cytoplasmic atp-sensing domains regulate gating

Citation: Bennetts, B, Rychkov, G, Ng, H, Morton, C, Stapleton, D, Parker, M and Cromer, B 2005, 'Cytoplasmic ATP-sensing domains regulate gating of skeletal muscle ClC-1 chloride channels', Journal of Biological Chemistry, vol. 280, no. 37, pp. 32452-32458.

CYTOPLASMIC ATP-SENSING DOMAINS REGULATE GATING OF SKELETAL MUSCLE CLC-1 CHLORIDE CHANNELS. Brett Bennetts1, Grigori Y. Rychkov2, Hooi-Ling Ng1, Craig J. Morton1, David Stapleton3, Michael W. Parker1* & Brett A. Cromer1* From: 1St. Vincent's Institute, 9 Princes St. Fitzroy, Victoria, 3065, Australia; 2 The University of Adelaide, Adelaide, South Australia, 5005, Australia 3 Bio21 Molecular Science and Biotechnology Institute, The University of Melbourne, Parkville, Victoria 3010, Australia. Running title; ClC-1 regulation by ATP Address correspondence to: Brett A. Cromer, St. Vincent's Institute, 9 Princes St. Fitzroy, Victoria, 3065, Australia. Tel:+61 3 9288 2480, Fax:+61 3 9416 2676, e-mail; [email protected]. *Joint senior authors ClC proteins are a family of chloride channels and transporters that are found in a wide variety of prokaryotic and eukaryotic celltypes. The mammalian voltage-gated chloride channel ClC-1 is important for controlling the electrical excitability of skeletal muscle. Reduced excitability of muscle cells during metabolic stress can protect cells from metabolic exhaustion and is thought to be a major factor in fatigue. Here we identify a novel mechanism linking excitability to metabolic state by showing that ClC-1 channels are modulated by ATP. The high concentration of ATP in resting muscle effectively inhibits ClC-1 activity by shifting the voltage-gating to more positive potentials. ADP and AMP had similar effects to ATP but IMP had no effect, indicating that the inhibition of ClC-1 would only be relieved under anaerobic conditions such as intense muscle activity or ischaemia, when depleted ATP accumulates as IMP. The resulting increase in ClC-1 activity under these conditions would reduce muscle excitability, thus contributing to fatigue. We show further that the modulation by ATP is mediated by cystathionine-β-synthase-related (CBS) domains in the cytoplasmic C-terminus of ClC1. This defines a function for these domains as gating-modulatory domains sensitive to intracellular ligands, such as nucleotides, a function that is likely to be conserved in other ClC proteins. Skeletal muscle has a high and variable demand for energy, in the form of ATP, and has elaborate systems to maintain the ATP supply. During intense exercise, however, ATP supply 1

may not keep up with demand and ATP concentration can decrease rapidly. In fast-twitch fibres ATP can drop to below 25% of resting concentration within 25 seconds (1), a rate of ATP consumption that, if it continued, would deplete all ATP within a further 10 seconds. As the majority of ATP is consumed by the sarcoplasmic reticulum (SR)1 Ca2+-ATPase pumping Ca2+ back into the SR after each Ca2+-activated contraction (2), complete ATP depletion would lead to a rise in cytoplasmic calcium, rigor and calcium-dependent damage (3,4). This does not normally occur because force generation and ATP consumption decrease during exercise, compromising short-term function but protecting cells from complete metabolic exhaustion. This process is well known as fatigue but the factors contributing to fatigue remain controversial. Direct reduction in force generation by the contractile apparatus is thought to be a factor early in fatigue (3,5) but a significant reduction in ATP consumption only occurs with a reduction in SR Ca2+ release (and consequent reuptake) that occurs late in fatigue, correlating with ATP depletion (3). Indeed, ATP depletion and the concomitant increase in cytoplasmic Mg2+ directly reduce the activity of the SR Ca2+ release channel and may contribute to fatigue (6). A reduction in the electrical excitation that stimulates SR Ca2+ release is also thought be a major contributor to fatigue (7). Activation, or excitation, of skeletal muscle is initiated at the neuromuscular junction by nicotinic acetylcholine receptor-mediated postsynaptic Na+ currents, leading to membrane depolarization. These currents are large so the threshold potential for voltage-gated Na+ channels is reached with high probability, leading to action potential initiation. Membrane repolarization

ClC-1 regulation by ATP

following an action potential is mediated by voltage-gated K+ channels and the voltage-gated Cl- channel, ClC-1 (8), both of which are activated by membrane depolarization. A significant proportion of ClC-1 channels remain open in resting muscle, acting as a barrier to depolarization that must be overcome for successful action potential generation. Action potentials are propagated along the surface membrane and into the internal network of membrane tubes (Ttubules) where they activate voltage sensors in the dihydropyridine receptor, stimulating Ca2+ release from the SR and muscle contraction. During intense exercise, run-down of Na+ and K+ gradients in the restricted volume of the T-tubules is thought to reduce action potential propagation and contribute to fatigue (7). ATP-sensitive K+ (KATP)1 channels (9) provide a mechanism for linking excitability to metabolic state but, although KATP channels reduce excitability and protect cardiac muscle cells during ischemia (10), they do not appear to contribute significantly to force reduction during fatigue in skeletal muscle (11). ClC-1 channels are important regulators of skeletal-muscle excitability, as highlighted by muscle hyper-excitability due to mutations in ClC1 in congenital myotonia (12) and aberrant splicing of ClC-1 in myotonic dystrophy (13,14). To date there is no evidence, however, that ClC-1 differentially regulates excitability during metabolic stress. Like other ClC family members, the ClC-1 protein comprises a large membrane-embedded channel domain followed by two cytoplasmic cystathionine β-synthase related (CBS)1 domains (15) (Fig. 1) and is thought to form homodimers with an ion conducting pore within each monomer, as revealed by the structure of a bacterial ClC (16). Voltage-dependent gating of ClC-1 has two components that are both activated by depolarization, fast or protopore gating that occurs independently in each pore and slow or common gating that operates on both pores simultaneously (17). Little is known about the mechanism of common gating but its temperature dependence (18,19) indicates that it involves substantial conformational changes. Perhaps not surprisingly, mutations of ClC-1 that lead to dominantly inherited myotonia (Thomsen’s disease) tend to affect the voltage dependence of common gating and cluster at the dimer interface (12). More surprisingly, modifications of the cytoplasmic CBS domains also affect common gating, or in some cases abrogate channel function (20-22). Although the functional role of CBS domains in ClC proteins

remains unclear their importance to channel function is underlined by disease causing mutations. In humans mutations in the CBS domains of ClC-1 underlie congenital myotonia (23), in ClC-2 generalized idiopathic epilepsy (24), in ClC-5 hypercalciuric nephrolithiasis (25), in ClC-Kb Bartter syndrome (26) and in ClC-7 infantile malignant osteopetrosis (27). In the present work we show that ATP modulates common gating of ClC-1, a mechanism that is likely to reduce the excitability of skeletal muscle and contribute to fatigue during severe metabolic stress. Furthermore, we show that the regulatory effect of ATP on channel-gating is mediated by the CBS domains of ClC-1, defining a functional role for these domains. MATERIALS AND METHODS Channel expression and site-directed mutagenesis - Human ClCN1 was expressed using a pCIneo (Promega) mammalian expression vector, as detailed previously (19,28). Human embryonic kidney (HEK293T)1 cells (American Type Culture Collection, Rockville, MD, USA) were transiently transfected with a mixture of the ClCN1 construct and pEGFP-N1 (Clontech) reporter plasmid at a molar ratio of 3:1 using Effectene (Qiagen), according to the manufacturer's specifications. Transfected cells were later identified by reporter-plasmid driven expression of the green fluorescent protein. Mutations were introduced into ClCN1 using the Quickchange (Stratagene) mutagenesis technique. The mutagenic primers were designed to incorporate silent restriction sites to aid in the identification of successfully mutated plasmid DNA. Mutant clones that were identified using restriction analysis were subsequently verified by sequencing. Electrophysiology - Patch-clamp experiments were conducted at room temperature (23 ± 1 °C) in the whole-cell configuration using an Axopatch 200B patch-clamp amplifier (Axon Instruments, Foster City, CA, USA) and associated standard equipment. Cells were continuously superfused with bath solution containing (mM): NaCl, 140; CsCl, 4; CaCl2, 2; MgCl2, 2; Hepes, 10; adjusted to pH 7.4 with NaOH. The standard pipette solution contained (mM): CsCl, 40; Caesium glutamate, 80; EGTA-Na, 10; Hepes, 10; adjusted to pH 7.2 with NaOH. Stock solutions of the following nucleotide salts: ATP di-sodium, ATP magnesium, ADP disodium, AMP di-sodium, IMP di-sodium, GTP 2


lithium and adenosine-3’,5’-cyclic-monophosphate (cAMP) 2-amino-2-(hydroxymethyl)-1,3propanediol (Tris) were stored at –20°C and diluted in pipette solution on the day of each experiment. Solutions of 5 mM adenine and adenosine in pipette solution were made fresh on the day they were used. Patch pipettes were pulled from borosilicate glass, and typically had resistance of 1-2 MΩ when filled with the above pipette solution. Series resistance did not exceed 4 MΩ and was 85-95 % compensated. After rupturing the cell membrane and achieving the whole-cell configuration, no less than 5 minutes was allowed for the pipette solution to equilibrate with the intracellular solution before current recordings were made. Currents obtained at a sampling frequency of 10 kHz were filtered at 5 kHz, and collected using Axograph (Axon Instruments, Foster City, CA, USA) software. Data analysis - Data were analyzed using Axograph 4.9 (Axon Instruments, Foster City, CA, USA) and GraphPad Prism 4.0 (GraphPad software, San Diego, CA, USA) software. The voltage dependence of channel inactivation was determined by measuring tail-current relaxation at a constant voltage of –100 mV for a period of 75 ms, after a test pulse at voltages ranging from –140 to +120 mV. Under certain conditions (for example in the presence of intracellular ATP), and for certain mutations current activation at positive potentials was slowed. For this reason the time of the test pulse was varied between 300 and 800 ms, depending on the degree of channel activation; a manipulation that in itself has previously been shown to have little effect on the determination of channel open probability (29). Between test sweeps the cell membrane was clamped at a holding potential of –30 mV for a period of 2 s. Current relaxations during the –100 mV tail pulse were fit with an equation of the form

# & # & I ( t ) = A1 exp% t " ( + A2 exp% t " ( + C $ 1' $ 2'

!

(1) where I is the whole-cell current, t is time, τ1 and τ2 are the time constants of the fast and slow components of current relaxation, respectively, and A1, A2 and C are the amplitudes of the fast, slow and steady-state components of the current, respectively. Instantaneous current amplitudes at the start of the –100 mV pulse (I(0)) were determined by extrapolation of this function. To obtain the voltage dependence of the apparent channel open-probability (Po), normalised

instantaneous current amplitudes were fit with a modified Boltzmann distribution of the form

Po (V ) = Pmin +

1 ' Pmin & # 1 + exp$ &$V 1 ' V #! k ! " " %% 2

(2) where Pmin is an offset, or a minimal open probability at very negative potentials, V is the membrane potential, V1/2 is the half-maximal activation potential, and k is the slope factor. To determine the voltage dependence of the slow or common gate that acts simultaneously on both protopores of a ClC-1 dimer, a 400 µs pulse to +170mV was inserted before the –100 mV tail pulse, to maximally activate the fast or protopore gate. Instantaneous tail-current amplitudes, determined as described above were fit with equation (2) to yield Po(V) for the common gate. Open probability of the protopore gate was determined by dividing the apparent channel open probability by the open probability of the common gate. This data was fit with equation (2) to yield Po(V) curves for the protopore gate. All data are expressed as means ± SEM. Homology modelling and nucleotide docking - The CBS domains from ClC-1 were individually modelled on the crystal structure of IMPDH from Streptococcus pyogenes (PDB entry 1ZFJ) using COMPOSER, a component of Sybyl 6.92 (Tripos Inc.). The individual domain models were then assembled with reference to the atomic coordinates of the IMPDH CBS dimer to form CBS 1/2 dimers in SwissPDB viewer (30), followed by energy minimisation using Sybyl6.92. Two alternative models were built based on different alignments in one strand of CBS 2. The quality of both models was assessed, and found to be satisfactory using Verify3D (31). The model with the best Verify3D score was chosen as the preferred model for in silico docking studies, and is presented here. AutoDock 3.0 (32) was used to explore the interaction of AMP and ATP with the entire surface of this ClC-1 CBS model. The Lamarckian genetic algorithm was used to produce 100 conformations of each ligand, which were clustered within a root mean square deviation of 1.8 Å. Docked conformations of the ligands were inspected using VMD version 1.8 (33).

RESULTS 3


Intracellular ATP regulates ClC-1 common gating - During whole-cell patch clamp measurements of heterologously expressed ClC-1 channels, we observed that the voltage dependence of ClC-1 gating shifts to more negative potentials over the first few minutes after rupturing the cell membrane, indicating that ClC-1 gating might be modulated by a diffusible factor present in the cytoplasm that is diluted upon membrane rupture. Given the high ATP turnover of active skeletal muscle, we postulated that this factor might be ATP or one of its metabolites, perhaps linking excitability to metabolic state via regulation of ClC-1 channels in a similar manner to regulation of KATP channels (9,34). We found that ATP did indeed have a marked effect on the voltage dependence of ClC-1 gating, predominantly affecting common gating with little effect on protopore gating (Fig. 2). Inclusion of 5 mM ATP, similar to the concentration in resting muscle, in the intracellular solution during whole-cell patch clamp recordings altered common gating such that the midpoint of the voltage sensitivity curve (V1/2) shifted to less negative potentials by approximately 50 mV and the fraction of channels that remained open at strongly negative potentials (Pmin) was reduced approximately 6-fold (Fig. 2). The nonhydrolysable ATP analogue, ATP-γs had a similar effect to ATP on common gating, indicating that this effect does not involve ATP hydrolysis and is likely to be due simply to ATP binding. Although ATP is often complexed with Mg2+, equimolar Mg2+ did not influence the effect of ATP on common gating. The effect of ATP on the voltagedependence of ClC-1 common gating was concentration dependent, with a half-saturation concentration of approximately 1 mM (Fig. 2B), similar to the concentration reached in fast-twitch fibres after 25 seconds of intense exercise (1). During moderate exercise, although ATP turnover increases markedly, a variety of systems maintain ATP at close to resting levels of 5-7 mM, minimizing any effect on ClC-1 voltage sensitivity. Slight ATP depletion during exercise does, however, show up as proportionally larger increases in its metabolites adenosine 5’diphosphate (ADP) and particularly adenosine 5’monophosphate (AMP). Differential regulation by these metabolites could be a factor in ClC-1 regulation as it is in other systems (34). We found, however, that AMP had a similar effect to ATP on ClC-1 common gating (Fig. 2), as did ADP. Furthermore 1 µM AMP, a high level physiologically, had no influence on the effect of 5

mM ATP on common gating, indicating that the ClC-1 common gating is not sensitive to the AMP:ATP ratio. We also investigated the effect on ClC-1 common gating of the metabolic breakdown products of AMP, adenosine and adenine. The effect of adenosine on common gating was similar to that of ATP or AMP, at both 5 mM and 1 mM (Fig. 2D). These results demonstrate that the presence of phosphates attached to adenosine are not important for the effect on common gating and reiterate that hydrolysis of a nucleotide highenergy phosphate bond is not necessary for this effect. The effect on common gating of 5 mM adenine was much less than that of adenosine, ATP or AMP (Fig. 2D), indicating that the ribose ring of adenosine is important for this effect. Although these data are useful in understanding ligand specificity, they do not indicate a physiological role for adenosine and adenine in regulating ClC-1 as the concentrations in skeletal muscle are well below the mM levels used here. Furthermore, their concentrations do not increase greatly during metabolic stress, due to the very high AMP deaminase activity that directs AMP breakdown to IMP rather than adenosine (35,36). Taken together, these data indicate that although ClC-1 common gating is clearly modulated by adenosine nucleotides it is unlikely to be affected by the small changes that occur in the concentration of these nucleotides during moderate exercise. Under the anaerobic conditions of severe metabolic stress such as intense sprint exercise or ischemia, however, ATP can be depleted to less than 1 mM (1,35), accumulating as inosine 5’monophosphate (IMP), the product of AMP deamination (37,38). We found that 5 mM intracellular IMP had no effect on ClC-1 common gating (Fig. 2A). Furthermore the effect of a mixture of 2.5 mM IMP and 2.5 mM ATP on common gating was essentially indistinguishable from 2.5 mM ATP alone. These data indicate that under conditions of severe metabolic stress when ATP is broken down to IMP, the drop in ATP concentration would lead to a shift in the voltage dependence of ClC-1 common gating towards more negative potentials. We also investigated the potential role of other nucleotides. As for IMP, inclusion of 5 mM guanosine 5’-triphosphate (GTP) in the intracellular solution did not affect the voltage dependence of ClC-1 common gating (Fig. 2A), indicating some specificity for adenosine nucleotides. Inclusion of 5 mM adenosine 3’, 5’cyclic monophosphate (cAMP) in the intracellular 4


solution reduced Pmin slightly, similar to sub-mM AMP, suggesting that AMP cyclisation reduces binding affinity (Fig. 2A). ATP modulation is mediated by CBS domains of ClC-1 - Having shown that intracellular ATP regulates ClC-1 activity, we then investigated whether this might occur by direct binding to the intracellular side of ClC-1. Tandem CBS domains in the carboxy-terminus of ClC-1 (15,39), seemed to be the best candidates for a potential ATPbinding site as they are the only defined cytoplasmic domains, modification of them affects common gating (20-22) and CBS domains occur in a number of nucleotide-sensitive proteins. Consistent with this postulate, Scott and colleagues (40) have shown recently that CBS domains of AMP-activated protein kinase and IMP dehydrogenas-1 bind nucleotides and regulate enzyme activity and that a piece ot the C-terminal tail of ClC-2, including the two CBS domains, is able to bind ATP. To examine the possibility that CBS domains of ClC-1 might bind ATP we built homology models based on the structure of a CBS dimer from bacterial inosine monophosphate dehydrogenase (IMPDH) (41) (Fig. 3). The validity of this structure as a modelling template is supported by the observation that CBS domains from mammalian IMPDH can substitute for those of ClC-1 while retaining channel function (21). In silico docking of AMP and ATP to this model identified a putative binding pocket between CBS 1 and 2 (Fig. 3B) lined by residues T636, P638 and S651 from CBS 1 and H847, L848 and V860 from CBS 2. The adenine base is predicted to bind deep within this pocket, close to L848, with the ribose ring at the top of the pocket, close to T636 and H847. Mutagenesis of individual residues lining the putative ATP-binding site reduced (T636A, P638A or H847A) or abrogated (L848A) the effect of ATP on the voltage dependence of common gating (Fig. 4, Table 1), supporting both our model of an ATP-binding site at the CBS domain dimer interface and the hypothesis that this site mediates ATP regulation of ClC-1 common gating. Mutations of other residues, A849 that faces away from the putative binding pocket and S651 that faces into the putative binding pocket but is not predicted to interact directly with bound ATP, had little effect on ATP sensitivity (Fig. 4, Table 1), consistent with the details of our model. The fact that functional ClC-1 channels were observed for all of these mutants indicates a specific effect on ATP modulation rather than non-specific

disruption of the CBS domain structure, as such disruption also appears to disrupt channel function (21). We also tested the effect of a myotonia associated mutation, G859D (12,23), in CBS 2 but functional ClC-1 channels were not observed, presumably due to disruption of the CBS domain structure. Interestingly, all of the mutations that reduced the effect of ATP also shifted the voltagedependence of common gating in the absence of nucleotide to be more like that of wt ClC-1 in the presence of ATP (Fig. 4). DISCUSSION We have demonstrated here a novel regulation of ClC-1 voltage sensitivity by intracellular adenosine nucleotides that would result in significant inhibition of ClC-1 activity by the high ATP concentration in resting and moderately exercising muscle. This inhibition is only likely to be alleviated during anaerobic conditions such as intense exercise or ischemia when marked ATP depletion occurs, accumulating as IMP (1). The physiological significance of ATP regulation of ClC-1 remains to be elucidated but the obvious prediction from our results is that, similar to KATP channels in cardiac muscle (10), increased ClC-1 activity due to ATP depletion during metabolic stress would stabilize the membrane potential and reduce skeletal muscle excitability. Given the significantly increased ClC1 activity at resting membrane potentials in the absence of ATP, excitability could be reduced by preventing initiation of action potentials as well as reducing their amplitude and time-course. Reduced excitability would protect cells both from the toxic effects of prolonged depolarization and, by reducing ATP demand, from complete ATP depletion and Ca2+-dependent damage. At the same time this reduced excitability is likely to contribute to fatigue. This mechanism may be most significant during intense sprint-type exercise in fast-twitch muscle fibres where ATP depletion is greatest (1) and expression of ClC-1 is also greatest (42). ClC-1 appears to be present primarily in the surface membrane of muscle fibres and is more densely clustered around the neuromuscular junction (42,43), a location where it will have the greatest effect on action potential initiation and propagation. ATP modulation of ClC-1 channels is one of many regulatory mechanisms that are likely to contribute to fatigue but, as with metabolic pathways, modulation of such an early step in the excitation-contraction 5


pathway of skeletal muscle may be the most effective and efficient means of regulating ATP consumption. Muscle acidosis, another consequence of anaerobic conditions that has long been thought to contribute to fatigue, has also been reported to increase ClC-1 activity (44) so could contribute to fatigue by reducing muscle excitability using the same argument as we have outlined above. This possibility and the general concept that acidosis contributes to fatigue are contradicted, however, by observations that acidosis preserves the excitability of depolarized muscle fibres and Ttubule membranes by decreasing the Clconductance of the muscle sarcolemma (45-48). The relative contribution of acidosis and ATP depletion to fatigue and the extent to which ClC-1 is involved clearly requires further investigation. Using molecular modeling and in silico docking, we have identified a putative ATPbinding site in a cleft between the two C-terminal cytoplasmic CBS domains of ClC-1. We found that mutation of residues predicted to interact with ATP at this site, reduced or abrogated the effect of ATP on ClC-1 voltage dependence, whilst mutation of residues in the local area of the proposed binding site that were not predicted to be important for binding had little or no impact on ATP modulation. These results clearly demonstrate the role of the CBS domains in mediating the modulatory effect of ATP and support our model of the ATP-binding site (Fig. 3). Scott and colleagues (40) have shown recently that CBS domains act as nucleotide-binding regulatory domains in a number of enzymes. They also showed that when expressed as a GST-fusion, a piece of the C-terminal tail of ClC-2, including the two CBS domains, can bind ATP but the functional effect of this binding was not tested. The results we have presented here show for the first time that CBS domains of a ClC channel regulate channel activity in response to binding of an intracellular ligand, in this case ATP. We have shown further that this regulation occurs by shifting the voltage dependence of common gating. It has been shown that mutations within the CBS domains of ClC proteins affect common gating (12,21) but physiologically relevant regulatory roles for these domains have not been identified previously (21). We have now identified such a role for the CBS domains of ClC-1. It is not clear whether CBS domains have a similar role in other ClC proteins but there are a couple of reports in the literature of adenosine nucleotides affecting ClC channel activity. Firstly,

Vanoye and George (49) showed that ATP was required for continued ClC-4 activity. A sizeable part of this effect was due to ATP hydrolysis suggesting a mechanism involving protein phosphorylation. It was unclear whether ATP affected ClC-4 gating, trafficking, or other factors. More recently, Niemeyer and colleagues (50) showed that replacement of intracellular ATP with AMP increased the rate of activation and deactivation for ClC-2. Whether this effect was due to ATP hydrolysis was not tested but the effect was reduced by an epilepsy-linked G715E mutation close to two defined protein kinase A phosphorylation sites, T655 and T691 (51), in a large loop between the two CBS domains. There is no direct evidence in either of the above cases that the effects of nucleotides on channel function involve the CBS domains or changes in the voltage dependence of gating. Thus it remains to be elucidated whether the underlying mechanism is related to the one we have described here for ClC1. As voltage can be sensed exclusively across the membrane, the involvement of the cytoplasmic CBS domains in the voltagedependence of ClC-1 common gating suggests intimate interactions between the CBS domains and the membrane inserted channel domain (21,52). ATP may exert its effect on common gating by allosterically modulating the interaction between the CBS and channel domains and hence gating within the channel domain itself. A similar allosteric mechanism could explain the effect of CBS mutations that shifted common gating in the absence of nucleotide to be more like wt in the presence of ATP, as well as reducing the effect of ATP (Fig. 4). Alternatively, ATP could exert its effect by competitively disrupting an interaction between the CBS domains and cytoplasmic loops from the channel domain that act as alternate ligands for the ATP-binding site. In this model a mutation in the ATP-binding site could simultaneously disrupt binding of both ligands, mimicking the effect of ATP binding at the same time as eliminating the effect of ATP, consistent with the mutational effects we have observed. In support of this model, mutations on the surface of CBS2 (21), close to but not directly involved in the putative ATP-binding site, and in one of the cytoplasmic loops of the channel domain (53,54) shift the voltage-dependence of common gating to more positive values, perhaps also by disrupting interactions between the CBS and channel domains. The veracity of this hypothesis will only become clear when detailed structural information 6


showing interactions between channel and CBS domains becomes available. Based on our results showing that the CBS domains of ClC-1 mediate ATP modulation of common gating, we propose that these domains can be thought of as channel regulatory domains sensitive to intracellular ligands, analogous to the variety of cytoplasmic regulatory domains associated with K+ channels (55). ClC proteins comprise a large family of Cl--selective ion channels and transporters (56) that are widespread in prokaryotic and eukaryotic cells and are involved in a diverse range of physiological functions (8) but CBS domains are conserved in most ClC proteins (Fig. 1). We speculate that the

function of these domains as channel regulators sensitive to intracellular ligands is also likely to be conserved and that whilst their ligand-sensitivity may vary, nucleotide sensitivity could be a recurring theme. The bacterial ClC homologues from Escherichia coli and Salmonella typhimurium for which high resolution structures have been solved lack CBS domains (16), but other prokaryotic ClC proteins have CBS domains (Fig. 1), making them prime targets for structural studies. Curiously some prokaryotic ClC proteins have additional tandem USPA domains, which have also been implicated in ATP binding (Fig. 1) (57).

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FOOTNOTES Acknowledgements - This work was supported by a grant from the National Health and Medical Research Council of Australia (NHMRC) to M.W.P. and B.A.C. M.W.P. is an NHMRC Senior Principal Research Fellow. G.Y.R. is an Australian Research Council Research Fellow. We thank Graham D. Lamb for constructive comments and advice on this work. 1

The abbreviations used are: SR, Sarcoplasmic Reticulum; KATP, ATP-sensitive K+ channels; CBS domain cystathionine -synthase related domain; HEK 293, human embryonic kidney 293. FIG. LEGENDS Fig. 1. Structural conservation in the ClC family. Schematic diagram illustrating domain organization of various ClC proteins, drawn to scale shown at bottom right of figure. Shown are the human isoforms of the mammalian ClC family members as well as homologues from various representative organisms. Stippled boxes indicate homology to the predominantly transmembrane ‘voltage-gated chloride channel’ motif, open boxes represent CBS domains and diagonally striped boxes represent USPA domains. Escherichia coli and Salmonella typhimurium ClC’s are those for which 3-dimensional crystal structures have been solved.

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Fig. 2. Nucleotide dependence of ClC-1 gating. ClC-1 open probability (A) overall and separated into (B) common gating and (C) protopore gating components, plotted as a function of membrane potential with (∆) standard intracellular solution, or including 5 mM () GTP, (◊) IMP, (◆) cAMP, () AMP or (●) ATP . Solid lines represent fits of equation 2 to the experimental data points. D, Half-saturation voltages (V1/2) for common gating plotted as a function of ligand concentration for () AMP, (●) ATP, (◊) adenosine or (◆) adenine. Solid lines represent fits of rectangular hyperbolae to the experimental data points that give half-saturation concentrations of 1.2 ± 0.5 mM and 1.1 ± 0.2 mM, and maximum ∆V1/2 of 52 ± 8 mV and 60 ± 4 mV for AMP and ATP respectively. E, Minimal open probability (Pmin) of common gating plotted as a function of nucleotide concentration. Symbols are the same as in panel D. Solid lines are straight lines connecting data points from the same data set. In all panels data points are means from n=4-6 cells, error bars are S.E.M. Fig. 3. Homology modeling of ClC-1 CBS domains. A, CBS domain sequence alignments. CBS domains from selected CBS-domain containing proteins are shown, including ClC-1. All proteins shown are human except for IMPDH, which is from Streptococcus pyogenes. ‘AMPK γ2’ represents AMPactivated protein kinase γ2 subunit and CBS represents Cystathionine β synthase. Residue numbers are; CBS 1 IMPDH 95-150, ClC-1 607-662, ClC-2 582-637, ClC-Kb 549-604, ClC-5 590-645, ClC-7 613673, AMPK γ2 274-328 and CBS 416-469; CBS 2 IMPDH 158-211, ClC-1 819-871, ClC-2 788-840, ClC-Kb 626-678, ClC-5 681-733, ClC-7 723-776, AMPK γ2 355-409 and CBS 481-544. Amino acid sequences were aligned with ClustalW and formatted as previously described(58). Shading is as follows; highly conserved residues, black; conserved nonpolar residues, green; conserved uncharged polar residues, orange; conserved positively charged residues, blue; conserved negatively charged residues, red. Boxed residues indicate residues mutated in human diseases. Red letters indicate residues mutated in this study. B, Model of ClC-1 CBS dimer built by homology with IMPDH CBS dimer using the alignment shown in a). AMP, docked as described in methods section, and residues lining the putative ATP/AMP binding pocket are shown as sticks coloured by atom. This image was prepared using VMD version 1.8 (33). Fig. 4. Voltage and nucleotide dependence of CBS domain mutants of ClC-1. A-F. Common gating of mutant ClC-1 with no nucleotide in the pipette solution (), with 5mM AMP () or with 5mM ATP () in the pipette solution. All abscissa are voltage in mV, with values given on panels E and F. All ordinate values are open probability. Mutants are A. T636A, B. P638A, C, S651A, D. H847A, E. L848A and F. A849V. In all panels data points are means from n = 3-6 cells, and error bars are S.E.M. Solid lines represent the fit of equation (2) to the experimental data points. Dashed lines and dot-dashed lines represent Po(V) for WT ClC-1 in the absence of nucleotide and with 5 mM ATP, respectively, in the pipette solution, as shown in Fig. 2A. G-H. Half-saturation voltages (V1/2) for common gating curves shown in panels A-F plotted against nucleotide concentration for intracellular G. AMP and H. ATP. Symbols are () WT ClC-1 and mutations () S651A, () T636A, (∆) P638A, (∇) L848A, () H847A and (●) A849V. Straight lines connect data points from the same data set. Data points are means from n=3-6 cells, error bars are S.E.M.

FIGURES.

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Fig. 1

Fig. 2

Fig. 3

Fig. 4