein-linked phospholipase C, resulting in hydrolysis of phosphatidylinositol 4,5-bisphosphate to the two potent stimulatory second messengers IP³and 1,2-diacylglycerol. 47,48IP³is rapidly metabolized to inositol 1,4-bisphosphate and inositol 1,3,4,5-tetraphosphate by IP³phosphatase and IP³kinase, respectively. Therefore, it is possible that propofol affects any point of the phosphatidylinositol-signaling pathway (e.g. , inhibition of G protein–phospholipase C or activation of IP³phosphatase–kinase) to decrease [IP³]ⁱ. Nagase et al.  49and Minami et al.  50reported that propofol had an inhibitory effect on receptor–G protein coupling of muscarinic and 5-hydroxytryptamine receptors, respectively; however, further studies are needed to clarify this point in uterine smooth muscle. There is the evidence that IP³also facilitates the influx of extracellular Ca²⁺through specific channels in the cell membrane. 47,48,51Therefore, inhibition of the increase in [IP³]ⁱby propofol might in itself account for the observed effects on both [Ca²⁺]ⁱand muscle contractility.

Using whole cell patch clamp techniques, we measured depolarization-induced inward Ba²⁺currents (I^Ba) in rat uterine smooth muscle cells. Based on their time and voltage dependences and their sensitivity to blockade by nifedipine, these currents are presumed to reflect the activity of L-type VDCCs. 52,53Young et al. , 52however, reported that freshly isolated human uterine smooth muscle cells showed two types of Ca²⁺currents (T and L types) that were analogous to those found in cardiac myocytes. Because it has been reported that the resting membrane potential is approximately −60 mV in the late stage of gestation, 52the L-type VDCCs might be primarily involved in increasing [Ca²⁺]ⁱby bulk Ca²⁺transport. 54 

Propofol significantly inhibited I^Bathrough L-type VDCCs of the uterine smooth muscle cells without an apparent change in the voltage dependence of I^Ba(fig. 5), suggesting that the anesthetic has no effect on membrane surface charge or on the voltage sensor of the channel. These data indicate one of the cellular effects of propofol that can account for the uterine smooth muscle relaxant effects of the anesthetic. 13To further evaluate the inhibitory action of propofol on VDCCs of uterine smooth muscle cells, we studied the effects of propofol on steady state, voltage-dependent inactivation of I^Ba. During prolonged depolarization (prepulse), a fraction of the VDCCs enters an unavailable or “inactivated” state. Propofol significantly shifted the inactivation curve to a more negative potential without changing the sigmoid shape of the curve (fig. 6). A qualitatively similar shift induced by a dihydropyridines-sensitive Ca²⁺antagonist, such as nifedipine, has been interpreted as evidence of a drug-induced stabilization of the inactivated state. 55We therefore speculate that propofol has a dihydropyridines-sensitive Ca²⁺antagonist-like inhibitory effect on VDCCs in uterine smooth muscle cells.

Two other possibilities for the inhibitory mechanisms of propofol on uterine smooth muscle contraction should be considered. Franks and Lieb 56reported that volatile general anesthetics activated a novel neuronal K⁺current. The resulting enhanced K⁺efflux can induce membrane repolarization or hyperpolarization, reduce the open-state probability of VDCCs, and in turn cause relaxation of uterine smooth muscle. In this study, the whole cell outward I^Kwas inhibited significantly by only a high concentration (10⁻⁴m) of propofol (fig. 7). This result suggests that mechanisms other than K⁺channel opening are likely to mediate propofol-induced relaxation.

Another possibility is protein kinase C. As previously described, stimulation of the oxytocin receptor can produce 1,2-diacylglycerol as well as IP³. Increased 1,2-diacylglycerol can activate protein kinase C. Although it has been known that protein kinase C has an inhibitory effect on IP³production and plays a role in generation of intracellular Ca²⁺oscillations in some cell types, 57,58there is no clear evidence of protein kinase C playing a role in uterine smooth muscle contractility. In airway smooth muscle, it is conceivable that protein kinase C plays a role in Ca²⁺-independent smooth muscle contraction. 20,28Further studies are needed to clarify the role of protein kinase C in uterine smooth muscle and the effect of propofol on the protein kinase C activity.

Propofol, ketamine, and pentobarbital each showed concentration-dependent inhibition of uterine smooth muscle contraction and [Ca²⁺]ⁱ(fig. 3). Based on the total solution concentration, ketamine was less potent than the other agents by a factor of approximate 3. Because propofol was added as an emulsion, it is likely that its free concentration was substantially less than the total concentration and that this drug is, in fact, more potent than pentobarbital for inhibition of muscle contraction. We suggest that the potency order of these agents is propofol > pentobarbital > ketamine. For the measurement of IP³concentrations and VDCC activity, a significant decrease and inhibition of these parameters were observed using the same range of concentrations as that used for investigation of contractions and [Ca²⁺]ⁱ(10⁻⁷–10⁻⁴m). Extrapolation of our data to the clinical situation must be viewed with caution because of possible species differences, in vivo –in vitro  differences, and the fact that our patch clamp experiments were conducted during nonphysiologic conditions of low (ambient) temperature and high (5 mm) extracellular Ba²⁺concentration.

Given that more than 95% of propofol (2–5 × 10⁻⁵m) in blood bound to plasma protein, 59the free propofol concentration of 4–10 × 10⁻⁷m in vitro , which has little effect on muscle tension, would be comparable to 9–10 μg/ml propofol in vivo . 7Because the therapeutic ranges of plasma propofol concentrations were 2–9 μg/ml, 59the effective propofol concentrations tested in this study were rather higher than the free concentrations observed clinically in serum. Whether the direct decrease in muscle tension produced by propofol would affect uterine atony–bleeding during cesarean section needs further study.

In conclusion, propofol at concentrations ranging from 10⁻⁷to 10⁻⁴m significantly reduces the oxytocin-induced contraction of uterine smooth muscle obtained from pregnant rats in isolated preparations. The inhibition of the contraction by propofol is, at least in part, caused by the decrease in [Ca²⁺]ⁱwithout inhibition of agonist-receptor binding. The decrease in [Ca²⁺]ⁱmay be mediated by a decrease in [IP³]ⁱand by an inhibition of VDCC activity.

The authors thank Noritsugu Tohse, M.D., Ph.D. (Professor and Chairman, Department of Physiology Section I, Sapporo Medical University School of Medicine, Sapporo, Japan) for his valuable review and comments on the manuscript; and Xiangdong Chen, M.D. (Visiting Scientist, Department of Anesthesiology, Sapporo Medical University School of Medicine, Sapporo, Japan), and Naoyuki Fujimura, M.D., Ph.D. (Instructor, Department of Anesthesiology, Sapporo Medical University School of Medicine, Sapporo, Japan), for technical assistance.