Membrane Phospholipid Control of Nucleotide Sensitivity of KATP Channels

See allHide authors and affiliations

Science  06 Nov 1998:
Vol. 282, Issue 5391, pp. 1138-1141
DOI: 10.1126/science.282.5391.1138


Adenosine triphosphate (ATP)–sensitive potassium (KATP) channels couple cell metabolism to electrical activity. Phosphatidylinositol phosphates (PIPs) profoundly antagonized ATP inhibition of KATP channels when applied to inside-out membrane patches. It is proposed that membrane-incorporated PIPs can bind to positive charges in the cytoplasmic region of the channel's Kir6.2 subunit, stabilizing the open state of the channel and antagonizing the inhibitory effect of ATP. The tremendous effect of PIPs on ATP sensitivity suggests that in vivo alterations of membrane PIP levels will have substantial effects on KATP channel activity and hence on the gain of metabolism-excitation coupling.

KATP channels (1) are formed from a sulfonylurea receptor (SURx) and an inward rectifier (Kir6.x) subunit (2). Evidence is accumulating that the Kir6.x subunit forms the pore and controls the hallmark inhibition by ATP (3), although the mechanism of this inhibition remains elusive. Recent reports indicate that membrane PIPs bind KATP and other Kirchannels, stabilizing them in an active conformation (4,5). Both the inhibitory effect of adenosine nucleotides and the stimulatory effects of PIPs increase with the number of phosphate groups in the molecule (1,6), which suggests that PIP activation and ATP inhibition may be related phenomena. We specifically hypothesized that PIPs and ATP might compete for binding to the KATP channel, stabilizing open and closed channels, respectively.

To test this hypothesis, we recorded currents in inside-out patches from COSm6 cells expressing cloned (Kir6.2+SUR1) KATP channels (7). After patch excision, a slow run-down of channel activity occurred over the following minutes, and this run-down could be avoided by addition of phosphatidyl inositol-4,5-bisphosphate (PIP2) (Fig. 1A). Open probability in the absence of ATP (P open) was ∼0.4 before application of PIP2 but increased to ∼0.85 (Fig. 1C), which is consistent with the approximate doubling of current in macroscopic patches that followed treatment with PIP2 (Fig. 1E) and demonstrates that PIP2 activates KATP current by increasing P open. ATP sensitivity (7) immediately after patch excision could be fit by a sigmoid relation with a half-maximal inhibition concentration (K 1/2) of 12.1 μM and a Hill coefficient (H) of 1.3 (n = 26), which is similar to previous reports (2), but it decreased by orders of magnitude after PIP2 application (Fig. 1, A and B). The rate of this decrease was variable from patch to patch (Fig. 1D), probably because of variable diffusion distances (8), but correlated with the time course of increase in ATP-independent channel activity (Fig. 1F). ATP sensitivity decreased 340 times (fromK 1/2 of 10.5 ± 0.1 μM to 3.6 ± 0.3 mM) after 10 min of exposure to PIP2 (5 μg/ml) (n = 18). This shift in ATP sensitivity is far greater than any previously reported effects of potassium channel–opening drugs (9). After PIP2 application, channel activity became very stable, with run-down time constants of >30 min, and the effect on ATP sensitivity remained, at least within 20 min of PIP2withdrawal (Fig. 2A). The general applicability of the phenomenon is illustrated in Fig. 2, C and D; prolonged exposure of KATP channels to PIP2 in either cardiac myocyte or β-cell (HIT-T15) membrane patches resulted in 100- to 700-fold increases of K 1/2.

Figure 1

(A) Representative WT (SUR1+Kir6.2) KATP channel currents in an inside-out patch (7). The patch was isolated at the arrow and exposed to differing ATP concentrations or to 5 μM PIP2, as indicated. The dotted line indicates zero current. (B) K 1/2 [estimated from fits of the Hill equationI rel = 1/{1 + ([ATP]/K 1/2)H}, withI rel being the current relative to the peak current after PIP2 and H fixed at 1.3; see inset] as a function of time for the record in (A). (C) Single WT KATP channel current (top) andP open (1-s bins) (bottom). The patch was isolated immediately before the onset of the record and exposed to 5 μM PIP2, as indicated. (D) K 1/2 from individual experiments like those in (A) versus time after onset of exposure to 5 μM PIP2. Averaged K 1/2 (±SEM) before application of PIP2 (Pre) is indicated by the larger symbol. (E) I rel versus time after onset of exposure to PIP2. (F)I rel [from (E)] versusK 1/2 [from (D)].

Figure 2

(A)K 1/2 as a function of time for an experiment like that in Fig. 1A. PIP2 was applied only during the period indicated. (B) K 1/2 from patches (n = 3 to 6) exposed to 5 μM PC, IP3, PI-4-P, PIP2, or PIP3 for 0 min (open bars) or 8 (7.3 to 9.1) min (solid bars). (C and D) Representative KATPchannel currents in inside-out patches from a HIT T15 cell (C) and a mouse ventricular myocyte (D). Patches were isolated at the arrows and exposed to ATP or to 5 μM PIP2, as indicated. In all experiments with HIT T15 cells and ventricular myocytes,K 1/2 increased from 9 ± 1 μM and 19 ± 4 μM to 2.1 ± 0.3 mM and 5.8 ± 0.8 mM, respectively (n = 4 and 7 patches), after 5.5 ± 0.3 min and 9 ± 0.5 min of PIP2 exposure.

Phosphatidyl inositol-4-phosphate (PI-4-P) also stimulated channel activity and reduced ATP sensitivity, although it did so less effectively than PIP2 (Fig. 2B). Phosphatidyl inositol-3,4,5-triphosphate (PIP3) was as effective as PIP2. Neither phosphatidyl choline (PC) nor inositol triphosphate (IP3) altered P open(4) or ATP sensitivity (Fig. 2B). Therefore, a negatively charged head and a lipid tail are necessary both to stimulate ATP-independent activity (4) and reduce ATP sensitivity. Polycations have been shown to inhibit KATP channels by screening negative charges (10), and this action is a substantial cause of channel run-down in excised patches. As shown inFig. 3A, application of polylysine or Ca2+ or spermine (11) after stimulation by PIP2 caused rapid reversal of both the increasedP open and the reduced ATP sensitivity.

Figure 3

(A) Representative WT current in an inside-out patch, isolated at the arrow and exposed to differing ATP concentrations or to 5 μM PIP2, as indicated. The gaps in the record are 2 and 3.5 min long. Polylysine (10 μg/ml, molecular weight ∼1000) was applied as indicated. (B) K 1/2 as a function of time for the record in (A). (C) Representative mutant Kir6.2 [R176A]+SUR1 currents in an inside-out patch, isolated at the arrow and exposed to differing ATP concentrations or to 5 μM PIP2, as indicated. Inset at left shows current immediately after patch excision, amplified 20 times.

There have been indications that the COOH-terminus of Kirchannels is involved in PIP2 activation (4,5) and in the ATP sensitivity of KATP channels (3). According to the above hypothesis, mutations that reduce ATP binding to Kir6.2 should reduce ATP sensitivity without altering P open. Neutralization of residue K185 in Kir6.2 reduces ATP sensitivity and may contribute to ATP binding (3, 12). Kir6.2[K185Q] mutant channels (13) are ∼30 times less sensitive to ATP than wild-type (WT) Kir6.2+SUR1 channels when first isolated, but have similarP open values (11). ATP sensitivity shifts in parallel for both WT and K185Q mutant channels after PIP2 treatment, with the K 1/2 of K185Q mutant channels rising to 39 ± 7 mM after 6.9 ± 1.0 min of PIP2 treatment (six patches were tested). Conversely, mutations that reduce PIP2 sensitivity should have a lowP open but show a saturatingP open and ATP sensitivity similar to those of WT channels after PIP2 treatment. Residues R176 and R177 of Kir6.2 are two positively charged amino acids that might contribute to PIP2 binding (4, 5); neutralization of residue R188 in Kir1.1 (equivalent to R177 in Kir6.2) reduces PIP2 binding to these channels (5). Both Kir6.2[R176A] and Kir6.2[177A] mutations (+SUR1) expressed considerably lower conductances in intact cells than did WT channels, as assessed by86Rb efflux (14) (21 ± 8% and ∼0%, respectively, in three separate paired transfections). In inside-out patches, no currents were detected from the R177A mutant, and very small currents were observed from the R176A mutant. The low channel activity makes accurate assessment of K 1/2 very difficult and reflects an extremely low P open, as confirmed by an enormous increase of current that followed PIP2 application (Fig. 3). The current in zero ATP increased >80-fold (the current before PIP2 was 1.2 ± 0.1% of the current after PIP2) and increased more slowly than in WT channels [half-time (t 1/2) = 76 ± 20 s and 32 ± 6 s, respectively], but K 1/2nevertheless reached comparable levels (2.1 ± 0.1 mM after 9.8 ± 0.3 min of PIP2 treatment; n = 6; Fig. 3C). These results are consistent with the R176A mutation lowering PIP2 affinity, thus underlying the reduced physiological activity of Kir6.2[R176A] channels and providing crucial evidence for the physiological role of PIP2 in maintaining normal channel activity.

To further examine interactions between PIP2, ATP, and the Kir6.2 subunit, we engineered a protein (Kir6.2-C) containing the Kir6.2 COOH-terminus (amino acids 170 through 390) fused to glutathione S-transferase (GST) (13). After PIP2 exposure, GST was without effect, but purified Kir6.2-C markedly inhibited channel activity (Fig. 4). When Kir6.2-C was removed, channel activity recovered only slowly; this recovery, however, was greatly accelerated by exposure to ATP. Kir6.2-C might block the channel, acting as a “ball”-like domain similar to the NH2-terminus ofShaker-like channels (15), but the following observations suggest an alternative hypothesis. Kir6.2-C inhibition was relieved in the presence of PIP2, which suggests that rather than directly blocking the pore, positive charges on Kir6.2-C may bind to the negatively charged PIPs in the membrane, screening them from the channel itself and effectively reducing the membrane PIP2 concentration that the channel itself sees. When Kir6.2-C is added in the presence of PIP2, Kir6.2-C is significantly bound to the micellar PIP2 in the solution rather than to PIPs in the membrane. These experiments were performed after prolonged pretreatment with PIP2. Under this condition, 1 mM ATP was without significant inhibitory effect (see Fig. 4B; ATP was added at the up arrow) but augmented the inhibitory effect of subsequently applied Kir6.2-C (Fig. 4, B and C), as would be expected if exogenous Kir6.2-C lowers the effective membrane PIP2 concentration by a charge-screening effect.

Figure 4

(A) Representative WT current in an inside-out patch. The patch had been exposed to 5 μM PIP2 for ∼10 min before the first trace, and the second trace was obtained ∼3 min after the first. The patch was exposed to 100 nM GST, 100 nM Kir6.2-C, or 5 mM ATP, as indicated. (B) Representative WT currents in another inside-out patch. The patch had been exposed to 5 μM PIP2 for ∼12 min before the first trace, and the traces were ∼5 min apart. The patch was exposed to Kir6.2-C with either 1 mM ATP or 5 μM PIP2, as indicated (down arrow). For the Kir6.2-C+ATP trace, 1 mM ATP alone was applied beforehand (up arrow). (C) Steady-state current (±SEM), relative to the current before treatment, during application of GST or Kir6.2-C, alone or with PIP2 or ATP from experiments (n = 3) like that in (B).

The present results show that ATP sensitivity can be changed over orders of magnitude by manipulation of the PIP content of the membrane, and they provide a mechanistic framework for understanding the hallmark inhibition of these channels by ATP. Although we do not expect exact overlap of the PIP2 and ATP binding sites, a negative heterotropic cooperativity is expected, so that individual residues may contribute to the binding of each ligand. Intact cell channel activity is reduced in the Kir6.2[R176A] mutant, demonstrating the physiological relevance of this finding. Membrane PIP composition may vary physiologically (16), and this may explain the wide variability in ATP sensitivity of native KATP channels (17). It has long been recognized that activation of KATP channels occurs under conditions where the cytoplasmic concentration of ATP is much higher than that required to inhibit channels in excised membrane patches (1, 6, 9). The profound effects of PIP2 on ATP sensitivity would suggest that as membrane PIP levels increase, KATP channels will be rendered insensitive to ATP, providing a mechanism for physiological activation.

  • * To whom correspondence should be addressed. E-mail: cnichols{at}


Stay Connected to Science

Navigate This Article