Phospholemman (PLM) regulates contractility and Ca2+ homeostasis in cardiac myocytes. We characterized excitation-contraction coupling in myocytes isolated from PLM-deficient mice backbred to a pure congenic C57BL/6 background. Cell length, cell width, and whole cell capacitance were not different between wild-type and PLM-null myocytes. Compared with wild-type myocytes, Western blots indicated total absence of PLM but no changes in Na+/Ca2+ exchanger, sarcoplasmic reticulum (SR) Ca2+-ATPase, α1-subunit of Na+-K+-ATPase, and calsequestrin levels in PLM-null myocytes. At 5 mM extracellular Ca2+ concentration ([Ca2+]o), contraction and cytosolic [Ca2+] ([Ca2+]i) transient amplitudes and SR Ca2+ contents in PLM-null myocytes were significantly (P < 0.0004) higher than wild-type myocytes, whereas the converse was true at 0.6 mM [Ca2+]o. This pattern of contractile and [Ca2+]i transient abnormalities in PLM-null myocytes mimics that observed in adult rat myocytes overexpressing the cardiac Na+/Ca2+ exchanger. Indeed, we have previously reported that Na+/Ca2+ exchange currents were higher in PLM-null myocytes. Activation of protein kinase A resulted in increased inotropy such that there were no longer any contractility differences between the stimulated wild-type and PLM-null myocytes. Protein kinase C stimulation resulted in decreased contractility in both wild-type and PLM-null myocytes. Resting membrane potential and action potential amplitudes were similar, but action potential duration was much prolonged (P < 0.04) in PLM-null myocytes. Whole cell Ca2+ current densities were similar between wild-type and PLM-null myocytes, as were the fast- and slow-inactivation time constants. We conclude that a major function of PLM is regulation of cardiac contractility and Ca2+ fluxes, likely by modulating Na+/Ca2+ exchange activity.
- cytosolic calcium concentration
phospholemman (PLM), a 72-amino-acid sarcolemmal phosphoprotein with a single transmembrane domain (20), belongs to the FXYD gene family of small ion transporter regulators (27). Studies in noncardiac tissues suggest that PLM can be a channel (15), a channel subunit, or an ion transport regulator (9) and is likely involved in regulation of cell volume (10). In adult rat cardiac myocytes, PLM overexpression altered contractility and cytosolic Ca2+ concentration ([Ca2+]i) transients (26). Specifically, at low (0.6 mM) extracellular Ca2+ concentration ([Ca2+]o), both contraction and [Ca2+]i transient amplitudes were larger in myocytes overexpressing PLM. At high (5.0 mM) [Ca2+]o, cell shortening and [Ca2+]i transient amplitudes were smaller in myocytes overexpressing PLM. The reduced dynamic range in response to increasing [Ca2+]o in PLM-overexpressed myocytes is similar to that observed in myocytes in which Na+/Ca2+ exchanger (NCX1) was downregulated (28). Conversely, PLM downregulation in adult rat myocytes resulted in enhanced dynamic range in response to increasing [Ca2+]o (18), mimicking the contractility and [Ca2+]i transient changes observed in NCX1-overexpressed myocytes (39). These observations provide circumstantial evidence that PLM regulates NCX1 activity. Indeed, PLM overexpression in adult rat myocytes (25, 38) and in a heterologous expression model system (2, 34) suppressed NCX1 current (INaCa), whereas PLM downregulation in adult rat myocytes enhanced INaCa (18). In native rat myocytes, PLM colocalized (38) and coimmunoprecipitated with NCX1 (2, 18). The present study was undertaken to further test the hypothesis that one of the major functions of PLM in cardiac tissues is to regulate NCX1, using a fundamentally different approach of genetic manipulation.
Generation of PLM-deficient mice and animal care.
A mouse line deficient in PLM was generated by replacing exons 3 to 5 of the PLM gene with lacZ and neomycin resistance genes, as described in detail previously (14). These mice grow to adulthood and are fertile. Studies were performed using mice backcrossed to a pure congenic C57BL/6 background. Homozygous adult littermates 3–6 mo old were used in the experiments. Mice were housed in ventilated racks in a barrier facility supervised by the Department of Comparative Medicine at the Pennsylvania State University College of Medicine. Standard care was provided to all mice used for experiments.
Isolation of adult murine cardiac myocytes.
Cardiac myocytes were isolated from the septum and left ventricular free wall of wild-type and PLM-null mice (25–37 g) according to the protocol of Zhou et al. (41). Briefly, mice were heparinized (1,500 U/kg ip) and anesthetized (pentobarbital sodium, 50 mg/kg ip). The heart was excised, mounted on a steel cannula, and retrograde perfused (100 cmH2O, 37°C) with Ca2+-free bicarbonate buffer followed by enzymatic digestion (collagenases B and D, protease XIV) as described (41). Isolated myocytes were plated on laminin-coated glass cover slips in a petri dish, and the Ca2+ concentration of the buffer was progressively increased from 0.05 to 0.125 to 0.25 to 0.5 mM in three steps (10 min interval each). The 0.5 mM Ca2+ buffer was then aspirated and replaced with minimal essential medium (MEM, Sigma M1018) containing 1.2 mM Ca2+, 2.5% FBS, and antibiotics (1% penicillin/streptomycin). After 1 h (5% CO2, 37°), media was replaced with FBS-free MEM. Myocytes were used within 2–8 h of isolation. The protocol for heart excision for myocyte isolation was approved by the Institutional Animal Care and Usage Committee.
Myocyte shortening measurements.
Myocytes adherent to cover slips were bathed in 0.6 ml of air- and temperature-equilibrated (37°), HEPES-buffered (20 mM, pH 7.4) medium 199 containing 0.6, 1.8, or 5.0 mM [Ca2+]o. Measurements of myocyte contraction (1 Hz) were performed as previously described (18, 25, 26, 28, 38, 39).
[Ca2+]i transient measurements.
Myocytes were exposed to 0.67 μM of fura-2 AM for 15 min at 37°C. Fura-2-loaded myocytes were field-stimulated to contract (1 Hz, 37°C) in medium 199 containing 0.6, 1.8, or 5.0 mM [Ca2+]o. [Ca2+]i transient measurements, daily calibration of fura-2 fluorescent signals, and [Ca2+]i transient analyses were performed as previously described (25, 26, 28, 37–39).
Ca2+ current measurements.
Whole cell patch-clamp recordings were performed at 30°C as previously described (18, 25, 28, 35, 38, 40). Briefly, fire-polished pipettes with resistances of 0.8–1.4 MΩ when filled with standard internal solution were used. Pipette solution consisted of (in mM) 120 cesium aspartate, 10 Na+-EGTA, 10 HEPES, 4 MgATP, and 4 MgCl2 (pH 7.2 with CsOH). External solution contained (in mM) 135 choline chloride, 1.8 CaCl2, 1 MgCl2, 10 HEPES, and 10 glucose (pH 7.4 with NaOH). Before myocyte stimulation was started, holding potential was changed from −70 to −40 mV to inactivate fast inward Na+ current. To ensure steady-state sarcoplasmic reticulum (SR) Ca2+ loading, six conditioning pulses (from −40 to 0 mV, 300 ms, 1 Hz) were delivered before arrival of each test pulse (from −30 to +60 mV, 10-mV increments, 400 ms). After the last test pulse at +60 mV, the myocyte was held at −40 mV for 1 s before being returned to a holding potential of −70 mV. Leak-subtracted inward currents were used in analysis for Ca2+ current (ICa) amplitudes and inactivation kinetics. We have previously demonstrated that inward currents measured under these conditions were completely blocked by 1 μM verapamil (35). ICa was normalized to membrane capacitance (Cm) before comparison between wild-type and PLM-null myocytes. Slope conductance was calculated from ICa data points (from +10 to +60 mV) for each myocyte by linear regression.
Action potential measurements.
Action potentials from wild-type and PLM-null mice were recorded using current-clamp configuration at 1.5× threshold stimulus and 4-ms duration (28, 38, 40). Pipette solution consisted of (in mM) 125 KCl, 4 MgCl2, 0.06 CaCl2, 10 HEPES, 5 K+-EGTA, 3 Na2ATP, and 5 Na2-creatine phosphate (pH 7.2). External solution consisted of (in mM) 132 NaCl, 5.4 KCl, 1.8 CaCl2, 1.8 MgCl2, 0.6 NaH2PO4, 7.5 HEPES, 7.5 Na+-HEPES, and 5 glucose, pH 7.4.
Measurement of SR-releasable Ca2+.
SR Ca2+ content was estimated by integrating forward INaCa induced by caffeine exposure as described previously (28, 39). The pipette solution consisted of (in mM) 100 cesium glutamate, 1 MgCl2, 30 HEPES, and 2.5 MgATP, pH 7.2. The external solution was composed of (in mM) 130 NaCl, 5 CsCl, 1.2 MgSO4, 1.2 NaH2PO4, 20 HEPES, and 10 glucose, pH 7.4, 30°C. [Ca2+]o was either 0.6 or 5 mM. Holding potential was −70 mV. At 200 ms after the 11th conditioning pulse (from −70 to 0 mV, 300 ms, 1 Hz), with membrane potential (Em) held at −70 mV, caffeine (5 mM, 2.4 s) was applied by puffer superfusion. Currents were digitized at 0.5 kHz and collected for 5 s.
PLM, NCX1, SERCA2, calsequestrin, and Na+-K+-ATPase immunoblotting.
Left ventricles were excised, rinsed in ice-cold PBS, and cut into small pieces. Approximately 60 mg of tissue were suspended in 700 μl of ice-cold lysis buffer containing (in mM) 50 Tris (pH 8.0), 150 NaCl, 1 Na+ orthovanadate, 1 phenylmethylsulfonyl fluoride, 100 NaF, 1 EGTA, and 0.5% Nonidet P-40. A Complete Mini protease inhibitor cocktail tablet (catalog no. 11836153001; Roche, Penzberg, Germany) was also added to 10 ml of lysis buffer. The tissue was homogenized with a glass dounce homogenizer (15–20 strokes) and placed on ice for 15 min before centrifugation at 20,800 g for 10 min at 4°C. The supernatant was snap-frozen with dry ice-ethanol and stored at −80°C.
Proteins in heart homogenates were subjected to 7.5% (NCX1, Na+-K+-ATPase, SERCA2, and calsequestrin) or 12% (PLM) SDS-PAGE under either nonreducing (10 mM N-ethylmaleimide for NCX1) or reducing (5% β-mercaptoethanol for PLM, Na+-K+-ATPase, SERCA2, and calsequestrin) conditions. The fractionated proteins were transferred to ImmunoBlot polyvinylidene difluoride membranes. Primary antibodies used were as follows: for PLM, polyclonal antibody C2Ab (1:10,000; see Ref. 26); for NCX1, polyclonal antibody π11–13 (1:500; Swant, Bellinzona, Switzerland); for calsequestrin, rabbit anti-calsequestrin antibody (1:5,000; Swant); and for SERCA2, polyclonal antibody BL337 (1:1,000; Bethyl Laboratories, Montgomery, TX). The secondary antibodies used were donkey anti-rabbit IgG (Amersham, Piscataway, NJ). Immunoreactive proteins were detected with an enhanced chemiluminescence Western blotting system. Protein band signal intensities were quantitated by scanning autoradiograms of the blots with a phosphorimager (Molecular Dynamics, Sunnyvale, CA).
For Na+-K+-ATPase immunoblotting, crude membranes from mouse left ventricles were prepared using a two-step centrifugation protocol described previously (25). Proteins in crude membranes were fractionated (7.5% SDS-PAGE, reducing conditions), transferred, and detected with polyclonal antibodies to the α1-subunit (1:750) of Na+-K+-ATPase (a generous gift of Dr. Robert Levenson, Pennsylvania State University).
All results are expressed as means ± SE. For analysis of a parameter (e.g., maximal contraction amplitude) as functions of group (wild type vs. PLM null), drug [forskolin or phorbol 12-myristate 13-acetate (PMA)] treatment, and [Ca2+]o, three-way ANOVA was used to determine statistical significance. Two-way ANOVA was used to analyze ICa as a function of group and membrane voltage. For analysis of protein abundance, Cm, cell lengths and widths, SR Ca2+ contents, slope conductances and fast and slow inactivation time constants of ICa, and action potential parameters, Student's unpaired t-test was used. A commercial software package (JMP version 4.05; SAS Institute, Cary, NC) was used. In all analyses, P < 0.05 was taken to be statistically significant.
Effects of PLM knockout on myocyte size and selected proteins.
Myocytes isolated from PLM-null hearts were similar in sizes when compared with those isolated from wild-type mice. Specifically, cell lengths were 128.6 ± 2.5 μm for wild-type myocytes (n = 34 cells from 4 hearts) and 130.5 ± 2.8 μm for PLM-null myocytes (n = 33 cells from 4 hearts; P > 0.63), whereas cell widths were 24.3 ± 0.8 and 25.9 ± 1.0 μm for wild-type and PLM-null myocytes, respectively (P > 0.22). In addition, whole cell capacitance Cm, a measure of cell surface area, was 166 ± 8 pF in PLM-null myocytes (n = 27) and statistically not different (P > 0.10) from that measured in wild-type myocytes (151 ± 5 pF; n = 29).
Western blots confirmed the absence of PLM in PLM-null hearts (Fig. 1). Absence of PLM did not affect protein levels of NCX1, SERCA2, calsequestrin, and the α1-subunit of Na+-K+-ATPase (Fig. 1 and Table 1).
Effects of PLM knockout on myocyte contraction.
In both wild-type and PLM myocytes, elevating [Ca2+]o resulted in the expected increase in contraction amplitudes (Fig. 2 and Table 2). However, the dynamic range in response to increasing [Ca2+]o was significantly increased in PLM-null myocytes. Specifically, at 0.6 mM [Ca2+]o, PLM-null myocytes shortened less than wild-type myocytes. In contrast, at 5.0 mM [Ca2+]o, PLM-null myocytes contracted more than wild-type myocytes. The differences in twitch amplitude were no longer apparent at intermediate (1.8 mM) [Ca2+]o levels. These conclusions are supported by insignificant group (wild type vs. PLM null, P > 0.16) but significant [Ca2+]o (P < 0.0001) and group × [Ca2+]o interaction (P < 0.0001) effects, indicating that the magnitude and/or direction of the effects of changing [Ca2+]o on cell shortening were different across the experimental groups.
To further analyze contraction dynamics, we measured maximal shortening and relengthening velocities. As a group, both maximal shortening and relengthening velocities were higher in PLM-null myocytes (group effect, P < 0.0015). Raising [Ca2+]o increased maximal shortening and relaxation velocities in both wild-type and PLM-null myocytes ([Ca2+]o effect, P < 0.0001). In addition, the group × [Ca2+]o interaction effect was highly significant (P < 0.0025), indicating that altering [Ca2+]o amplified the inherent differences in shortening and relengthening velocities between wild-type and PLM-null myocytes.
Effects of PLM knockout on [Ca2+]i transients.
Differences in myocyte contractility between wild-type and PLM-null myocytes may be because of changes in [Ca2+]i homeostasis associated with PLM deficiency. Indeed, compared with wild-type myocytes, systolic [Ca2+]i values in PLM-null myocytes were lower at 0.6, similar at 1.8, but higher at 5.0 mM [Ca2+]o (Fig. 3 and Table 3). This conclusion is supported by two-way ANOVA that indicated insignificant group (P > 0.06) but highly significant group × [Ca2+]o interaction (P < 0.0003) effects. As expected, elevating [Ca2+]o significantly increased systolic [Ca2+]i in both groups ([Ca2+]o effect, P < 0.0001). There were no differences in diastolic [Ca2+]i between wild-type and PLM-null myocytes (group effect, P < 0.12; group × [Ca2+]o interaction effect, P < 0.15).
The percent increase in the fura-2 fluorescence intensity ratio is an accurate reflection of [Ca2+]i transient amplitude. Compared with wild-type myocytes, [Ca2+]i transient amplitudes in PLM-null myocytes were lower at 0.6, not different at 1.8, but higher at 5.0 mM [Ca2+]o (Table 3). Two-way ANOVA indicated insignificant group (P > 0.64) but highly significant [Ca2+]o (P < 0.0001) and group × [Ca2+]o interaction (P < 0.0004) effects.
The half-time (t1/2) of [Ca2+]i decline, an indicator of SR Ca2+ uptake activity (25), showed no significant differences between wild-type and PLM-null myocytes (Table 3; group effect, P < 0.43; group × [Ca2+]o interaction effect, P < 0.17). Elevating [Ca2+]o, which increased the amplitudes of [Ca2+]i transients, significantly lowered the t1/2 of [Ca2+]i decline in both wild-type and PLM-null myocytes ([Ca2+]o effect, P < 0.0001; Table 3). This observation is consistent with that of Bers and Berlin (7) who demonstrated the kinetics of [Ca2+]i decline were dependent on peak [Ca2+]i.
Effects of PLM knockout on SR-releasable Ca2+ contents.
Changes in contraction and [Ca2+]i transient amplitudes observed in PLM-null myocytes may be because of alterations in SR Ca2+ content. The time integral of forward INaCa induced by caffeine (inward current in Fig. 4) was an estimate of SR Ca2+ content (28, 39). At 0.6 mM [Ca2+]o, SR-releasable Ca2+ (normalized to cell size; fmol/fF) was smaller in PLM-null myocytes than in wild-type myocytes (Table 4). In contrast, at 5 mM [Ca2+]o, SR-releasable Ca2+ was larger in PLM-null than wild-type myocytes. Two-way ANOVA indicated significant group (P < 0.038), [Ca2+]o (P < 0.0001), and group × [Ca2+]o interaction (P < 0.0003) effects, indicating that changing [Ca2+]o affected the inherent differences in SR Ca2+ contents between wild-type and PLM-null myocytes.
Effects of PLM knockout on action potential.
We have previously demonstrated that INaCa was larger in PLM-null when compared with wild-type myocytes (34). Altered Na+/Ca2+ exchange activity may affect action potential morphology (3). Therefore, we measured action potential in wild-type and PLM-null myocytes (Fig. 5). Resting Em (P > 0.75) and action potential amplitude (P > 0.38) were similar between wild-type and PLM-null myocytes (Table 5). Both action potential duration at 50% (P < 0.038) and 90% (P < 0.008) repolarization were significantly prolonged in PLM-null myocytes.
Effects of PLM knockout on ICa.
Altered contractility, [Ca2+]i transients, and action potential morphology in PLM-null myocytes may partly be because of changes in L-type ICa and its kinetics. We therefore compared ICa measured in wild-type and PLM-null myocytes. Figure 6 shows the current-voltage relationships of ICa measured in wild-type and PLM-null myocytes. There were no significant differences in maximal ICa density (P > 0.99), fast (P > 0.62) and slow inactivation time constants (P > 0.82), and slope conductance (P > 0.56) between wild-type and PLM-null myocytes (Table 6). The test potential at which maximal ICa occurred was ∼10 mV for both wild-type and PLM-null myocytes.
Effects of forskolin on contraction in wild-type and PLM-null myocytes.
In a second series of myocyte contraction experiments, we measured the effects of protein kinase A (PKA) activation on contraction in wild-type and PLM-null myocytes. In the absence of forskolin, PLM-null myocytes displayed enhanced dynamic range in response to increases in [Ca2+]o compared with wild-type myocytes (Table 7; group × [Ca2+]o effect, P < 0.026), similar to what we observed in our initial experiments (Table 2). Addition of forskolin resulted in increases in maximal contraction amplitudes at all three [Ca2+]o, in both wild-type (P < 0.0002) and PLM-null (P < 0.0001) myocytes (Table 7). However, the differences in contraction amplitudes between unstimulated wild-type and PLM-null myocytes measured at 0.6 and 5.0 mM [Ca2+]o were no longer apparent after forskolin treatment (P < 0.35). Notably, the forskolin-induced percent increase in contraction amplitude was larger in PLM-null myocytes at 0.6 mM [Ca2+]o but smaller at 5.0 mM [Ca2+]o when compared with wild-type myocytes (P < 0.043).
Forskolin accelerated both maximal shortening and relaxation velocities in both wild-type (P < 0.0004) and PLM-null (P < 0.0001) myocytes (Table 7). The increase in shortening and relaxation velocities by forskolin progressively decreased as [Ca2+]o was increased (forskolin × [Ca2+]o interaction effect, P < 0.02). There were no differences in either maximal shortening (P < 0.48) and relaxation (P < 0.50) velocities between wild-type and PLM-null myocytes after forskolin treatment.
Effects of PMA on contraction in wild-type and PLM-null myocytes.
In a third series of myocyte contraction experiments, we evaluated the effects of protein kinase C (PKC) activation on contraction in wild-type and PLM-null myocytes. In the absence of PMA, PLM-null myocytes tended to contract less at 0.6 mM [Ca2+]o but more at 5 mM [Ca2+]o when compared with wild-type myocytes (Table 8), similar to what we observed in the first 2 series of contraction experiments (Tables 2 and 7). Stimulation of PKC by PMA resulted in significant (P < 0.035) decreases in maximal contraction amplitudes and shortening and relengthening velocities in both wild-type and PLM-null myocytes (Table 8). In the presence of PMA, there were no differences in maximal myocyte contraction amplitudes or shortening and relaxation velocities between wild-type and PLM-null myocytes (P < 0.52).
PLM (FXYD1) and other members of the FXYD gene family, including γ-subunit of the Na+-K+-ATPase (FXYD2; see Ref. 29), channel-inducing factor (FXYD4; see Ref. 5), and FXYD7 (6), are known to be regulators of Na+-K+-ATPase. In addition, a 15-kDa homologue of phospholemman isolated from shark rectal glands, PLMS, also associated with and regulated the activity of the α-subunits of Na+-K+-ATPase (17). PLM, however, is the only FXYD family member to have a consensus sequence for phosphorylation by PKA (RRXS), PKC (RXXSXR), and “never in mitosis” A kinase (FRXS/T; see Ref. 27). In addition, PLM is the only FXYD member shown to date to regulate important sarcolemmal ion transporters other than Na+-K+-ATPase. Specifically, PLM colocalized with (38), coimmunoprecipitated with (2, 18), and inhibited NCX1 (18, 25, 38) in rat cardiac myocytes. In addition, the mechanisms by which PLM regulate Na+-K+-ATPase and NCX1 appear to be quite different. Based on analogy of phospholamban inhibition of SERCA2 and experimental observations on the effects of PLM on shark Na+-K+-ATPase (17), the current working model is that the Na+ pump is inhibited by unphosphorylated PLM. On phosphorylation of PLM, inhibition of Na+-K+-ATPase is relieved. This hypothesis has been given strong support by the observation that the maximal velocity (Vmax) of sarcolemmal Na+-K+-ATPase was increased threefold after acute cardiac ischemia, in association with increased PLM phosphorylation by >300% (12). In addition, comparison of β-adrenergic effects on Na+ pump function between wild-type and PLM-null myocytes supports the notion that the inhibitory effects of PLM on Na+-K+-ATPase are relieved by phosphorylation (11). Finally, Silverman et al. (24) demonstrated that the Na+ pump current was directly increased in association with PLM phosphorylation in response to forskolin. By contrast, based on the assumption that the serine-68 to alanine (S68A) and serine-68 to glutamic acid (S68E) mutants of PLM faithfully mimic the unphosphorylated and phosphorylated forms of PLM, respectively, and the observations that S68A mutant had no effect on INaCa, whereas S68E mutant inhibited cardiac INaCa much more effectively than wild-type PLM (25), we proposed that the phosphorylated PLM form inhibits the NCX. Follow-up experiments using a heterologous expression system unambiguously proved that PLM, when phosphorylated at serine-68, inhibited NCX1 (34). These observations suggest a coordinated paradigm in which PLM, on phosphorylation by PKA (serine-68) or PKC (serine-63 and serine-68; see Refs. 30 and 36), enhances Na+-K+-ATPase but inhibits NCX1 activities in cardiac myocytes. Thus PLM is unique among FXYD gene family members in that not only does it regulate two important sarcolemmal ion transporters but the same phosphorylation mechanism activates one but inhibits the other ion transporter.
The first major finding is that, at the cellular level, PLM-null mice backcrossed to congenic C57BL/6 background did not exhibit myocyte hypertrophy at 3–6 mo of age or changes in levels of some proteins intimately associated with excitation-contraction coupling. This is in contrast to our previous report on PLM-null mice with a mixed C57BL/6 129/SvJ genetic background in which the cardiac mass-to-tibial length ratio was 24% larger in PLM-null mice than their wild-type counterparts, and cardiomyocytes of PLM-null animals appeared to be larger than those from wild-type mice (14). The differences may be the result of different techniques used in estimating cell sizes (counting nuclei per unit area in photomicrographs of heart tissue vs. measurement of individual cell length and width and whole cell capacitance) or because of phenotypic variation from animals with a mixed genetic background. Indeed, in a recent study utilizing mice backcrossed to pure congenic C57BL/6 background, whole cell capacitance was not different between wild-type and PLM-null myocytes at 3–4 mo of age (11). Another possibility to account for the differences may be that cardiac hypertrophy would not be manifest until later in life.
Previous measurements on α-subunits of Na+-K+-ATPase in myocytes isolated from PLM-null mice indicated decreased expression when compared with their wild-type littermates, based on ouabain binding (25%) and myocyte Western blot (20%, using the isoform nonspecific α5-antibody; see Ref. 11). Although Jia et al. (14) also reported a statistically insignificant 19% decrease in α-subunits of Na+-K+-ATPase (using α-isoform pan-specific antibody), more detailed analysis using α1- and α2-specific antibodies demonstrated that α2, but not α1, was significantly decreased in heart sarcolemma of PLM-null mice. Our observation that α1-subunit levels of Na+-K+-ATPase were unchanged in heart sarcolemma of PLM-null mice is thus similar to the results of Jia et al. (14). Taken together, it appears that downregulation of α-subunits of Na+-K+-ATPase in PLM-null hearts was largely the result of decreased expression of α2 isoforms.
The second major finding is that the contractile behavior of PLM-null myocytes clearly displayed an enhanced dynamic range in response to increases in [Ca2+]o. This contractile phenotype is similar to that observed in rat myocytes in which PLM was downregulated (18) and reminiscent of the contractile behavior in NCX1-overexpressed rat myocytes (39). By contrast, overexpressing PLM (26) or downregulating NCX1 (28) in cultured adult rat myocytes resulted in a reduced dynamic range in response to altered [Ca2+]o: exactly opposite to that observed in PLM-null myocytes. The results from these fundamentally different experimental approaches indicate that: 1) altering NCX1 levels (and by implication, activity) results in changes in myocyte contractile behavior and 2) one of the physiological functions of PLM in cardiac tissues is regulation of cardiac contractility, plausibly by modulating NCX1 activity (2, 25, 34, 38).
Changes in NCX1 activity in PLM-null myocytes would be expected to alter Ca2+ fluxes during excitation-contraction coupling. Indeed, compared with wild-type myocytes, [Ca2+]i transient amplitudes were lower at 0.6, not different at 1.8, and higher at 5.0 mM [Ca2+]o in PLM-null myocytes. This pattern of altered [Ca2+]i transients is similar to that observed in PLM-downregulated (18) or NCX1-overexpressed (39) rat myocytes and opposite to that observed in PLM-overexpressed (26) or NCX1-downregulated (28) rat myocytes. Altered [Ca2+]i transient amplitudes in PLM-null myocytes are most likely the result of differences in SR Ca2+ load as a result of enhanced NCX1 activity (28, 39). Indeed, SR Ca2+ contents in PLM-null myocytes were significantly lower at 0.6 but higher at 5 mM [Ca2+]o, mirroring the changes in [Ca2+]i transient and contraction amplitudes. Note also that, despite significant changes in [Ca2+]i transient amplitudes and SR Ca2+ contents, there were no differences in L-type ICa density, SERCA2 protein levels, or SR Ca2+ uptake activity (as reflected by t1/2 of [Ca2+]i transient decline) in PLM-null myocytes. It should also be recalled that resting intracellular Na+ concentration ([Na+]i) was similar between wild-type and PLM-null myocytes (12.5 ± 1.8 vs. 12.0 ± 1.5 mM; see Ref. 11), suggesting that altered [Ca2+]i transients, SR Ca2+ contents, and contractility in PLM-null myocytes were the result of direct effects of PLM on NCX1 rather than indirect effects because of changes in [Na+]i from Na+-K+-ATPase inhibition.
Although there were no differences in t1/2 of [Ca2+]i transient decline between wild-type and PLM-null myocytes, maximum relengthening velocity was faster in PLM-null myocytes (especially at higher [Ca2+]o), suggesting that factors in addition to [Ca2+]i transient recovery contribute to myocyte relaxation. Indeed, in rabbit, rat (4), and canine (16) cardiac myocytes, relaxation was complete well before the end of the [Ca2+]i transient. For example, in rat myocytes stimulated at 1.0 mM [Ca2+]o and 22°C, t1/2 from a twitch was 0.08 ± 0.01 s, whereas the time constant of [Ca2+]i transient decline was 0.194 ± 0.10 s (4). At 5.0 mM [Ca2+]o and 37°C, t1/2 from a twitch was 120 ± 3 ms (32), but t1/2 of [Ca2+]i transient decline was 197 ± 5 ms in adult rat myocytes (33). The fact that myocyte relaxation was complete well ahead of [Ca2+]i transient decline suggests that mechanisms in addition to SR Ca2+ uptake and forward Na+/Ca2+ exchange contribute to myocyte mechanical relaxation.
Direct measurements of NCX1 currents in PLM-null myocytes demonstrated that both forward and reverse Na+/Ca2+ exchange were increased compared with wild-type myocytes (34). The conditions used in our INaCa measurements were carefully designed to ensure that the thermodynamic parameters ([Ca2+]o, [Ca2+]i, extracellular Na+ concentration, [Na+]i) that determined the equilibrium potential (ENaCa) of INaCa, and hence its driving force (Em − ENaCa), were similar between wild-type and PLM-null myocytes (34). There were also no differences in NCX1 protein levels between wild-type and PLM-null myocytes. Thus the differences in INaCa between wild-type and PLM-null myocytes could be unambiguously assigned to the effects of PLM deficiency.
Another major finding is that, despite similar Em and action potential amplitude, the action potential duration was dramatically prolonged in PLM-null myocytes. Prolongation of action potential plateau was not because of changes in ICa, since both the current density, slope conductance, and inactivation time constants were similar between wild-type and PLM-null myocytes. Alteration in NCX1 activity may theoretically change action potential duration (3). In rat and mouse cardiac myocytes with short action potential duration, there is a net Ca2+ efflux via NCX1 at the shoulder of the action potential. This is because the action potential is already repolarizing before the [Ca2+]i transient reaches its peak, and INaCa is predominantly inward during this phase of the action potential (23). Therefore, increases in NCX1 activity in PLM-null myocytes would be expected to prolong the action potential duration, which is what we observed. The measured differences in INaCa between wild-type and PLM-null myocytes at the shoulder/plateau of the action potential (approximately −40 mV), however, appeared to be quite modest and would not be expected to have such a profound effect on action potential duration. It should be recalled, however, that the conditions we used for measuring INaCa (5 mM [Ca2+]o, heavily buffered [Ca2+]i) were biased to measure predominantly outward INaCa (Ca2+ influx) and to rigorously control the thermodynamic driving force for INaCa (34) and therefore quite different from those present during a normal mouse action potential. Changes in other repolarizing currents, e.g., transient outward currents (40) or other K+ currents, may also contribute to prolonged action potential in PLM-null myocytes. The present study does not go into sufficient detail to delineate the ionic mechanisms of action potential prolongation in PLM deficiency.
The magnitude of ICa (∼6 pA/pF or ∼900 pA/cell) measured in our wild-type murine myocytes is similar to the value of 764 ± 63 pA/cell reported by Zhou et al. (41). There were no differences in ICa magnitude, slope conductance, or kinetics between wild-type and PLM-null myocytes, although NCX1 activity was increased in PLM-null myocytes. Although complete absence of Na+/Ca2+ exchange activity in cardiac-specific NCX1 knockout myocytes resulted in 50% reduction in ICa (13), less severe perturbations in NCX1 levels (and by implication, activity) were not associated with detectable changes in ICa. For example, neither heterologous overexpression of NCX1 in mice (∼200% increase in NCX1 levels; see Refs. 1 and 31) nor NCX1 downregulation in adult rat myocytes (∼30% decrease in NCX1 levels after 72 h; see Ref. 28) resulted in changes in ICa densities. Thus the 22–30% increase in INaCa observed in PLM-null murine myocytes or the ∼25% increase in INaCa associated with PLM downregulation in rat myocytes (18) would not be expected to induce significant changes in ICa density or inactivation kinetics.
Because PLM is known to regulate cardiac Na+-K+-ATPase (9, 11, 12, 14, 24, 36), it is tempting to explain the observed changes on excitation-contraction coupling and NCX1 activity in PLM-null myocytes simply by changes in [Na+]i due to alterations in Na+-K+-ATPase activity, thereby altering the driving force for NCX1. This is unlikely based on the following observations. First, inhibition of Na+-K+-ATPase by PLM in wild-type myocytes would theoretically increase [Na+]i, thereby decreasing the driving force for forward Na+/Ca2+ exchange (Ca2+ efflux) and increasing the driving force for reverse Na+/Ca2+ exchange (Ca2+ influx). Both of these actions will increase [Ca2+]i and result in enhanced myocyte contractility at all [Ca2+]o. This is not observed, especially at high [Ca2+]o where wild-type myocytes actually had lower contraction amplitude than PLM-null myocytes. In addition, measured [Na+]i levels were similar between wild-type and PLM-null myocytes (11), suggesting that altered contractility and [Ca2+]i transients were not the result of [Na+]i changes with resultant indirect effects on NCX1 activity. On the other hand, increase in NCX1 activity in PLM-null myocytes would result in increased dynamic range of contractility and [Ca2+]i transients, as we have observed in rat myocytes in which NCX1 was overexpressed (39). Second, in our INaCa measurements, Na+-K+-ATPase activity was eliminated because of absence of K+ and presence of ouabain, yet we still detected significant differences in INaCa between wild-type and PLM-null myocytes (34). Third, although phosphorylation relieved inhibition of Na+-K+-ATPase by PLM (11, 12, 24), PLM phosphorylated at serine-68 is the PLM form that inhibited NCX1 (2, 25, 34). The different mechanisms by which PLM inhibits Na+-K+-ATPase (dephosphorylation) and NCX1 (phosphorylation) argue that PLM exerts 9 direct effect on NCX1. Fourth, in HEK 293 cells expressing exogenous PLM and NCX1, PLM inhibited Na+-dependent Ca2+ uptake in the presence of ouabain (2), indicating a direct effect of PLM on NCX1.
Activation of PKA resulted in increased phosphorylation of serine-68 in PLM (30, 36) but had no effect on INaCa in adult murine myocytes (34). We previously speculated that PKA stimulation in intact cardiac myocytes would simultaneously increase phosphorylation of both NCX1 (stimulatory; see Ref. 21) and PLM (inhibitory), plus or minus other unknown effects on protein phosphatase 1 (PP1) such that the net effect would be no measurable changes in INaCa (34). In the unstimulated state, INaCa was higher in PLM-null myocytes, and this was correlated with the increased dynamic range of contraction in response to increases in [Ca2+]o. After PKA activation, there were no differences in INaCa between wild-type and PLM-null myocytes (34), and this was associated with no differences in contraction amplitudes at all three [Ca2+]o examined. The large increases in contraction amplitudes after PKA stimulation observed in both wild-type and PLM-null myocytes were because of multiple effects, including but not limited to increases in ICa and SR Ca2+ uptake. Because forskolin-treated wild-type and PLM-null myocytes had similar contraction amplitudes, the differences in SR Ca2+ content observed in unstimulated wild-type and PLM-null myocytes (because of different NCX1 activities) were likely abolished by enhanced Ca2+ entry and SR Ca2+ uptake after PKA activation.
Activation of PKC increased both phosphorylation of phospholemman (30, 36) and INaCa in intact cardiac myocytes (34). Despite larger increases in INaCa in response to PMA in PLM-null myocytes (34), contraction amplitudes of both wild-type and PLM-null myocytes were depressed to similar extents by PMA treatment. Our observations that PKC activation resulted in suppression of myocyte contractility are in agreement with those published in the literature (8, 22). The mechanism by which stimulation of PKC-α, the predominant PKC isoenzyme expressed in murine hearts, affects contractility was thought to be due in part to increased PP1 activity. Enhanced phosphatase activity leads to phospholamban hypophosphorylation and thereby promotes greater inhibition of SERCA2 and decreased SR Ca2+ content (8).
Over a decade ago, Nuss and Houser (19) proposed that one of the physiological functions of NCX1 is to modulate SR Ca2+ load. In wild-type and PLM-null myocytes in which INaCa were different but SERCA2 activities and ICa densities were similar, different SR Ca2+ contents observed are consistent with the Nuss and Houser hypothesis. When SERCA2 activity was depressed by PKC-α activation (8), contractility in wild-type and PLM-null myocytes suffered to similar extent despite unequal increases in INaCa induced by PMA (34). By contrast, when SERCA2 but not NCX1 activities were enhanced by PKA activation, contractility in both wild-type and PLM-null myocytes improved to similar extents. These observations suggest the primacy of SERCA2 over NCX1 in terms of controlling SR Ca2+ content, a major determinant of myocyte contraction amplitude.
There are limitations to the present study. The first is that we did not measure Na+-K+-ATPase activity (although α1-subunit levels were similar between wild-type and PLM-null myocytes). This is because reduction in Na+-K+-ATPase activity has been previously reported in PLM-null myocytes on a mixed C57BL/6 129/SvJ background (14). In PLM-null myocytes on a pure congenic C57BL/6 background, Vmax of the Na+ pump was similar, but Km was lower when compared with wild-type myocytes (11). In addition, the contractile phenotype of enhanced dynamic range in response to increasing [Ca2+]o observed in PLM-null myocytes was more compatible with alterations in NCX1 than Na+-K+-ATPase activities, as we have discussed in detail previously (18, 25, 26). Another limitation is that we did not perform studies in sufficient detail to characterize the ionic basis for action potential prolongation in PLM-null myocytes. Finally, we have totally ignored the osmoregulatory properties of PLM (10, 15). Cell length, cell width, and whole cell capacitance, however, were not different between wild-type and PLM-null myocytes, suggesting that PLM deficiency had no detectable effects on steady-state cell volume.
In summary, we demonstrated that PLM deficiency affected cardiac contractility, [Ca2+]i transients, SR Ca2+ contents, Na+/Ca2+ exchange activity, and action potential duration. Expression of NCX1, SERCA2, calsequestrin, and α1-subunit of Na+-K+-ATPase was unaffected by PLM knockout. ICa density, ICa slope conductance and inactivation kinetics, resting Em, action potential amplitude, and SR Ca2+ uptake were similar between wild-type and PLM-null myocytes. PLM deficiency did not result in myocyte hypertrophy when evaluated at 3–6 mo of life. PKA stimulation resulted in enhanced myocyte contraction to similar extents between wild-type and PLM-null myocytes. PKC activation depressed myocyte contraction equally in wild-type and PLM-null myocytes. We conclude that one of the physiological functions of PLM in the heart is regulation of contractility, likely by modulating Ca2+ fluxes via NCX1.
This study was supported in part by National Institutes of Health Grants HL-58672 and HL-74854 (to J. Y. Cheung), DK-46678 (to J. Y. Cheung, co-investigator), HL-70548 and GM-64640 (to J. R. Moorman), GM-69841 (to L. I. Rothblum), and HL-69074 (to A. L. Tucker); American Heart Association, Pennsylvania Affiliate, Grants-in-Aid 0265426U (to X. Q. Zhang) and 0355744U (to J. Y. Cheung); American Heart Association, Pennsylvania Affiliate, Postdoctoral Fellowship 0425319U (to B. A. Ahlers); and grants from the Geisinger Foundation (to J. Y. Cheung and L. I. Rothblum). This investigation was conducted in a facility constructed with support from Research Facilities Improvement Grant no. C06 RR-15428–01 from the National Center for Research Resources, National Institutes of Health.
We thank Caitlin Custer and Janelle Roman for assistance in preparation of the manuscript.
↵* A. L. Tucker and J. Song contributed equally to this study.
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- Copyright © 2006 by the American Physiological Society