1We have investigated the possible role of A-kinase anchoring proteins (AKAPs) in protein kinase A (PKA) signalling to ATP-sensitive K+ (KATP) channels of rat isolated mesenteric arterial smooth muscle cells using whole-cell patch clamp and peptides that inhibit PKA-AKAP binding.
2Intracellular Ht31 peptide (20 μm), which inhibits the PKA-AKAP interaction, blocked KATP current activation by either dibutyryl cAMP or calcitonin gene-related peptide. Ht31-proline (20 μm), which does not inhibit PKA binding to AKAP, did not block KATP current activation.
3Ht31 reduced KATP current activated by pinacidil and also prevented its inhibition by Rp-cAMPS, effects consistent with Ht31 blocking steady-state KATP channel activation by PKA. However, Ht31 did not prevent KATP current activation by the catalytic subunit of PKA.
4An antibody to the RII subunit of PKA showed localization of PKA near to the cell membrane. Our results provide evidence that both steady-state and receptor-driven activation of KATP channels by PKA involve the localization of PKA by an AKAP.
ATP-sensitive K+ channels (KATP channels) of arterial smooth muscle play roles in controlling arterial diameter and so blood flow, and are modulated by vasodilator and vasoconstrictor transmitters (Dart & Standen, 1993; Quayle et al. 1997; Clapp & Tinker, 1998). Several vasodilators act at receptors coupled to the G-protein Gs to cause channel activation via adenylyl cyclase and cyclic AMP-dependent protein kinase (PKA) (Quayle et al. 1994; Kleppisch & Nelson, 1995; Wellman et al. 1998), and we have recently shown that PKA also causes steady-state activation of arterial KATP channels in the absence of receptor agonists (Hayabuchi et al. 2001). In recent years subcellular targeting of PKA through association with A-kinase anchoring proteins (AKAPs) has been shown to underlie the specificity of PKA phosphorylation in a number of cellular processes (Rubin, 1994; Pawson & Scott, 1997; Colledge & Scott, 1999). Each AKAP has a conserved helical region that binds the regulatory (RII) subunits of the PKA holoenzyme at a site distinct from that involved in cAMP binding (Hausken et al. 1996), and a specialized anchoring domain that tethers the PKA-AKAP complex to specific intracellular locations close to its substrate (Colledge & Scott, 1999). Membrane targeting of PKA by AKAPs is involved in PKA-mediated phosphorylation and regulation of several types of ion channel, including glutamate channels, calcium channels, and calcium-activated and inwardly rectifying potassium channels (Rosenmund et al. 1994; Wang & Kotlikoff, 1996; Gao et al. 1997; Westphal et al. 1999; Zhong et al. 1999; Dart & Leyland, 2001). However, a role for AKAPs in signalling to KATP channels has not been reported to our knowledge, despite the fact that PKA plays an important role in regulating these channels, particularly in smooth muscle.
Much of the evidence for the role of AKAPs in signalling in native cells has come from the use of synthetic Ht31 peptides derived from human thyroid anchoring protein. Ht31 peptides contain an amphipathic helix domain and bind to the RII subunit of PKA with nanomolar affinity, thus competing for PKA with native AKAPs and disrupting PKA anchoring within cells (Carr et al. 1992; Vijayaraghavan et al. 1997). In this study, we have used Ht31 peptide and its inactive analogue Ht31-proline (Ht31-P) to investigate the potential role of AKAPs in KATP channel regulation in native arterial smooth muscle cells, and immunocytochemistry to study the localization of PKA. Our findings provide the first evidence for the involvement of an AKAP in the localization of a kinase pathway signalling to KATP channels, and suggest that such an anchoring protein is important for maintaining steady-state channel activation by PKA.
Dissociation of arterial smooth muscle cells
Single smooth muscle cells were isolated enzymatically as we have described previously (Hayabuchi et al. 2001) from small branches of mesenteric artery from male adult Wistar rats rendered unconscious by exposure to a rising concentration of CO2 and killed by exsanguination. Cells were stored at 4 °C, and used on the day of preparation. Cells had membrane capacitances of 15.9 ± 0.4 pF (n= 94).
Solutions and chemicals
The intracellular solution for conventional whole-cell recording of KATP current contained (mm): 110 KCl, 30 KOH, 10 Hepes, 10 EGTA, 1 MgCl2, 1 CaCl2, 1.0 Na2ATP, 0.1 ADP, 0.5 GTP; adjusted to pH 7.2. The free [Ca2+] calculated using the program Maxchelator (http://www.stanford.edu/~cpatton) was 20 nm. The 6 mm K+ extracellular solution contained (mm): 134 NaCl, 6 KCl, 1 MgCl2, 0.1 CaCl2, 10 Hepes, 10 glucose; adjusted to pH 7.4. To separate KATP currents, we recorded at −60 mV to minimize the activation of voltage-dependent K+ channels, and raised [K+]o to 140 mm to give a substantial driving force for K+, as we have described previously (Kubo et al. 1997). The 140 mm K+ extracellular solution contained (mm): 140 KCl, 1 MgCl2, 0.1 CaCl2, 10 Hepes, 10 glucose; pH 7.4. The external solution was changed by continuous perfusion of the experimental chamber (volume 0.4 ml); the dead time for the solution to reach the bath was about 30 s and complete exchange took about 2 min. Pinacidil, angiotensin II, dibutyryl cAMP (db-cAMP), calcitonin gene-related peptide (CGRP) and glibenclamide were from Sigma. Rp-cAMPS (the Rp isomer of adenosine 3′,5′-cyclic monophosphorothioate triethylammonium salt) and the catalytic subunit of PKA were from Calbiochem (UK). Ht31 peptide, sequence DLIEEAASRIVDAVIEQVKAAGAY, and Ht31-P in which the isoleucines at positions 10 and 15 are replaced by prolines, were from Promega and were added to the intracellular solution at 20 μm. Pinacidil and glibenclamide were dissolved in dimethylsulphoxide (DMSO). The final concentration of DMSO was less than 0.2 %.
Freshly dissociated mesenteric smooth muscle cells were plated onto poly-lysine-coated coverslips (0.01 % solution, Sigma) and fixed and permeabilized by immersion in methanol-acetone (1:1) at −20 °C for 10 min. Fixed cells were washed in PBS (Gibco) and incubated in blocking buffer (10 % (v/v) normal goat serum in PBS) for 10 min at room temperature to suppress non-specific antibody binding. Rabbit polyclonal antibodies reactive with the type IIα regulatory subunit (RIIα) of PKA (Santa Cruz Biotechnology, Inc.) were diluted 200-fold in blocking buffer and incubated with the cells for 2 h at room temperature followed by overnight incubation at 4 °C. As a control, a 5-fold excess of PKA RIIα blocking peptide (Santa Cruz Biotechnology, Inc.) was incubated with the primary antibody for 2 h at room temperature before addition to some of the coverslips. Following overnight incubation, coverslips were washed for 5 × 10 min in PBS, then incubated in blocking buffer containing fluorescein (FITC)-conjugated goat anti-rabbit secondary antibody (200-fold dilution; Jackson ImmunoResearch Laboratories, Inc.) for 2 h at room temperature. Coverslips were washed in PBS and mounted onto microscope slides using fluorescent mounting media (Dako Ltd). Confocal images were obtained using a Perkin-Elmer UltraView imaging system with a ×60, NA 1.4 objective lens. Image analysis was carried out using UltraView software.
Data recording and analysis
Whole-cell K+ currents were recorded from single smooth muscle cells in the conventional configuration of the patch clamp technique. Membrane currents were recorded, and voltage was controlled, using an Axon interface and Axopatch 200 amplifier (Axon Instruments). Patch pipettes were made from thin-walled borosilicate glass (Clark Electromedical, Pangbourne, Berks, UK) using a pp-83 puller (Narishige, Tokyo, Japan) and coated with sticky wax (Kemdent, Swindon, UK) to reduce capacitance. Currents were filtered at 5 kHz. Electrode resistance before sealing was 3–5 MΩ, and seals were > 10 GΩ. Experiments were done at 20–25 °C. Results are expressed as means ±s.e.m. Intergroup differences were analysed using ANOVA followed by the Student-Newman-Keuls test and P < 0.05 was considered statistically significant.
Disruption of PKA anchoring with Ht31 peptide prevents KATP channel activation by db-cAMP or CGRP
Arterial KATP currents are activated by cAMP, applied either in the intracellular solution or in membrane-permeant form (Quayle et al. 1994; Kleppisch & Nelson, 1995; Wellman et al. 1998; Hayabuchi et al. 2001). To see whether such activation requires PKA localization by an AKAP, we investigated the effects of intracellular Ht31 peptide on the activation of KATP current by the membrane-permeant cAMP analogue db-cAMP. Ht31 was applied to the inside of the cell by dialysis from the pipette solution. Whole-cell current was recorded at −60 mV and [K+]o was increased to 140 mm so that K+ currents were inward at this potential. The change in [K+]o from 6 to 140 mm is indicated by the arrows in Fig. 1A and subsequent figures, and caused a small increase in inward current. Figure 1A and B shows that in control cells (without intracellular peptide) db-cAMP (0.5 mm) activated a substantial KATP current (−83.0 ± 17.9 pA, n= 8), which was blocked by the sulphonylurea KATP channel inhibitor glibenclamide (10 μm). However in cells dialysed with Ht31 (20 μm), KATP current activation by db-cAMP was greatly reduced (to −11.7 ± 3.9 pA, n= 10, P < 0.005vs. control). To test whether this effect depended on the ability of Ht31 to disrupt PKA anchoring to an AKAP, we used the inactive analogue peptide of Ht31, Ht31-P. The introduction of proline residues within the RII-binding helical region of Ht31 abolishes its ability to interact with RII and therefore prevents it from disrupting PKA binding to native AKAPs. Ht31-P (20 μm) did not significantly affect activation of KATP current by db-cAMP (n.s. vs. control, Fig. 1A and B).
We also investigated whether Ht31 peptide could inhibit KATP channel activation by a physiological agonist that acts at a Gs-coupled receptor, using the potent vasodilator CGRP, which activates arterial KATP channels through the adenylyl cyclase-PKA pathway (Wellman et al. 1998). Our results are shown in Fig. 1C and D. Like db-cAMP, CGRP (10 nm) activated a substantial KATP current, and this activation was greatly reduced in cells dialysed with Ht31 (P < 0.005vs. control), but was unaffected by dialysis with Ht31-P (n.s. vs. control, Fig. 1C and D). These results suggest that PKA-induced KATP channel activation triggered either by application of a cAMP analogue or by activation of CGRP receptors requires binding and localization of PKA by an AKAP.
Ht31 peptide blocks steady-state KATP channel activation by PKA
Arterial KATP channels are subject to steady-state activation by PKA, even in the absence of stimulation of receptors that act through adenylyl cyclase (Hayabuchi et al. 2001). If such activation involves the anchoring of PKA close to the channel by an AKAP, steady-state activity should be reduced by disruption of PKA anchoring by Ht31. Because KATP currents of arterial smooth muscle are small, corresponding to a low density of KATP channels (Quayle et al. 1997), and because of the variability between cells, it is difficult to resolve significant differences in steady-state current. However, in the presence of the KATP channel opener pinacidil, KATP currents are increased in size and a component that depends on steady-state activation by PKA is revealed (Hayabuchi et al. 2001). We therefore measured KATP current in the presence of 10 μm pinacidil. Figure 2B shows that this current was smaller in cells dialysed with Ht31 (−114.1 ± 12.3 pA, n= 8) than in control cells (−228.5 ± 36.5 pA, n= 8), while dialysis with Ht31-P did not significantly reduce KATP currents compared to control.
Steady-state KATP channel activation by PKA can also be demonstrated by a reduction in the pinacidil-activated current on addition of a PKA inhibitor (Hayabuchi et al. 2001). We therefore investigated the ability of Rp-cAMPS, which inhibits PKA by binding to its regulatory subunit (Rothermel & Parker-Botelho, 1988), to reduce pinacidil-activated current in cells with or without disruption of PKA anchoring. Figure 2A and C shows that Rp-cAMPS reduced KATP current in cells dialysed with control solution (or with Ht31-P), but was much less effective in cells dialysed with Ht31, as would be expected if steady-state channel activation by PKA was already greatly reduced in those cells. These results suggest that steady-state PKA activation of KATP requires PKA localization by an AKAP.
The catalytic subunit of PKA activates KATP current in the presence of Ht31
The results of the experiments described above show that Ht31 blocks the stimulation of KATP current in response to the activation of PKA, presumably because the localization of PKA close to the channel is disrupted by this peptide. The catalytic subunit of PKA, applied to the inside of cells, has been shown to activate channels of both rabbit mesenteric and pig coronary arterial smooth muscle (Quayle et al. 1994; Wellman et al. 1998). Intracellular PKA catalytic subunit should be able to interact directly with the KATP channel, so its action should not depend on the localization of endogenous PKA and therefore should not be affected by Ht31 peptide. To test this, we examined the effects of intracellular application of purified PKA catalytic subunit in the absence and presence of Ht31. Figure 3A (upper trace) shows that dialysis of control intracellular solution without PKA and subsequent application of glibenclamide revealed a small basal KATP current that averaged −18.8 ± 3.7 pA (n= 10, Fig. 3B). Inclusion of PKA catalytic subunit (100 units ml−1) in the intracellular solution led to the development of an approximately 6-fold greater current (Fig. 3A, middle trace). Ht31 did not prevent KATP activation by PKA catalytic subunit; when both this subunit and Ht31 (20 μm) were included in the pipette solution, KATP current activation by PKA was unaffected (n.s. compared to PKA alone, Fig. 3).
Membrane localization of PKA
The results of the functional studies outlined above suggest that an AKAP facilitates PKA-induced modulation of KATP channel activity, presumably by anchoring a pool of PKA close to the channel. This suggests that PKA should be targeted to the plasma membrane in these cells. To test this, we incubated permeabilized mesenteric smooth muscle cells with a polyclonal antibody directed against the RIIα subunit of PKA, and visualized the location of the antibody within the cells through addition of FITC-conjugated secondary antibodies. Figure 4A and B shows confocal images of antibody-treated mesenteric smooth muscle cells, which revealed that PKA RIIα is indeed predominantly located at or near the plasma membrane. No staining was observed in cells exposed to anti-PKA RIIα that had been pre-incubated with blocking peptide (see Methods; not shown). The extent of PKA localization in these cells is clearly illustrated by looking at the intensity of fluorescence along a line drawn across the cell at a point away from the nuclear region (Fig. 4C and D). These plots show clear peaks in the fluorescence intensity corresponding to the bright PKA-rich regions near the plasma membrane.
In this paper we have shown that Ht31 peptide blocks the activation of arterial KATP channels by db-cAMP or CGRP. The inactive analogue peptide Ht31-P was ineffective, confirming that the effect of Ht31 occurs through disruption of PKA anchoring by PKA-AKAP binding. In addition, the application of exogenous PKA catalytic subunit still activated KATP current in the presence of Ht31, ruling out any non-specific effect of the peptide in preventing channel activation by PKA. Using anti-PKA RIIα antibodies we have also shown that the regulatory subunit of the kinase is located predominantly near the plasma membrane, presumably in close physical proximity to the ion channel it regulates. The lack of availability of reliable antibodies to putative smooth muscle KATP channel subunits prevented us from investigating their co-localization with RII, but such studies should become possible in the future. Together, our results provide strong evidence that the activation of KATP channels by PKA involves localization of the kinase by an AKAP. Ht31 also blocked the steady-state PKA activation of arterial KATP channels that occurs in the absence of receptor stimulation. Steady-state activation may be important for the contribution of these channels to vasodilatation and so to blood flow that has been demonstrated in several vascular beds (Samaha et al. 1992; Quayle et al. 1997; Goto et al. 2000; Duncker et al. 2001). Our present findings suggest that AKAP localization of PKA close to the KATP channel is essential for the tonic drive exerted by PKA on the channel.
AKAPs form a family of around 70 functionally related proteins (Dodge & Scott, 2000). The prototypic AKAP, AKAP79, has been shown to bind both protein kinase C (PKC) and protein phosphatase 2B (calcineurin) in addition to PKA (Klauck et al. 1996), thus AKAPs can assemble multi-unit enzyme complexes that co-ordinate the phosphorylation state of cellular substrates. These AKAP complexes may themselves be part of larger signalling complexes that can involve both upstream activators and downstream targets (Dodge & Scott, 2000). Arterial KATP channels can be modulated by both PKC and calcineurin in addition to PKA (Bonev & Nelson, 1996; Kubo et al. 1997; Wilson et al. 2000). Thus it is an attractive hypothesis that the functions of these enzymes in regulating arterial KATP channels are integrated by AKAP anchoring into a signalling complex. The nature of the AKAP involved remains to be determined, as do possible scaffolding links between receptors for vasoactive agonists, AKAP and the KATP channel.
We thank Diane Everitt for skilled technical assistance, and the Wellcome Trust, Royal Society and British Heart Foundation for support.