© 1997 American Heart Association, Inc.
Articles |
Lysophosphatidylcholine Modulates Cardiac INa via Multiple Protein Kinase Pathways
From the Department of Medicine, Division of Cardiology and Department of Physiology, University of Maryland, Baltimore.
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Abstract |
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Abstract Lysophosphatidylcholine (LPC) is a naturally occurring intracellular phospholipid metabolite that has been implicated in arrhythmogenesis during ischemia. LPC has been shown to affect the cardiac Na+ current (INa), but the mechanism of modulation remains undescribed. Recently, low concentrations of LPC have been shown to activate protein kinase C (PKC) independent of the receptor-delineated pathway. The purposes of this study were to describe the effects of intracellularly introduced LPC on INa and to determine if these effects were mediated by kinases. Modulation of INa was studied in ventricular cells with LPC (1 nmol/L to 1 µmol/L) internally applied using whole-cell patch-clamp techniques. Intracellular LPC caused a dose-dependent depolarizing shift of steady state inactivation that was accompanied by a change in slope factor. The development of resting inactivation from closed states was delayed 40%, whereas the recovery from inactivation was significantly accelerated. These results were mimicked by another bioactive lipid, lysophosphatidylethanolamine, or by a peptide analogue of PKC, which is a potent stimulator of endogenous PKC activity. Maximal recruitable current was significantly increased by LPC but not by PKC activation. Some of the effects of LPC on INa could be partially inhibited by the specific PKC inhibitor chelerythrine chloride or by downregulation of PKC with phorbol ester pretreatment. However, genistein, a specific tyrosine kinase inhibitor, completely inhibited all the modulation of INa caused by LPC. These data suggest that LPC modulates INa in cardiac myocytes by a pathway that involves both PKC-dependent and tyrosine kinase dependent phosphorylation.
Key Words: cardiac Na+ current • lysophosphatidylcholine • protein kinase C • protein tyrosine kinase
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Lysophosphatidylcholine is formed intracellularly by phospholipase A2 hydrolysis of a prevalent membrane phospholipid, phosphatidylcholine. LPC accumulates during myocardial ischemia1 but is also produced intracellularly during normal phospholipid turnover. In vitro biochemical studies and in vivo investigations have demonstrated that low concentrations (<10 µmol/L) of LPC activate PKC in some cell types, including vascular tissue.2 3 4 5 6 7 Interestingly, higher concentrations (>10 µmol/L) of LPC inhibit PKC activity.5 LPC and some PKC isoforms are universal species; ie, they are not tissue specific. Therefore, it would be expected that LPC would also activate PKC in cardiomyocytes, although this relationship has not been investigated in these cells.
Previous studies of the actions of LPC demonstrated that high concentrations applied extracellularly are associated with conduction abnormalities in isolated hearts8 9 10 and cause changes in INa amplitude and inactivation kinetics.11 12 13 Similar concentrations of LPC applied to excised patches have been shown to reduce peak INa and affect activation and inactivation of the cardiac INa.14 At these concentrations, LPC may cause a change in membrane phospholipid packing when it is inserted into the lipid bilayer. Accordingly, it is possible that the changes in INa observed were due to membrane deformation, which favors certain thermodynamic states of the Na+ channel,15 rather than PKC activation. Therefore, the aims of the present study were to describe the modulation of INa caused by intracellular application of low concentrations of LPC in intact cells and to investigate the possible role of PKC in this modulation.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Cell Preparation
Isolated ventricular cells were prepared from the hearts of 1-day-old Sprague-Dawley rats by enzymatic dissociation.16 The rat pups were decapitated, and the hearts were excised and minced in a 0.9% saline solution. The tissue was dissociated in 50 mL of Joklik MEM supplemented with (mmol/L) HEPES 10, sodium pyruvate 10, nicotinamide 1, L-ascorbate 0.4, adenosine 1, D-ribose 1, MgCl2 1, taurine 1, DL-carnitine 2, KHCO3 26, and L-glutamine 5, pH 7.6, along with 10 mg/mL gentamycin, 50 µmol/L CaCl2, and 0.5 mg/mL collagenase (Worthington Scientific). After six changes of the medium, the cells were centrifuged and suspended in medium enriched with fetal bovine serum, plated, and incubated at 37°C. For these studies, cells were used 18 to 36 hours after plating (after they had attached to the coverslip glass and before they lost their spherical shape). All recordings were performed at room temperature (20°C to 22°C).
Experimental Solutions and Protocols
The pipette solution contained (mmol/L) CsCl 120, CsOH 5,
NaCl 5, MgCl2 5, EGTA 1, Na2-ATP 5, and MOPS
10, adjusted to pH 7.2 with CsOH. The extracellular solution contained
(mmol/L) NaCl 130, NaHCO3 5, KCl 5.4,
CaCl2 1, MgCl2 1, CoCl2 1, glucose
10, and HEPES 5, adjusted to pH 7.4 with NaOH. Cobalt was included in
the bath solution to block L- and T-type Ca2+ current.
Since it was present in all bath solutions, any shift in activation
caused by this divalent ion or voltage-dependent block of
INa17 should be constant across
treatment groups and cells. To ensure that cobalt had no effect on the
modulation of INa by LPC, control experiments
were performed substituting 1 mmol/L amiloride for 1
mmol/L CoCl2. In this concentration, amiloride has
been shown to block T-type Ca2+ current18
without significantly affecting
INa.19 The V1/2 of
activation and the V1/2 of inactivation assessed in
cobalt-free amiloride-containing solution did not differ significantly
(n=4).
LPC was added directly to the pipette or bath solution from a freshly made stock solution. LPE, chelerythrine chloride, and PMA were dissolved in DMSO in concentrations that allowed DMSO concentration to remain at <1:10 000 in all experiments. Control cells were exposed to the same concentration of DMSO, although in this concentration DMSO had no affect on INa based on comparisons with other cells not exposed to DMSO. PKCP (Sigma Chemical Co) was dissolved in 0.05 mol/L acetic acid stock solution and frozen. This peptide was diluted to a final concentration of 2 to 4 nmol/L and added to the pipette solution the day of the experiment. The measured pH of the pipette solution was unchanged by this dilution of PKCP. No data were collected for the first 2 minutes after attaining access to the intracellular space, ie, after entering the whole-cell patch-clamp mode, to allow equilibration of the pipette solution with the cytoplasm. Since LPC, PKCP, and LPE were in the pipette solution, the data reported are from cells different from control.
For inhibition experiments, PKC was downregulated by exposing the cells overnight to 100 nmol/L PMA. In other experiments, the inhibitors chelerythrine chloride and genistein were added to the bath solution 5 to 20 minutes before recording INa or introducing LPC intracellularly.
Electrophysiological Recording
Procedures
Borosilicate glass capillaries (World Precision Instruments,
Inc) were used to make pipettes, and conventional pulling techniques
were performed. Electrode resistance in the extracellular solution
ranged from 1 to 4 M
, and seal resistances were 1 to 3 G.
Voltages were corrected for the liquid junctional potential. Whole-cell
currents were recorded with an Axopatch 200A amplifier (Axon
Instruments). Series resistance and cell capacitance were measured
using the series resistance/whole-cell capacitance compensation circuit
of the amplifier. The accuracy of this measure of cell capacitance (C)
was assessed in 23 cells by comparing this value with the total charge
(Q) movement in response to a 10-mV voltage step (V): Q=VC. Q was
determined by integrating the area under the capacitance spike. These
measures of cell capacitance were highly correlated (r=.95,
P<.001), with no significant difference observed
(P=.40). Whole-cell capacitance averaged 8 pF and remained
unchanged throughout the experiments, unless the cell lost its
morphological integrity. Data from such cells (<2%) were discarded.
Series resistance averaged 
3 M, resulting in voltage errors of
<6 mV for clamping peak INa. Series resistance
was followed throughout the experiment, and data from the cell were
discarded if series resistance changed by >10% over the course of the
recordings.20 Ionic currents were eliminated in a
dose-dependent manner by tetrodotoxin, indicating that
INa was the only current present under these
experimental conditions.
I-V relationships were determined in all cells before any other experimental protocols to test the adequacy of the voltage clamp. Activation was assessed with 20-ms pulses from a holding potential of -100 mV to various test potentials. The pulse amplitude was incremented to +70 mV in 5- or 10-mV steps with 2-second interpulse intervals. Data from cells were discarded if there was any lack of smoothness in the negative limb of the I-V curve, because this indicated a lack of voltage control,20 or if the slope of activation was <6.0 mV.21 Steady state inactivation was determined using a standard two-pulse protocol, with a holding potential of -100 mV and a test potential of 0 mV. One-second conditioning pulses from -125 to -5 mV were applied 1 ms before the test potential to assess INa availability. Reactivation was assessed using a standard double-pulse method. A 200-ms conditioning pulse was followed at varying intervals by a 20-ms depolarizing pulse from -90 or -110 mV to 0 mV. The development of resting inactivation was measured using a two-pulse protocol, with a conditioning pulse of variable duration (1 to 200 ms) to -65 mV, followed by a 20-ms test pulse to 0 mV.22 23
Data Analysis
Data were collected and analyzed using pClamp software
(Axon Instruments). Kinetic data were fit with exponential curves using
a least-squares algorithm. Discrimination between single- and
double-exponential fits of the data was done visually. If the standard
deviation of the fits differed by >30%, then the second-order fit was
selected. Steady state inactivation and activation were fit with a
standard Boltzmann equation:
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.05 was considered
significant. |
Results |
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INa Activation
To assess the action of LPC on the activation of INa, families of currents were recorded in control cells and in cells exposed to LPC intracellularly. I-V relationships, evoked from a holding potential of -100 mV to test potentials between -80 and +70 mV, reversed at potentials near the value calculated by the Nernst equation (+ 57 mV), with no differences between treatments. Conductance transformations of peak current were performed and fit to a Boltzmann equation. V1/2 of activation of INa was -37.7±1.4 mV in control cells (n=14) and -34.9±1.7 mV in cells with 1 µmol/L LPC (n=10) (P=NS); the slopes of activation were 7.3±0.3 and 7.5±0.5 mV, respectively (P=NS). Representative currents with this activation protocol from a control cell and an LPC-treated cell are shown in Fig 1
. Also shown in Fig 1 is an I-V curve
representing mean current amplitudes for control and LPC.
Although the values for the voltage dependence of activation were very
similar in the two groups, currents were larger in LPC-treated
cells.
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) (14 cells) and 1 µmol/L LPC (
) (10
cells). Test pulses of 20 ms were given at 10-mV intervals.
Maximal Recruitable INa
The above studies indicate that intracellular LPC increases
INa amplitude. However, at a holding potential
of -100 mV, resting inactivation is greater in control cells (see
below), which may confound this result. Accordingly, to assess whether
maximal recruitable INa is influenced by
intracellular LPC, peak current amplitude was measured from a holding
potential of -125 mV, a voltage potential at which steady state
inactivation is minimal in all conditions. A significant increase of
peak current was still noted in cells treated with LPC (198.3±21.6
pA/pF [n=15] versus 245.1±22.5 pA/pF [n=20], P=.04), as
shown in Fig 2.
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Current Availability
Steady state inactivation was measured in control cells and those
intracellularly treated with LPC. Using a standard two-pulse protocol,
resting inactivation was reduced in a concentration-dependent manner in
LPC cells. There was an 8-mV depolarizing shift of V1/2
with 1 µmol/L intracellular LPC exposure (Table 1). Examples of these data are shown in
Fig 3A. Increasing LPC concentration
between 1 and 5 µmol/L did not result in a further
voltage shift of inactivation, suggesting that 1 µmol/L
was a saturating concentration for this effect.
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), 1 nmol/L LPC (
), and 1 µmol/L LPC (x) data
points fit with Boltzmann equations. In these examples, the
V1/2 of inactivation was -83.1 mV for control, -77.0 mV
for 1 nmol/L LPC treatment, and -71.4 mV for 1 µmol/L LPC
treatment. The voltage protocol used for the study of steady state
inactivation is shown in the insert. B, The effects of LPC (x) were
mimicked by PKC (Previous studies in this cell preparation have shown that the V1/2 of steady state inactivation is stable over an average recording time (10 to 20 minutes).24 To reconfirm the temporal stability of half-inactivation, consecutive measurements of V1/2 were made in 11 cells. As with all cells used in the present study, there was a 2-minute equilibration period after achieving intracellular access before measurements were made. The average recording time in these 11 cells was 12 minutes. The mean V1/2 of inactivation of the final trial varied by <1 mV from initial measurements. We presume that any voltage shifts of inactivation occur within the first 2 minutes, during the rapid diffusion of the pipette solution into the intracellular space of these small cells. This results in an equilibrium state with a stable V1/2 of steady state inactivation during the recording period. Serial measurements of V1/2 from cells dialyzed with LPC showed similar results. The constancy of half-inactivation in this preparation and the concentration dependence of the LPC effect on current availability indicate that LPC causes a true shift in the voltage-dependent current availability.
There was also an increase in slope factor with LPC (Table 1
). This
change in slope may reflect a change in the voltage dependence of
resting inactivation, or LPC may be inducing slow inactivation during
the 1000-ms conditioning pulse. Additional experiments were performed
with the conditioning pulse increased to 5000 ms. If a slowly
inactivated state was affected by LPC after 1000 ms, then
its effect should be even greater after a conditioning pulse of five
times that duration. However, neither the slope nor the
V1/2 of inactivation (7.0 and -72.3 mV) differed from that
of the 1000-ms conditioning pulse, indicating that LPC changes the
voltage dependence of inactivation.
To assess if the LPC-induced increase in current availability is
mediated through activation of the PKC phosphorylation
system, two strategies were adopted. First, we internally applied
another bioactive lysophospholipid known to activate PKC, LPE.
The effects of 1 µmol/L LPC were mimicked by the same
concentration of LPE, which caused a similar depolarizing shift of
V1/2 of 8 mV and an increase of slope factor (see Table 1). The action of LPC on steady state inactivation was then compared
with the direct effects of PKC activation. To activate PKC, a
peptide fragment of the catalytic domain of PKC was added to the
pipette solution. This peptide binds to the regulatory subunit of
native PKC, exposing the active site, which results in a potent
activation of PKC.25 The specificity of this peptide
ensures that any effect is attributable to PKC activation. The effect
of PKC on resting inactivation mimicked that of LPC. In 14
PKC-activated neonatal cardiac cells, the V1/2 of
inactivation was -74.4±1.6 mV. Both LPC and PKC caused a similar
increase in current availability and a change in voltage dependence, as
shown in Fig 3B.
Recovery From Inactivation
Reactivation was assessed after a 200-ms conditioning pulse to 0
mV. In these cells, reactivation is best fit as a double-exponential
process. At a holding potential of -90 mV, both reactivation time
constants were decreased by LPC treatment in a concentration-dependent
manner. The ratio of the amplitudes of 
f to
s remained unchanged between treatments. Examples of
normalized currents from the reactivation protocol in control cells and
in 1 nmol/L LPC– and 1 µmol/L LPC–treated cells
are shown in Fig 4A. Treatment with
1 µmol/L LPE also significantly accelerated reactivation
at this holding potential. These results are summarized in Table 2.
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), and 1 µmol/L LPC (x)
data fits. Under normal conditions (control) at a holding potential of
-90 mV,
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At a holding potential of -90 mV, the shift in steady state
inactivation caused by LPC could contribute to an apparent difference
in the rate of reactivation. Accordingly, identical reactivation
protocols were performed from a more hyperpolarized holding potential
(-110 mV). This is the most negative voltage at which these cells can
be maintained for long periods of time. Reactivation remained a
double-exponential process, and both time constants were reduced
50% by LPC. f decreased from 9.2±1.4 ms in control
cells (n=8) to 5.1±0.8 ms in LPC-treated cells (n=6,
P<.01), and s decreased from 81.4±13.1 to
37.9±8.7 ms (P<.01). Examples of reactivation from a
holding potential of -110 mV are shown in Fig 4B.
Development of Resting Inactivation
At normal resting membrane potentials, current availability is
inversely related to the number of channels in the
inactivated state. The number of inactivated
channels in turn is due both to the time course of reactivation from
the preceding action potential and to the development of inactivation
from resting states. As noted above, the time course of reactivation is
accelerated by LPC. To assess the development of resting inactivation,
we measured this transition using a conditioning pulse of incrementally
increased durations to -65 mV, followed by a test pulse to 0 mV to
measure the recruitable current. No current is elicited with a pulse to
-65 mV (Fig 1A), so inactivation develops from the resting state. In
the presence of LPC, INa amplitudes were larger
after conditioning pulses, reflecting less inactivation.
Analysis of the time course of this process confirmed the
longer time constant of development of inactivation with LPC-treated
cells (110.0±11.6 ms, n=7) compared with control cells (67.3±11.6 ms,
n=8, P<.01). Examples of normalized current amplitudes fit
with monoexponential equations for LPC are shown in Fig 5.
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INa Decay and Persistent
INa
Macroscopic current decay and the amount of persistent, slowly
inactivating INa were measured at a test
potential of -40 mV from a holding potential of -100 mV.
INa decay, measured from the peak current
amplitude to 16 ms, was best fit as a single exponential in both
control cells and with 1 µmol/L LPC treated cells in
the pipette. The time constants of decay did not differ significantly
(2.98±0.3 ms in control, n=12; 3.07±0.31 ms with LPC, n=10;
P=.83). Persistent INa was evaluated
by normalizing the amplitude of the current at 16 ms to the peak
amplitude of the current at -40 mV. LPC did not cause a significant
increase in the amplitude of persistent current (2±1% in control,
n=12; 3±1% with LPC, n=10; P=.3).
Extracellular LPC
The effects of LPC described so far involved direct intracellular
application of low concentrations of this lysophospholipid. Previous
studies of INa modulation evaluated high
concentrations of extracellular LPC.12 13 14 However, LPC has
been shown to have nonlinear concentration-dependent effects,
particularly with regard to activation of PKC. To determine if the
effect of intracellular LPC on INa was a
function of intracellular application or the concentration, additional
experiments evaluating the effects of 1 µmol/L LPC in the
bath solution were undertaken in nine myocytes. The effects of 1
µmol/L LPC extracellularly on INa are
comparable to those of 1 nmol/L LPC delivered intracellularly.
Extracellular LPC depolarized resting inactivation by 3 mV compared
with control (-77.6±2.5 mV versus -80.9±1.1 mV, P=.05),
whereas 1 nmol/L intracellular LPC depolarized steady state
inactivation by an average of 2 mV. The change in slope factor was also
comparable to 1 nmol/L intracellular LPC (-6.3±0.3 versus
-6.0±0.2 mV). Reactivation during 1 µmol/L LPC
superfusion was accelerated relative to control (f,
16.4±6.2 ms; s, 158.8±82.7; and Af,
0.70±0.1). In summary, extracellular LPC had the same effects as far
lower concentrations of intracellular LPC. This may be due to
incomplete transfer of LPC through the cell membrane.
Inhibition of LPC Modulation of INa
If the effects of LPC on INa were mediated
solely through PKC, inhibition of PKC should reverse the effects. Cells
were superfused with one of two PKC inhibitors, each having
different molecular modes of inhibition. Chelerythrine chloride
inhibits PKC by binding at the catalytic domain,26 whereas
overnight incubation with phorbol ester downregulates native PKC.
During intracellular LPC application, both PKC inhibitors
partially reversed the depolarizing shift of steady state inactivation
(Fig 6A). V1/2 shifted 3 mV
in the hyperpolarizing direction in cells treated with chelerythrine
chloride and 4 mV after PMA incubation, with no significant change in
slope from LPC-treated cells (Table 1). Neither chelerythrine chloride
nor PMA caused a significant hyperpolarizing shift in steady state
inactivation in control cells (PMA, -82.4±2.5 mV, n=6;
chelerythrine chloride, -84.6±1.2 mV, n=2). In contrast to the
partial inhibition of the action of LPC, chelerythrine chloride and PMA
completely reversed the modulation of INa by the
PKC catalytic subunit. This implies that LPC modulation of
INa involves more than the PKC pathway.
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Since PKC has been shown to induce tyrosine kinase
phosphorylation,27 28 29 30 31 we investigated the
possibility that the modulation of INa by LPC
also involved a protein tyrosine kinase. Cells were superfused with 50
or 16 µmol/L of the specific tyrosine kinase
inhibitor genistein32 ; the pipette solution
contained 5 µmol/L LPC. As reported earlier, this
concentration of LPC does not further alter INa
amplitude or kinetics but was used to ensure a saturating
concentration. Both concentrations of genistein were equally effective
in inhibiting LPC. The results of the measurement of steady state
inactivation are shown in Fig 6B. Genistein completely inhibited the
depolarizing shift in V1/2, without a reversal of the
voltage-dependent effects of LPC. In three cells, genistein was
superfused after recordings with intracellular LPC. In each
case, genistein shifted the V1/2 of resting inactivation in
LPC-treated cells by an average of 8 mV within 10 minutes (-76.0±0.35
versus -84.5±0.32 mV; P<.001, paired t test)
(Fig 6C). In these same cells, genistein accelerated the development of
inactivation from closed states by 33% (Fig 6D). LPC accelerated the
rate of recovery from inactivation at both
physiological and hyperpolarized holding
potentials. Interestingly, this effect could be prevented by overnight
incubation with PMA or with genistein but not with superfusion of
chelerythrine chloride, as summarized in Table 2.
In contrast to the variable effects of inhibitors
on the kinetics and voltage dependence of inactivation, the change in
maximal available INa was inhibited equally well
by PMA, chelerythrine chloride, or genistein. Each of these
inhibitors reduced the current amplitude of LPC-treated
cells to control values, as illustrated in Fig 2.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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This is the first study to evaluate the effects of low concentrations of intracellularly applied LPC on INa in cardiac myocytes. The major findings of the present study are that intracellular LPC is a potent modulator of INa availability and inactivation gating. Under these conditions, LPC functions as a bioactive lipid capable of initiating a complex phosphorylation cascade that appears to involve both PKC and tyrosine kinase.
In electrophysiological studies, LPC has been studied as a metabolite produced during myocardial ischemia that interferes with phospholipid packing of the membrane, deforming the sarcolemma and affecting membrane curvature.33 34 This alone can modulate the activity of membrane proteins.35 Once incorporated into the membrane, LPC remains primarily in the outer leaflet without transferring to the inner leaflet.36 Therefore, extracellular application of LPC may derange the phospholipids of the outer leaflet and diffuse slowly into the cytoplasmic space. Intracellularly, in concentrations well below those found in pathophysiological conditions, LPC induces phosphorylation.2 3 4 5 6 7 Physiologically, then, LPC may have multiple mechanisms of action, including activating PKC intracellularly at low concentrations and destabilizing the membrane at high concentrations.
Neonatal rat cardiac myocytes were used in these studies in the
whole-cell recording mode, a preparation that has the potential
to disrupt protein phosphorylation. However, the rapid
effect on INa of PKCP, which has no activity of
its own but activates endogenous
PKC,25 implies that the PKC
phosphorylation pathway remained intact. In rats,
neonatal cardiomyocytes express the same PKC isoforms (with
the addition of the 
isoform) that are expressed in adult
cardiomyocytes37 ; the same protein tyrosine
kinases are also expressed in both neonatal and adult
cardiomyocytes.38 The small size and favorable
geometry of the cells allowed rapid intracellular transfer of the
pipette solution. This, in conjunction with the universality of the
kinases expressed, made the neonatal cardiac myocyte an excellent model
for these studies.
The ability of LPC to activate PKC has been documented in other
cell types6 39 40 41 (for reviews see References 4 and 54 5 ),
although the biomolecular mechanism of LPC-PKC interaction has not been
well described.4 In our investigations of neonatal
ventricular cells, the effects of intracellular LPC on
INa were mimicked in many respects by PKCP, a
peptide analogue of the catalytic domain of PKC. This peptide was
chosen because of its specificity. Phorbol esters, which are commonly
used to activate PKC, are not specific for PKC and have
receptor sites on cytoplasmic proteins with other cellular actions,
such as 
-chimerin42 and phospholipase
D.43 44 45 This may account for the contradictory findings
regarding PKC modulation of INa (see Reference
4646 for review). PKCP caused quantitatively similar depolarizing shifts
of steady state inactivation as LPC (Fig 2). We have shown previously
in this same preparation that PKC also increased the rate of
reactivation and delayed the development of resting
inactivation.47
The present results on the effect of 1 nmol/L intracellular LPC on INa were similar to those noted with extracellular application of 1 µmol/L of this agent. Previously, extracellular LPC has been shown to reduce peak INa, slow reactivation, and reduce current availability.11 12 13 14 However, it is important to note that concentrations of 10 to 25 µmol/L were used in these studies. In vitro biochemical studies have demonstrated that the effect of LPC on PKC are biphasic and concentration dependent, with low concentrations of LPC (<10 µmol/L) activating PKC and high concentrations (>10 µmol/L) inhibiting PKC.2 5 Thus, it is likely that previous studies using extracellular application of high concentrations of LPC involved changes of membrane packing as well as the possible inhibition of PKC activity.
The biophysical effects of LPC on INa in the
present study were primarily due to modulation of inactivation
gating. Both time constants of reactivation were faster, without a
change in the proportion of current that recovered with each time
constant. This suggests that LPC did not promote a change in the
relative emptying of inactivated states but caused a simple
acceleration of normal reactivation. This effect was not voltage
dependent, as it also occurred at hyperpolarized holding potentials.
Resting inactivation was significantly reduced by LPC in a
concentration-dependent manner. This suggests that in the presence of
LPC fewer channels reside in the inactivated state. The
decrease in resting inactivation and the increased rate of reactivation
could be explained by assuming that LPC caused the
inactivated state of the Na+ channel to become
thermodynamically or conformationally unstable, thus favoring a
transition to either open or closed states. This interpretation is
supported by measurements of the time course of the development of
resting inactivation (Fig 5). During intracellular LPC treatment, there
was more recruitable INa after depolarizations
to -65 mV, a voltage that would inactivate channels from a
resting state without passing through an open
state.22 23
The 
subunit of the Na+ channel contains an
intracellular loop between domains III and IV, which is believed to
function as an "inactivation gate," and also contains consensus
sites for PKC phosphorylation.48 The
effects of LPC reported in the present study were primarily on
inactivation gating and involved both the PKC and tyrosine kinase
phosphorylation cascades. Although our results are
consistent with the explanation that intracellular LPC promotes
phosphorylation of this cytoplasmic loop, which is
important in inactivation, it is also possible that
phosphorylation of other membrane or cytoskeletal
proteins that affect INa inactivation could be
involved.49 Protein tyrosine kinase
phosphorylation is known to regulate delayed rectifier
K+ channels,28 including a human cardiac
isoform,50 by direct phosphorylation of
the channel on a tyrosine residue.29 50 Since there is
significant homology between the K+ and Na+
channels,51 the cardiac Na+ channel is a
potential substrate for tyrosine kinase
phosphorylation.
Although there are many similarities between the actions of LPC and PKC on INa, some notable differences are present. LPC significantly increased INa peak amplitude, which did not occur with PKC activation.47 Also, the effects of LPC are only partially inhibited by chelerythrine chloride and PMA, inhibitors that fully reverse the effects of the peptide activator of PKC. These results imply that LPC modulation of INa involved a mechanism more complex than simple activation of the PKC phosphorylation system.
Recent investigations of intracellular signaling pathways have shown that PKC can induce tyrosine protein kinase phosphorylation.31 52 Tyrosine kinases, a family of cytoplasmic and receptor-associated enzymes that phosphorylate substrates at tyrosine residues, are distinct from the PKC enzymes that phosphorylate serine/threonine residues. Little is known regarding how these pathways interact in cardiac myocytes. The observation that the effects of LPC on INa were largely mimicked by PKC activation but were effectively inhibited only by a tyrosine kinase inhibitor suggests that tyrosine kinase phosphorylation occurs after PKC activation. However, the partial inhibition of the effects of LPC by PKC inhibitors implies concurrent activation of PKC and tyrosine kinase by a common effector. Intracellular Ca2+ is an attractive candidate for this effector, since LPC causes an increase in cytosolic free Ca2+.53 54 Elevated intracellular Ca2+, in conjunction with bioactive lipids, can activate PKC (see Reference 44 for review) or tyrosine kinase.28 29
Although the precise signal transduction pathway responsible for the action of low concentrations of LPC cannot be elucidated with certainty from these data, we conclude that even at nanomolar concentrations, INa can be dramatically modulated by intracellular LPC via pathways that include both PKC and tyrosine kinase activation. Intracellular LPC may have important effects on the cardiac action potential, because tyrosine phosphorylation also decreases outward K+ current.28 50 The increase of INa availability at depolarized potentials and decreased K+ current would prolong the action potential and delay repolarization. Regional changes of LPC activity could then cause a dispersion of repolarization that is associated with life-threatening arrhythmias.
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Selected Abbreviations and Acronyms |
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Acknowledgments |
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This study was supported by a Fellowship and Grant from the American Heart Association.
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Footnotes |
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Reprint requests to Michael R. Gold, MD, PhD, University of Maryland Medical Center, Division of Cardiology N3W77, 22 South Greene St, Baltimore, MD 21201.
Received November 15, 1996; accepted June 26, 1997.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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1. Creer MH, Dobmeyer DJ, Corr PB. Amphipathic lipid metabolites and arrhythmias during myocardial ischemia. In: Zipes DP, Jalife J, eds. Cardiac Electrophysiology From Cell to Bedside. Philadelphia, Pa: WB Saunders; 1990; pp 417-433.
2.
Oishi K, Raynor RL, Charp PA, Kuo JF.
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