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1Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, Missouri 63110; 2Institut National de la Santé et de la Recherche Médicale Unitè 533, Physiopathologie et Pharmacologie Cellulaires et Moléculaires, Faculté de Médecine, 44035 Nantes, France; 3Department of Physiology, University of Michigan Medical School, Ann Arbor, Michigan 48109; and 4Department of Pharmacology, University of Alberta, Edmonton T6G 2H7, Alberta, Canada
Submitted 25 July 2003 ; accepted in final form 2 December 2003
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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N2–30,K185Q)] under transcriptional control of the 
-myosin heavy chain promoter. Strikingly, myocyte contraction amplitude was increased in TG myocytes (15.68 ± 1.15% vs. 10.96 ± 1.49%), even though KATP channels in TG myocytes are very insensitive to inhibitory ATP. Under normal metabolic conditions, steady-state outward K+ currents measured under whole cell voltage clamp were elevated in TG myocytes, consistent with threshold KATP activation, but neither the monophasic action potential measured in isolated hearts nor transmembrane action potential measured in right ventricular muscle preparations were shortened at physiological pacing cycles. Taken together, these results suggest that there is a compensatory remodeling of excitation-contraction coupling in TG myocytes. Whereas there were no obvious differences in other K+ conductances, peak L-type Ca2+ current (ICa) density (–16.42 ± 2.37 pA/pF) in the TG was increased compared with the wild type (–8.43 ± 1.01 pA/pF). Isoproterenol approximately doubled both ICa and contraction amplitude in wild-type myocytes but failed to induce a significant increase in TG myocytes. Baseline and isoproterenol-stimulated cAMP concentrations were not different in wild-type and TG hearts, suggesting that the enhancement of ICa in the latter does not result from elevated cAMP. Collectively, the data demonstrate that a compensatory increase in ICa counteracts a mild activation of ATP-insensitive KATP channels to maintain the action potential duration and elevate the inotropic state of TG hearts.
ATP-sensitive K+ channels; contractility; action potential
The physiological role of one cardiac ion channel, the ATP-sensitive K+ (KATP) channel in the sarcolemma (21), has thus far remained elusive. Under normal metabolic conditions, KATP channels are not significantly open and thus do not contribute to AP repolarization and excitation-contraction coupling (5, 19, 27, 28). When myocytes are exposed to a severe metabolic stress (anoxia, hypoxia, metabolic inhibition, ischemia, etc.), KATP channels do open, causing AP shortening and contractile failure (13, 27, 28). However, the precise trigger for KATP channel activation, the timing of channel opening, and physiological consequences in vivo remain murky, and nature offers no clues because there are no known KATP mutations linked with cardiac disease.
The coexpression of two gene products, Kir6.2 and SUR2A, in heterologous expression systems produces channels with the essential properties of the cardiac sarcolemmal KATP channel (2) and, in vitro, these two proteins physically associate (15). Thus it is now widely accepted that KATP channels are heteromeric protein complexes composed of four Kir6.2 and four SUR2A subunits. The Kir6.2 subunits form the potassium-selective pore and contain determinants of ATP inhibition, phosphatidylinositol 4,5-bisphosphate activation, and inward rectification, whereas the SUR subunits contribute additional determinants of channel regulation (24).
To further probe the function of the cardiac sarcolemmal KATP channel, we recently engineered several TG mouse lines that express an ATP-insensitive Kir6.2 mutant [Kir6.2 (N30,K185Q)] under the transcriptional control of the -myosin heavy chain promoter (10). Four independent TG lines were established, and two have been extensively characterized. In one line (line 4 TG), all isolated myocytes express mutant Kir6.2 subunits, and KATP channels are very insensitive to inhibition by ATP (K1/2 = 2.7 ± 0.01 mM) compared with wild-type (WT) myocytes (K1/2 = 51 ± 2 µM) (10). In another line (line 2 TG), the expression level varies from cell to cell, and the sensitivity of the sarcolemmal KATP channels is not as dramatically altered (K1/2 = 514 ± 5 µM) (10). Because computer modeling studies (18, 25) and experimental simulations (9, 20, 29) predict that activation of only 1% of the KATP channels in a ventricular myocyte should be sufficient to significantly shorten the AP duration (APD) at 90% repolarization (APD90) and reduce contractility, it was expected that, in these TG hearts, the AP would be shortened and contractility would be reduced.
Remarkably, there is no significant electrophysiological impairment in these TG mice (10, 22). Although there is a mild reduction of heart rate and ventricular ectopy as well as an increased incidence of atrioventricular block, electrocardiographic waveforms are similar between WT and TG mice (10). In addition, the AP recorded in isolated cardiac myocytes at room temperature is not shortened (10). Contractility is also not reduced. In fact, left ventricular developed pressure is significantly elevated in isolated TG hearts compared with WT hearts (22).
In the present study, we expand on these initial findings to further examine the electrophysiological and contractile properties of these TG mice. Here, we elucidate one compensatory mechanism that may underlie the altered electrical and contractile function, demonstrating that there is stimulation of the contraction trigger, L-type Ca2+ channel current (ICa), which can lead to increased contractility and contribute to the absence of AP shortening in TG myocytes.
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Solutions. Wittenberg isolation medium (WIM) contained the following (in mM): 116 NaCl, 5.3 KCl, 1.2 NaH2PO4, 11.6 glucose, 3.7 MgCl2, 20 HEPES, 2.0 L-glutamine, 4.4 NaHCO3, and 1.5 KH2PO4, with 1x essential vitamins (GIBCO-BRL catalog no. 12473-013) and 1x amino acids (GIBCO-BRL catalog no. 11120-052); pH 7.3–7.4.
Normal Tyrode solution contained the following (in mM): 137 NaCl, 5.4 KCl, 0.16 NaH2PO4, 10 glucose, 1.8 CaCl2, 0.5 MgCl2, 5.0 HEPES, and 3.0 NaHCO3; pH 7.3–7.4.
Krebs-Henseleit (KH) solution contained the following (in mM): 118 NaCl, 4.7 KCl, 25 NaHCO3, 1.2 NaH2PO4, 1.2 MgSO4, 1.2 CaCl2, and 10 glucose and was saturated with a 95% O2-5% CO2 gas mixture; pH 7.3–7.4.
Intracellular K+ solution contained the following (in mM): 140 KCl, 5 K2ATP, 10 HEPES, and 10 EGTA; pH 7.3–7.4.
Intracellular Cs+ solution contained the following (in mM): 120 CsCl, 20 TEA-Cl, 5 ATP, and 10 HEPES; pH 7.3–7.4.
Isolation of cardiac myocytes. Mice were housed in the animal facility at Washington University and fed a standard diet and water ad libitum. All animal procedures were approved by the Animal Studies Committee at Washington University School of Medicine. Mice were anesthetized by subcutaneous injection with a ketamine-xylazine-acepromazine cocktail. The heart was rapidly excised, and the aorta was cannulated. The heart was retrogradely perfused with a Ca2+-free solution for 5 min, followed by perfusion with a digestion solution containing 0.8 mg/ml collagenase (type 2, Worthington Biochemical) and 0.01 mg/ml protease (type XIV, Sigma). Left ventricular cells were gently dispersed by manual trituration using a pasteur pipette. Cells were stored at room temperature in WIM.
Cellular electrophysiology. Isolated myocytes were transferred into a recording chamber containing either normal or cesium-containing (replacement of KCl with 5.4 mM CsCl) Tyrode solution. Isoproterenol (Iso; 1 µM) and tetrodotoxin (TTX; 10 µM) were added to the bathing medium where indicated. Macroscopic currents in isolated ventricular myocytes were recorded using standard whole cell voltage-clamp recording techniques. Patch-clamp electrodes (1–3 M
when filled with electrode solution) were fabricated from soda lime glass microhematocrit tubes (Kimble 73813). Cell capacitance and series resistance were determined using a 5- to 10-mV hyperpolarizing square pulse from a holding potential of –80 mV after establishment of the whole cell recording configuration. The series resistance was electronically compensated by 60–80%. pCLAMP 6.0 software and a DigiData 1200 converter were used to generate command pulses and collect data. The specific voltage-clamp protocols are described in the figures and text where appropriate. Data were filtered at 5 kHz.
Single cell contraction measurements. To assess contractility, unloaded cell shortening of isolated ventricular myocytes was measured using a video edge detection system (Crescent Electronics). Cells were placed in the recording chamber, bathed with normal Tyrode solution, and stimulated with a bipolar stimulating electrode placed near the cell. Electrodes were constructed by immersing Ag/AgCl wires in Tyrode solution within a 
-glass capillary tube pulled to a fine tip.
Monophasic AP recording. Monophasic APs (MAPs) were measured from freshly isolated hearts. After excision, hearts were retrogradely perfused through the aorta with KH solution at a constant pressure of 75–85 cmH2O and submerged in a glass tissue bath that was continuously bathed with KH solution, maintained at 37°C. After an initial equilibration period (15–30 min), MAPs were recorded using a DP-304 differential amplifier (Axon Instruments) and digitized at 3.3 kHz with a Digidata 1322A and pCLAMP 8.0 software. MAP electrodes were constructed from Teflon-coated silver wire (0.01 in. diameter). The exposed tip of the wire was polished with fine sandpaper and plated with AgCl. APD was determined as the time between the maximum upstroke velocity and 30% (APD30), 60% (APD60), or 90% of full repolarization (APD90).
Microelectrode studies. Transmembrane recordings were obtained as previously described (4). Briefly, the heart was quickly removed and immersed in a cool modified Tyrode solution containing (in mM) 108 NaCl, 25 NaHCO3, 1.8 NaH2PO4, 27 KCl, 1 MgCl2, 0.6 CaCl2, and 55 glucose, which was saturated with a 95% O2-5% CO2 gas mixture (pH 7.3). The right free ventricular wall was dissected from the heart and sliced to obtain the experimental preparations (1–1.5 mm wide and 
3 mm long). Preparations were mounted in a Lucite tissue chamber with the epicardial surface facing down, so that the endocardial surface could be impaled with a microelectrode. The preparation was superfused with oxygenated (95% O2-5% CO2) Tyrode solution warmed to 37 ± 0.5°C containing (in mM) 120 NaCl, 27 NaHCO3, 1.2 NaH2PO4, 5.4 KCl, 1.2 MgCl2, 1.8 CaCl2, and 10 glucose (pH 7.4). The flow rate in the tissue chamber was 10 ml/min. The tissue was allowed to recover for at least 1 h before the experiment. During this period, the tissue was paced at a cycle length of 200 ms by bipolar stimulation through Teflon-coated silver wire electrodes. The stimulus pulse width was 1.5 ms, and the amplitude was twice threshold. AP characteristics were measured at the steady state for each pacing cycle length. APDs were measured at APD30, APD60, and APD90.
Radioimmunoassay to detect cAMP. Hearts were rapidly excised and retrogradely perfused for 10 min with Ca2+-free WIM (±1 µM Iso). After perfusion, the ventricles were immediately removed and submerged in liquid nitrogen. The tissue was homogenized in 0.3 M perchloric acid. After centrifugation, the supernatant was neutralized with K2HPO4, and the cAMP concentration was assessed by radioimmunoassay (Amersham) following the manufacturer's protocols.
Real-time RT-PCR. The relative expression of Cav1.2 and Kir6.2 mRNA in TG compared with WT hearts was assessed using real-time RT-PCR (3). Total RNA was isolated from cardiac ventricles using TRIzol (Invitrogen) following the manufacturer's protocols. The isolated RNA was then further purified using a silica-based column protocol (RNeasy, Qiagen). The RNA concentration was determined spectrophotometrically (Nanodrop Technologies). RT and PCR were carried out in a single tube in an ABI Prism 7000 sequence detection system (Applied Biosystems); 100 ng of template RNA were used in all reactions. after the RT reaction (30 min, 48°C), 40 cycles of PCR were carried out. Double-stranded DNA was fluorescently labeled with SYBR green (Applied Biosystems). Gene-specific primers for Cav1.2 and Kir6.2 were designed using PrimerExpress software (ABI) and purchased from Integrated DNA Technologies. 
-Actin control primers were obtained from Ambion. Reactions with each primer pair and template were performed in triplicate or quadruplicate. After baseline correction, a fluorescence threshold was established, and the cycle when this threshold was crossed (Ct) was determined for each reaction. To control for variability in RNA quantity, the normalized value, (Ct) for each sample was determined using the formula Ct = Ct(actin) – Ct(Cav1.2/Kir6.2). The relative expression in TG tissue (normalized to WT tissue) was then determined using the following relationship: gene expression = 2–Ct, where Ct = Ct (WT) – Ct (TG).
Data analysis. All data were analyzed using ClampFit and Microsoft Excel software, and (except where noted in the figures) results are presented as means ± SE. Statistical tests and P values are denoted in the figures where appropriate.
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RESULTS |
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Contractility is enhanced in TG myocytes. The increased contractility detected in isolated TG hearts (22) was also observed in isolated myocytes in this study. To assess contractile function, the contraction amplitude of single isolated ventricular cells was measured using video edge detection. Typical results are shown in Fig. 3A. The fractional shortening of line 4 TG myocytes was largely independent of the stimulus frequency over the range of 0.5–2 Hz, similar to previously reported studies (1, 26, 30), and was significantly greater (15.68 ± 1.15%, n = 14) than that of control cells (9.86 ± 0.96%, n = 27) at a stimulus frequency of 1 Hz. We also examined the dependence of contraction amplitude on extracellular Ca2+ concentration ([Ca2+]o). The contraction amplitude was assessed in normal Tyrode solution containing 0.3–3.6 mM Ca2+, and the data were fit with a modified Hill equation (see Fig. 3). As shown in Fig. 3B, the [Ca2+]o dependence of the contraction amplitude was significantly shifted in line 4 TG myocytes (K
= 0.89 ± 0.08 mM) compared with control cells (K = 1.50 ± 0.09 mM).
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Frequency and calcium dependence of contraction was also studied in myocytes from line 2 TG myocytes. Fractional shortening also tended to be increased in the line 2 TG myocytes; however, this increase was not significantly different (P > 0.05) from either WT or line 4 TG myocytes, but lay between the two (Fig. 3A). The calcium dependence of line 2 TG myocytes was also intermediate (K = 1.18 ± 0.17 mM) between the WT and line 4 TG extremes (Fig. 3B). Because the overall expression of the transgene is lower and more variable in line 2 TG mice, these data suggest that the magnitude of the phenotype is commensurate with the level of transgene expression and not simply an artifact of the transgene insertion point.
Compensatory increase in ICa can account for increased myocardial contractility and maintained APD. The above results collectively suggest that excitation-contraction coupling is significantly altered as a consequence of the altered KATP channel substrate in TG hearts. Several compensatory mechanisms could conceivably explain the remodeling of excitation-contraction coupling. We began by examining two alternative, but not mutually exclusive, hypotheses that might explain the observed consequences for both the AP waveform and myocardial contractility. First, an alteration in either the amplitude or the kinetics of voltage-dependent K+ currents could prolong the APD. Such a mechanism has previously been proposed to account for the AP prolongation and increased Ca2+ transient amplitude induced after myocardial infarction (7, 23). Alternatively, an increase in ICa could produce both an increase in APD and myocyte contractility. Therefore, we assessed both outward K+ conductances and ICa in isolated cardiac myocytes using the whole cell voltage-clamp technique.
Representative families of macroscopic currents elicited by 4.5-s depolarizing voltage-clamp pulses (from –50 to +50 mV in 10-mV increments) and the corresponding mean current-voltage relationships for both the peak and steady-state currents are shown in Fig. 4. There were no differences in the inactivation time constants of the fast and slow components of the transient outward K+ current. Both peak and steady-state K+ current were slightly increased in TG myocytes at positive holding potentials. The difference current (IDIFF = ITG – IWT; Fig. 4, C and D) revealed a time-independent, weakly inward rectifying current. Such a current is consistent with the predicted activation of KATP channels given that KATP channels in TG mice are very insensitive to inhibition by intracellular ATP (10). Because deletion of the NH3 terminal of Kir6.2 abolishes the channels sulfonylurea sensitivity (12), we cannot conclusively demonstrate pharmacologically that this current is the KATP channel current (IKATP). Regardless, because net outward K+ currents are not decreased, but are actually increased, changes in K+ currents could not explain the lack of AP shortening and enhanced contractility. Collectively, the data suggest that voltage-gated K+ conductances do not differ in WT and TG myocytes.
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In contrast to the lack of effect on voltage-gated K+ channels, peak ICa was increased in both line 4 and line 2 TG myocytes. Representative families of macroscopic Ca2+ currents are shown in Fig. 5A. Peak currents (membrane potential = 5 mV) were increased from –10.46 ± 1.01 pA/pF in control myocytes (n = 31) to –14.72 ± 1.94 pA/pF in line 4 TG myocytes (n = 23). The increase was significant at all voltages tested between –25 and +15 mV (Fig. 5B). Ca2+ current in line 2 TG myocytes also tended to be elevated (Fig. 5B). As with the functional measurements of cell contractility, ICa was not significantly different from either WT or line 4 TG, but again fell between the two. Collectively, the data demonstrate that there is a compensatory increase in ICa in TG myocytes that likely contributes to both maintaining APD and enhancing contractility in TG myocytes.
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Calcium channel mRNA expression is not elevated. We examined the possibility that the increase of ICa resulted from an increased expression of the Ca2+ channel gene using real-time RT-PCR. The transcription of Cav1.2, Kir6.2, and -actin were assessed in RNA samples obtained from WT (n = 3) and line 4 TG hearts (n = 5). The results of these experiments are summarized in Fig. 6. In contrast to the Kir6.2 transcript, which was significantly elevated (156-fold), the expression of Cav1.2 transcripts were not significantly different between the WT and TG hearts, suggesting that an upregulation of channel mRNA does not explain the increased ICa in TG myocytes.
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ICa is prestimulated. Because the expression does not appear to be upregulated, we considered the possibility that ICa is hyperactivated in TG mice. As expected, the application of 1 µM Iso to non-TG myocytes caused an increase in the peak amplitude of both contraction (Fig. 7, A and B) and ICa (Fig. 7C). In contrast, there was no increase in either ICa or contraction in myocytes isolated from TG mice. Because Iso is without effect on the TG myocytes, increased PKA-dependent channel phosphorylation appears a likely mechanism for increased basal ICa in TG myocytes. To test the possibility that elevated cAMP underlies this stimulation, isolated hearts were perfused with or without Iso, and [cAMP] in tissue lysate was measured using radioimmunoassay. As shown in Fig. 7D, there was no difference in either basal or Iso-stimulated [cAMP] in WT or TG hearts. While elevated cAMP is thus excluded, the data are consistent with the notion that the chronic activation of sarcolemmal KATP channels causes a prestimulation of ICa (through an as-yet-unknown mechanism) that maintains APD and myocardial contractility despite an increase in IKATP.
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The molecular mechanisms that link increased IKATP to ICa remain elusive. RT-PCR analysis suggests that expression of the L-type Ca2+ channel is not altered. We have not directly determined the Ca2+ channel protein level in TG compared with WT hearts; however, the results with Iso argue against an increase in channel protein because the maximum attainable current in WT myocytes (i.e., the Iso-stimulated current) was not significantly different from the basal current in line 4 TG mice (which was not stimulated by Iso). Because the mRNA levels are certainly not increased and the protein levels are not likely to be increased, the parsimonious conclusion is that the function of the Ca2+ channel is altered. The -adrenergic agonist Iso failed to elicit an increase in either ICa or contractility in TG myocytes, suggesting that ICa is prestimulated. However, the basal and Iso-stimulated cAMP concentrations (and presumably PKA activation) are equivalent in WT and TG hearts, implying that the adrenergic-cAMP-PKA axis of signaling is unaltered in the TG heart. We cannot distinguish at this point between an elevated downstream action (e.g., enhanced PKA activity independent of cAMP) or stimulation through a separate convergent pathway.
It should also be noted that there is no evidence for cardiac hypertrophy, as assessed by the heart weight-to-body weight ratio (22) or myocyte capacitance (data not shown) in TG animals. It has recently been demonstrated that TG mice overexpressing the L-type Ca2+ channel -subunit develop an age-dependent hypertrophy, resulting from a sustained increase in Ca2+ entry and activation of PKC (16). At present, it is unknown what mechanisms protect the cell from hypertrophy despite the increased ICa in our TG mice, but the absence of hypertrophy might suggest that activation of KATP channels could rescue an ICa-dependent hypertrophy.
KATP overactivity versus KATP suppression in the heart and pancreas. A number of recent studies of the cardiac phenotype of Kir6.2–/– mice have recently been reported; however, none have demonstrated evidence of myocardial excitation-contraction coupling remodeling (14, 27, 28, 31). Neither APD nor myocardial contractility differ between WT and knockout animals (27). The principal phenotype of knockout mice is an altered electrophysiological response to metabolic inhibition and ischemia (14, 28) as well as abolition of ischemic preconditioning (28). There is some evidence that KATP channels may play a role in the Ca2+ cycling during periods of enhanced or stressful activity in both skeletal (6) and cardiac muscle (31). This finding, combined with the data presented here, may suggest a central role for KATP channels in maintaining Ca2+ homeostasis.
The striking lack of pathological consequences of cardiac KATP channel gene manipulation remains in stark contrast to the effect of similar perturbations in the pancreas. Whereas the transgene described in the present study does not significantly impair cardiac function, a similar ATP-insensitive Kir6.2 transgene expressed in -cells severely disrupts glucose-induced insulin secretion, causing a lethal diabetic phenotype (11). The reasons for such disparate effects of similar transgenes at present are unclear but may reflect differences in SUR subunit composition of the heteromeric channels.
In summary, we generated TG mice that overexpress a cardiac-restricted, ATP-insensitive mutant of Kir6.2. Contrary to predictions, these mice display enhanced contractile function that results from a remodeling of the molecular substrates of excitation-contraction coupling. A marked increase in peak ICa density likely contributes to counteracting the effects of transgene expression, maintaining the APD and causing increased myocyte contractility. The generality of such excitation-contraction coupling remodeling, as a counterbalance to the negative inotropic effects of increasing K+ currents, remains an open question.
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ACKNOWLEDGMENTS |
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GRANTS
This work was primarily supported by National Institutes of Health Grant HL-45742 (to C. G. Nichols), Cardiovascular Training Grant HL-07275 (fellowship support of T. P. Flagg), Grant HL-69052 (to A. Lopatin), and Grant KO8 DK-60086 (to D. Enkvetchakul).
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FOOTNOTES |
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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.
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. Circulation 103: 140–147, 2001.This article has been cited by other articles:
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where K
slow and 
