Intracellular ATP binding is required to activate theslowly activating K+ channel IKsYang Lia, Junyuan Gaob, Zhongju Lub, Kelli McFarlanda, Jingyi Shia, Kevin Bocka, Ira S. Cohenb, and Jianmin Cuia,1

aCenter for the Investigation of Membrane Excitability Disorders, and Cardiac Bioelectricity and Arrhythmia Center, Department of Biomedical Engineering,Washington University in St. Louis, St. Louis, MO 63130; and bDepartment of Physiology and Biophysics and the Institute for Molecular Cardiology, StateUniversity of New York, Stony Brook, NY 11794

Edited by Richard W. Aldrich, The University of Texas at Austin, Austin, TX, and approved October 11, 2013 (received for review August 24, 2013)

Gating of ion channels by ligands is fundamental to cellularfunction, and ATP serves as both an energy source and a signalingmolecule that modulates ion channel and transporter functions.The slowly activating K+ channel IKs in cardiac myocytes is formedby KCNQ1 and KCNE1 subunits that conduct K+ to repolarize theaction potential. Here we show that intracellular ATP activatesheterologously coexpressed KCNQ1 and KCNE1 as well as IKs incardiac myocytes by directly binding to the C terminus of KCNQ1to allow the pore to open. The channel is most sensitive to ATPnear its physiological concentration, and lowering ATP concentra-tion in cardiac myocytes results in IKs reduction and action potentialprolongation. Multiple mutations that suppress IKs by decreasing theATP sensitivity of the channel are associated with the long QT (in-terval between the Q and T waves in electrocardiogram) syndromethat predisposes afflicted individuals to cardiac arrhythmia and sud-den death. A cluster of basic and aromatic residues that may forma unique ATP binding site are identified; ATP activation of the wild-type channel and the effects of the mutations on ATP sensitivity areconsistent with an allosteric mechanism. These results demonstratethe activation of an ion channel by intracellular ATP binding, andATP-dependent gating allows IKs to couple myocyte energy state toits electrophysiology in physiologic and pathologic conditions.

ischemia | heart failure

Significant energy is required to sustain both the electrical andcontractile events that accompany each heart beat, suggest-

ing that the level of ATP is one key to normal cardiac functions.Not surprisingly, a reduction in ATP concentration ([ATP]) playsa key role in the pathogenesis and progression of ischemic heartdiseases, including heart failure. Intracellular ATP is importantnot only in providing energy (1) and as a substrate for proteinkinases (2), but also as signaling molecules to bind and modulateproteins. Only a handful of results have shown that intracellularATP serves as a signal for membrane channels and transporters(3). The best-studied example is the KATP channel (4, 5). Thischannel is inhibited in physiologic conditions by [ATP]s of 5–10mM (6), but when the ATP levels drop to submillimolar concen-trations, as in cardiac ischemia, the KATP channels open, shorteningthe action potential duration and providing metabolic protectionagainst the insult of ischemia (7). However, at normal physiologicconditions, whether and how ATP serves as a signal connecting theenergetic state of the cell to membrane excitability is still unknown.The slowly activating K+ channel IKs plays an important role

in controlling the action potential duration (APD) in cardiacmyocytes; it opens in response to depolarization to conduct po-tassium ions out of the cell, which contributes to repolarizationof the membrane, terminating the cardiac action potential andthereby the myocyte contraction. The IKs channel consists ofpore-forming KCNQ1 subunits and the single-transmembraneauxiliary subunits KCNE1 (8, 9). Loss-of-function mutations in ei-ther KCNQ1 or KCNE1 lead to prolongation of ventricular actionpotentials and long QT syndrome (LQTS) that manifests as QT(interval between the Q and T waves in electrocardiogram) pro-longation in the electrocardiogram. LQTS predisposes patients tocardiac arrhythmias that lead to syncope and sudden death (10).

Previous studies show that ATP is required for activation of IKschannels (11), but the molecular mechanism and the physiologicfunction of this ATP modulation is not known. It was the purposeof this study to investigate the mechanism by which ATP regulatedIKs. Our results show that ATP directly binds to the KCNQ1 proteinto regulate channel function at concentrations ([ATP]) close to thephysiologic intracellular [ATP] in cardiac myocytes. Our studiesalso reveal a unique ATP binding site and mechanism for modu-lation of ion channel function. Further, we found that several LQT-associated mutations alter IKs function by reducing ATP sensitivity.These results demonstrate that ATP regulation is vital for IKschannel function; a disruption of these regulations predisposes tolife-threatening cardiac arrhythmias.

ResultsATP-Dependent Activation of IKs Channels. We studied heterolo-gously expressed human KCNQ1/KCNE1 currents (hIKs) in inside-out membrane patches from Xenopus oocytes with application ofvarious intracellular ATP concentrations ([ATP]). Upon patchexcision, the current ran down in low [ATP] (Fig. 1A), consistentwith previous findings that the loss of native ATP in cytosolresulted in reduced channel activity (11). However, the current ranup in high [ATP], suggesting that the open probability of channels(Po) is not maximal with the native cytosolic [ATP] that is in-sufficient to saturate channel activation. Thus, a reserve of the IKschannels open in higher applied [ATP], resulting in currents largerthan those observed at patch excision (Fig. 1A). The steady-statecurrent amplitude increased with [ATP] with the EC50 at 1.7 mM(Fig. 1B), which is close to the physiological [ATP] in cardiacmyocytes (6). This result suggests that cardiac IKs could be sensitiveto the cellular energetic state, and fluctuations of [ATP] as in is-chemia could alter electrical properties via regulating IKs.

Significance

We show that intracellular ATP at physiological concentrationsacts as a signaling molecule to activate the slowly activating K+

channel IKs that regulates heart rate adaptation. ATP binding tothe pore-forming α-subunit of IKs, KCNQ1, allows the channelto open. Congenital mutations that reduce ATP binding orsubsequent opening of the IKs channel are associated with car-diac arrhythmias in human patients. Electrical abnormalities areoften the cause of fatality in cardiovascular diseases, includingischemia and heart failure, in which ATP level is reduced in car-diac cells. Our results open up new possibilities to study andmanage these diseases, and the ATP site provides a unique tar-get for therapies.

Author contributions: Y.L., J.G., Z.L., I.S.C., and J.C. designed research; Y.L., J.G., Z.L., K.M.,J.S., and K.B. performed research; Y.L. contributed new reagents/analytic tools; Y.L., J.G.,Z.L., and J.C. analyzed data; and Y.L., I.S.C., and J.C. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.1To whom correspondence should be addressed. E-mail: jcui@wustl.edu.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1315649110/-/DCSupplemental.

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The physiological importance of the ATP sensitivity of IKs issupported by our subsequent finding that a LQT-associated mu-tation Q357R in KCNQ1 (12) reduces ATP sensitivity, as shownby an increased EC50 of the response to [ATP] and the largerfraction of the current activated by applied high [ATP] (Fig. 1B).Consistent with previous studies (13), we found that Q357Rcoexpressed with KCNE1 (Q357R hIKs) showed a smaller currentamplitude, a slower activation time course, and a shift of the voltagedependence of activation toward more depolarized potentialscompared with the WT hIKs measured from whole-cell currents.Each of these changes in channel properties would decrease theability of IKs to participate in cardiac repolarization, resulting inprolongation of APD. Furthermore, application of a high [ATP](20 mM) during inside-out patch-clamp recordings of Q357R hIKsrestored the WT channel characteristics. Specifically, the currentamplitude increased three- to fivefold, sufficient to account for all ofthe reduction in the whole-cell current (SI Appendix, Fig. S1), andthe voltage dependence of channel activation shifted back towardless-depolarized voltages to superimpose on that of the WT hIKs(Fig. 1C). These results suggest that a decrease in ATP sensitivity ofthe IKs channel due to mutation Q357R can lead to LQT syndrome.KCNQ1 expressed alone without KCNE1 shows a similar

dose–response to [ATP] (Fig. 1D), indicating that the ATP de-pendence is an intrinsic property of KCNQ1 and not altered byKCNE1 association. IKs channels also require phosphatidylinositol4,5-bisphosphate (PIP2) for function (11, 14, 15) and are modulatedby calmodulin (CaM) (16, 17) and phosphorylation of residuesS27 and S92 in KCNQ1 by PKA (18, 19). However, the response ofhIKs currents to [ATP] did not change with reduced [PIP2], en-hanced [Ca2+], or mutations S27D/S92D that mimic phosphoryla-tion (14, 18) (Fig. 1D), indicating that ATP activates the channelindependently from these other intracellular regulatory molecules.

ATP Level Changes Affect Action Potentials in Cardiac MyocytesThrough Regulating IKs. To directly examine the role of ATP sensi-tivity of IKs in cardiac myocytes, we studied guinea pig ventricularmyocytes using whole-cell patch-clamp techniques. IKs currentsincreased with increasing [ATP] in pipette solutions (0–25 mM);a dose–response curve yielded an EC50 of 1.4 mM (Fig. 2 A and B).These results demonstrate that [ATP] regulates IKs in cardiacmyocytes as it does hIKs in oocytes with a similar EC50. To examinethe effects of ATP on action potentials (APs), we recorded APswith pipette [ATP] ranging from2 to 10mM (Fig. 2C andD, black).TheAPD shortened as pipette [ATP] was increased. Application ofchromanol 293B, an IKs-specific blocker, lengthened the APD (Fig.2 C and D, red); the lengthening was greater when pipette [ATP]was higher, from less than 10% at 2-mM pipette [ATP] to more

than 40% when pipette [ATP] is 10 mM, consistent with the ideathat ATP activates IKs to shorten the action potential. The channelactivity and APD are specifically dependent on [ATP] becauseATP added as a salt of K+ or Mg2+ had the same effects (Fig. 2).Taken as whole, these results demonstrate that variations in myo-cyte [ATP] can significantly alter APD by changing IKs magnitude.

ATP Binding in KCNQ1. ATP could activate the IKs channel byserving as the substrate for phosphorylation, binding to an as-sociated protein or directly binding to the channel proteins. Todistinguish these mechanisms, we first measured the ability ofvarious nucleotides to prevent hIKs current run-down due towashout of the native ATP after inside-out membrane patchexcision (11). GTP and a nonhydrolyzable ATP analog, 5′-ade-nylyl-β−γ-imidodiphosphate (AMP–PNP), in the intracellularsolution can sustain channel function similarly as ATP, and,furthermore, the dose–response curves of channel function onGTP and AMP–PNP are superimposed with that on ATP (SIAppendix, Fig. S2), whereas the rundown of hIKs currents becameprogressively faster when ADP and AMP were applied (Fig. 3A).Thus, ATP is not unique in activating the channel and phos-phorylation is not required. Furthermore, an ATP analog biotinphotoprobe, 2-azidoadenosine 5′-triphosphate 2′,3′-biotin-longchain-hydrazone (AB11) (20) can prevent channel run-down (SIAppendix, Fig. S3) and be photo–cross-linked to the KCNQ1protein (Fig. 3B), indicating that the nucleotide directly binds toKCNQ1 to modify hIKs channel function. We found that thechannels formed by coexpression of KCNQ2 and KCNQ3 do notrequire ATP for function (Fig. 3 B and C and SI Appendix, Fig.S4), and, correspondingly, AB11 cannot be photo–cross-linkedto the KCNQ2 or KCNQ3 proteins (Fig. 3B).To locate the ATP binding site in KCNQ1, we first studied

chimeric channels by transplanting the cytosolic C terminus toKCNQ2 and KCNQ3 to form Q2ctQ1 and Q3ctQ1 (SI Appendix,Fig. S5). Similar to hIKs, channels formed by the coexpression ofQ2ctQ1/Q3ctQ1 ran down after inside-out membrane patchexcision (Fig. 3C), but intracellularly applied ATP slowed therun-down (SI Appendix, Fig. S6), suggesting that the chimerasacquire ATP sensitivity and the C terminus of KCNQ1 containsthe ATP binding site. Because the potency of nucleotides inactivating hIKs correlates with the number of phosphates (Fig.3A), the channel may associate with ATP through electrostaticinteractions between basic residues and the negatively chargedphosphates of ATP. Interestingly, neutralization mutations ofseveral of the cytosolic basic residues are associated with LQTS.We performed a mutational scan to neutralize each of the basicresidues to Ala, LQT-associated mutation, or deletion in the N

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Fig. 1. ATP-dependent activation of IKs channels.(A) KCNQ1 + KCNE1 (hIKs) currents in inside-outpatches run down or up in 0 (red), 0.5 (green), and5 mM (blue) [ATP] from that immediately afterpatch excision (black, I0). Voltage pulses were +80mV from a −80-mV holding potential. (Right) It, I0tail current amplitudes. (B) ATP dose–response ofWT (black) and Q357R (red, also scaled to the WTcurrents at 20 mM ATP) IKs, and fits to the Hillequation (solid curves) with Hill coefficient 1 and1.1, and EC50 1.66 and 9.60 mM, respectively. (C )Currents of WT (Left) and Q357R (Center) hIKsrecorded from inside-out patches and G–V rela-tions after patch excision in solutions containingvarious [ATP]. Solid curves are fits of Boltzmannequation (Right; Materials and Methods) with V1/2

(mV) at 0.5, 5, and 20 mM [ATP], respectively: WT,23.7 ± 3.5 (black), 25.6 ± 3.2 (purple), 25.4 ± 5.0(blue); Q357R, 53.2 ± 2.5 (red), 30.8 ± 2.6 (green),and 26.2 ± 3.8 (cyan). (D) Normalized ATP dose–response of WT hIKs channel activation in controlsolution [PIP2] = 100 μM (black), 5 μM PIP2 (green),100 μM Ca2+ (red), and S27D/S92D hIKs in control solution (blue; Materials and Methods). Five and 100 μM [PIP2] are 50% and 100% of saturation for IKschannel activation, respectively (14). Dashed curves in B and C are the fittings of the model in Fig. 7A.

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terminus and the C terminus of KCNQ1 to examine which ofthese residues affected ATP sensitivity (Fig. 3D). We useda simplified ATP dose–response assay to examine ATP sensi-tivity of the mutant channels. We chose three ATP concen-trations, 0.5, 1.5, and 20 mM, at which the WT IKs channels areactivated to 20, 50, and 100% of saturation, respectively (Fig. 3D,Inset). If a mutation reduces ATP sensitivity, the ATP dose–re-sponse curve is expected to shift to higher ATP concentrationssuch that the fractional increments of channel activation at 0.5, 1.5,and 20 mM differ from those of the WT IKs. A more completedose–response curve would be measured once the mutation isidentified as positive for confirmation and further characterization.The results revealed three mutations, R380S, K393M, and

R397W, that are all associated with LQT and reduced the ex-pression of macroscopic hIKs currents and ATP sensitivity of thechannel (Figs. 3D and 4A and B; SI Appendix, Fig. S7). ResidueR380 is located in helix A downstream from the S6 gate of thechannel, whereas K393 and R397 are located in the linker be-tween helix A and helix B (Fig. 4B and SI Appendix, Fig. S8).Though each of these mutations reduced ATP sensitivity, acombined mutation R380S/R397W eliminated ionic current al-together, although channel expression in the plasma membranecould still be detected (Fig. 4 C and D). Furthermore, R380S/R397W also eliminated photo–cross-linking of the ATP analogAB11 (Fig. 4D). These results suggest that these three residuesare part of the ATP binding site; whereas each individual mu-tation reduces ATP binding, the combined mutations disrupt

ATP binding, resulting in the loss of channel function. Likewise,mutating each of these residues to negative charges, which couldrepel ATP, also eliminated hIKs and AB11 photo–cross-linking(Fig. 4 C and D). The known ATP binding sites in other proteinsalso contain aromatic residues to coordinate the adenine group(21). A mutational scan of each aromatic residue in helices Aand B and the A–B linker (SI Appendix, Figs. S8 and S9) iden-tified one mutation, W379S, that eliminated ionic current andAB11 photo–cross-linking of hIKs (Fig. 4 C and D), suggestingthat W379 also participates in ATP binding. Similar to R380S,K393M, and R397W, W379S is also associated with LQT syn-drome (22–25), further revealing the physiological importance ofATP modulation of IKs.

The Mechanism of ATP Regulation of IKs Channels. As a member ofthe Kv channel family, KCNQ1 is comprised of a voltage-sensingdomain (VSD) and a pore-gate domain (PGD; SI Appendix, Fig.S10). The IKs channel contains four KCNQ1 subunits with theVSDs surrounding a central pore across themembrane; in responseto membrane depolarization, voltage sensors move to trigger poreopening (26). Does ATP binding affect voltage sensor movements,pore opening, or the coupling between the two processes?We usedvoltage clamp fluorometry (VCF) to answer this question.Fluorescence signals from a fluorophore (Alexa 488 C5 Mal-

eimide) attached to the VSD (Materials and Methods; SI Appendix,Fig. S10) were recorded to monitor VSD movements, and ioniccurrents were simultaneously measured to show pore opening

Fig. 2. ATP activates IKs and shortens APs in cardiacmyocytes. (A) Voltage protocol and current traces atdifferent [ATP]s in the patch pipette in the absence(red) and presence (black) of chromanol 293B (10μM). (B) IKs vs. [ATP]. n = 5, error bars are SD.Smooth curve: fitting to the data from 1 to 25 mM[ATP] with Hill equation; Hill coefficient = 1 andEC50 = 1.4 mM. The 0 [ATP] data were not includedin the curve fitting because 0 [ATP] inside the cellcould not be achieved in live myocytes; the pre-dicted value of [ATP] for that magnitude of IKs (grayopen circle) is 0.64 mM. Potassium ATP at a concen-tration of 3 mM and 10 mM (red open circles) werealso used in the pipette solution to substitute forMgATP. (C) AP traces in the absence and presenceof chromanol 293B, with intracellular infusion ofATP (2–10 mM). (D) Summary of APD90 at various[ATP]. (Upper) The APD for each cell is plotted in theabsence and presence of chromanol 293B. (Lower)Average percentage increase in APD induced by chromanol 293B. Note a significantly larger APD prolongation induced by chromanol 293B at 5 or 10 mM ATPcompared with that at 2 or 3 mM ATP (P < 0.01, individual t tests). Potassium ATP at two different concentrations (2 mM and 10 mM, in gray) were used tosubstitute for MgATP in the pipette solution.

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Fig. 3. ATP binding to KCNQ1. (A) hIKs tail current amplitudeelicited by voltage pulses from −80 to +80 mV in intracellularsolutions containing 1.5 mM of various nucleotides: ATP (blue),GTP (green), AMP–PNP (pink), ADP (red), AMP (purple), and nonucleotide control (black). (B) Western blot of channel proteinspulled down with avidin beads after UV light photo–cross-linkingof the biotin-containing AB11 (Top) or after biotin treatment ofintact cells to detect expression in the membrane (Middle). Gbeta

is a cytoplasmic protein. Antibodies against KCNQ1 (for lanes ofuninjected, hIKs, and KCNQ1), KCNQ2 (KCNQ2), KCNQ3 (KCNQ3),and Gbeta (low bands) were used. (C) Whole-cell currents ofWT and chimeric KCNQ1 and KCNQ2/KCNQ3 channels (Upper)and time dependence of current amplitude after inside-outpatch excision without application of intracellular ATP (Lower).WT hIKs (black), WT KCNQ2/KCNQ3 (blue), and Q2ctQ1/Q3ctQ1(red). (D) Mutation scan of all cytosolic basic residues in KCNQ1.(Inset) Experimental design for ATP sensitivity screen. ATP dose–response curves of WT (red) and a hypothetical mutant hIKs withreduced ATP sensitivity (green). Three concentrations of ATP areused: 0.5, 1.5, and 20 mM.

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(15, 26) (Fig. 5A). The mutations that disrupt ATP binding,W379S and R380S/R397W (Fig. 4D), eliminated ionic currentsof KCNQ1 but did not abolish ΔF/F signals; moreover, thefluorescence–voltage (F–V) relationship and fluorescence signalare superimposed with that of the WT KCNQ1 (Fig. 5 B and C),indicating that ATP binding is not required for VSD movements.Next we tested if ATP is required for the coupling between theVSD movements and pore opening by measuring if opening thepore by the mutation L353K affected VSD activation (15).L353K is located in the S6 gate and the mutation locks thechannels constitutively open (15, 27), such that instantaneouscurrents were observed at every applied voltage, and these cur-rents were not inhibited by disrupting ATP binding (Fig. 6A andSI Appendix, Fig. S11). The opening of the pore facilitated theVSD activation via the VSD–pore coupling, resulting in a left-ward shift in the F–V relation by comparing the F–V relation ofWT and that of locked open L353K channels (Fig. 6B). Ourrecent study showed that this VSD–pore coupling is mediated byPIP2 bound to the channel, and depletion of PIP2 abolishes thecoupling and thus the L353K-induced F–V shift (15). However,when we depleted ATP binding by mutations, the F–V relationshipremained to be left-shifted by L353K, indicating that, unlike PIP2,ATP is not required for the coupling between VSD activation andpore opening. These results (Figs. 5 and 6) lead to a conceptualmodel for ATP-dependent activation of KCNQ1 and IKs channels(Fig. 7A and SI Appendix, Fig. S12) such that ATP binding to thechannel PGD is a prerequisite for the pore to open, but does notaffect VSD activation either directly or via the VSD–pore cou-pling. We were unable to detect fluorescence signals of the mutantchannels in the presence of KCNE1, but the model can fit theexperimental data of the hIKs - [ATP] dose response (Fig. 1B) andconductance–voltage (G–V) relationship of hIKs at various [ATP]s(Fig. 1C), suggesting that the coexpression of KCNE1 does notalter the fundamental mechanism of ATP-dependent activation.The lack of influence of KCNE1 on ATP-dependent activation isalso supported by the result that both KCNQ1 and hIKs channelsshowed a similar response to [ATP] (Fig. 1D).The mutation Q357R also does not alter the fundamental

mechanism of ATP-dependent activation, because the disruptionof ATP binding in the background of Q357R does not affect theF–V relation (Fig. 7B). Q357 is located just downstream from

the S6 gate toward the C terminus in KCNQ1 and away from theamino acid cluster that is important for ATP binding (Fig. 4B), yetit causes a reduction in ATP sensitivity of hIKs activation (Fig. 1B).Perhaps more strikingly, although the WT hIKs activationrequires ATP binding, the properties of hIKs activation, includingsteady-state G–V relation (Fig. 1C) as well as the time courseof voltage-dependent activation and deactivation (SI Appendix,Fig. S13), do not change with [ATP], whereas Q357R renders theG–V relation dependent on [ATP] (Fig. 1C). Although Q357Ralters the slope of the F–V relation compared with the WTKCNQ1 (Figs. 5C and 7B), the mutation does not change thekinetics of fluorescence signals (Fig. 7C) or properties of the hIKsG–V relation at high [ATP] (Fig. 1C), suggesting that the changein the ATP dependence of G–V relations may not be due to achange in VSD movement. It is also unlikely that all thesechanges are brought about by a direct influence of the mutationon ATP binding because, unlike Q357R, mutations that directlyaffect ATP binding do not cause a shift of the G–V relation todifferent voltages at the native [ATP] (Fig. 7D). Interestingly,a simple change in the equilibrium constant of pore opening inthe proposed model (Fig. 7A) can recapitulate the mutation-caused changes in ATP sensitivity (Fig. 1B) and ATP dependenceof G–V relations (Fig. 1C). These results suggest that the lack of[ATP] dependence in the WT hIKs activation properties (Fig. 1Cand SI Appendix, Fig. S13) is due to a strong mass reaction pullingthe VSD-activated and ATP-bound channels to the open state(Fig. 7A), whereas a reduction of the equilibrium constant for thistransition by the mutation Q357R unmasks the ATP dependenceof activation properties of the channel.

DiscussionIKs currents repolarize ventricular action potentials in the heart,and the IKs channel is an important regulator of heart rhythm.Here we show that intracellular ATP at physiologic concen-trations (0.5–7.5 mM) acts as a signaling molecule to activate IKschannels and shorten cardiac action potentials (Figs. 1 and 2).We also identify a cluster of residues important for ATP bindinglocated in the C terminus of KCNQ1 (Figs. 3 and 4) and themechanism by which ATP binding alters channel function (Figs.5–7). These results indicate that ATP binding to the cytosolicdomain promotes pore opening via an allosteric mechanism.The IKs ATP binding site has a low affinity for ATP (EC50 =

1.7 mM; Fig. 1) and does not select between ATP and GTP (Fig.

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Fig. 4. ATP binds in the C terminus of KCNQ1. (A) ATP dose–response curve ofWT and key mutant IKs channels. (B) The KCNQ1 motifs important for ATP in-teraction (Upper) and EC50 of ATP dose–response of mutant hIKs channels (Lower).Due to insolubility of ATP above 20mM, dose–response for somemutants did notreach saturation, leading to underestimated EC50. Asterisk indicates no currentexpression. (C) Whole-cell currents of mutant hIKs. (D) Western blot to detectAB11 labeling of mutant IKs. (Upper) AB11 labeling of mutant IKs and Gβ in thewhole-cell lysate to indicate similar inputs. (Lower) Western blot probing forbiotinylated mutant IKs and Gβ in the membrane. Gβ is a cytoplasmic protein.

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Fig. 5. ATP is not required for VSD movement. (A) VCF recordings of the WTand ATP-binding disruptive mutant KCNQ1 channels. Whole-cell current(Upper) and fluorescence signal (Lower) of WT (black), W379S (red), andR380S/R397W (green) in response to increasing voltages. (B) Superimposedfluorescence signal (Left) of WT (black), W379S (red), and R380S/R397W(green) in response to a series of voltage pulses with increasing voltages. Thecurrents activation time constants in response to voltages (Right). (C) Steady-state fluorescence changes vs. voltage (F–V).

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3 and SI Appendix, Fig. S2), but in the cell, ATP is present ata much higher concentration than GTP so that in physiologicconditions it can be considered an ATP binding site. Consistentwith the low affinity and nonselective features, sensitivity tonucleotides with different phosphates and mutagenesis (Figs. 3and 4) suggest that ATP is primarily coordinated by a cluster ofbasic charges, R380, K393, and R397, via electrostatic interactionwith phosphates, whereas hydrophobic residues, including W379,stabilize the nucleoside moieties. We made a mutation scanningon aromatic residues in a part of the cytosolic domain flankingthe basic residues to identify W397 (SI Appendix, Figs. S8 and S9).Our study cannot rule out the possibility that other residues mayalso contribute to ATP binding. Our search of the sequence ofKCNQ1 did not find the conserved ATP binding motifs WalkerA [GXXXXGK(T)] or Walker B (R/KXXXGXXXLhhhhD),where X represents any of the amino acids and h denotes a hy-drophobic amino acid (28). KATP channels are formed by theinward rectifier channel subunit Kir 6.2/6.1 and regulatory sul-phonylurea receptor subunit (29), and ATP binds to the Kirsubunit. Both N and C termini for ATP binding have beenidentified by mutagenesis. Positively charged residues play animportant role interacting with negatively charged phosphategroups of ATP, with K185 and R201 in the C terminus of onesubunit and R50 in the N terminus of another directly interactingwith ATP (30). In other known ATP-binding ion channels, thecystic fibrosis transmembrane conductance regulator containsWalker motifs for ATP binding (31), whereas P2X receptors bindATP with extensive hydrophilic interactions from amino acids lo-cated in two different structural domains (32). Unlike in Kir6.2 orP2X, the putative ATP interacting residues in KCNQ1 are locatedin a close cluster (Fig. 4). The 3D structure of the cytosolic domainof the KCNQ1 channel at the putative binding site has not beensolved; it is not clear how the ATP binding site relates to the poreor other parts of the channel.In addition to voltage, IKs channels require both ATP and PIP2

to fully activate (11) (SI Appendix, Fig. S14), but our current andprevious studies (14, 15) show that these two signaling moleculesactivate the channel with distinct molecular mechanisms. First, inthis study, 100 μM PIP2, which is saturating for IKs channels(EC50 5 μM) (14), was always present in intracellular solutionsduring patch-clamp recordings to prevent reduced channel ac-tivities due to PIP2 level variations. GTP and nonhydrolysable

ATP analog AMP–PNP could activate the channels with thesame doses as ATP (Fig. 3A and SI Appendix, Fig. S2), ruling outthe possibility of ATP activating the channel via hydrolysis for thesynthesis of PIP2. Second, the PIP2 binding site was identified inbetween the VSD and the PGD (15), whereas the putative ATPbinding site is located in the C terminus toward the S6 gate (Fig. 4).The mutations of the residues in the ATP binding site reduceATP sensitivity but have no effect on the PIP2 sensitivity of IKsactivation (SI Appendix, Fig. S15), supporting the distinctness ofthe two binding sites. Third, PIP2 activates the channels by medi-ating the coupling between VSD movements and pore opening(15), whereas ATP activates the channels by opening the pore(Figs. 5–7). The last distinctive properties between PIP2 andATP-dependent activation is that PIP2 sensitivity of the channelsis modulated by the association of the KCNE1 subunit, the EC50of PIP2-dependent activation decreased more than 100-fold withKCNE1 association (14), whereas ATP-dependent activation isnot affected by whether KCNE1 is present (Fig. 1).Our results show that the IKs channel is a voltage- and ligand-

activated K+ channel, with voltage sensor and ATP binding in-dependently regulating pore opening. Likewise, HCN (33) andBK (34) channels are also voltage- and ligand-activated channels,both with voltage-sensing domains and intracellular ligand bindingsites. A unique property of the IKs channel is that both voltage andATP are required for the pore to open, whereas BK channels canbe activated by either voltage or Ca2+ binding (35), and HCNchannels can be activated by voltage alone (33). Nevertheless, it isnot known if at extreme voltages the IKs channel can be activatedin the absence of ATP. For both BK and HCN channels, ligandbinding shifts the voltage dependence of channel opening (33, 35).However, ATP binding only enhances current amplitude withoutaltering the G–V relation of WT IKs channels (Fig. 1).Heart rate changes with physical activity and emotion. The mod-

ulation of the IKs channel by ATP at physiologic concentrations di-rectly links electrical activity and heart rhythm to the energetic stateof cardiac myocytes. In normal physiologic conditions, the cellularATP level in cells is tightly maintained, but submembrane ATPconcentration has been shown to fluctuate with Na+-K+ ATPaseactivity during action potential firings (36). In cardiac myocytes the

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Fig. 6. ATP is not required for VSD–PGD coupling. (A) VCF recordings of WTKCNQ1 (black), W379S (red), and R380S/R397W (green) with and withoutL353K. (B) Steady-state fluorescence change vs. voltage (F–V). Solid curvesare fits of a Boltzmann equation with V1/2. WT (black solid): −53.5 ± 1.1 mV;W379S (red solid): −56.2 ± 0.8 mV; R380S/R397W (green solid): −50.3 ± 0.9mV; L353K (black open): −72.3 ± 0.8 mV; W379S/L353K (red open): −76.9 ±0.9 mV; R380S/R397W/L353K (green open): −70.9 ± 1.1 mV.

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Fig. 7. ATP is required for pore opening. (A) The scheme of voltage andATP-dependent activation of hIKs channels. Voltage sensor movements aresimplified as one ATP-independent transition between the resting (V black)and activated (V red) state; the transition of the pore from closed (P black) tothe open (P red) state can happen only after voltage sensor activation andATP binding. K1: 300 M−1, L(V): 4.2 × 10−4exp(0.94VF/RT) (V, voltage; F,Faraday constant; R, gas constant; and T, absolute temperature), and K2:1,340 and 360 for WT and Q357R IKs, respectively, are obtained from fittingsto ATP dose–responses (Fig. 1B) and G–V relations at various [ATP] (Fig. 1C).(B) VCF recordings and F–V relations of Q357R and Q357R/R380S/R397W. (C)Activation (Upper) and deactivation (Lower) kinetics of fluorescence for bothWT and Q357R. (D) G–V relations of hIKs WT (black), R380S (green), K393M(blue), and R397W (red). Smooth curves in B–D are Boltzmann fits to the data.

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change of APD and frequency is accompanied by changes inNa+-K+

ATPase and Ca2+ pump activity (37), which may also alter sub-membrane ATP concentrations. Therefore, ATP may act as a con-tinuous regulator of heart rhythm during normal physiologic events.In the failing human heart, ATP concentration is decreased by

35–40% (38). Complex ventricular arrhythmias are presented inabout half of heart failure patients, and sudden death is common(39). Studies showed prolongation of the ventricular action po-tential both in animal models and in the human heart failure (40,41). Many ion currents underlie these changes in APD, andamong them IKs was found to be decreased in a canine model ofheart failure, whereas IKr remained unchanged (40). However,the mechanism of this IKs reduction has not been determined.Our results provide a possible mechanism. In addition, whereasduring severe ischemia, ATP concentration falls to submillimolarlevels and combined with the increase in ADP levels open KATPto reduce APD and prevent excessive depolarization, thus main-taining excitability, in acute ischemia the ATP level is reduced by50% (41) but still well above the range where activation of theKATP channel would be expected to occur (42), and a prolongedQT interval is observed (43). This phenomenon, which suggeststhe involvement of repolarization currents, could be explained bya reduction of the IKs channel activity. The increase of APD inacute ischemia would allow L-type Ca2+ channels to open fora longer time to compensate for the reduced Ca2+ channel ac-tivities (44) and maintain contraction. Finally, the activation of IKsby ATP also presents a unique therapeutic opportunity such that

at physiologic [ATP] a reservoir of IKs remains unopened (Fig. 1);inducing these remaining closed channels to open would provideadditional repolarizing currents that should shorten the APD inall circumstances in which a congenital or acquired LQT syn-drome exists, independent of its origins.

Materials and MethodsAll mutations were generated using overlap-extension PCR (14) and verifiedby sequencing. Xenopus laevis oocytes were injected with 0.05–20 ng ofcRNA per oocyte, and macroscopic currents were recorded from wholeoocytes or inside-out patches in 2–4 d (14). Voltage-clamp fluorometry andWestern blot experiments were as previously described (15). Guinea pigsingle left ventricular myocyte were isolated and recorded for APs and IKscurrents as described (45).

The relative conductance was determined by measuring tail currentamplitudes at indicated voltages. The G–V and F–V relationships were fittedwith the Boltzmann equation: G

Gmax= 1

1+exp−zeðV−V1=2 Þ=kT, where z is the numberof the equivalent charges, V1/2 is the voltage at which the channel is 50%activated, e is the elementary charge, k is Boltzmann’s constant, and T is theabsolute temperature.

More detailed experimental procedures and data acquisition and analysescan be found in SI Appendix, Materials and Methods.

ACKNOWLEDGMENTS. We thank Mark Zaydman for technical help in the VCFrecording. This study was funded by National Institutes of Health Grants R01-HL70393 and R01-NS060706 (to J.C.), and R01-HL094410 and HL111401 (to I.S.C.).

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