Opposite Regulation of Slick and Slack K⁺ Channels by Neuromodulators

Slo2.1 and Slo2.2 channels have similar primary sequences

Four high-conductance potassium channels of the Slo family are encoded in the mammalian genomes. All four genes are located on different chromosomes, suggesting an ancient origin in evolution. These channels share a common feature with the extended gene family of voltage-dependent potassium channels in having six homologous membrane-spanning domains per α subunit (S1 through S6), which surround the permeation pathway of the channel. (Note that Slo1 and Slo3 have an additional membrane-spanning domain, S0, as described below.) However, Slo family channels differ in having a much larger carboxyl extension, which is thought to be entirely cytoplasmic and involved with modulation of channel function by cytoplasmic factors (Schreiber and Salkoff, 1997; Vergara et al., 1998; Moss and Magleby, 2001; Bao et al., 2002; Shi et al., 2002; Tang et al., 2004; Zeng et al., 2005). All four of the encoded peptides are similar in length. The four known genes fall into two subgroups defined by the closely related paralogs, Slo2.1/Slo2.2 and Slo1/Slo3. These two groupings are evident in the hydrophobicity plots shown in Figure 1 in which Slo2.1/Slo2.2 clearly show greater similarity than Slo1/Slo3.

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Figure 1.

A, Hydrophobicity plot of Slo family channels showing that the four Slo family channels fit into two distinct subgroups. Slo2.1 and Slo2.2 lack the extra membrane spanning segment (S0) found in Slo1/Slo3. In both grouped pairs, the highest similarity (black shading) is in the S1–S6 membrane-spanning domains and in the immediately adjacent downstream region. Hydrophobicity analysis used the Kyte–Doolittle program (Kyte and Doolittle, 1982). Arrow indicates site of exchange in the construction of the C1 chimera (see Results). B, Alignment of residues in the S4 regions of the Slo2 channel from C. elegans, Slo2.2 (Slack) from humans and rodents, Slo1 from humans and rodents, and the voltage-sensitive Shaker channel from Drosophila. A string of positively charged amino acids occurring every three residues is found in the S4 region of voltage-dependent channels. Such charges at the appropriate interval are shaded in red and indicated in the voltage (and calcium) dependent Slo1 channel and in the voltage-dependent Shaker (Kv1) channel. Such a string of positive charges at the appropriate intervals is absent in Slo2 channels, and the overall region is six residues shorter than in voltage-dependent channels.

Two main structural features distinguish Slo2.1/Slo2.2 from Slo1/Slo3. Slo2.1/Slo2.2 channels lack the typical arrangement of positively charged residues in S4 associated with voltage-dependent gating, which is found in all voltage-gated potassium channels, including Slo1/Slo3 (Fig. 1B). Indeed, the S4 region in Slo2 channels appears to have six fewer residues overall compared with S4 in voltage-sensitive channels and, thus, may have a radically different orientation in the channel protein. Also, Slo2.1/Slo2.2 lack the extra membrane spanning domain (S0) near the N-terminal of Slo1/Slo3 that results in placing the N-terminal outside of the cell (Meera et al., 1997; Schreiber et al., 1998). Slo2.1/Slo2.2 paralogs have high amino acid sequence similarity (74% identity in humans, which contrasts with the lower Slo1 and Slo3 identity of ∼43%), with highest similarity in the membrane-spanning domains (Fig. 1). Slo2.1/Slo2.2 have highly conserved mammalian orthologs. Rat and human Slo2.1 channels have >95% identity, and rat and human Slo2.2 channels have >91% identity (Bhattacharjee et al., 2003; Yuan et al., 2003). Interestingly, the main sequence divergence between Slo2.1/Slo2.2 channels resides in the C terminus (Fig. 1), which we will show to be important for their modulation by PKC.

Slo 2.1 and Slo2.2 share basic functional properties

Expression of Slo2.1 and Slo2.2 channels was tested in Xenopus oocytes using two-electrode voltage clamp, 3–5 d after cRNA injection (Fig. 2). Both channels produced noninactivating outward K⁺ currents, with tails reversing at approximately −70 mV. Currents were blocked by extracellular tetraethylammonium in a concentration range between 70 and 150 mm. Typically, Slo2.1 current amplitudes were smaller than those of Slo2.2 (Fig. 2A). Slo2.1 channels were also more difficult to express than Slo2.2 channels, being observed in only 25% of injected oocytes, thus making single-channel recordings difficult to obtain. Consistent with a previous report (Bhattacharjee et al., 2003), Slo2.1 currents showed faster kinetics of activation than those of Slo2.2 (Fig. 2A). Also consistent with previous reports (Bhattacharjee et al., 2003; Yuan et al., 2003), both channels showed Na dependence and were activated by the application of 80 mm Na⁺ to the intracellular surface of inside-out patches. The Na dependence for the human clone of Slo2.1 was similar to that reported for rat Slo2.1 expressed in CHO cells (Fig. 2B) (Bhattacharjee et al., 2003) and similar to that reported for Slo2.2 (Yuan et al., 2003). However, the mean single-channel conductance for hSlo2.1 was 60 pS, which was smaller than the 140 pS described for rSlo2.1. This discrepancy might be partially explained by the fact that we used 80 mm symmetrical K⁺, instead of 130 mm used previously (Bhattacharjee et al., 2003). Both currents showed very weak voltage dependence, reflected in the slope values obtained from fitted I–V curves in Figure 2C, a fact consistent with the lack of a conventional S4 voltage-sensing region (Fig. 1B). Low voltage sensitivity is also evident in ramps obtained from cell-attached patches. Single-channel openings can be clearly distinguished at potentials even more negative than the resting potentials of many cells (Fig. 2D). Control recordings in uninjected oocytes did not show any currents with the characteristics described above. The low sensitivity to voltage permits Slo2 channels to affect the resting potential of a cell, potentially controlling basal electrical excitability.

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Figure 2.

Functional properties of Slo2 channels expressed in Xenopus oocytes. A, Whole-cell currents of Slo2 channels. Records were obtained with two-microelectrode voltage clamp. V_h was −80 mV, in 10 mV voltage steps. B, Human Slo2.1 cRNA expresses a high-conductance Na-sensitive K⁺ channel. Bi, Single-channel Slo2.1 currents from an inside-out patch held at +50 mV and perfused with 80 mm Na or 0 Na, as indicated. Symmetric (80 mm) K⁺ was used in this experiment. Bii, Mean current–voltage relationship of single channels from multiple patches (n = 5). The single-channel conductance in 80 mm symmetrical K⁺ is 59 ± 0.001 pS (mean ± SE). C, Slo2 channels have shallow voltage dependence. Average current–voltage relationship for whole-cell Slo2 currents at V_h of −60 mV (n = 23 for Slo2.1 and n = 13 for Slo2.2). Open circles represent Slo2.1, and open triangles represent Slo2.2. Solid lines are the best fit of I = g × (V_m − V_k), where g = g_max/(1 + exp(−(V_m − V_1/2/K))). Parameters for the best fit were as follows: for Slo2.1, V_1/2 = −23 ± 9 mV, K = 51 ± 5 mV, g_max = 0.087 ± 0.002 mS; for Slo2.2, V_1/2= −15 ± 8 mV, K = 48 ± 6 mV, g_max = 0.14 ± 0.02 mS. Note that the slope factor K is quite high, indicating low intrinsic voltage sensitivity. D, Slo2 channels are active at negative potentials near the resting potential. Records elicited by ramps at indicated voltages are shown. Because records were obtained in the cell-attached configuration, the resting potential (V_r of approximately −80 mV) had to be taken into account to estimate the true membrane potential. [K⁺] in the pipette was 80 mm. Note that both channels are active at potentials even more negative than the usual cell resting potentials.

Slo2.1 and Slo2.2 channels are oppositely modulated by PMA

Because Slo2.1 and Slo2.2 have only slight voltage sensitivity, we considered the possibility that these channels might be modulated by factors in addition to Na⁺ and Cl⁻, possibly involving second-messenger systems. In considering this possibility, we noticed that several putative PKC phosphorylation sites were identified previously that are unique to one or the other channel (Bhattacharjee et al., 2003). Hence, we investigated whether Slo2.1 and Slo2.2 channels were subject to PKC modulation. Both Slo2 channels were expressed in Xenopus oocytes and exposed to the specific and potent PKC activator PMA. Figure 3 shows that the application of very low (nanomolar) concentrations of PMA had profound and opposite effects on the amplitude of Slo 2.1 and Slo2.2 whole-cell currents. Slo2.1 whole-cell currents were reduced by ∼90% after addition of 60 nm PMA to the bath solution. In contrast, Slo2.2 whole-cell currents were substantially increased after application of PMA (Fig. 3A). The inhibition of Slo2.1 currents occurred at all voltages (Fig. 3B). Notably, oocyte resting potentials were depolarized by ∼20 mV after application of PMA (from approximately −50 to −30 mV), suggesting that the Slo2.1 current contributed to the cells resting conductance. In contrast, Slo2.2 currents were increased approximately fivefold by 160 nm PMA, and the application of PMA to Slo2.2-expressing oocytes caused an increase in the cell resting potential of approximately −15 mV. Slo2.1 currents were inhibited by PMA with an IC₅₀ of ∼20 nm, whereas the increase in Slo2.2 currents had an IC₅₀ of ∼50 nm (Fig. 3C). The inhibition of Slo2.1 currents at a concentration of 20 nm PMA reached a maximal effect in 6–10 min (time constant, ∼2 min; voltage test pulse at +20 mV) (Fig. 3D). Stimulation of Slo2.2 took ∼4 min to half-saturation. The time course for PMA action observed here is consistent with reports in the literature for the effects of PKC on several membrane proteins and channels (Covarrubias et al., 1994; Boland and Jackson, 1999). The experiments described above were undertaken in the Xenopus oocyte expression system, but similar results were seen when HEK-293 cells were used as the heterologous system. In nystatin perforated patch-clamp recordings, bath application of PMA (100 nm) produced a rapid decrease in Slick whole-cell K⁺ currents and an increase in Slack currents (Fig. 3E).

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Figure 3.

Phorbol ester has opposite effects on Slo2.1 and Slo2.2 channels expressed in Xenopus oocytes and HEK cells. A, Effect of PMA on whole-cell records of Slo2.1 and Slo2.2. Top traces, Currents recorded before the addition of PMA; bottom traces, currents recorded after the addition of 60 nm PMA for Slo2.1 and 160 nm PMA for Slo2.2. B, PMA oppositely affects Slo2 currents at all voltages. Open symbols, Average current–voltage relationship before PMA exposure. Filled symbols, Current–voltage relationship after treatment with 60 nm PMA for Slo2.1 (n = 9) and 160 nm PMA for Slo2.2 (n = 10). C, Dose–response curves for PMA. Current is plotted as a function of PMA concentration. Note the semilog scale for Slo2.1 (left) and Slo2.2 (right). Test pulses were at 20 mV. Solid lines show the best fit (sigmoidal) of dose–response curves. IC₅₀ was 20 nm for Slo2.1 and 50 nm for Slo2.2. The Hill coefficient was similar in both cases (1.6). Although both channels are sensitive to PMA, Slo2.1 is more sensitive than Slo2.2. D, Response of Slo2 channels to PMA develops in minutes. Left, Onset of inhibition for Slo2.1 exposed to 20 nm PMA; right, onset of current enhancement for Slo2.2 exposed to 60 nm PMA. Test pulses were at 20 mV. E, PMA decreases the amplitude of Slick (Slo 2.1) currents and increases the amplitude of Slack (Slo 2.2) currents in HEK-293 cells. In nystatin perforated patch-clamp recordings, whole-cell K⁺ conductances were evoked by voltage steps between −120 and +120 mV in 20 mV increments from a holding potential of −70 mV. Bath application of PMA (100 nm) produced a rapid decrease in Slo2.1 and increase in Slo2.2 currents (currents started to change in 3–5 min after PMA application).

In Xenopus oocytes, the PMA effects on Slo2.1 and Slo2.2 were not reversed after 10 min of wash, suggesting a long-lasting effect of phosphorylation by PKC. PMA effects on both channels were almost completely eliminated by preincubation (8–12 h) in 1 μm staurosporin, a kinase blocker highly effective for PKC (n = 15). Also, modulatory effects were not seen in control experiments using 4-αPMA, an inactive analog of PMA, in concentrations up to 200 nm (n = 2). Additional evidence implicating PKC was achieved by experiments using the membrane permeant analog of diacylglycerol (DAG), 1-oleoyl-2-acetyl-sn-glycerol (OAG). Results of these experiments adding 30–40 μm OAG to the bath instead of PMA produced essentially the same results as PMA (data not shown). Because DAG is a second-messenger activator of PKC, this is additional evidence of the involvement of PKC. We thus conclude that the activation of PKC had opposite modulatory effects on Slo2 channels, activating Slo2.2 and inhibiting Slo2.1. Considering that Slo2.1 and Slo2.2 are high-conductance channels, even a minor fraction of these channels open at normal cell resting potentials of cells might permit PKC regulation of action potential threshold and basal cell excitability. The differential modulation by PKC of these two channels would thus have opposite functional implications; inhibition of Slo2.1 would increase excitability, whereas the stimulation of Slo2.2 would decrease it.

Transfer of inhibition by PMA to Slo2.2 by swapping Slo channel tails

Modulation of potassium channels by intracellular factors often occurs through the interaction of such factors with structural domains on the carboxyl oriented “tail” of the channel. The tail of the channel describes the cytoplasmic region that follows the membrane-spanning domains of the pore-forming “core” of the channel (Wei et al., 1994; Meera et al., 1997; Quirk and Reinhart, 2001). We noticed that several putative PKC phosphorylation sites were identified previously as unique to Slo2.1 and present on the tail of the channel. If the PMA-induced inhibition of Slo2.1 involved this region, a revealing demonstration to support this would be to construct a chimera, transplanting the tail of Slo2.1 onto the core of Slo2.2. If the hypothesis were correct, the action of PMA should inhibit rather than enhance the currents produced by this chimera. We therefore constructed the chimera (C1) that had the core of Slo2.2 and the tail of Slo2.1 ligated in-frame at a junction point immediately after the core (Figs. 1, 4A, arrows). The C1 construct expressed large currents similar to currents from Slo2.2 channels, albeit with faster activation kinetics (Fig. 4B). Like Slo2.2, C1 displayed shallow voltage dependence, sensitivity of gating to intracellular Na, and high unitary conductance (Fig. 4B,C). However, the effect of PMA on C1 was strikingly similar to that of Slo2.1 and unlike that of Slo2.2 (Fig. 5). Application of 20–50 nm PMA to oocytes expressing C1 currents caused significant inhibition (Fig. 5A). Application of 20 nm PMA reduced the C1 current ∼75% (Fig. 5B). Similar results were obtained from cell-attached patches (Fig. 5C). The cell-attached patch recordings shown in Figure 5C show a patch with three channels at −40 mV. After addition of 40 nm PMA, the activity diminished dramatically. The diminished current was mainly the result of lowered open-state probability (NPo); single-channel conductance was unaffected. When currents were evoked by linear ramps from −80 to +80 mV, PMA reduced C1 single-channel openings at all voltages. Similar results were obtained in several cell-attached patches, consistent with experiments using the two-microelectrode technique. Cell-attached experiments done as a control with Slo2.2 showed an opposite result; currents greatly increased after 1 min of PMA application (Fig. 5D). As with Slo2.1 and Slo2.2, incubation of the C1-injected oocytes in staurosporine prevented the action of PMA (n = 6). The inactive PMA analog 4-αPMA, had no modulatory effect on C1 currents (n = 2). Thus, all results support the conclusion that the PMA sensitivity by Slo2.1 is mediated by its C-terminal tail. The converse chimera (Slick core and Slack tail) was synthesized but had very low and inconsistent expression, and we were unable to determine whether PMA affected the level of current. However, we constructed a second chimera whose functional properties further supported the general hypothesis that the cytoplasmic tail altered modulation by PMA. This construct had the core of Slack (Slo2.2) with the tail of Slo1 ligated at the same restriction site used to construct the C1 chimera. This construct expressed large currents (Fig. 5E). However, these currents showed no response to PMA, neither increasing nor decreasing after PMA application (Fig. 5E). Thus, to summarize, the Slo2.1 channel is negatively regulated by PMA, the Slo2.2 channel is positively regulated by PMA, the chimera having the core of Slo2.2 and tail of Slo2.1 is negatively regulated by PMA, and, finally, the chimera having the core of Slo2.2 and tail of Slo1 is insensitive to PMA. Together, these results support the hypothesis that domains in the tail markedly influence the modulatory response of the channel to PMA and that negative regulation by PMA is likely to be attributable to domains in the Slo2.1 tail.

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Figure 4.

Slo2.2/Slo2.1 chimeric channels express well and retain basic properties. A, Chimera (C1). C1 was constructed with the Slo2.2 core and the Slo2.1 tail. Site of ligation is indicated by arrow (see Materials and Methods). B, C1 has weak voltage dependence. Currents elicited by 10 mV step pulses from a holding potential of −60 mV are shown to the left. Average current–voltage relationship is shown on the right (n = 9). Best-fit parameters were similar to those of Slo2.1 and Slo2.2. Current expression levels were usually much higher than those of Slo2.1 or Slo2.2 channels. C, C1 chimera is a high-conductance Na-activated K⁺ channel. Ci, Currents from an inside-out patch expressing C1 channels. Single-channel currents were continuously recorded at +40 mV in the presence or absence of 80 mm intracellular Na. Cii, Mean I–V relationship from n = 14 patches. The unitary conductance was 79 ± 0.002 pS (mean ± SE), similar to Slo2.2 (88 pS).

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Figure 5.

PMA inhibits C1 currents but stimulates Slo2.2. A, C1 whole-cell currents before and after addition of 20 nm PMA. B, Average current–voltage relationship before and after addition of 20 nm PMA (n = 8). C, Inhibition of C1 single channels by PMA. Continuous recordings from cell-attached patches were obtained at −40 mV before and after the application of 40 nm PMA. Records are shown at compressed (i) and expanded (ii) time scales. Before exposure to PMA, three active channels can be seen in the patch indicated as O₁–O₃. PMA caused inhibition in ∼2 min. NPo (measured for 10 s intervals) decreased from 1.2 to 0.028. Ciii, Single-channel recordings obtained with ramps from −80 to +80 mV from a holding potential of 0 mV before (left) and after (right) 40 nm PMA. D, Slo 2.2 single-channel activity is increased by PMA. Cell-attached recordings at +10 mV are shown at compressed (i) and expanded (ii) time scales. After applying 40 nm PMA for 2 min, additional Slo2.2 channels are active. Before the application of PMA, four levels of unitary conductance can be seen (O₁–O₄) and NPo was 1.1. After treatment with PMA, five levels of unitary conductance were observed (O₁–O₅) and NPo increased to 3.8. The recruitment of Slo 2.2 channels by PMA is also illustrated with histograms obtained for 40 s intervals (iii). E, Control experiments with Slo1 tail grafted to the Slack core show no effect of added PMA. Current traces show the Slack/Slo1 chimera currents in control conditions and in the presence of 60 nm PMA as labeled. Currents were elicited from a V_h of −90 mV, with test pulses ranging from −110 to +70 mV in 10 mV steps. The effect of PMA (60 nm) was measured after 6 min of the application of the drug. The graph at the right shows the mean normalized current–voltage relationships of the Slack/Slo1 chimera in control conditions (filled squares) and in the presence of 60 nm PMA (open squares) for five oocytes in each condition.

To determine whether PKC phosphorylation plays a major role in C1 inhibition and consequently in Slo2.1 inhibition, we took advantage of the high levels of single-channel expression of C1 and compared the effect of PMA with that of PKC directly applied to the intracellular surface of the channel. Figure 6Ai–Aiii illustrates an experiment in which we applied PKM (a constitutively active form of PKC) (Pontremoli et al., 1990) to the cytoplasmic face of the channel in inside-out patches. First, we demonstrated single-channel activity from C1 in a cell-attached patch, with ramps from −80 to +80 mV. Two channels are clearly seen in the patch (Ai). The patch was then excised to a bath solution with low concentrations of Na⁺ and Cl⁻ (20 and 40 mm, respectively), similar to those present in the egg cytoplasm (Cooper and Fong, 2003) (Aii). Under these conditions, the two channels remained active at a level comparable with that seen in the cell-attached configuration. Finally, the intracellular side of the patch was perfused with 0.1 U/ml PKM (containing 200 μm ATP-Mg). After treatment and return to the bath solution, channel activity in the patch diminished dramatically (Aiii). This result could be seen clearly in a continuous recording obtained from a test voltage of +40 mV (Fig. 6B). After 1–2 min of perfusion with PKM, the activity of the channel decreased by six times (NPo diminished from 0.6 to 0.1). Inhibition of single-channel activity by PKM was also seen in three other patches. In such experiments, channel rundown is a potential factor to be considered. However, we noted that rundown, when it occurred, usually developed very fast (<1 min). Thus, we took care to use membrane patches that had stable recording properties for several minutes, with no apparent rundown. Control experiments with direct application of PMA to the intracellular surface of excised patches did not show any effect.

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Figure 6.

Comparison of the effects of PMA and PKM on C1 channels. A, The activity of C1 chimeric channels in inside-out patches is reduced by PKM. Records were obtained with the ramp protocol shown at the bottom of the figure. Ai, Cell-attached patches. Aii, Cell-attached patches excised in low NaCl solution. Aiii, Reduced single-channel activity after perfusion of the cytoplasmic surface with 0.1 U/ml PKM. B, Reduction of single-channel activity after PKM application seen in continuous recordings from inside-out patches held at +40 mV. Effects of PKM were clearly seen after 2 min of perfusion of 0.1 U/ml PKM. NPo values are 0.6 before and 0.1 after 2 min of application of PKM. C, Reduced activity of C1 channels from oocytes treated with PMA persists in excised patches. Records were obtained with the ramp protocol shown in the figure. Ci, Single-channel activity in cell-attached patches. Cii, Oocytes were treated with 40 nm PMA, and inhibition of channels in cell-attached patches was observed. Ciii, Single-channel activity observed in excised patches of PMA-treated oocytes remains low, indicating that PMA promotes a long-lasting modification of the channel, independent of possible cytosolic factors.

Direct action of PMA-stimulated PKC on the channel may produce a long-lasting effect. Presumably, after PMA treatment of an oocyte, C1 channels would be phosphorylated and remain inhibited, even after patch excision. To test this hypothesis, single-channel activity was first recorded in the cell-attached patch configuration. The oocyte was then exposed to 40 nm PMA. After 1–2 min of exposure, channel activity was almost completely inhibited (Fig. 6C). This low state of activity remained significantly lower than in the pretreated cell-attached condition and persisted for at least 5 min. After excision of the patch, the low level of activity persisted at a similar level (Fig. 6Ciii). These experiments suggest that, at least in part, the inhibitory effect of PKC may be attributable to a direct phosphorylation of the Slo2.1 tail. However, we cannot rule out the possibility that there is an accessory protein associated with the channel that is the target of PKC-stimulated phosphorylation.

Slo2.1 and Slo2.2 channels are modulated by neurotransmitters

Activation of PKC is an integral part of the signaling cascade elicited by neurotransmitter stimulation of Gα_q-coupled protein receptors. We noted that Slo2 channels are expressed in many regions of the brain in which Gα_q-coupled protein receptors are present (Pontremoli et al., 1990; Hoffman and Johnston, 1998; Marinissen and Gutkind, 2001; Brady and Limbird, 2002; Cooper and Fong, 2003; Kroeze et al., 2003; Breitwieser, 2004; Rashid et al., 2004). Examples include hippocampal neurons of the CA1 region, in which Slo2.1 channels are abundant (Bhattacharjee et al., 2003). Conceivably, in some cells, Slo2 channels and Gα_q-coupled receptors may be coexpressed and functionally coupled (Fisahn et al., 2002). To test whether such functional coupling was possible, we coexpressed Gα_q receptors and Slo2 channels in the Xenopus oocyte expression system. Examples of such experiments are shown in Figure 7, in which we reconstituted the functional coupling between both Slo2 channels and the M₁ receptor. As might be expected from the results obtained with PMA, stimulation of the M₁ receptors by 5–40 μm oxotremorine (a specific agonist of the M₁ receptor) inhibited Slo2.1 currents, whereas it augmented Slo2.2 (Fig. 7A). The average current–voltage relationships are shown in Figure 7B. Similar experiments were also performed with AT1B and mGluR1. In each case, the results were consistent with those obtained with the M₁ receptor (data not shown). (Angiotensin II was used to stimulate the AT1B receptors, whereas mGluR1 receptors were stimulated by the agonist 1-aminocyclopentane-1,3-dicarboxylic acid.) Receptors of these types are known to be expressed in hippocampus, in which they are likely to modulate neuronal excitability (Mannaioni et al., 2001; Ireland and Abraham, 2002). In control experiments, application of the above agonists to oocytes injected only with Slo2 channels did not have any effect. Also, the expression of the receptor alone either with or without the agonists did not produce currents larger than endogenous background currents.

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Figure 7.

Opposite modulation of Slo2.1 and Slo2.2 channels by M₁ receptor activation in Xenopus oocytes. A, Slo2.1 and Slo2.2 currents recorded from oocytes coinjected with channel and M₁ receptor cRNA. Currents are shown before (reference) and after the addition of 30 μm oxotremorine. Slo2.1 (left) is reduced whereas Slo2.2 (right) is enhanced by oxotremorine application. Effects are attributable to stimulation of the M₁ receptors by oxotremorine because no effects by oxotremorine were observed in oocytes injected with the Slo2 channels alone (data not shown). B, Average current–voltage relationship in oocytes coinjected with Slo2.1 and Slo2.2 cRNAs and the M₁ receptor. Slo2.1 currents are shown by circles (n = 4), and Slo2.2 currents are shown by triangles (n = 4). Reference conditions are shown by open symbols, and 30 μm oxotremorine is shown by filled symbols.

Slo2 channels and Gα_q-coupled protein receptors are coexpressed in the CNS

To gain additional evidence for the hypothesis that Slo2 channels are regulated through activation of the M₁ and mGluR1α receptors in vivo, as well as in our reconstituted system, we undertook coimmunostaining experiments on rat brain sections, as well as on primary cell cultures of neurons derived from the hippocampus. The goal of these colabeling experiments was to verify that both receptors and channels could be expressed in the same types of identified cells. Both Slo2.1 and Slo2.2 are very widely expressed throughout the CNS (Bhattacharjee et al., 2002, 2005). Figure 8 shows some examples of colocalization. For example, Slo2.1 is expressed in neurons of the prefrontal cortex, in which mAChR1 receptors are also strongly expressed (Levey et al., 1991). Clear colocalization of receptors and Slo2.1 channels can be observed (Fig. 8A). The Slo2.2 channel is less widely expressed in cortex but is found in certain neurons of frontal cortex (Bhattacharjee et al., 2002). Figure 8B shows colocalization of Slack (Slo2.2) with mAChR1 in a subset of neurons of layer V of frontal cortex. In hippocampus, the Slick (Slo2.1) channel is expressed in both CA1 and CA3 regions, in which it colocalizes with the mAChR1 receptor (Fig. 8C,D) (Levey et al., 1995; Bhattacharjee et al., 2005). Finally, mGluR1 receptors are expressed in hippocampus (Shigemoto et al., 1997). Figure 8E demonstrates coimmunolabeling of Slo2.1 with mGluR1α in cultures of hippocampal neurons. The observed pattern of staining is consistent with colocalization of Slo2.1 and mGluR1 at the plasma membrane. In general, in this and previous studies, Slo2.1 and Slo2.2 channels appear to be most prominently localized in the soma and initial segments in which they conceivably could play an important role in the integrative properties of neurons.

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Figure 8.

Colocalization of Slack and Slick with Gα_q-coupled receptors. Top 12 panels show colocalization of Slick or Slack with mAChR1 in sections of rat cortex (A, frontal cortex; B, layer V of frontal cortex) and in hippocampus (C, CA1 region; D, CA3 region). E shows colocalization of Slick with mGluR1α in cultures of hippocampal neurons. Slick and Slack immunoreactivity was localized with Cy3 (red), and receptor immunoreactivity was visualized using Alexa Fluor 488 secondary antibody (green). On the overlaid images, regions of colocalization appear orange to yellow. Scale bars, 20 μm.