Alterations in Ca2+ cycling by lysoplasmenylcholine in adult rabbit ventricular myocytes

Shi J. Liu1, Richard H. Kennedy1, Michael H. Creer2, and Jane McHowat2

1 Departments of Pharmaceutical Sciences and Pharmacology and Toxicology, University of Arkansas for Medical Sciences, Little Rock, Arkansas 72205; and 2 Department of Pathology, St. Louis University Medical School, St. Louis, Missouri 63104


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ABSTRACT
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We previously reported that lysoplasmenylcholine (LPlasC) altered the action potential (AP) and induced afterdepolarizations in rabbit ventricular myocytes. In this study, we investigated how LPlasC alters excitation-contraction coupling using edge-motion detection, fura-PE3 fluorescent indicator, and perforated and whole cell patch-clamp techniques. LPlasC increased contraction, myofilament Ca2+ sensitivity, systolic and diastolic free Ca2+ levels, and the magnitude of Ca2+ transients concomitant with increases in the maximum rates of shortening and relaxation of contraction and the rising and declining phases of Ca2+ transients. In some cells, LPlasC induced arrhythmias in a pattern consistent with early and delayed aftercontractions. LPlasC also augmented the caffeine-induced Ca2+ transient with a reduction in the decay rate. Furthermore, LPlasC enhanced L-type Ca2+ channel current (ICa,L) and outward currents. LPlasC-induced alterations in contraction and ICa,L were paralleled by its effect on the AP. Thus these results suggest that LPlasC elicits distinct, potent positive inotropic, lusitropic, and arrhythmogenic effects, resulting from increases in Ca2+ influx, Ca2+ sensitivity, sarcoplasmic reticular (SR) Ca2+ release and uptake, SR Ca2+ content, and probably reduction in sarcolemmal Na+/Ca2+ exchange.

calcium; lipid metabolites; excitation-contraction coupling; heart


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INTRODUCTION
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AN INCREASE IN membrane-associated, Ca2+-independent phospholipase A2 (PLA2) activity in mammalian cardiac ventricular myocytes occurs during short intervals of hypoxia (20) or after exposure to interleukin-1beta , a proinflammatory cytokine (19). Activation of membrane-associated, Ca2+-independent PLA2 results in selective hydrolysis of membrane plasmalogen phospholipids and accumulation of lysoplasmenylcholine (LPlasC; see Ref. 20). Lipid metabolism is altered within seconds of myocardial ischemia, and significant accumulation of amphiphilic metabolites such as LPlasC and the structurally similar compounds lysophosphatidylcholine (LPC) and palmitoylcarnitine have been demonstrated within 2 min (15, 21). Under these conditions, the ischemic heart often exhibits electrophysiological abnormalities, including ventricular arrhythmias within 1-2 min (15, 21). Perfusion of normoxic cardiac myocytes with LPC and palmitoylcarnitine causes changes in the action potential (AP) configuration, suggesting that these amphiphilic metabolites are potentially arrhythmogenic. Moreover, inhibition of the accumulation of these lipid metabolites in the ischemic myocardium is associated with a decrease in arrhythmogenesis (8). Recently, we have demonstrated that LPlasC also exerts an arrhythmogenic effect on normoxic ventricular myocytes by altering the AP configuration, thereby inducing early and delayed afterdepolarizations at concentrations significantly lower than those reported previously for LPC and palmitoylcarnitine (20).

The arrhythmogenic effect of LPC has been attributed to its ability to exert a nonspecific effect on the biophysical properties of membrane phospholipids, resulting in an inhibition of most cardiac currents (for review, see Refs. 21 and 27). Similar to LPC, palmitoylcarnitine blocks inward-rectifier K+ channels (IK1; see Ref. 22), the Na+ current (23), the L-type Ca2+ channel current (ICa,L; see Ref. 30), and the Na+/K+ pump current (25) in mammalian ventricular myocytes. In contrast, palmitoylcarnitine was also shown to activate a slow-inactivating Na+ current followed by an increase in a transient inward current (31) but lacked effects on IK1 and transient outward K+ currents (32). Nevertheless, these LPC- and palmitoylcarnitine-induced electrophysiological changes could not satisfactorily account for the observed concomitant positive inotropic effect (1, 25) and the increased intracellular Ca2+ (Cai; see Ref. 33). In contrast to LPC and palmitoylcarnitine, little is known about the cellular mechanism underlying LPlasC-induced changes in cardiac contractile and electrical function.

Disturbance of Ca2+ handling such as that elicited by increasing Ca2+ influx or Ca2+ release from sarcoplasmic reticulum (SR), and/or by decreasing Ca2+ efflux, leads to Cai overload that is often associated with cardiac arrhythmias (for review, see Ref. 7). Thus, in this study, we investigated the effects of LPlasC on contraction, Ca2+ transients, and membrane currents to assess its actions on excitation-contraction coupling in adult rabbit ventricular myocytes. We found that LPlasC exerted potent, distinct effects on contractile and electrical functions in ventricular myocytes. These effects could be accounted for by LPlasC-induced alterations in Cai handling.


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Myocyte isolation. The protocol for the use of animals in this study conformed with the National Institutes of Health approved Guide for the Care and Use of Laboratory Animals and was approved by the Institutional Animal Care and Use Committee, University of Arkansas for Medical Sciences.

Single adult ventricular myocytes were isolated from the hearts of adult rabbits (either sex, 2-3 kg), as described previously (20). Isolated ventricular myocytes were harvested and plated in 60-mm culture dishes (Falcon) for 2 h or overnight in culture medium composed of 60% medium-199 (GIBCO, Grand Island, NY) and 36% Earle's balanced salt solution containing (in mM) 116 NaCl, 4.7 KCl, 0.9 NaH2PO4, 0.8 MgSO4, 26 NaHCO3, 5.6 glucose, and 4% FBS (pH 7.40 in 5% CO2-95% air at 37°C; GIBCO), as described previously (17). Rod-shaped cells with clear striations were used for experiments, and there was no significant difference in the response to lipid metabolites between freshly isolated and primary cultured myocytes (within 24 h). All experiments were carried out at 35-37°C.

Measurement of cell shortening. Unloaded cell shortening (CS) or contraction of myocytes was elicited in normal Tyrode solution containing (in mM) 140 NaCl, 5.4 KCl, 1 CaCl2, 0.8 MgCl2, 10 HEPES/Tris, and 5.6 glucose (pH = 7.40 at 37°C; 290 mosmol/kgH2O) through field stimulation with bipolar platinum electrodes at a frequency of 0.5 Hz with 1- to 2-ms voltage pulses. Cells were then superfused with normal Tyrode solution containing ethanol, followed by solutions with different concentrations of LPlasC. The stimulating electrode was placed near the suction pipette in the perfusion chamber to minimize its damage to cells during long periods of stimulation. The video signal was fed into the video motion detector (Crescent Electronics, Sandy, UT) connected to a video monitor through a charge-coupled device video camera mounted to a microscope. The analog voltage output was calibrated to indicate actual micrometers of cell motion and recorded on a personal computer using pClamp software (Axon Instruments). Measured parameters of contractile function in myocytes included peak magnitude of CS, maximum rates of contraction (+dL/dtmax) and relaxation (-dL/dtmax), and rise and decay times between 10 and 90% of peak CS.

Measurement of intracellular free Ca2+ concentration. Ventricular myocytes seeded on 25-mm coverslips in culture medium were loaded for 30 min in a culture incubator at 37°C with 2 µM fura-PE3-AM (TEFLABS, Austin, TX), a cell-permeable form of fura-PE3 that is a new analog of fura 2 and is retained inside the cell longer than fura 2. Myocytes were then transferred to a recording/perfusion chamber (Harvard Apparatus, Holliston, MA) on the stage of an inverted microscope (model TE300; Nikon, Irving, TX) and superfused with normal Tyrode solution. Fluorescent measurements were made through a ×40 long-working-distance ultraviolet (UV) objective (Nikon Fluor with numerical aperture of 1.3). Fura-PE3-loaded cells were alternately excited with UV light at 340- and 380-nm wavelengths via a filter wheel, controlled by a spectrophotometry unit (Cairn Research) at 60-75 Hz. The emitted fluorescence signal at 510 nm was collected through an adjustable diaphragm and a photomultiplier tube (Cairn) to the spectrophotometer control unit. The signals were sampled at 200-300 Hz using pClamp software (Axon Instruments) and stored in a personal computer for later calibration and analysis. After subtraction of the background signal, fluorescent signals were recorded as the ratio (R or f340/f380) of the fluorescent intensity when excited at 340 nm (f340) to that when excited at 380 nm (f380). Because of difficulties with the in vivo calibration procedure, many results were represented as f340/f380. The measured parameters of the Ca2+ transient included peak magnitude, maximum rates of the rising phase (+dR/dtmax) and the declining phase (-dR/dtmax), and the rise and decay times between 10 and 90% of peak amplitude. In some experiments with successful in situ calibrations, cytosolic free Ca2+ concentrations were determined using the equation (13) [Ca+]i = Kd × beta  × (R - Rmin)/(Rmax - R) where Kd is the apparent dissociation constant of 224 nM at 37°C and beta  is the ratio of f380,free to f380,bound measured under Rmin and Rmax conditions, respectively. Rmin is the minimum fluorescent ratio in Ca2+-free solutions containing 3 µM ionomycin and 10 mM EGTA, and Rmax is the maximum intensity ratio in perfusion buffer solution containing 3 µM ionomycin and 2 mM CaCl2. In some experiments, myocyte contraction was recorded simultaneously with Ca2+ transients when the cells were illuminated with a halogen lamp (Nikon) through a long-wavelength pass filter (640 nm; Chroma).

Electrophysiological measurements. Ventricular myocytes were perfused with normal Tyrode solution and patch-clamped using perforated-patch (16) or conventional whole cell patch techniques (14) with a patch-clamp amplifier (Axopatch 200A; Axon Instruments; see Refs. 17 and 18). APs of myocytes were measured in normal Tyrode solution and K+-rich pipette solutions only in perforated-patch clamp configurations, as described previously (18, 20). The current-voltage (I-V) relationship of the steady-state membrane current was obtained by applying 300-ms voltage step pulses to potentials between -120 and +80 mV from the holding potential in 20-mV increments at 0.2 Hz or a voltage ramp between -120 and +80 mV at a rate of 1 V/s.

Whole cell ICa,L was measured as described previously (17, 18). The I-V relationship of peak ICa,L was constructed using 25- or 250-ms voltage pulses to potentials between -60 and +70 mV from the holding potential of -40 mV in 10-mV increments. The kinetics of ICa,L inactivation in response to a voltage pulse to +10 mV were analyzed using a double-exponential equation: y(t) A0 + A1e-t/tau f + A2e-t/tau s, where tau f and tau s are the fast and slow time constants, respectively, and Ax is the amplitude scalar. In other experiments, to isolate ICa,L more effectively, ICa,L or Ba2+ current (IBa) was measured in myocytes voltage-clamped at -70 mV under Na+- and K+-free conditions, as described previously (17). The voltage dependency of steady-state inactivation and activation were determined using a gapped double-pulse protocol one time every 2 s; a 250-ms prepulse to potentials between -90 and +50 mV was followed by a 5- or 10-ms return to the holding potential and then a fixed 250-ms test pulse to +10 mV. Raw data from each experiment of the voltage dependency of steady-state inactivation and activation of ICa,L were curve fit by a Boltzmann equation using the Marquardt-Levenberg nonlinear least-squares curve-fitting algorithm included in the Origin program (OriginLab, Northampton, MA).

Synthesis of LPlasC. LPlasC was prepared by alkaline hydrolysis of bovine heart choline glycerophospholipids, as described previously (9). The LPlasC product was isolated by column chromatography on a 2.5 × 60-cm column of silica using a stepwise gradient elution procedure. According to our previous study (20), the majority of experiments in this study used 1 µM LPlasC, which elicited apparent cardiac effects, unless otherwise indicated.

Chemicals. Most reagents were purchased from Sigma Chemical (St. Louis, MO) and were added directly when needed. Stock solutions of lipid metabolites (10-2 M) were prepared in 100% ethanol. The final concentration of ethanol in extracellular solutions was <0.01% and had no significant effect (<10%) on measured parameters.

Statistics. In all experiments, data in response to LPlasC were compared with the steady-state control before treatment in each individual cell and thus expressed as a ratio or percentage of each control value before combining for statistical analysis. Values are presented as means ± SE. Statistical significance was evaluated by the two-tailed Student's paired t-test or, when more than two conditions were compared, by one- or two-way ANOVA with Duncan's multiple-range test. Differences with P < 0.05 were considered significant.


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Effects of LPlasC on contraction of intact ventricular myocytes. We first examined the effect of LPlasC on the unloaded contractile function of intact ventricular myocytes under physiological conditions. Figure 1 shows that LPlasC increased contractility, which reached a plateau in 5 min and was followed by a small decrease. In combined data, LPlasC elicited a maximum positive inotropic effect (2.59 ± 0.44-fold increase, n = 25) in 3-8 min (varied from cell to cell), followed by a decline (83.9 ± 4.3% of the maximum, n = 18). After the baseline to the control level was offset, Fig. 1C shows superimposed CS before, during, and after exposure to 1 µM LPlasC, as shown in Fig. 1B, each of which was obtained by averaging five to six shortening traces. The first derivative of contraction traces (Fig. 1C, inset) shows that +dL/dtmax and -dL/dtmax were increased dramatically during exposure to LPlasC. Combined data in Fig. 1E show that LPlasC caused a 3-fold and 4.3-fold increase in +dL/dtmax and -dL/dtmax, respectively, concomitant with reductions in the rise time and decay time of contraction. Normalized traces (to each peak amplitude) as shown in Fig. 1D confirmed that LPlasC had a more profound effect on -dL/dtmax than +dL/dtmax, and combined data show 41 and 18% increases in -dL/dtmax and +dL/dtmax, respectively (Fig. 1E). Thus LPlasC elicited positive inotropic and lusitropic effects in adult rabbit ventricular myocytes. Figure 1 also shows that, after removal of LPlasC, a rebound stimulation of contraction was observed before returning to the control level. Nine of twenty-five tested myocytes showed the rebound activation of contraction (1.32 ± 0.11-fold of the amplitude before LPlasC removal), whereas nine cells did not (also see Fig. 2). The remaining cells did not survive during LPlasC exposure or washout because of arrhythmia and/or contracture.


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Fig. 1.   Effect of lysoplasmenylcholine (LPlasC) on contraction in adult ventricular myocytes. A: time course of inotropic effects of 1 µM LPlasC in a myoycte in normal Tyrode solution. Upon LPlasC removal, a rebound stimulation of contraction was observed before returning to the control level. B: traces of steady-state contraction as indicated in A on an expanded time scale. C: superimposed traces (average of 5 steady-state contractions) in three conditions, as shown in B. D: superimposed scaled traces (normalized to peak amplitude) from C. Insets in C and D, the first derivative of individual traces before and during LPlasC exposure. E: combined data of LPlasC-induced changes in the parameters of contraction from 25 cells. +dL/dtmax, maximum rate of contraction; -dL/dtmax, maximum rate of relaxation. Data are means ± SE. LPlasC elicited significant effects on all parameters (P < 0.005, Student's paired t-test).



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Fig. 2.   Effects of LPlasC on Ca2+ transients in ventricular myocytes. Cell shortening (CS; A) was recorded simultaneously with Ca2+ transients (B) in one myocyte in response to LPlasC stimulation. The disruption of CS recordings when switching to LPlasC-containing solution was caused by a temporary loss of video contrast resulting from air bubbles. f340/f380, ratio of fluorescence at 340 to 380 nm. C and D: steady-state CS and Ca2+ transients traces obtained where indicated in A and B, respectively. In C, an aftercontraction (*) was developing in the presence of LPlasC. D, inset, the first derivative of the Ca2+ transient before and during LPlasC exposure. E: combined data of LPlasC-induced relative changes in parameters of Ca2+ transients from 29 cells. LPlasC elicited significant effects on all parameters (P < 0.005, except for the decay time: P < 0.05).

Effects of LPlasC on the Ca2+ transient in fura-PE3-loaded ventricular myocytes. The effect of LPlasC on Cai handling in fura-PE3-loaded myocytes was examined under conditions similar to those described for CS. Figure 2 shows results from simultaneous measurements of CS (Fig. 2A) and the Ca2+ transient (Fig. 2B) in a myocyte. LPlasC caused rapid increases in systolic and diastolic Ca2+ levels, and the magnitude of the Ca2+ transient, accompanied by a fourfold increase in CS, an inotropic response similar to that observed in non-fura-PE3-loaded cells. The time course for the increase in Ca2+ transients appears to be more rapid than that of the positive inotropy (tau : 63.8 vs. 91.2 s), suggesting that LPlasC increases Ca2+ cycling and alters the Ca2+ sensitivity of contractile machinery. Upon removal of LPlasC, Ca2+ transients and contraction partially recovered without a rebound activation. Steady-state Ca2+ transients and contraction before and during LPlasC exposure were superimposed and shown in Fig. 2, D and C, respectively. It is noticeable in Fig. 2C that an aftercontraction was developing 8 min after exposure to LPlasC compared with the control. In addition, Fig. 2D, inset, shows that LPlasC increased +dR/dtmax and -dR/dtmax of the Ca2+ transient. The LPlasC-induced increases in systolic and diastolic f340/f380 were 0.041 ± 0.004 and 0.019 ± 0.002 (n = 29), respectively, which approximated 340 and 80 nM of free Cai, respectively. Combined data in Fig. 2E show that LPlasC increased the magnitude of the Ca2+ transient approximately twofold and doubled +dR/dtmax and -dR/dtmax. It is also worth mentioning that the declining phase of the Ca2+ transient in the presence of LPlasC was better fit by a biexponential function, whereas it was best fit by a single exponential in control conditions. The LPlasC-associated initial rapid phase (i.e., -dR/dtmax) and later slow phase of the f340/f380 decline were more rapid and slower than the decay time constant in control, respectively. As a consequence, LPlasC increased the area under the Ca2+ transient and the decay time (Fig. 2E).

A phase-plane plot of contraction as a function of the simultaneously measured f340/f380 (using data in Fig. 2, C and D) shows a hysteresis relationship between CS and free Cai (or fluorescence ratio). Figure 3 shows that LPlasC shifted the hysteresis relationship upward and to the right. When this relationship was replotted using relative changes in CS vs. f340/f380 (normalized to each peak magnitude), a left shift in the contraction-Cai trajectory during the early phase of relaxation is revealed, suggesting an increase in the myofilament Cai sensitivity (Fig. 3, inset), as described previously by others (26). Meanwhile, the slope of the relaxation in this hysteresis relationship in LPlasC displayed two phases, consistent with the biexponential process of the decline phase in the Ca2+ transient described previously.


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Fig. 3.   LPlasC-induced change in the relationship between CS and intracellular free Ca2+. A hysteresis relationship between CS and intracellular free Ca2+ (fluorescence ratio) in the absence and presence of 1 µM LPlasC was obtained from a phase-plan plot of CS as a function of simultaneously recorded Ca2+ transients obtained from data shown in Fig. 3. Inset: superimposed plots of the same data prescaled to each peak amplitude.

The role of SR Ca2+ load in the LPlasC-induced increase in systolic and diastolic Cai was also examined in fura-loaded myocytes using caffeine (10-15 mM for ~5 s) pulses that have been used to estimate SR Ca2+ content (4, 12). Figure 4A shows that, in the presence of LPlasC, caffeine induced a greater Ca2+ transient. Figure 4B shows that LPlasC slowed the decay of the Ca2+ transient in the presence of caffeine, an indirect index of sarcolemmal Na+/Ca2+ exchange activity (2), compared with control. Combined data in Fig. 4F show that LPlasC increased the time constant of the decay of the caffeine-induced Ca2+ transient ~65%. Figure 4, D and E, summarizes data showing that LPlasC caused a 40-60% increase in the magnitude of the caffeine-induced Ca2+ transient when estimated using either the resting level (termination of electrical stimulation) or diastolic level as baseline. Results also show that the fractional Ca2+ release from SR in the steady-state twitch (i.e., a ratio of the magnitude of steady-state systolic Ca2+ transients to that of the caffeine-induced Ca2+ transient) was greater in the presence of LPlasC than in control. Note that the ~40% fractional Ca2+ release observed in control was consistent with the 43% fractional SR Ca2+ release reported previously by others using rabbit ventricular myocytes (4).


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Fig. 4.   Effect of LPlasC on sarcoplasmic reticulum (SR) Ca2+ content in ventricular myocytes. A: SR Ca2+ content monitored by applying 10 mM caffeine before (a) and during (b) exposure to 1 µM LPlasC. B: caffeine-induced Ca2+ transients (the second peaks) measured under control (a) and LPlasC (b) conditions from A on an expanded time scale. The decline phase of the caffeine-induced Ca2+ transient was best fit with a single exponential function; fit curves (dashed lines) were superimposed with respective traces. C: recovery of systolic Ca2+ transients upon caffeine removal in control (open circle ) and LPlasC () conditions, which were also curve-fit with a single exponential function. D and E: summarized data of LPlasC effects on SR Ca2+ content and fractional Ca2+ release, estimated by the magnitude of the caffeine-induced Ca2+ transients using a resting level (D) or the steady-state diastolic level (E) as reference (insets). The relative amplitude of SR Ca2+ content in the presence of LPlasC was normalized to the amplitude in control in each cell (filled bars). Fractional SR Ca2+ release during regular contraction was obtained by dividing the amplitude of the steady-state Ca2+ transient (SS) by the amplitude of the caffeine-induced Ca2+ transient before (open bars) and during (hatched bars) exposure to LPlasC. Nos. in bars are no. of experiments. F: LPlasC-induced changes in the time constant (tau ) of the decline phase of the caffeine-induced Ca2+ transient (hatched bar), the amplitude of the first electrically elicited Ca2+ transient (filled bar) upon caffeine removal, and the tau  of systolic Ca2+ recovery (open bar). Values presented were normalized to each control. * P < 0.01; ** P < 0.005.

The time course of recovery of the systolic Ca2+ transients after removal of caffeine (postcaffeine) was shown in Fig. 4C. Results show that 1) the magnitude of the first postcaffeine Ca2+ transient in response to electrical stimulation during exposure to LPlasC was greater than that in control, suggesting an increased Ca2+ influx, and 2) the rate of the recovery to a new steady state was best fit with a single exponential with a time constant of 8.6 and 9.5 s in the presence of LPlasC and in control, respectively. Summarized data in Fig. 4F show that LPlasC increased the magnitude of the first postcaffeine Ca2+ transient by ~50%, whereas the rate of the recovery of Ca2+ transients from caffeine exposure did not differ significantly.

Arrhythmiogenic effect of LPlasC. In many cells, LPlasC induced arrhythmias with different patterns and severity that varied from cell to cell. For example, 1 µM LPlasC elicited mild arrhythmias in one cell (Fig. 5A), whereas it induced more severe arrhythmias in another cell (Fig. 5B). In Fig. 5A, inset, potentiated contractions were preceded by early aftercontractions and oscillations. In contrast, primarily delayed aftercontractions were observed in the cell shown in Fig. 5B. Figure 5C shows Ca2+ transients during LPlasC-induced arrhythmias; an apparent increase in systolic Cai was followed by Ca2+ transients in a pattern consistent with early and delayed aftercontractions 2 min after exposure to LPlasC. Figure 5C also shows that SR Ca2+ load was reduced after the arrhythmia but gradually returned to the control level upon removal of LPlasC. When the concentration of LPlasC was increased to 10 µM, severe arrhythmias and contracture occurred in <1 min in all four cells tested (data not shown).


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Fig. 5.   LPlasC-induced arrhythmias in ventricular myocytes. A: LPlasC-induced mild arrhythmias in a myocyte. B: a more severe arrhythmia induced by 1 µM LPlasC in another myocyte. C: Ca2+ oscillations occurred after increases in systolic and diastolic free Ca2+ (*) in the presence of 1 µM LPlasC. The caffeine-induced Ca2+ transient on LPlasC removal was initially decreased before its recovery. Insets in A, B, and C reveal different patterns of arrhythmias (*) on an expanded time scale.

Effects of LPlasC on ICa,L in patch-clamped ventricular myocytes. We next examined whether LPlasC-elicited increases in systolic Cai and postcaffeine Ca2+ influx result from an enhancement of ICa,L. In a perforated-patch configuration, ICa,L was monitored before, during, and after exposure to LPlasC in normal Tyrode solution. Figure 6A shows that the I-V curve of peak ICa,L was increased ~10% (measured at +10 mV) in 20 s and reached a maximum (~20%, measured at 0 mV) after 80-130 s of exposure to 1 µM LPlasC. In addition, it seemed that LPlasC had a profound stimulatory effect at potentials between -20 and 0 mV; e.g., it shifted the maximum peak ICa,L in response to depolarizing pulses from +10 to 0 mV. Figure 6A also shows that LPlasC gradually increased peak outward current (measured at +60 mV) and caused a leftward shift in the zero-current potential. The LPlasC-induced changes in ICa,L were reversible after washout (data not shown); however, this was preceded by a transient increase in current amplitude (rebound stimulation of ICa,L; Fig. 6A). Superimposed selected current traces (measured at -10, 0, and +10 mV) recorded before, during, and after exposure to LPlasC are shown in Fig. 6B. Summarized data show that LPlasC increased ICa,L by 22 ± 3% (n = 5) within 1-2 min, whereas its rebound during LPlasC removal was 34 ± 6% (n = 3) above the control level. The LPlasC-induced increase in ICa,L and rebound stimulation are consistent with its inotropic effects, as shown in Fig. 1A.


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Fig. 6.   Effects of LPlasC on the current-voltage (I-V) relationship of peak ICa,L. A: I-V relationships of ICa,L obtained before, during, and after exposure to LPlasC by applying 25-ms voltage pulses to potentials between -50 and +60 mV from a holding potential of -40 mV using a perforated-patch configuration. The use of 0.2 mM Cd2+ to block ICa,L was for clarification of ICa,L in normal Tyrode solution. B: superimposed current traces elicited by voltage pulses to potentials of -10, 0, and +10 mV before (C), during (L), and after (W) exposure to 1 µM LPlasC. Cell membrane capacitance: 94 pF. C: I-V curves of ICa,L obtained before (open circle ), during (), and shortly after (triangle ) removal of 1 µM LPlasC using a conventional whole cell configuration. I-V curves were constructed by scaling maximum ICa,L to 1 in each cell before exposure to LPlasC.

Measurement of ICa,L and its rebound using the perforated whole cell patch-clamp configuration was often very difficult because of cell contracture and death. Thus some experiments were carried out in the conventional whole cell configuration. Figure 6C shows the I-V relationships of ICa,L (normalized to the peak amplitude of ICa,L at +10 mV before LPlasC exposure) before, during, and after exposure to 1 µM LPlasC. Similar to findings obtained using the perforated-patch configuration, the results show that LPlasC increased ICa,L by 35 ± 8% in 2 min (n = 7, P < 0.002) and induced a rebound activation by 54 ± 23% (n = 4) upon its removal. In contrast to results in the perforated-patch recordings, no significant shift in the I-V curve of ICa,L (between -20 and 0 mV) was observed in the whole cell configuration.

LPlasC-induced changes in ICa,L were also examined in Na+- and K+-free solutions to minimize the contamination by currents associated with Na+ and K+. Results showed that LPlasC increased ICa,L 1.07 ± 0.03 (n = 3)-, 1.12 ± 0.02 (n = 11)-, and 1.23 ± 0.06 (n = 5)-fold at 0.1, 1, and 2 µM, respectively. The increased ICa,L was then followed by a reduction to 68.6 ± 4.6% (n = 7) of the control level in the continued presence of 1 µM LPlasC. Subsequent removal of LPlasC also induced a transient rebound activation of ICa,L by 37.1 ± 8.6% (n = 6), similar to that observed in the presence of Na+ and K+, as shown in Fig. 6. The LPlasC-induced changes in the I-V relationship of ICa,L in Na+- and K+-free conditions were also comparable to that observed in normal Tyrode solutions. Meanwhile, outward currents were increased by +83.7 ± 8.3% (n = 8, measured at +60 mV) during a long exposure (>2 min) to LPlasC. We also used Ba2+ as the charge carrier to replace Ca2+ in the perfusion solution to minimize Ca2+-activated changes in current measurements and further confirm the LPlasC-induced changes in ICa,L. Under these conditions, 1 µM LPlasC elicited a transient increase in IBa followed by a 30% decline after 3 min. Removal of LPlasC caused a 27% rebound activation of IBa above the control level.

The effects of LPlasC on the kinetics and steady-state activation and inactivation of ICa,L were then examined using the conventional whole cell configuration. ICa,L was more stable in normal Tyrode solution during exposure to LPlasC; thus its inactivation was analyzed in normal Tyrode solution. The inactivation of ICa,L was best characterized as a double-exponential function, consistent with findings reported previously in Na+- and K+-free solutions containing 2 mM Ca2+ or Ba2+ (17). Results show that LPlasC accelerated the fast time constant (tau f, from 4.2 ± 0.3 to 3.7 ± 0.4 ms, n = 6, P < 0.05, paired t-test) but slowed the slow time constant (tau s, from 19.6 ± 0.8 to 24.3 ± 0.8 ms, n = 6, P < 0.005) of ICa,L inactivation in response to a voltage pulse to +10 mV. These results are consistent with those observed in perforated-patch recordings (see Fig. 6B). In contrast, LPlasC increased both tau f and tau s of IBa inactivation by 15 ± 2 and 5 ± 2%, respectively (n = 4). Such discrepancy in effects on the tau f of ICa,L and IBa inactivation probably results from the initial increase in ICa,L induced by LPlasC, which enhances Ca2+-dependent inactivation.

Effects of LPlasC on the voltage dependency of steady-state inactivation (finfinity ) and activation (dinfinity ) of ICa,L were determined in Na+- and K+-free solutions before and during exposure for >3 min to 1 µM LPlasC when ICa,L reached a quasi-steady state. Figure 7 shows that LPlasC caused a 5-mV depolarizing shift in holding potential (Vh) of finfinity (from -17.1 ± 0.8 mV, n = 6, in control, to -12.3 ± 0.8 mV, n = 5, P < 0.005, paired t-test) without a significant change in the slope factor (k; control: 5.1 ± 0.2 mV; LPlasC: 5.8 ± 0.4 mV). Interestingly, LPlasC decreased the state of inactivation from 0.25 to 0.46 at +50 mV (i.e., after a prepulse potential of +50 mV a greater ICa,L was observed in LPlasC). Results also show that LPlasC elicited a 6-mV depolarizing shift in Vh of dinfinity (from -1.7 ± 1.5 mV, n = 5, in control, to +4.5 ± 1.8 mV, n = 5, P < 0.001) with a significant increase in k values (control, 6.4 ± 0.1 mV; LPlasC, 7.2 ± 0.3 mV, P < 0.05). The LPlasC-induced depolarizing shift in dinfinity resulted in an increase in ICa,L window current.


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Fig. 7.   LPlasC-induced alterations in the voltage dependency of steady-state inactivation and activation of ICa,L. Steady-state inactivation (finfinity ) and activation (dinfinity ) of ICa,L were determined using a gapped double-pulse protocol (see MATERIALS AND METHODS) and curve-fit by Boltzmann equations as follows: I/Imax = 1/[1 + exp(V - Vh)/k], where Vh is the half-maximum inactivation potential and k is the slope factor for finfinity , and G/Gmax = 1/{1 + exp[-(V - Vh)/k]}, where Vh is the half-maximum activation potential for dinfinity , G is conductance, and Gmax is maximal conductance. Data represent means ± SE from 5-6 experiments.

Effects of LPlasC on steady-state membrane currents. Figure 8A demonstrates that nearly identical quasi-steady-state I-V relationships were generated using voltage-pulse or voltage-ramp protocols in a ventricular myocyte. With the use of the voltage-ramp protocol, Fig. 8B shows I-V relationships of the steady-state membrane current before, during, and after exposure to 1 µM LPlasC. Results show that, after 5 min of exposure, LPlasC caused a 60-80% increase in the steady-state outward current in the voltage range between +40 and +80 mV without altering IK1 (measured between -70 and -110 mV). Similar to its effect on the peak outward current, the effect of LPlasC on the steady-state outward current was irreversible. Comparable results were obtained using the perforated-patch configuration in which 1 µM LPlasC increased the steady-state outward current (measured at +60 mV) 2.1 ± 0.2-fold without significant change in IK1 (n = 4).


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Fig. 8.   Effect of LPlasC on steady-state membrane currents in adult ventricular myocytes. A: in conventional whole cell patch-clamp recordings, currents were elicited in normal Tyrode solution using a voltage-step pulse (inset on right) and ramp (inset on left; see MATERIALS AND METHODS) protocol. The I-V curve was constructed by plotting the current amplitude at the end of test pulses (open circle ) vs. corresponding test potentials or the current magnitude in response to a hyperpolarizing voltage ramp (indicated by arrow). B: steady-state whole cell membrane currents elicited using a voltage-ramp protocol (as shown in the inset of A) before (open circle ), during (), and after (triangle ) exposure to 1 µM LPlasC for 5-7 min. EtOH, ethanol. Data represent means ± SE.

Effects of LPlasC on AP and cell contraction in patch-clamped ventricular myocytes. Figure 9 shows selected traces obtained from continuous, simultaneous recordings of AP (A) and contraction (B) in a myocyte before, during, and after exposure to LPlasC. Exposure for 3 min to 0.1 µM LPlasC caused 35 and 16% prolongations of AP duration at 25 and 75% of repolarization, respectively (Fig. 9A), concomitant with a small increase in systolic shortening (Fig. 9B). A subsequent increase in the concentration of LPlasC to 1 µM resulted in a substantially prolonged AP duration accompanied by a ~10-mV depolarization of diastolic membrane potential and a 20% increase in the magnitude of contraction at 40 s. When the diastolic membrane potential depolarized dramatically toward -20 mV in the presence of LPlasC (e.g., an afterdepolarization was observed at 45 s), contraction became smaller. The basal (diastolic) level of contraction was increased gradually with time during exposure to LPlasC, consistent with CS measured in intact cells (Fig. 1A) and the increased diastolic Cai (Fig. 2B). Upon rapid removal of LPlasC, AP partially recovered, whereas cell contraction was transiently increased by 134% before returning to the control level, similar to that shown in Fig. 1A. Similar results were observed in three other experiments.


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Fig. 9.   LPlasC-induced changes in action potential (AP) and simultaneously recorded contraction. Selected AP (A) and CS (B) traces, which were elicited and acquired at 1 Hz, were obtained at indicated time points before, during, and after exposure to LPlasC in one experiment of simultaneous and continuous recordings of AP and CS. Continuous traces of the whole experiment were recorded on chart paper. Because the acquisition duration for each trace was set ~255 ms, some APs with a longer duration were only partially acquired.

Although the time course and exact pattern of LPlasC-induced changes in AP configuration varied from cell to cell, LPlasC-induced changes in AP duration were consistent with our previous findings (20). The occurrence of diastolic membrane potential depolarization (toward approximately -20 mV) and afterdepolarization was 68 and 53%, respectively, in 1 µM LPlasC (n = 19) and increased to 100 and 82%, respectively, during exposure to 2-10 µM LPlasC (n = 11). The LPlasC-induced membrane depolarization was also concentration and time dependent, e.g., depolarization to -29.5 ± 5.2 mV (n = 9), -22 ± 4.0 mV (n = 5), and -13.3 ± 1.4 mV (n = 3) was observed at 1, 3, and 10 µM, respectively. The onset of membrane depolarization induced by 10 and 1 µM LPlasC occurred within 15 s and 3 min, respectively. Afterdepolarizations (without sustained depolarization) were observed at 0.1 µM LPlasC in only one out of nine experiments. When afterdepolarization and membrane depolarization occurred, cells tended to round up and die unless LPlasC was removed immediately.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

LPlasC accumulates after activation of a membrane-associated, Ca2+-independent PLA2 in ventricular myocytes during short intervals of hypoxia and is a potent arrhythmogenic lipid metabolite (20). With the use of multiple technical approaches, the present study demonstrated that 1) LPlasC elicits positive inotropic, positive lusitropic, and arrhythmogenic effects in adult ventricular myocytes, 2) LPlasC-induced changes in contractile function are paralleled by its effects on intracellular free Ca2+ concentration and SR function, 3) LPlasC increases Ca2+ influx and ICa,L, and 4) LPlasC-induced changes in contractile function are accompanied by anticipated effects on AP. The increased Ca2+ influx, SR Ca2+ content, and SR Ca2+ release lead to AP prolongation and augmented Ca2+ transients, thereby increasing contractility and/or inducing arrhythmias.

LPlasC increases the systolic and diastolic state of cell contraction with maximum effects in 5-6 min, followed by a small decrease. Simultaneous recordings in fura-PE3-loaded cells showed that LPlasC-induced increases in systolic and diastolic free Ca2+ levels precede its effect on contraction. LPlasC increases ICa,L, and the first electrically elicited Ca2+ transient after SR Ca2+ has been emptied by caffeine, supporting our hypothesis that LPlasC-induced inotropic effects are initiated by an increase in Ca2+ influx via ICa,L. In addition, the time course for CS to reach a quasi-steady state was longer than that for the Ca2+ transient during exposure to LPlasC (Fig. 2A), suggesting an incremental increase in the Ca2+ sensitivity of contractile proteins. The LPlasC-induced left shift of the CS-Cai trajectory during early relaxation supports this suggestion and is consistent with an increase in myofilament Ca2+ sensitivity demonstrated previously by others using adult rat ventricular myocytes (26).

Our data also show that LPlasC increased ICa,L by ~12%, smaller than the ~100% increase in the Ca2+ transient of regular twitch and the 43% increase in the first postcaffeine electrically evoked Ca2+ transient. Several possibilities could account for this difference in the LPlasC-induced changes in these parameters. First, ICa,L might be underestimated because the observed concomitant increase in outward currents masks the true magnitude of the increased ICa,L. Second, LPlasC-induced prolongation of ICa,L inactivation, increases in channel availability and ICa,L window currents, and reduction in ICa,L steady-state inactivation during depolarization would enhance Ca2+ influx and thereby increase Ca2+ transients to values greater than the measured increase in ICa,L. Third, LPlasC amplifies calcium-induced calcium release by increasing available SR Ca2+ release and/or the Ca2+ content of SR (11). This is supported by our data showing increases in the magnitude of caffeine-induced Ca2+ transient, fractional Ca2+ release, and +dR/dtmax elicited by LPlasC. Fourth, LPlasC reduces Ca2+ efflux via sarcolemmal Na+/Ca2+ exchange, thereby increasing net Ca2+ gain in each cycle (11). This possibility is supported by our data showing that LPlasC increases the diastolic Ca2+ and shortening level. Fifth, LPlasC decreases Cai buffering power, i.e., the same Ca2+ influx and SR Ca2+ release results in a greater increase in Cai. Studies in rat ventricular myocytes have suggested that a decrease in Ca2+ buffering power at elevated Cai, such as that induced by caffeine, increases the decay rate of free Ca2+ (10). This seems unlikely because our data showed that LPlasC slows the decay rate of the caffeine-induced Ca2+ transient. Taken together, LPlasC increases Ca2+ influx via L-type Ca2+ channels, SR function, and myofilament Ca2+ responses.

The LPlasC-induced elevation in diastolic Cai could account for the increased diastolic state of contraction. The return of Cai to baseline during diastole depends primarily on SR Ca2+ uptake (contributing 70% and being a fast component) and the normal mode of Na+/Ca2+ exchange (28% and a slow component) in rabbit ventricular myocytes (2, 3). Our data showed that LPlasC increases -dCa2+/dtmax, attributable to an enhanced SR Ca2+ uptake, but prolongs the slow phase of the Ca2+ transient, attributable to a reduced Na+/Ca2+ exchange activity. These changes were paralleled by a LPlasC-induced shift in the decay phase of the Ca2+ transient from a single to a double exponential function. In addition, LPlasC caused a 60% increase in the time constant of the decay phase of the caffeine-induced Ca2+ transient, an indirect measure of sarcolemmal Na2+/Ca2+ exchange resulting from the absence of SR uptake function (2). Moreover, in the presence of 10-15 mM caffeine, the steady state of free Cai during LPlasC exposure was ~20 nM higher than that in control (Fig. 4B). Thus these results support the notion that LPlasC decreases Ca2+ efflux via sarcolemmal Na+/Ca2+ exchange.

An intriguing finding was rebound stimulation of contraction (Figs. 1 and 9) and ICa,L (Fig. 6) observed upon removal of LPlasC in some myocytes. The phenomenon of rebound ICa,L stimulation has been shown during withdrawal of ACh (28, 29) and carbachol (CCh), a muscarinic agonist (5). This rebound activation of ICa,L was suggested to be responsible for the initiation of delayed afterdepolarizations in cat atrial myocytes (29) and to result from an increase in cAMP that is mediated by nitric oxide-induced cGMP-mediated inhibition of phosphodiesterase (29). In contrast, a study in ferret right ventricular myocytes showed that a cGMP-dependent pathway is not involved in the rebound ICa,L stimulation observed upon removal of CCh (5). In the present study, the rebound activation of contraction and ICa,L upon LPlasC removal was paralleled by a rapid recovery of AP duration from the preceding shortened AP duration. Interestingly, we have not observed any rebound stimulation of Ca2+ transients under the same experimental conditions. Comparable changes in ICa,L were obtained using perforated- and conventional patch-clamp recordings; however, the role of intracellular second messengers in LPlasC removal-induced rebound could not be completely excluded and requires further investigation.

LPlasC at 1 µM also elicits arrhythmias in intact myocytes, fura-loaded cells, and patch-clamped cells. The pattern of arrhythmia is consistent with a combination of early and delayed afterdepolarizations. LPlasC-induced arrhythmias apparently result from an increase in free Cai and membrane potential depolarization. Both early and delayed afterdepolarizations can be elicited by Cai overload (for review, see Ref. 7). Our data show that LPlasC increases SR Ca2+ release and could thereby cause depolarization of membrane potential and delayed afterdepolarization, as demonstrated by others (24). Thus LPlasC-induced Cai overload could account for its arrhythmogenic effect.

It is also worth mentioning that the LPlasC-induced increase in ICa,L was detected in 1-2 min and followed by a decline, whereas contraction and Ca2+ transients continue to rise to a plateau in 5-6 min, followed by a reduction. The discrepancy in the time course of LPlasC-induced changes in these two parameters could have resulted from a disturbance of the cell membrane in patch-clamped myocytes, which becomes more severe during LPlasC exposure. For example, the arrhythmias induced by LPlasC are more severe and occur more often in perforated-patch-clamped cells than in those with the conventional whole cell configuration or in intact myocytes. Combined effects of lipid metabolites and ionophores (nystatin and amphotericin B) could account for the disruption of membrane stability. In some cells, increases in peak and steady-state outward currents were observed during prolonged exposure to LPlasC (Fig. 6, A and B). This could have been mediated by Cl- currents and/or a nonselective current because the zero-current potential was shifted from -70 mV to potentials between -20 and 0 mV with a relatively linear I-V relationship, a current similar to that reported by others (6). Thereafter, myocytes rarely recovered. In addition, the glibenclamide-sensitive ATP-sensitive K+ current (IK,ATP) was observed in some cells during and after exposure to LPlasC and could have been responsible for the observed inexcitability of myocytes with an extremely short AP duration and a membrane potential of approximately -80 mV (unpublished data). LPlasC-associated changes in AP configuration are determined by the net balance of its effects on all membrane currents. Because of the high membrane resistance that exists at the negative slope between -60 and -20 mV of the I-V curve in rabbit ventricular myocytes, a small increase in inward currents (e.g., ICa,L or nonspecific current) can depolarize membrane potential. On the other hand, an increased outward K+ current (e.g., IK,ATP) can drive the membrane potential toward K+ equilibrium potential. This could account for the observed unstable AP configurations and delayed afterdepolarizations induced by LPlasC. Lipid metabolites, including LPlasC, have been suggested to alter the lipid microenvironment of ion channels (15). The distinct physical and chemical properties of LPlasC and its distribution in membrane phospholipids could alter lipid-protein interactions, thereby leading to alterations in gating properties of ion channels and/or transporters (15). Although measurements of membrane currents and APs in patch-clamped cells provide important information, lipid metabolite-induced disturbances in cell membrane structure and function make the correlation with physiological measurements in intact cells more difficult.

In comparison with other amphiphilic metabolites, LPlasC exerts distinct effects on cardiac contractile and electrical properties at 1 µM, a concentration comparable to those observed in the plasma of animal models of myocardial ischemia or in coronary sinus effluent from patients with ischemic myocardium (21). Similar to LPlasC, palmitoylcarnitine (but not LPC) has been reported to elicit a transient increase in ICa,L in guinea pig ventricular muscle measured in normal Tyrode solution, consistent with its positive inotropic effect (1). In contrast, a study in rabbit ventricular myocytes suggested that 5 µM palmitoylcarnitine decreases ICa,L 10 min after exposure (30). LPlasC (1 µM) appears to have more profound effects on ICa,L than palmitoylcarnitine (5 µM). In contrast to other studies, we found that most myocytes could not survive in LPlasC at concentrations of >3 µM for >= 3 min in patch-clamp experiments. Thus LPlasC appears to be more potent at causing arrhythmias and cell death than LPC (6) and palmitoylcarnitine (30).

In summary, the present data show that LPlasC causes increases in ICa,L, intracellular free Ca2+, and myofilament Ca2+ sensitivity, prolongs AP duration, and augments contractility. Such changes are often followed by early and/or delayed afterdepolarizations and sustained membrane depolarization, resulting from an abnormal AP duration that is probably mediated by intracellular Ca2+ overload. Myocytes would eventually become inexcitable because of substantial depolarization or hyperpolarization, probably resulting from LPlasC-induced nonspecific currents or IK,ATP, respectively. Such rapid, dramatic electrophysiological and mechanical changes in the heart function could occur under many pathophysiological conditions, including ischemia/reperfusion, hypoxia, cytokine-related cardiac dysfunction, and sudden cardiac death.


    ACKNOWLEDGEMENTS

We thank Meei-Yueh Liu and Kerrey A. Roberto for excellent technical assistance.


    FOOTNOTES

This work was supported in part by the American Heart Association/Heartland Affiliate and National Heart, Lung, and Blood Institute Grant R01HL-62226.

Address for reprint requests and other correspondence: S. J. Liu, Dept. of Pharmaceutical Sciences, Univ. of Arkansas for Medical Sciences, 4301 West Markham St., MS 522-3, Little Rock, AR 72205 (E-mail: sliu{at}uams.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

First published November 27, 2002;10.1152/ajpcell.00465.2002

Received 3 October 2002; accepted in final form 22 November 2002.


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ABSTRACT
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MATERIALS AND METHODS
RESULTS
DISCUSSION
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