(Received for publication, December 3, 1996, and in revised form, December 27, 1996)
From the Department of Medicine, University of Wisconsin, Madison, Wisconsin 53792 and University of Chicago, Chicago, Illinois 60637
The ATP-sensitive potassium channel (KATP)
controls insulin release in pancreatic -cells and also modulates
important functions in other cell types. In this study we report that
anionic phospholipids activated KATP in pancreatic
-cells, cardiac
myocytes, skeletal muscle cells, and a cloned KATP composed of two
subunits (SUR/Kir6.2) stably expressed in a mammalian cell line. The
effectiveness was proportional to the number of negative charges on the
head group of the anionic phospholipid. Screening negative charges with
polyvalent cations antagonized the effect. Enzymatic treatment with
phospholipases that reduced charge on the lipids also reduced or
eliminated the effect. These results suggest that intact phospholipids
with negative charges are the critical requirement for activation of
KATP, in distinction from the usual cell signaling pathway through
phospholipids that requires cleavage. Mutations of two positively
charged amino acid residues at the C terminus of Kir6.2 accelerated
loss of channel activity and reduced the activating effects of
phospholipids, suggesting involvement of this region in the activation.
Metabolism of anionic phospholipids in plasmalemmal membrane may be a
novel and general mechanism for regulation of KATP and perhaps other ion channels in the family of inward rectifiers.
The ATP-sensitive potassium channel (KATP)1 is a highly regulated channel type important in the physiology and pathophysiology of pancreas, heart, vascular smooth muscle, and perhaps other tissues as well (1-5). KATP activity can be reconstituted by co-expression of an inwardly rectifying channel (Kir6.2) (6, 7), and a sulfonylurea receptor (SUR) (8, 9). Major regulatory mechanisms are postulated to reside in the nucleotide binding domains on SUR (8-10). Activation of KATP generally requires a reduced ATP concentration. This regulation, including inhibition of KATP activity by other nucleotides, does not require phosphorylation. Activation of KATP, however, also depends on another less well characterized regulatory process, because KATP gradually inactivates when the integrity of the cell is disrupted by, for example, the excised patch method of voltage clamp (11-13). This property is shared not only by most KATP channels from different tissues, but also by other channels of the inward rectifier potassium (Kir) superfamily (for reviews, see Refs. 14 and 15). Despite intense investigation, the mechanisms for maintaining activation of KATP have not been fully identified and characterized, although previous work has focused on protein phosphorylation processes on the channel or related subunit (for a review, see Ref. 16). We found that anionic phospholipids, especially phosphoinositides, are necessary and sufficient to activate and maintain KATP activity in native and cloned KATP and in two other Kir channels. We also propose and test a novel hypothesis involving the cytoplasmic tail of the pore-forming subunit to account for this regulation. Modulation of channel activity through the composition and phosphorylation of membrane phospholipids may represent an important regulation for KATP in particular, and for channels in the Kir superfamily in general.
Single pancreatic -cells were
enzymatically (0.05% trypsin) dispersed from isolated islets of Wistar
rats collected by centrifugation through a discontinuous gradient of
Ficoll (17). Single ventricular myocytes were isolated from rabbit
hearts by an enzymatic (0.1% collagenase, Williton) dissociation
method (17, 18). Single skeletal muscle cells were dissociated
enzymatically (0.3% collagenase, Williton) from the flexor digitorum
brevis muscles of the hind legs of Wistar rats (13). Isolated single
skeletal muscle cells were suspended in Dulbecco's modified Eagle's
medium until they were used. Membrane vesicles were formed by washing
the isolated cells with high K+ solution.
The pipette solution (extracellular side, 140-K+-pipette) contained (in mM) KCl (140.0), CaCl2 (1.8), MgCl2 (0.5), HEPES (5.0), and glucose (5.5), pH 7.4. The pipette solution for membrane vesicles from skeletal muscle cells and cloned KATP (SUR/Kir6.2) contained (in mM) NaCl (140.0), CaCl2 (1.8), MgCl2 (0.5), KCl (10.0), HEPES (5.0), and glucose (5.5). The bath solution (cytoplasmic side, 140-K+-bath) contained (in mM) KCl (142.0), HEPES (5.0), glucose (5.5), and EGTA (2.0), pH 7.4. EGTA was included unless indicated otherwise. For whole cell clamp experiments, the pipette solution (140-K+ with Mg-ATP) contained (in mM) KCl (140.0), EGTA (2.0), HEPES (5.0), glucose (5.5), and Mg-ATP (5.0), pH 7.3. The bath solution (5.4-Tyrode solution) contained (in mM) KCl (5.4), NaCl (137.0), CaCl2 (1.8), MgCl2 (0.5), and HEPES (5.0), and the pH was adjusted to 7.4. K2-ATP (Sigma) was dissolved into the bath solution just before use, and the pH of the solution was readjusted. DNase I and cytochalasin B were products of Sigma. Polyvalent cations were prepared in stock solutions and added to the solution prior to use. The concentration of free cations was calculated (19). A 2-ml bath was perfused with a solution of dispersed lipids at concentrations of 0.005-1 mg/ml. PPIs (a mixture of PI(4,5)P2, PI(4)P (15-20%), PI, PS, and PC, from bovine brain; Sigma) were used. Other lipids were as follows: PI(4,5)P2 (Boehringer Mannheim), PI(4)P (sodium salt, from bovine brain; Sigma), PI (from bovine liver; Avanti Polar Lipids), PS (from brain; Avanti Polar Lipids), PC, PE (from bovine heart; Avanti Polar Lipids), IP3 (potassium salt, from bovine brain; Sigma), DOG (8:0 DG), and DPG (16:0 DG; Sigma). Anionic phospholipids were dispersed in the solution with 30-min sonication (half setting) on ice. The following uncharged lipids were dissolved first in dimethylsulfoxide and then sonicated for 30 min: PLC-PI (from Bacillus cereus; Sigma), PLC (Type IV, from Bacillus cereus; Sigma), PLA2 (Sigma), and PLD (Sigma). All enzymes were dissolved directly into reaction solution. Experiments were done at room temperature (21-24 °C).
cDNA Expression and MutagenesisThe wild type recombinant complementary DNAs encoding the rat SUR (rSUR), and rat Kir6.2 (rKir6.2) were cloned from a cDNA library of insulinoma beta cell tumor line, and a rat islet cDNA library (7), respectively. A polymerase chain reaction-based site-directed mutagenesis kit (ExSite, Stratagene) was used to generate the site-directed point mutations, which were confirmed by sequencing. The cDNA of SUR (in a pcDNA3 vector, Invitrogen) and wild type Kir6.2 or its mutants (in a pCR3 vector, Invitrogen) were co-transfected and heterologously expressed in mammalian cells either transiently (tsA201 cell line (20)) or stably (a human embryonic kidney cell line, HEK) using a calcium-phosphatide transfection kit (Life Technologies, Inc.). The transfected cells were cultured in Dulbecco's modified Eagle's medium for 12 h with the precipitated cDNA. The transiently transfected cells were kept in culture for another 24-48 h before being used in electrophysiological experiments. Stably transfected cells were selected by growth in media containing 400-600 µg/ml active G418 (Life Technologies, Inc.) for at least 1 week, and single colonies surviving were isolated and grown to harvest in the continuous pressure of selection with 200-300 µg/ml active G418.
Electrophysiological Recordings and Data AnalysisCurrents were recorded using inside-out patch clamp (cytoplasmic membrane exposed to the bath) or whole cell clamp (external membrane exposed to bath) through a patch clamp amplifier (Axopatch 1-D or Axopatch 200A, Axon Instruments) and filtered through a built-in low-pass filter at 1 kHz, unless indicated otherwise. Leak current was compensated electronically on-line or subtracted off-line. Data were acquired by digitizing at 2 kHz and analyzed by a pClamp 6.0 (Axon Instruments) and graphic plotting software running on a PC-compatible computer (Gateway 2000).
Currents of native KATPs were recorded from inside-out patches of
pancreatic -cells (Fig. 1A), cardiac
ventricular myocytes (Fig. 1B), and sarcomeric membrane
vesicles from skeletal muscle cells (Fig. 1C). KATP current
expressed in HEK cells stably transfected by cloned cDNA of rSUR
and rKir6.2 was also studied (Fig. 1D). Open channel
activity was increased within 1-2 s to a maximal value after excising
the patch membrane into an ATP-free solution; this was followed by
decreased channel activity for pancreatic cells, cardiac cells, and
transfected HEK cells. In the vesicles from skeletal muscle cells, the
patch was initially devoid of KATP activity in most cases (Fig.
1C), presumably because these channels had inactivated
during the vesicle formation process. An anionic phospholipid-rich
tissue extraction containing PPIs (0.05-1 mg/ml) consistently
activated KATP in cells from all three tissues and also the cloned
KATP. Maximal effects were reached within 30 s to 2 min and did
not change with time if the perfusion of the lipid-containing solution
was maintained. After wash-out of PPIs from the bath solution, the
activation effect remained for a time but diminished slowly with a time
course that depended upon the previous concentration of PPIs. PPIs had
no effects on single channel current amplitudes or ATP sensitivity. A
non-KATP inwardly rectifying K+ channel, presumably the
cardiac inward rectifier, or IRK1, noted coincidentally in Fig.
1B (small conductance channel observed before excision) also
lost activity and was then activated by PPIs (small conductance
apparent after ATP suppressed KATP). In separate experiments, PPIs
activated currents expressed from clones of mouse inwardly rectifying
K+ channels mIRK1 and the ATP-regulated inward
rectifier hROMK1 (Fig. 2).
The preparation of PPIs was a mixture of PI(4,5)P2, PI(4)P,
PI, PS, and PC; to evaluate the contribution of the individual components to the activation effect, we individually tested purified PI(4,5)P2 (5 negative charges), PI(4)P (3 negative
charges), PI (1 negative charge), PS (1 negative charge), PC (neutral),
and another neutral phospholipid, PE, using the same
protocol. All negatively charged phospholipids tested
activated KATP; Fig. 3A shows an example for
PI(4,5)P2, and results for all lipids tested are summarized
in Fig. 3D. Similar to PPIs, the activation effect also
remained after washing the lipids from the bath solution. The most
effective anionic phospholipid was PI(4,5)P2, and others were less effective. The effectiveness sequence correlated with the
number of negative charges in the head groups, suggesting a role for
electrostatic force. Uncharged PC (1 mg/ml) had no effect on KATP
activity, while uncharged PE (1 mg/ml) induced a slight depression of
KATP activity. In intact cells, phospholipase C hydrolyzes
phospholipids such as PI(4,5)P2 into the cascade products,
soluble IP3 and membrane-delimited diacylglycerol, which as
important second messengers, may affect channel function. Although IP3 has 6 negative charges/molecule, it had no effect on
KATP activity (0.1 mg/ml) (Fig. 3B). The diacylglycerols
(DPG and DOG) actually reduced activity of KATP (results summarized in
Fig. 3D). This indicates that an intact unit of a
hydrophobic tail and a charged hydrophilic head are a basic structural
requirement for the anionic phospholipids to activate KATPs. Other
cascade products of phospholipids, arachidonates, had inconsistent
effects on KATPs in cardiac cells (21) and insulinoma cells (22). The
effect of intact phospholipids to activate KATP was also tested by
using phospholipase-treated lipids (Fig. 3C). PLC-PI, PLC, or PLA2 was incubated with PI(4,5)P2, PI(4)P,
and PS. The enzyme-treated lipids were much less potent than the
untreated lipids (Fig. 3E). Interestingly, PLD treatment of
PC increased KATP activity (Fig. 3E). PLD adds an anion to
the head group of neutral PC, making this observation consistent with
the effectiveness sequence depending upon net negative charge. Finally,
exogenous PLA2 decreased KATP activity in inside-out
patches, consistent with a recent report (23). This effect of
PLA2 was reversed by subsequent application of PPIs.
We further tested the hypothesis that the negative charges in the
anionic heads of the anionic phospholipids are the functional group
critical to the activation of KATP by screening those charges with
polyvalent cations. As an example (Fig. 4A),
the polyvalent cation antibiotic neomycin decreased KATP activity, and
PPIs could reverse and antagonize this effect. Divalent cations such as
Ca2+ and Mg2+ are already known to accelerate
the decrease of KATP activity (24, 25) by an unknown mechanism. We
found that trivalent cations such as Gd3+ and
La3+ (ED50 = 70 nM and 0.3 µM, respectively; ED50 defined as 50%
inhibition at 1 min) were even more potent inhibitors of KATPs than
divalent cations (Fig. 4B). Other polyvalent cations
including aminoglycoside antibiotics and polyamines used as negative
charge chelators for phospholipids also have the similar effect.
Neomycin (ED50 = 20 µM), gentamicin
(ED50 = 60 µM), and spermine
(ED50 = 0.94 mM) inhibited KATP activity
irreversibly (Fig. 4B). A decreased potency for the bigger
molecules may reflect the influence of the molecular sizes in chelating
negative charges of the phospholipids for these polyvalent cations.
Application of exogenous PPIs activated KATPs previously inhibited by
polyvalent cations, and application of polyvalent cations with PPIs
reduced the effect of PPIs.
The effects of phospholipids on KATP were rapid, sustained, and general
to all anionic phospholipids and all types of KATPs tested. The
mechanism for the effect of these lipids appeared less likely to
involve a very specific or complex cellular process requiring many
steps. We hypothesized that PPIs directly interact with KATPs or one of
its subunits, mediated by electrostatic force, to maintain the channel
in a functional state. We noted a segment of Kir6.2 composed of a high
concentration of positively charged residues at the beginning of the
cytoplasmic C terminus (Fig. 5A). This
positioning of a cluster of positive charges has been shown to be an
important determinant of protein topology in other membrane proteins
(26, 27). To test our hypothesis, two mutants were made in this region:
1) adjacent positively arginine residues at positions 176 and 177 were
mutated to neutral alanines (R176A,R177A), and 2) the arginine at 176 was mutated to the negatively charged glutamic acid (R176E). Wild type
Kir6.2 co-transfected with SUR expressed a peak KATP current of
1.59 ± 0.3 nA (n = 8) recorded by whole cell
clamp at 0 mV with no ATP in the pipette. The current was blocked by
glibenclamide and also exhibited the usual loss of activity with time.
The R176A,R177A mutant co-transfected with rSUR also expressed a
glibenclamide-sensitive K+ current, but the current was
much smaller, with a mean value of 0.32 ± 0.07 nA
(n = 4, p = 0.008, compared with wild
type) at 0 mV at maximum. Loss of channel activity was much faster than wild type, and the current showed more fluctuation. Examples of whole
cell wild type and R176A,R177A currents are shown in Fig. 6, panels A and B, respectively.
In inside-out patches, exogenous application of PPIs also activated the
channels expressed from the R176A, R177A mutant, but activation was
less effective than for wild type, as demonstrated by a lower open
probability and by the transient nature of the activation. Single
channel recordings (Fig. 7A) from excised
patches showed that the lower open probability in the mutant was caused
by a decrease in open times (Fig. 7B) with a possible
increase in the closed times. No glibenclamide-sensitive current could
be recorded from the cells co-transfected with the R176E mutant and
rSUR, suggesting that either this mutant did not express or that the
mutation rendered it incapable of opening.
Gradual inactivation of KATP
with time was recognized with the first description of KATP (1) using
the inside-out patch clamp technique, a method that severely disrupts
the cytoplasmic environment. This inactivation, sometimes called
run-down, has been reported or addressed in nearly all research on
KATP. Loss of activity is a consistent feature of all native and cloned
KATP (Fig. 1, and Refs. 11-13 and 16). Despite much study, the
mechanism required to maintain KATP activity is still unclear. Mg-ATP
(28, 29) was found to transiently and partially activate and maintain KATP in cardiac, pancreatic, and skeletal muscle cells. Some anions (F, VO
) (30, 31) were also shown to have
similar effects. Trypsin treatment was found to dramatically maintain
KATP activation (18, 32). Mg2+ was found to be necessary
for ATP activation of KATP; therefore, many researchers speculated that
phosphorylation/dephosphorylation was the key to maintenance of KATP
channel activity (16). Mg-ATP, however, was often ineffective in
activating KATP in rat skeletal muscles (13). In addition, intensive
searches for protein kinases and the protein phosphorylation targets
have not been successful (32). These results suggest that although
phosphorylation may be a part of the regulatory process, additional key
elements are involved. In rat skeletal muscle cells, KATP was found to
be activated by high concentrations (>50 mM) of gluconate,
with the effect proportional to the length of the carbon chain (13).
This finding suggested to us the hypothesis that native anionic
phospholipids might be an endogenous mediator of this effect, because
like gluconate they have an anionic head with a hydrophobic tail.
Activation of KATP in cardiac cells by PI(4,5)P2 and Mg-ATP/PI has been recently reported (33, 34). All negatively charged intact phospholipids we tested possess the ability to activate KATP, including PS, PI, PI(4)P, and PI(4,5)P2, directly, and in the absence of magnesium and ATP. Mg-ATP and PI, therefore, are not critical requirements to activate the channel. Although Mg-ATP is not required to mediate activation by anionic phospholipids, the anionic lipid hypothesis can account for the well known Mg-ATP activation effect on KATPs. It has previously been reported that nonphosphorylating analogs of ATP, AMP-PNP and AMP-PCP, are incapable of increasing KATP activity, suggesting that Mg-ATP acts as a phosphoryl group donor in kinase-mediated phosphorylation reaction, although such kinases have not yet been identified (29). Our effectiveness sequence of PI(4,5)P2 > PI(4)P > PI suggests the possibility that in vivo, Mg-ATP potentiates the activity of KATPs by serving as a substrate for the phosphoinositide kinases (35) and aminophospholipid translocases that maintain sufficient membrane concentrations of PPIs to maintain KATP by increasing net negative membrane charge. PI(4,5)2 and phosphorylation of PI may perhaps be the important physiological regulators of KATP activity in this mechanism, a subject that requires further study.
Possible Molecular Mechanisms for the Anionic Lipid EffectDo
anionic phospholipids activate KATPs through an intermediate structure
or process or through a direct structural interaction with the channel?
Some components of PPIs, especially PI(4,5)P2, are known to
directly regulate various cellular proteins. One of the most important
proteins interacting with the anionic phospholipids is protein kinase
C, which uses Ca2+ as part of its signaling pathway. In our
cell-free patch clamp experiments, however, ATP was not included in the
perfusion solution to the cytoplasmic side of the membrane, and
Ca2+ was chelated by EGTA to less than 5 × 1010 M. Indeed, regardless of the presence of
ATP, raising the cytoplasmic Ca2+ concentration decreased,
rather than increased, the effect of PPIs. Thus, ATP and
Ca2+ are not required for the phospholipid effect on KATP,
making it unlikely that the protein kinase C signaling pathway is
involved in these effects of anionic lipids. Our observation that the
protein kinase C activator diacylglycerol was ineffective in activating KATPs is consistent with noninvolvement of protein kinase C. Likewise, the immediate requirement of other phosphorylation processes in maintaining KATP activity is unlikely because of the ATP-free conditions of our study. Another important protein group known to
interact with PI(4,5)P2 is the cytoskeletal regulating
proteins such as gelsolin. The possibility of involvement of
cytoskeletal structure in the KATP activity has been suggested (33).
The effects of PPIs remained even after application of high
concentrations of the cytoskeleton disrupters DNase I and cytochalasin
B (data not shown). Also, PS, which does not interact with most
cytoskeletal proteins (36), was effective in activating KATPs in our
experiments. Participation of the cytoskeleton network is evidently not
a requirement for the effect, although a modulatory role has not been
excluded.
The apparent importance of electrostatic interactions suggested by our data lead us to propose a molecular/physical model involving the channel protein (Fig. 5B). Kir channels that are not thought to associate with SUR (mIRK1, but hROMK1) were also activated by anionic phospholipids. This caused us to focus on the channel structure (Kir6.2) rather than SUR for the mechanisms of the effect. In our model, the positively charged residuals at the beginning of the C terminus of Kir6.2 are anchored or tethered by the electrostatic force of the anionic phospholipid head groups at the cytoplasmic face of the membrane. A decrease in the concentration of anionic phospholipids causes a release or conformation change of the tethered portion of the C terminus, triggering the formation of a gate that closes off the inner vestibule of the channel pore. This model is highly speculative, but protein-membrane electrostatic interactions at this portion of membrane proteins have been proposed previously (26, 27). The model also has value because it can account for the known experimental data such as the inhibitory effects of pH and di- and trivalent ions (by charge screening or neutralization), the anionic charge effectiveness sequence, and suggests possible explanations for such effects as trypsin (cleaving the gate) and Mg-ATP (increasing net negative membrane charge by phosphorylation of membrane phospholipids). This model was tested by the Kir6.2 mutant R176A,R177A, which reduced positive charge on the cytoplasmic tail. As the model predicted, the mutant channel activity was reduced and less sensitive to PPIs.
How widely might this protein-lipid interaction mechanism apply to ion channel activity in general? To date, KATP has been cloned from pancreatic cells (6, 7) and cardiac cells (9, 37); both require a sulfonylurea receptor and Kir6.2 to function. Kir6.2 is one member of the Kir family with functional and structural similarities, including a concentration of positive charges at the beginning section of the C terminus (Fig. 5A). Loss of channel activity has been generally found in most members of the Kir family, in both native channels (Refs. 38-40, for example) and expressed channels (41, 42). Our demonstration that PPIs activated native non-KATP inward rectifier (Fig. 1B) and also two cloned non-KATP inward rectifiers (Fig. 2) indicates that this mechanism may be general to all channels of this family. The higher effectiveness for more highly charged phospholipids suggests a role for lipid phosphorylation in this regulation. Thus, the PPIs, especially PI(4,5)P2 and phosphatidylinositol 3,4,5-triphosphate, along with enzymes maintaining them such as phosphoinositide kinases (35), may be important cellular regulators by maintaining a channel protein-anionic lipid interaction required to activate channels.
We thank Dr. Y. Tokuyama for providing the rat
pancreatic -cells; Drs. Y. Tokuyama and H. Yano in Dr. G. Bell's
lab for rKir6.2, rSUR, and hROMK1 clones; Drs. J. Kyle and L. Phillipson for the mIRK1 clone, B. Ye for technical assistance, Dr. C. T. January for reading and commenting on the manuscript, and D. Pittz
for secretarial help.