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1Department of Pharmacology, Faculty of Medicine, National University of Singapore, 117597 Singapore; and 2Departments of Medicine and Molecular Pharmacology, Albert Einstein College of Medicine, Bronx, New York 10461
Submitted 5 February 2004 ; accepted in final form 28 June 2004
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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1A-adrenergic receptors produced a reduction in current amplitude of the rapidly activating delayed rectifier K+ current (the current carried by ERG protein) from rabbit ventricular myocytes. The -adrenergic-induced current reduction was accentuated by PKC blockers and also unmasked a depolarizing shift in the voltage dependence of activation, which supports the conclusion that receptor activation of PLC results in PIP2 consumption that alters channel activity. These results support a physiological role for PIP2 regulation of the rapidly activating delayed rectifier K+ current during autonomic stimulation and localize a site of interaction to the COOH-terminal tail of the HERG K+ channel.
human ether-à-go-go-related gene; phosphatidylinositol 4,5-bisphosphate; delayed rectifier K+ current; channel; phospholipids; G protein-coupled receptor; mutagenesis; phospholipase C
-adrenergic receptor pathways (6, 7, 16, 30). In this study, we sought to investigate the -adrenergic affects on HERG and IKr to better understand the complex control of this current during physiological stresses. A signaling pathway that is set in motion upon -adrenergic stimulation involved G protein-mediated activation of PLC. Activated PLC generates diacylglycerol (DAG) and 1,4,5-inositol trisphosphate (IP3) as by-products from the consumption of the endogenous phospholipid phosphatidylinositol 4,5-bisphosphate (PIP2). Changes in PIP2 concentration ([PIP2]) have been shown to regulate a variety of K+ channels (3, 9, 14, 17, 21, 26, 33, 34). Heterologously expressed HERG was the first voltage-gated channel shown to be responsive to changes in [PIP2] (4). The other LQT-linked delayed rectifier K+ current, IKs, which is composed of the KCNQ1 channel (with the KCNE1 subunit), has also been shown to be regulated in cardiac myocytes and in a heterologous expression system (18). We now show that native IKr in cardiac myocytes is responsive to PIP2 changes. Moreover, we have employed mutagenesis to map a site within the channel protein that binds to PIP2 and is responsible for much of the PIP2 responsiveness of HERG and IKr. |
MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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1,500-bp fragment was purified and ligated into the FseI-XhoI sites within HERG in HERG-myc-pCI-Neo. The final plasmid encoded a myc-tagged mutant HERG protein (for both D- and M-HERG).
Cell culture and transfection.
Chinese hamster ovary (CHO) and human embryonic kidney (HEK)-293 cell lines (from American Type Culture Collection) were cultured in Ham's F-12 or RPMI 1640, respectively, and supplemented with L-glutamine, 10% fetal calf serum (Hyclone), and penicillin-streptomycin (GIBCO). Cultured cells were maintained in 5% CO2-95% humidified air at 37°C. We transiently cotransfected 5 µg of WT-HERG, D-HERG, or M-HERG cDNA to CHO cells together with 2 µg of GFP cDNA via electroporation. Cells were washed with cytomix buffer (in mM: 120 KCl, 0.15 CaCl2, 10 K2HPO4-KH2PO4, 25 HEPES, 2 EGTA, and 5 MgCl2, pH 7.6) and centrifuged at 300 g for 5 min. Cell suspension (400 µl) in complete cytomix buffer that contained 2 mM ATP and 5 mM glutathione was electroporated in a 2-mm gap cuvette using a BTX ECM600 electroporation system with the following settings: capacitance, 180 µF; resistance, 72 
; and voltage, 225 V. After electroporation, the cells were plated sparsely and grown on sterile glass coverslips in 100-mm tissue culture dishes. Cells were used for electrophysiological studies 24–72 h after electroporation. For biochemical analyses, transient transfections were performed with Lipofectamine 2000 (Invitrogen) as previously described (6, 16).
Isolation of rabbit cardiac myocytes. Male rabbits (1–2 kg) were anesthetized with an injection of 55 mg/kg im ketamine and 7 mg/kg im xylazine. Heparin (1,000 IU iv) was administered to prevent coagulation during heart removal. The heart was quickly excised, mounted on a Langendorff apparatus, and perfused in a retrograde fashion via the aorta with nominally Ca2+-free Tyrode solution at 37°C. The Tyrode solution contained (in mM) 137 NaCl, 5.4 KCl, 1 MgCl2, 10 HEPES, and 10 glucose, pH 7.4. After 5 min, the perfusion solution was changed to the Tyrode solution that contained 1 mg/ml type I collagenase (Sigma) and 0.28 mg/ml type XIV protease (Sigma) and was perfused for 25–30 min. The perfusion solution was then changed to Ca2+-Tyrode solution that contained 0.2 mM CaCl2 without proteases for 5 min. The ventricular tissue was then cut into small pieces in a petri dish with 20 ml of prewarmed Ca2+-Tyrode solution and was shaken gently inside the solution for the dispersion of dissociated cardiac myocytes. A 250-µM mesh screen was used to separate the isolated cardiac myocytes from cardiac tissue. The cells were washed three times in Ca2+-Tyrode solution and collected by centrifugation at 100 rpm for 1 min. Isolated myocytes were resuspended in Ca2+-Tyrode solution and plated onto laminin (Sigma)-coated glass coverslips. Cells were allowed to attach for 2 h and then were used for electrophysiology studies.
Patch-clamp recordings.
Cells on coverslips were taken directly from the cell culture incubator and placed in an acrylic-polystyrene perfusion chamber (Warner Instruments) for electrophysiological measurements. For the rabbit cardiac myocytes, rod-shaped myocytes with clear striations were selected for electrical recordings. Patch pipettes were pulled and polished to obtain a tip resistance of 2–3 M in the patch-clamp solutions. Pipette offset potential in our study solutions was compensated to zero just before gigaseal formation. The junction potential in the experimental solutions was estimated to be 3–4 mV (by pCLAMP 9 analysis software) and was not corrected for analysis. Whole cell capacitance was compensated electronically through the amplifier. The series resistance in the whole cell configuration was between 9 and 10 M. All experiments were carried out at room temperature (20–22°C). Cells were studied on an inverted microscope equipped with electronic patch-pipette micromanipulators and epifluorescence optics for GFP (transfected cells). Axopatch 200B patch-clamp amplifiers (Axon Instruments) were used for voltage-clamp measurements. Voltage-clamp protocols were controlled via personal computer using pCLAMP 9 acquisition and analysis software. To elicit HERG K+ currents, depolarizing voltage pulses were applied at various levels from a holding potential of –70 mV for 4.5 s and were followed by stepwise repolarization to –40 mV and then to –120 mV to measure outward and inward tail currents, respectively. Signals were analog filtered at 2,000 Hz and sampled at 5–10,000 Hz. Voltage-dependent activation data were fitted to the Boltzmann relation I = 1/{1 + exp[(Vh – V)/k]}, where I is the relative tail current amplitude, V is the applied membrane voltage, Vh is the voltage at half-maximal activation, and k is the slope factor (22). To compare the effects before and after the administration of reagents, current amplitude was normalized to the control group before application of drugs.
For whole cell voltage-clamp measurements, the pipette solution consisted of (in mmol/l) 126 KCl, 2 MgSO4, 0.5 CaCl2, 5 EGTA, 4 Mg-ATP, and 25 HEPES (pH 7.2; osmolality, 280 ± 10 mosmol/kg). The external bath solution consisted of (in mM) 150 NaCl, 1.8 CaCl2, 4 KCl, 1 MgCl2, 5 glucose, and 10 HEPES (pH 7.4; osmolality, 320 ± 10 mosmol/kg). To increase the cellular [PIP2] directly, 10 µM PIP2 was included in the pipette solution. For IKr recordings in the cardiac myocytes, the extracellular solution contained (in mM) 150 N-methyl-D-glucamine (NMDG), 1.8 CaCl2, 4 KCl, 1 MgCl2, 5 glucose, and 10 HEPES (pH 7.4; osmolality, 320 ± 10 mosmol/kg) at room temperature (1). To buffer the increased intracellular Ca2+ concentration ([Ca2+]i) upon 1A-receptor stimulation, 15 mM BAPTA was added to the pipette solution (with appropriate pH adjustment).
In vitro PIP2 binding assay. HEK-293 cells were transfected with WT-HERG-myc, D-HERG-myc, M-HERG-myc, or Kir2.1-myc. At 48 h after transfection, cells were washed with ice-cold PBS and lysed in ice-cold buffer that contained 150 mM NaCl, 25 mM Tris·HCl, pH 7.5, 5 mM EDTA, 1% Nonidet P-40, 0.4% deoxycholic acid, and EDTA-free protease inhibitor cocktail tablets (Roche Pharmaceuticals). Lysates were cleared by centrifugation at 13,200 rpm for 10 min at 4°C. The supernatant was incubated with 20 µl of rabbit polyclonal A14 anti-myc antibody (Santa Cruz) and 60 µl of protein G-agarose (Pierce) for 4 h at 4°C. Precipitated proteins were washed with 0.25% Nonidet P-40 buffer that contained 150 mM NaCl, 25 mM Tris·HCl, pH 7.5, and 0.25% Nonidet P-40 and were then washed with PBS. A 1:1 mixture of [3H]PIP2 in chloroform-methanol-buffered saline (sp. act., 8 µCi/nmol; American Radiolabeled Chemicals) was dried by vacuum at 4°C and sonicated in 100 µl of PBS to form pure [3H]PIP2 liposomes. The [3H]PIP2 liposomes were incubated with the immobilized proteins for 2 h at 4°C with or without varying concentrations of unlabeled PIP2. After precipitates were washed twice with PBS, 10% were dissolved in SDS gel-loading buffer, and 90% of the proteins were counted in a scintillation counter (Beckman) using a window for 3H. The SDS-PAGE was probed with mouse monoclonal 9E10 anti-myc antibody and subjected to immunoblot analysis. Specific binding was obtained by subtracting the background binding value obtained from protein G antibody applied to samples from nontransfected HEK-293 cells.
To determine PIP2 affinity, different concentrations of unlabeled PIP2 (10–7 to 10–3 M) were used to compete with 0.5 µM [3H]PIP2 for binding. The competition data were analyzed using the equation (binding of radiolabeled ligand) = Bmax([3H]PIP2)/([3H]PIP2) + [(unlabeled PIP2) + Kd].
Materials. PIP2, bisindolymaleimide I (BSM), and chelerythrine were purchased from Calbiochem. Collagenase type I, protease type XIV, laminin, and BAPTA were from Sigma. Chromanol 293B was obtained from Tocris. For preparation of PIP2, the lipid was first dispersed in water (at 0.5 mM concentration) by sonication for 30 min on ice and then divided into aliquots and kept at –80°C. For each experiment, a new aliquot was thawed and used only once. PIP2 was diluted to 10 µM in the electrophysiology pipette solution and sonicated again for 10–30 min. This procedure results in the formation of a suspension of mostly small micelles of PIP2 that can be readily absorbed by lipid membranes (10). Chelerythrine and BSM were first dissolved in DMSO as stock solutions and then used at the desired final concentrations in the bath solution such that the final DMSO concentration was <0.5%.
Statistics. Values presented are means ± SE. Student's t-test was used to compare the differences between two groups. Significance level was set at P < 0.05.
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RESULTS |
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PIP2-dependent regulation of HERG inactivation and activation. We previously showed that the current augmentation in response to PIP2 elevation was in part due to deceleration of inactivation kinetics of HERG channels (4). Voltage-dependent inactivation accounts for the negative slope conductance in the isochronal I–V curve of HERG, which is the only situation where PIP2 appeared to alter D-HERG activity. Accordingly, we investigated whether PIP2 altered the inactivation kinetics of D-HERG. We used voltage-clamp protocols described by Smith et al. (27) to specifically measure voltage dependence of steady-state inactivation. After a prolonged depolarizing pulse to +20 mV to fully activate the channels, we briefly stepped the membrane voltage to various test potentials and then returned to +20 mV (Fig. 4, A–C). During the brief step, the inactivation process rapidly reversed to the steady-state level appropriate to the test potential. The outward tail current observed on returning to +20 mV gives the relative fraction of open channels, which is determined by the degree of steady-state inactivation remaining at the test potential (27). The fraction of channels that deactivate during the brief hyperpolarizing steps was corrected (Fig. 4, A–C, inset) using the procedure described by Smith et al. (27). Our results show that increased [PIP2] significantly enhanced the current amplitude upon release of inactivation in both WT- and D-HERG (Fig. 4, A–C). By normalizing the curves, we see that there is a small but not significant leftward shift. During an inactivation protocol, the time course for the decline in outward current is a measure of the rate of inactivation and is best fit to a single exponential function (Fig. 4, D–F). Increasing the [PIP2] caused a comparable and significant slowing of voltage-dependent inactivation in WT- and D-HERG (Fig. 4, D–F). This suggests that regions of the channel other than the COOH-terminal site (amino acids 883–894) may be affected by changes in PIP2 that result in altered inactivation.
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0.48 µM.
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1A-adrenergic receptor stimulation on IKr in ventricular myocytes.
To investigate the IKr properties of rabbit heart, we performed whole cell patch-clamp recordings of isolated rabbit ventricular myocytes. In the whole cell configuration, cells were depolarized from a holding potential of –70 mV to voltages ranging from –40 to 50 mV for 4.5 s. Upon repolarization to –40 mV, an outward current characteristic of IKr was observed (Fig. 7A). Application of the specific IKr blocker E-4031 (1 µM) to the external solution abolished this tail and thereby confirmed its identity as IKr (Fig. 7B). Moreover, 1 µM E-4031 produced a significant prolongation in the action potential duration at 90% repolarization from 501.4 ± 31.2 to 593.0 ± 27.8 ms in cardiac myocytes (Fig. 7C). This is consistent with the previous findings of cardiac IKr (5, 15, 20). To exclude contamination of IKr by coexisting IKs, we also measured IKr in the presence of the IKs inhibitor Chromanol 293B. Figure 7D shows that administration of 10 µM Chromanol 293B had no significant effect on IKr, which indicates the lack of contribution of IKs to the tail currents.
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1A-adrenergic agonist phenylephrine (PE) was applied 3 min after the whole cell configuration was established. Application of 10 µM PE resulted in a consistent decrease in the current amplitude (Fig. 8B). The same result was also found in cells pretreated with 10 µM Chromanol 293B (data not shown). To test whether the effect of PE is mediated by 1A-receptor activation, the selective 1A-receptor antagonist 5-methylurapidil was used. Pretreatment with 1 µM 5-methylurapidil for 1 h abolished the effect of PE (Fig. 8C), which suggests a receptor-mediated effect. Internal application of PIP2 (10 µM) abolished the inhibitory effect of PE on IKr (Fig. 8D), which suggests that PIP2 may be involved in this effect.
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-adrenergic stimulation, we pretreated cells with PKC inhibitors. Pretreatment of cells with the PKC inhibitor chelerythrine (1 µM) for 1 h before patch clamping enhanced the effects of -adrenergic stimulation and resulted in an even greater PE-induced inhibition of IKr amplitude (Fig. 8E). When PKC was inhibited, subsequent PE treatment produced a reduction in current density and a depolarizing shift in voltage dependence of activation (Vh: before PE treatment, 0.03 ± 0.49; after treatment, 6.51 ± 0.96 mV). Comparable results occurred using another PKC inhibitor, 100 nM BSM (data not shown). These data suggested that PIP2 consumption and PKC activation may alter cardiac IKr in competing directions during -adrenergic stimulation and that the PIP2 effect is not due to activation of PKC.
The other second messenger generated by PLC activation is IP3, which upon binding to its receptor releases Ca2+ from the endoplasmic reticulum. Alterations of intracellular Ca2+ may also alter channel behavior. To examine the possibility that -adrenergic-mediated regulation of IKr is due to IP3-mediated [Ca2+]i elevation, the Ca2+ chelator BAPTA (15 mM) was included in the whole cell pipette solution to increase the intracellular buffering capacity. In cells pretreated with chelerythrine and BAPTA in the pipette, PE still produced a comparable reduction in current amplitude (Fig. 8F) and a small right shift of the voltage dependence of activation (Vh: before PE treatment, 0.83 ± 1.05; after treatment, 4.47 ± 0.35 mV). These data support the hypothesis that stimulation of cardiac GPCRs that activate PLC can alter endogenous [PIP2] sufficiently to modify the behavior of IKr. Moreover, inclusion of a neutralizing anti-PIP2 monoclonal antibody in the whole cell pipette solution produced a significant reduction of HERG K+ current density (Fig. 8G). When the data were normalized to unity, a 9–10-mV depolarizing shift in voltage-dependent activation was seen (Vh: control, 5.1 ± 1.6; anti-PIP2 monoclonal antibody, 15 ± 2.2 mV; n = 7).
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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q subunits from G
subunits. Activated Gq protein in turn stimulates the activity of phosphatidylinositol-specific PLC. The substrate for PLC is PIP2, a membrane phospholipid that is hydrolyzed to IP3 and DAG. Thus as these GPCRs are activated, there is an increase in the concentrations of second messengers IP3 and DAG with a simultaneous reduction of [PIP2] within the plasma membrane. We previously showed that activation of 1A-receptors in cells that heterologously express the receptor and HERG with the selective agonist PE altered HERG channel activity in a fashion consistent with reduction of PIP2. These effects were not prevented by internal Ca2+ buffering or PKC inhibition, which is in support of a mechanism whereby GPCR stimulation of PLC results in the consumption of endogenous PIP2 (4). In the present study, we show that stimulation of cardiac -adrenergic receptors produced a response in cardiac IKr similar to that seen with heterologously coexpressed HERG and 1A-adrenergic receptors. One particular difference was noted, however. In cardiac myocytes, the -adrenergic-mediated effects on IKr were accentuated by prior block of PKC, which suggests that PKC activation in this situation has an opposing regulation of the channel. The effect of PKC on HERG and IKr, however, remains controversial. Stimulation of the Gq-coupled thyrotropin-releasing hormone receptor, when coexpressed with HERG in Xenopus oocytes, produced kinetic changes that reduced current density; this effect was mimicked by phorbol ester and prevented by the PKC inhibitor FG109203X (2). These findings were also seen in pituitary GH3 cells (11, 24). HERG expressed in Xenopus oocytes was also suppressed by PKC activation via a mechanism that may not involve direct phosphorylation of the channel itself (31). In contrast with these findings, PKC activation in guinea pig cardiac myocytes enhanced IKr through a reduction in C-type inactivation (12). These conflicting effects of PKC on HERG and IKr may result from different signaling systems in different tissues including different differential expression of PKC isoforms or anchoring proteins.
A change in [Ca2+]i may also alter HERG and IKr function. Heath and Terrar (13) found that that the use of Ca2+ buffers reduced -adrenergic regulation of IKr. In the present study, we found that any changes in [Ca2+]i caused by stimulation of -adrenergic receptors were unlikely to play a role in IKr regulation given that 15 mM BAPTA did not alter the PE-induced effects on channel behavior.
HERG is the first voltage-gated channel shown to be affected by changes in [PIP2]; however, the site(s) of interaction with PIP2 has not been determined. More progress has been made in characterizing PIP2 interactions with IKr channels. Clustering of positively charged side-chain amino acids along NH2- or COOH-terminal cytoplasmic channel portions has been proposed as a potential PIP2 interaction site (9). The mechanism of interaction may be an electrostatic attraction between these side chains and the negatively charged polar heads of the phospholipid. We found that the polycationic region from amino acid 883 to 894 is responsible for much of the PIP2 regulation of HERG activity. Despite the elimination of most PIP2 binding by this mutation, there remained some effects on voltage-dependent inactivation. These data suggest that another, possibly lower-affinity binding site for PIP2 may exist within the HERG protein. Multiple PIP2 binding sites in Kir2.1 channels have been proposed in several COOH-terminal loci (25, 28). In the HERG K+ channel, another polycationic segment (672: RYHTQMLRVREFIRFHQIPNPLRQRL697) exists in the cytoplasmic COOH terminus; this may represent an additional site for PIP2 interaction that affects inactivation.
In the IKr channels, the constitutively active Kir2.1 and Kir1.1 interact with PIP2 with high affinity, whereas the G protein-activated Kir3.1/3.4 channels show weaker interactions with PIP2. When exogenous PIP2 is applied, the channels are dynamically activated in the reverse order as follows: Kir3.1/3.4 > Kir2.1 > Kir1.1 (14, 21, 34). It is hypothesized that weaker binding to Kir3.1/3.4 channels allows for appreciable off-rates at physiological concentrations of PIP2, whereas the higher affinity of Kir2.1 (Kd, 0.5 µM; Ref. 14)and Kir1.1 (Kd, 0.58 µM; Ref. 17) allows PIP2 to be constitutively bound (21). Weaker PIP2 binding therefore may produce a situation where physiological consumption by PLC may more readily regulate the channel. Our study showed that the affinity of HERG for PIP2 is comparable to Kir2.1 or Kir1.1, which suggests that the in vivo regulation of HERG by PIP2 may be more similar to these channels. A small reduction or change in the voltage dependence of IKr by changes in PIP2 may have significant impact on repolarization in vivo particularly during periods of extreme stress and autonomic stimuli.
Taken together, our present study demonstrates that alteration of endogenous [PIP2] of cardiac myocytes regulates IKr. The polycationic region (amino acids 883–894) in the COOH terminus of HERG is a physical and functional binding site for PIP2. These findings provide an additional link between cardiovascular stresses, autonomic stimulation, and arrhythmias (both hereditary and acquired). This novel response supports a model where autonomic stimulation of cardiac G protein-coupled receptors may alter IKr-dependent repolarizing forces that may impact propensity toward ventricular arrhythmias.
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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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) and 3 min after whole cell configuration and internal perfusion of 10 µM PIP2 (
). A and B: representative current tracings in response to a series of membrane depolarizations in a Chinese hamster ovary (CHO) cell expressing HERG. Currents were elicited by 4.5-s depolarizing steps to various levels (between 50 and –40 mV) followed by repolarizing steps to –40 mV then to –120 mV (B, bottom). C: current-voltage (I–V) relation curve plotted from current measured at the end of depolarizing test pulses (relative current amplitude, I/Imax; during depolarizing steps plotted against the step potential, Imax is the maximal current before PIP2 perfusion). D: voltage-dependent activation curves plotted from peak tail currents during a repolarizing step to –40 mV after depolarization to various voltages. Data were normalized to maximal current (top inset). Time course of the effect of PIP2 on current amplitude at 0 mV (bottom inset) is shown. Values are shown as means ± SE; n = 8 cells; *P < 0.05 vs. data in same voltage at 0 min. E: alignment of WT-HERG shows a clustering of positively charged amino acids (arginines and lysines) in the COOH terminus.
, time constant of onset of inactivation. E and F: summary of voltage dependence of rates of inactivation shows that PIP2 slows the rate of inactivation in a voltage-dependent fashion in either WT-HERG (E) or D-HERG (F). Symbols represent means ± SE; n = 8–13 cells; *P < 0.05.
