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We explored the structural basis of voltage sensing in HCN1 hyperpolarization-activated channels by examining the relative orientation of the voltage-sensor and pore domains. The opening of channels engineered to contain single cysteine residues at the extracellular ends of the voltage-sensing S4 (V246C) and pore-forming S5 (C303) domains is inhibited by formation of disulfide or cysteine:Cd2+ bonds. As Cd2+ coordination is promoted by depolarization, the S4-S5 interaction occurs preferentially in the closed state. The failure of oxidation to catalyze dimer formation, as assayed by Western blotting, indicates the V246C:C303 interaction occurs within a subunit. Intriguingly, a similar interaction has been observed in depolarization-activated Shaker Kv channels at depolarized potentials but such an intrasubunit interaction is inconsistent with the X-ray crystal structure of Kv1.2, wherein S4 approaches S5 of an adjacent subunit. These findings suggest channels of opposite voltage-sensing polarity adopt a conserved S4-S5 orientation in the depolarized state that is distinct from that trapped upon crystallization.
The HCN cation channels are unusual among voltage-gated cation channels in that they activate upon membrane hyperpolarization. This enables them to open following repolarization of the action potential, where they generate an inward Na+ current that contributes to the spontaneous pacemaker depolarization in cardiac myocytes and certain neurons (41, 46). Despite their atypical polarity of gating, HCN channels are homologous to the six-transmembrane-segment (S1-S6) depolarization-activated Kv channels. Thus, in both HCN (34, 45) and Kv channels (1, 6, 28, 30, 32, 37) the S4 transmembrane helix shows a highly conserved pattern of positively charged amino acids at every third residue; this positive charge is critical in sensing transmembrane voltage changes that lead to channel activation. Similarly, there are notable sequence similarities between HCN and Kv channels in the S5, S6 and reentrant P loop motif that form the ion-conducting pore (3, 19, 34, 45). The basis of the opposite polarity of voltage gating in the face of this overall conservation of architecture is, thus, intriguing.
The motions of the positively charged S4 transmembrane helices during voltage sensing that lead to channel gating are still controversial, despite over two decades of research on voltage-sensitive channels [inclusive of ion channels across the super-families of Kv, NaV and CaV; for reviews see (17, 49)]. From this wealth of data three quite different putative models of voltage-sensing are emerging, broadly grouped according to the translational motion of S4 through and relative to the transmembrane electric field. The ‘transporter model’ involves limited translational motions (~2-4 Å), often accompanied by a twist of the S4 helix, through a narrow, focused electric field created by deformations of, and aqueous crevices into, the lipid membrane around the S4 helix (2, 6, 7, 13, 14, 20, 39, 48, 51, 52). In a second model type, the ‘helical screw model’, the S4 helix translates ~5-14Å (depending on the modeled angle of tilt of S4) perpendicular to the plane of the lipid bilayer, a motion often accompanied by a 180° rotation of the helix (4, 9, 12, 22, 26, 38, 53). Finally, the ‘paddle model’, largely derived from a series of crystal structures and functional current analyses from the MacKinnon lab, depicts S4, in concert with transmembrane helix S3b (together forming the paddle structure), moving from a prone/tilted position to an upright/vertical position in the membrane electric field, with a translational motion of ~15-20Å (23, 33, 43).
Even amongst the limited members of the hyperpolarization activated HCN channel family no single, consensus model fits the different S4 motions observed, and data exemplifying characteristics of each of the three models of S4 voltage sensing movements have been described. Thus, in the sea urchin spHCN channel S4 motions resemble the helical screw model (35); whilst motions in the plant hyperpolarization-activated KAT1 channel (21, 24) suggest a paddle model (although S4 translational distances are shorter at ~12-15Å). In both the bacterial MVP channel (47) and mammalian HCN channels (5)(50) the data are most readily explained by the transporter model wherein S4 undergoes a limited translational motion in the confines of aqueous crevices.
One approach that can shed light on dynamic rearrangements in channels is through the assay of disulfide bond formation or metal ion coordination between pairs of cysteine residues localized in different regions of the channel. Depending on the thermal energy present, Cys-Cys disulfide bridges only arise if the Cys residues are within 15 of each other (10), while Cd2+ coordination between 2 or more Cys residues further restricts the molecular proximity to <6 (11, 42). With this approach, it has been shown that the extracellular end of the Kv S4 lies within 6 of the extracellular end of S5 of a neighboring subunit, suggesting a close intersubunit proximity (8, 25, 36). However, there is also evidence that similar residue positions on Kv S4 and S5 within the same subunit interact to coordinate Cd2+, suggesting a close intrasubunit proximity. For example, Elliott et al. (18) reported a state-dependent change in the orientation of S4 relative to S5. Using tandem dimers of concatenated Shaker channel subunits to fix the stoichiometery of inserted Cys residues on S4 and S5, they showed that both intra- and intersubunit S4 to S5 disulfide bonds could form upon exposure to the catalytic agent CuPhen. Further, by inference from these tandem dimer experiments, they showed that in the resting state of the Kv channel, when the membrane is hyperpolarized, the top of S4 lies near the top of S5 in an adjacent subunit, enabling the formation of an intersubunit Cd2+ coordination site. In contrast, following channel activation in response to depolarization, the orientation of S4 changes so that its extracellular end now lies near the extracellular end of the S5 segment in the same subunit, permitting the formation of an intrasubunit Cd2+ coordination site. Furthermore, using the hyperpolarization-activated KAT1 channel, the Jan laboratory performed elegant protein-helix-packing studies and provided evidence for close atomic proximities for three sets of residue pairings between S4 and S5 (21, 24). They suggest that two of these pairings are intrasubunit whereas the third is intersubunit. However, these S4 to S5 intra- and intersubunit interactions are inferred from a model based on the Kv1.2 crystal structure (31) and thus do not provide direct evidence to discriminate between intra- and intersubunit interactions.
Here we explore the proximity of the extracellular ends of S4 and S5 in a minimal HCN1 channel (HCN1-R) engineered to remove all but one endogenous cysteine residue (C303) that is located near the extracellular end of S5 and is essential for channel function (5). We find that when a cysteine was substituted for valine 246 at the external, N-terminal side of the HCN1-R S4 segment, the resulting channels demonstrate robust formation of disulfide bonds or Cys:metal ion bridges. Generation of either of these linkages stabilizes the closed state of the channel, shifting voltage-gating to more hyperpolarized potentials. Consistent with this, the rate of modification of gating by Cd2+ is enhanced when the membrane is depolarized so that the channels are in the deactivated, closed state. Biochemical experiments support the hypothesis that these functional effects are mediated via formation of intrasubunit interactions between V246C and C303. These results suggest HCN and Kv channels (c.f 18) can adopt similar conformations at depolarized potentials, in which the externally facing ends of S4 and S5 on the same subunit lie in close proximity. This implies that the relative orientation of the voltage-sensor and pore domains in the deactivated conformation of HCN channels resembles the orientation of these domains in the activated conformation of Kv channels.
Our experiments were performed in the background of a near cysteine-free channel, HCN1-R, which has a single endogenous Cys residue at position 303, as previously described (5). In brief, HCN1-R has a truncation at the end of S6, resulting in the deletion of the entire cytoplasmic C-terminus and removal of 6 out of 12 endogenous Cys residues in full-length HCN1. Five of the six remaining Cys were replaced using the following substitution mutations: C55S, C298I, C318S, C347S and C374T. Mutation of C303 in the S5 segment to any one of the other 19 amino acids failed to yield functional channels. HCN1-R displayed normal hyperpolarization-activated gating and was resistant to modification by a variety of sulfhydryl-reactive reagents (for more information see 5). An individual Cys substitution, V246C was made at the external NH2-terminal end of the S4 region of HCN1-R using PCR mutagenesis. All constructs were subcloned into the pGEM-HE vector (29) and verified using dideoxy chain termination sequencing. RNA was transcribed from NheI-linearized DNA using a T7 RNA polymerase (Message Machine, Applied Biosystems). RNA concentration was measured from the optical density ratio at 260 nM and 280 nM wavelengths (44) using a DU640 spectrophotometer (Beckman, CA).
Xenopus laevis oocytes were harvested according to a Columbia University approved protocol (PI#366G CU#2928). 50 nl of cRNA (0.5 µg/ul) were injected into each oocyte and cells were maintained in Barths Solution (Specialty Media) in the absence or presence of 0.15 mM DTT as indicated. Media was changed every 24 hours.
Two-electrode voltage clamp (TEVC) recordings were routinely performed 48 hours after oocyte injection. Data was acquired using an OC-725C voltage-clamp amplifier (Warner Instrument Corp.), filtered at 1 kHz using an 8-pole low pass Bessel filter (Frequency Devices) and sampled at 2 kHz using an ITC16 interface (InstruTech Corp.) controlled by Pulse software (HEKA Elektronik). No linear leak subtraction was applied to current recordings. In representative sweeps, the capacity transients are truncated for clarity. The extracellular solution contained (in mM): 112 KCl, 2 MgCl2, 10 HEPES, pH 7.4 (KOH). Microelectrodes were filled with 3 M KCl and had resistances of 0.5-3 MΩ. Ag-AgCl grounding wires were isolated from bath solutions by 3M KCl / 2% agar bridge electrodes. All recordings were performed at room temperature (21-24° C).
In all voltage clamp experiments, the holding potential was -20 mV. To measure steady-state gating parameters, tail current amplitudes were determined upon repolarization to the holding potential after stepping for 3 s to hyperpolarizing test potentials applied in −10 mV intervals from −25 mV. The relationship between peak tail current and test potential was plotted and the resulting current voltage (I-V) relationship was fitted with the Boltzmann equation (equation 1):
where A1 is an offset due to the holding current, A2 is the maximal tail current amplitude, V1/2 is the midpoint activation voltage and s is the slope factor of the relation (mV). Mean tail current activation curves were obtained by subtracting the offset (A1) from individual current-voltage curves and then normalizing by the maximal amplitude (A2). The normalized data were then averaged among different experiments and the mean normalized tail I-V relation fitted with the Boltzmann equation.
To determine the state dependence of the kinetics of inhibition by Cd2+, current amplitudes were monitored in response to 3 s steps to −125 mV applied at different frequencies during the application of Cd2+. A protocol in which a strong 3 s hyperpolarization was applied once every 4 s was used to determine the rate of Cd2+ block when channels were largely in the open state cycle (channels open for ~75% of the duty cycle). A protocol in which a 3 s hyperpolarization was applied once every 24 s was used to measure the rate of Cd2+ block when channels were biased towards the closed state (channels closed for 87.5% of the duty cycle). Peak currents were measured before, during and after application of Cd2+ (at the indicated concentrations) in the bath solution and plotted against the duration of the Cd2+ application. The measured currents were normalised to the control current prior to Cd2+ perfusion. The Cd2+ inhibition (coordination) time constant was determined by fitting a single exponential to the current decay on application of Cd2+. The Cd2+ unblock (recovery) time constant was determined by fitting a single exponential to the growth of current (Cd2+ unblock) on wash-off of applied Cd2+.
Analyses were performed using PulseFit (v8.5, HEKA Elektronik) and Microcal Origin software (v6.0, Microcal Software, Inc.). Statistical analysis was performed using paired or unpaired Student's t-tests and one-way ANOVA as indicated. Differences were considered significant if p was less than 0.05.
Forty-eight hours after cRNA injection, batches of 20-50 oocytes were washed with 2×5 ml of DTT-free ice-cold Barths solution, incubated at room temperature for 3 min in Barths containing either no addition or CuPhen (30 µM, prepared as described below), followed by 2×5 ml washes in ice cold Barths. Cells were then immediately disrupted under a nitrogen atmosphere by sonication (CP130 Ultrasonic processor, Cole Parmer) in 20 µl per cell of ice cold, nitrogen purged (bubbled with N2 for 2 hr on ice before use) CHAPS lysis buffer that was prepared fresh on day of use and contained: 1% CHAPS, 20 mM NEM, 2 mM EGTA, 2 mM EDTA, 100 mM NaF, 10 mM Na2MoO4, 50 mM Tris base, 2 mM Na3VO4, 10 mM Na4P2O7, 1 mM PMSF, pH adjusted to 7.0 with HCl and with one “CompleteMini” protease inhibitor tablet per 7.5 ml final volume. Following sonication the lysate was mixed at 4 °C for 90 min on a rotary shaker. Solubilized protein was recovered by centrifugation (4×5 min at 13000 RPM in an Eppendorf microfuge). Lysates were stored at −80 °C until analyzed by PAGE and Western Blotting.
Aliquots of lysates corresponding to 0.3 oocytes were denatured in PAGE loading buffer (62.5 mM Tris-HCl pH6.8, 1mM EDTA-Na2 5 % sucrose, 0.0125 % bromophenol blue, 2.5 % SDS) at 95-100 °C for 2 min then separated on 12% polyacrylamide gels using standard Laemmli electrophoresis buffers. To determine whether any higher MW bands on the gels represent multimeric proteins formed by cross-linking of monomers through disulfide bonds we ran reducing gels in which the loading buffer was supplemented with 10 mM DTT. After separation, proteins were transferred to PVDF membranes using an alkaline (10 mM 3-[cyclohexylamino]-1-propane-sulfonic acid pH 11 plus 0.01 % SDS), low current (90 mA, 20 hours), transfer protocol. All electrophoresis steps were performed using a mini-protean 3 system (Biorad).
Following transfer, the membranes were washed 3 times in 1× TBS followed by 15 s in methanol. After air-drying for 15 min, the positions of the Dual Color molecular weight markers were recorded using a rabbit antigen pen (Alpha Diagnostics). Membranes were rewetted by immersion in methanol (15 s) followed by 3 washes in 1× TBS. Membranes were then blocked overnight at 4 °C followed by a further 90 minutes at room temperature. All subsequent incubations and washes were at room temperature. Blocking and binding was performed in 1×TBS-T 2%BSA (1× TBS supplemented with 0.05 % Tween-20 and 2 % BSA). Washes were performed with either 1× TBS or 1× TBS-T 0.2% BSA (1×TBS supplemented with 0.05 % Tween-20 and 0.2 % BSA) as indicated below.
Membranes were incubated with the rabbit anti-HCN1 primary antibody (APC-056 obtained from Alomone Labs and applied at 100 ng/ml for 60 min) and with the HRP-conjugated goat anti-rabbit secondary antibody (20320 from Alpha Diagnostics applied at 2 ng/ml) with 6 washes in 1× TBS-T 0.2% BSA following each antibody application. The blot was developed using SuperSignal West Femto (Pierce Biotechnology) and imaged using Labworks version 184.108.40.206 and an Epichem3 Darkroom (UVP BioImaging Systems). Sequential images were collected between 0.1 and 3000 s at approximately half log intervals of time using a 2×2 binning and 0× gain. The longest exposure that did not result in saturation was chosen for densitometric analysis using LabWorks. The density of bands in sample lanes were corrected for background by subtracting the density of size and position matched areas in lanes where uninjected lysate samples were run. Images for display were exported from LabWorks at a bin range of 0-255 with no other image processing then imported into Adobe Illustrator version 10.0.3 (Adobe Systems) where they were scaled and the contrast inverted but the image not otherwise altered.
A sequence alignment between the Kv1.2-Kv2.1 chimera (33) and mHCN1 was generated based on a combination of a primary protein sequence alignment program (Clustal W), transmembrane topology predictions (TMHMM, TM-pred, SOSUI, http://ca.expasy.org/tools/), and visual inspection. Modeler (8v1, http://salilab.org/modeller/) was used to generate the homology model of the HCN1 channel, based on the sequence alignment and the crystal structure of Kv1.2-Kv2.1 chimera (2R9R) (33).
Unless otherwise indicated, all reagents were from Sigma Aldrich and were of the highest available purity. Cadmium chloride was stored at 4 °C as a 1M stock made with double distilled water. Copper (II)-phenanthroline (CuPhen) was prepared fresh on day of use from stock solutions of CuSO4 and Phenanthroline that were each stored at −20 °C. After combining the reagents to final concentrations of 30 µM CuSO4 and 90 µM Phenanthroline the solution was incubated at room temperature for at least 10 min then used within 4 hr. DTT, NEM, Na4P2O7 and “CompleteMini” protease inhibitors (Roche Diagnostics) were dissolved on the day of use into water, ethanol, water and lysis buffer, respectively. All other reagents were prepared as stock solutions and kept at 4 °C except for PMSF and Na3VO4 which were kept at −20 °C. SDS (BP1311-1 nuclease and protease free) and methanol (BP1105-4 Biotech Grade) were from Fisher Scientific. Dual color molecular weight markers were from Biorad. Fraction V protease-free BSA (03-117-332-001) was from Roche.
We focused our studies on the HCN1-R channel, in which all 12 endogenous cysteine residues were conservatively mutated (to either serine, threonine or isoleucine), except for Cys303 which is located in the S5 transmembrane helix and is necessary for channel function (see Methods and 5). Exposure of oocytes expressing HCN1-R to the reducing agent DTT for 3 min produced no functional changes in HCN1-R currents. Thus, the midpoint voltage of activation (mean V1/2 = -77.6 mV, n = 4; see Figure 1a) and slope factor (mean s = 7.9 mV) of the tail current activation curve before exposure to DTT are not different (p>0.05 – see legend to Figure 1) from the respective values after exposure to the reducing agent (V1/2 = -79.3, s = 8.3 mV, n = 4; see Figure 1a). The failure of DTT to alter channel properties suggests that C303 may fail to form intersubunit disulfide bonds spontaneously.
V246C channels (containing two Cys residues, V246C and C303, per subunit) displayed a more negative V1/2 (mean V1/2 = -90.6 mV, n = 10) and reduced steepness of activation (mean s = 13.4 mV) compared to HCN1-R under control conditions. Importantly, and in contrast to HCN1-R, application of DTT significantly enhanced V246C channel opening, shifting the V1/2 to more positive potentials by approximately +10 mV and increasing the steepness of the activation curve (V1/2 = -80.7 mV and s = 10.1, n = 10; p<0.001 for both parameters; Figure 1b). These results suggest that V246C channels spontaneously form a disulfide bond that inhibits channel opening, and that the reduction of this bond by DTT reverses this inhibitory effect. Indeed the voltage dependent gating of V246C channels in the presence of DTT is very similar to that of HCN1-R channels, indicating that the main effect of the V246C mutation is to promote disulfide bond formation, rather than to alter the energetics of channel gating by itself.
To investigate further the ability of V246C channels to form a disulfide bond, we applied Copper (II)-Phenanthroline (CuPhen), a weak oxidizing reagent known to promote Cys-Cys disulfide bridges in proteins where unbridged Cys residues lie in sufficiently close proximity. We first pretreated HCN1-R and V246C channels with DTT (0.15 mM in culture medium - see Methods) to reverse any spontaneous disulfide bridges and then applied CuPhen (30 µM, for 2 minutes). HCN1-R channels showed no response to the reagent, with the mean V1/2 and slope under control conditions (-73.2 mV and 8.8 mV, n = 4) similar to values determined after exposure to CuPhen (-73.3 mV and 8.5 mV, n = 4; p>0.05; Figure 2a), suggesting that C303-C303 intersubunit disulfide bonds could not form, even upon addition of the catalytic agent. In V246C channels, however, CuPhen produced a robust inhibitory effect on channel opening, shifting the V1/2 by approximately 20 mV to more hyperpolarized potentials, the opposite of the effect seen with DTT (mean V1/2 before and after exposure to CuPhen were -75.1 mV and -95.9 mV, n = 5; p<0.01; Figure 2b). Evidence that this effect of CuPhen is due to disulfide bond formation comes from the finding that subsequent perfusion of DTT was able to reverse the effects of CuPhen on channel activation, with the V1/2 shifting back to close to its original control value (mean V1/2 = -82 mV, n = 5; Figure 2b).
The above observations suggest that one or more pairs of Cys residues in V246C channels must lie in close proximity when the channels are closed. Furthermore, the fact that CuPhen treatment of previously reduced channels produces a larger voltage shift (-20 mV) than that seen upon acute DTT treatment of channels maintained in the absence of DTT (+10 mV) suggests that the level of spontaneous disulfide bond formation in V246C channels is incomplete.
Although the formation of a disulfide bond suggests the reactive sulfhydryls are in close proximity, cross linking between residues wherein the mean center to center distances are up to 15 apart have been observed depending on the intrinsic flexibility and mobility of the protein due to thermal fluctuations (10). To characterize more precisely the proximity of Cys residues in V246C we examined the effects of extracellular Cd2+ on channel function (Figure 3), as high-affinity coordination of metal ions by two or more Cys sulfhydryl groups occurs only when the distance between Cys β-carbons is <6 (11, 42).
HCN1-R currents were unaltered by a >2 minute application of 30 µM Cd2+, with no change in the tail current activation curve (mean control V1/2 = -75.9 mV, n = 5 versus mean with Cd2+ V1/2 = -74.1, n = 5; non-significant difference p>0.05 with paired Student's t-test; Figure 3a). This lack of effect of Cd2+ suggests that the C303 residues in each of the four HCN1-R subunits do not come into sufficient proximity to trap and coordinate Cd2+, in agreement with the failure of these channels to form disulfide bonds. In contrast, V246C channels, were very sensitive to Cd2+, showing a significant ~20 mV hyperpolarizing shift in their activation curves in response to a 2 minute application of 2 µM Cd2+ (mean control V1/2 = -78 mV, n = 4, versus mean with Cd2+ V1/2 = -97.1mV, n = 4; significant difference p<0.01 with paired Student's t-test), a concentration 15-fold lower than those having no effect on HCN1-R (Figure 3b). The hyperpolarizing effect of Cd2+ on channel activation is similar to the effect seen with disulfide bond formation. Thus, our data are consistent with the view that V246C channels contain pairs of Cys residues in close proximity and that either formation of disulfide bridges or Cd2+ coordination between these residues has an inhibitory effect on channel opening.
Does the distance between the Cys residues that coordinate Cd2+ change during channel gating? To investigate this question we compared the rate of Cd2+ action when the membrane was held either at a negative voltage for most of a 24 second duty cycle where channels are largely open or at a more depolarized potential where channels are largely closed (see Methods for details). With the “open state” paradigm the mean Cd2+ inhibition (coordination) time constant (measured with 10 µM Cd2+) was 115 s (n = 6). With the “closed state” paradigm the modification time constant shortened to 68 s in the presence of 2 µM Cd2+ (n = 6; significantly faster than with the “open state” paradigm, p=0.02) and was too fast to quantify at 10 µM Cd2+. We did not observe any significant difference in unblock time constants upon Cd2+ removal at the two voltages (mean Cd2+ unblock time constant for the open state was 24.6 s, n = 5, versus a closed state time constant of 60.6 s, n = 6; p>0.05).
In principle, an effect of voltage on the time course of Cd2+ action could reflect a direct electrostatic effect of membrane voltage on the strength of Cd2+ binding to a site within the membrane field. However, an electrostatic effect should lead to an enhanced rate of Cd2+ binding from the external solution upon membrane hyperpolarization, the opposite to the hyperpolarization-dependent slowing we observe experimentally. Thus, we conclude that the more rapid binding to the channels of Cd2+ at positive potentials is caused by a conformational change associated with channel closing that brings the pair of cysteine residues that form the Cd2+ coordination site in close proximity. This idea is consistent with our finding that Cd2+ (and disulfide bond formation) stabilizes the closed state of the channel relative to the open state (Figure 3).
There are three potential arrangements of disulfide bond formation involving V246C and C303: 1. An intersubunit disulfide bond forms between V246C residues in adjacent subunits; 2. An intersubunit disulfide bond forms between V246C and C303 in adjacent subunits; and 3. An intrasubunit disulfide bond forms between V246C and C303 in the same subunit. A fourth possibility, intersubunit bonds between C303 residues, is judged to be unlikely due to the lack of effect of the various cysteine reactive reagents in HCN1-R channels. These possibilities could, in principle, be distinguished by coexpressing subunits that contain a single cysteine at position 246 with subunits that contain a single cysteine at residue 303. However, such experiments are precluded as the endogenous cysteine at position 303 cannot be replaced (by any of the 19 other amino acids) without loss of channel function (for further information see 5).
To gain insight into the nature of the disulfide bond, we have used an alternative biochemical approach to ask whether CuPhen treatment promotes formation of intersubunit bond formation in the V246C HCN1-R channels (containing C303). This approach is predicated on the following considerations: 1. Redox sensitive formation of homotypic or heterotypic intersubunit disulfide bridges (between V246C-V246C and C303-V246C, respectively) will result in generation of subunit dimers that will run at a higher molecular weight than monomeric subunits when analysed on a non-reducing PAGE gel; 2. A redox sensitive formation of a disulfide bond between C303 and V246C within a channel subunit will not markedly alter migration of the channels on a non-reducing PAGE gel.
To assess dimer formation, we initially maintained oocytes in the presence or absence of 0.15 mM DTT. 48 hr after cRNA injection, we washed out the DTT and exposed a fraction of the DTT-treated oocytes to CuPhen, using incubation conditions comparable to the DTT/CuPhen protocols used in the electrophysiology experiments shown in Figure 2. Cells were then disrupted under a nitrogen atmosphere using a nitrogen-purged lysis buffer that contained CHAPS and NEM at a pH of 7. These conditions were chosen to minimize the post-lysis formation of disulfide bonds. Thus, CHAPS is a small micelle (4-14 molecules) dissociating detergent that should separate uncrosslinked monomers. The use of pH 7 will retard disulfide bond formation during the initial homogenization while the high concentration of NEM will further suppress post-lysis bridge formation by irreversibly alkylating free sulphydryls.
Figure 5a shows a Western blot analysis of samples extracted from oocytes from three separate donor frogs. The band on the left (ΔC) shows a control experiment using HCN1-ΔCterm, which contains six endogenous Cys residues. There is a predominant band at ~40 kD representing the monomeric HCN1-ΔCterm species. In addition, there is a significant amount of staining at ~90 kD, representing disulfide cross-linked HCN1-ΔCterm multimers, indicating that our assay is capable of detecting intersubunit disulfide bond formation. In contrast, V246C HCN1-R subunits are found predominantly as a monomeric species irrespective of the redox treatments prior to cell lysis, suggesting that V246C and C303 participate in an intrasubunit disulfide bond.
The above conclusion is somewhat tempered by our finding that a V246C dimer band can be detected upon overexposure of the gels, although the dimers are present at a much lower level of abundance relative to the monomer band (see the longer exposure shown in the lower panel). The low abundance of the dimer band is consistent with the view that it is a minor protein species, possibly formed under non-physiological post-lysis conditions, and that the relevant disulfide bond formed under physiological conditions in intact oocytes is indeed intrasubunit. Alternatively, the minor band of dimers could represent the species relevant to our physiology experiments if only a small fraction of the V246C HCN1-R channels are properly assembled and trafficked to the surface membrane. In this case the large fraction of monomers could reflect an intracellular pool of immature V246C channels that fail to form disulfide bonds due to the normal reducing environment of the oocyte.
To probe the significance of the V246C dimer band, we examined its sensitivity to treatment with DTT or CuPhen under conditions similar to those used in the electrophysiology experiments. If the minor band of dimers does indeed represent the fraction of channels that are present in the surface membrane, then we expect that the fraction of subunits migrating in the high M.W. band should be decreased by DTT treatment and increased by CuPhen treatment, similar to the ability of these reagents to regulate channel function in our physiological recordings. We therefore measured the density of the dimer band as a percentage of the total channel protein signal (monomer plus dimer). This analysis revealed that pretreatment of oocytes with DTT for 48 hours prior to lysis did indeed reduce the fraction of protein in the dimer band by about 50%, compared to the fraction of dimers observed when oocytes were maintained in the absence of DTT. This finding indicates that some intersubunit disulfide bridges in V246C channels do form during the 48 hour incubation and so are not simply an artifact of dimer formation during the extraction procedure. However, this result still does not indicate whether the minor fraction of dimers is the relevant species responsible for the physiological effects of the cysteine-reactive reagents.
To address this issue, we next treated intact oocytes with CuPhen and asked whether the catalysis of disulfide bond formation with this reagent is able to increase the fraction of channel protein in the dimer band. If V246C and C303 residues do indeed form an intersubunit bond that is responsible for the dimer band on the gels, then we would expect that CuPhen should increase the fraction of channel protein that appears in this dimer band, similar to the ability of this reagent to alter the functional properties of the channel in our electrophysiological experiments on intact oocytes under similar conditions. However, exposure of oocytes pretreated with DTT to CuPhen had no effect on the density of the dimer band relative to the density observed in the paired, DTT-treated, samples, despite the ability of this reagent to produce a large 20 mV in channel gating under similar conditions. These results are thus consistent with the view that CuPhen catalyzes the formation of an intrasubunit disulfide bond and, therefore, does not enhance the fraction of protein that appears in the dimer band, which likely represents a physiologically irrelevant channel species. However, we cannot definitively rule out the possibility that CuPhen also catalyzes the formation of intersubunit bonds in some very small fraction of channel protein that is undetectable on our gels
Our electrophysiological and biochemical results for HCN1 channels indicate that the extracellular end of S4 comes into close apposition with the adjacent S5 helix in the same subunit when the channels are in the resting, closed state. Thus, we observed that disulfide crosslinking and Cd2+ coordination between an engineered cysteine at residue 246 in the S4 helix and an endogenous cysteine 303 in the S5 helix stabilize the deactivated, closed state of the channel. Moreover, Cd2+-coordination between these residues occurs preferentially when the membrane is depolarized, suggesting preferential coordination in the closed state of the channel. Importantly, biochemical studies show that disulfide bond formation is not associated with formation of dimers or higher molecular weight species on a non-reducing gel, suggesting that the disulfide bond is formed between residues within the same subunit. These results provide new information about the state-dependent arrangements of the transmembrane segments in the HCN channel family and suggest both similarities and differences with the architecture of depolarization-activated Kv channels.
Electrophysiological recordings of V246C channels, which contain two Cys residues per subunit (V246C in S4 and C303 in S5), demonstrate robust and complementary effects of reagents that interact with sulfhydryl groups. Treatment with the reducing agent DTT enhanced channel opening by shifting its voltage-dependence of activation to more positive potentials, providing evidence for spontaneously formed disulfide bonds that inhibit channel opening. In contrast, application of CuPhen, which catalyzes the formation of disulfide bonds, inhibited channel function by shifting channel activation to more negative potentials. For channels pretreated with DTT to reduce spontaneous disulfide bonds, CuPhen produced a hyperpolarizing shift of ~20 mV. Application of Cd2+ also inhibited channel function by shifting the voltage-dependence of activation by a similar amount. None of these effects were seen with HCN1-R channels that contain only the endogenous C303 residue. Thus, V246C is likely to lie within 6 of either C303 or a second V246C residue in a neighboring subunit. Moreover, the metal coordination site involving V246C is preferentially formed when the membrane is depolarized and channels are in the closed state.
The electrophysiology data alone do not allow us to determine whether the Cys-Cys interactions represent an intersubunit bond between two V246C residues, an intersubunit bond between V246C and C303 in neighboring subunits, or an intrasubunit bond between V246C and C303. Western blot analysis of extracted channel protein under non-reducing conditions showed that the predominant species is monomeric (although a small fraction of total protein does exist as dimers, see below). The lack of a prominent dimeric species argues that the majority of disulfide bonds in functional channels must occur in an intrasubunit manner between V246C and C303.
As this conclusion is based on a negative result, we need to consider alternative explanations for the lack of dimer formation. Because we find that the distance between V246C and C303 is increased when the channels are in the open state (membrane is hyperpolarized), disulfide bond formation might not occur in the biochemical experiments if the channels were predominantly in the active state. However, we find that disulfide bonds form spontaneously during incubation conditions similar to those used in the biochemical experiments, based on our functional data that DTT treatment enhances channel opening. Moreover, the majority of V246C channels should be in the closed state at the typical oocyte resting potential (-30 to -50 mV) under our post-injection incubation conditions (Figure 1).
A second potential complicating factor could occur if only a minor fraction of the total V246C protein on the Western blots represents functional channels expressed in the surface membrane. Any pool of intracellular channel protein might be unable to form disulfide bonds due to the reducing environment of the oocyte. As a result, only a small fraction of the total protein would be able to form dimers, even if V246C participates in an intersubunit disulfide bond. Although it is difficult to address this problem definitively, we can readily detect significant multimer formation in HCN1-ΔCterm, which contains six endogenous Cys residues and is the parent construct for HCN1-R. Importantly, we also found that the dimer signal in the V246C channel Western blots is unaffected by pretreatment of oocytes with CuPhen (Figure 5), despite the ability of this reagent to catalyze disulfide bond formation in V246C channels expressed in the surface membrane. This suggests that the small amount of dimer formation in these channels represents a rare event that is not representative of the conformation of the fully assembled channel in the surface membrane, e.g. possibly an artifactual intersubunit proximity of Cys-Cys residues arising during oocyte homogenization and protein extraction. However, it is important to point out that our results do not rule out the formation of physiologically relevant intersubunit bonds that are below the limits of detection of our biochemical experiments.
Both the state-dependence of Cd2+ binding and the effect of Cd2+ and disulfide bonds to inhibit channel opening suggest that V246C and C303 within the same subunit are in close proximity when the membrane is depolarized and the channels are closed. Previous data suggests that there are minimal vertical, transmembrane motions of the S4 helix during voltage sensing and gating of mammalian HCN channels and that V246C remains readily accessible to the external aqueous environment in both closed and open states of the channel (5, 50). These results together suggest that protein motions during membrane depolarization that account for the state-dependent interaction of V246 and C303 may involve a lateral translation or rotation in the plane of the membrane, bringing the top of S4 in closer proximity to S5 of the same subunit. Such a lateral motion could be explained by tilting of S4 relative to S5, consistent with the previous S4 accessibility studies on mammalian HCN channels (5). Alternatively, a transmembrane motion of S5 might move C303 in closer proximity to S4 during membrane depolarization. Presumably, the curtailing of such motions by disulfide bridging or Cd2+ coordination between the S4 and S5 helices give rise to the observed inhibition of channel opening and shift of gating to more negative potentials.
Hyperpolarization-activated HCN channels are members of the Kv channel superfamily and, like other family members, contain six transmembrane segments, a positively charged S4 segment that is part of the S1-S4 voltage-sensing domain, and a pore-forming S5-S6 region. Previously we found that the voltage-sensing mechanism in mammalian HCN channels (5) shares some key elements with the transporter model for voltage sensing (7, 14, 48), one of three putative gating models hypothesized for depolarization-activated Kv channels (see Introduction for a summary). Thus, HCN channels appear to display a highly focused electric field due to aqueous crevices and membrane deformation around S4 and exhibit small transmembrane motions (<3 Å) of S4.
The state-dependent changes in the S4-S5 arrangement seen here for HCN1 channels show intriguing similarities and differences with Shaker Kv channels (8, 18, 25, 36). Thus, certain pairs of Shaker residues near the extracellular ends of S4 and S5 form intersubunit disulfide bonds and Cd2+ coordination sites (8, 25, 36). In contrast, Elliott et al. (18) found that a different pair of Shaker residues near the external ends of S4 and S5 form an intrasubunit Cd2+ coordination site preferentially when the channels are activated by depolarization. Of particular interest, this same pair of Shaker residues forms an intersubunit interaction when the channels are in the resting state (membrane hyperpolarized). From these results, Elliott et al. (18) concluded that the top of S4 lies near the top of S5 of an adjacent subunit in the resting state; during channel activation there is a conformational change that causes the top of S4 to lie near the top of S5 in the same subunit. Furthermore, studies in the hyperpolarization activated KAT1 channel suggest the possibility of three residue pairings along the lengths of S4 and S5, including possibly both intra- and intersubunit interactions, which arise during channel gating (21, 24).
The above results demonstrate that in HCN1, KAT1 and Shaker channels S4 and S5 within the same subunit may move into close proximity during voltage sensing motions when the membrane is depolarized, even though Shaker channels are in the open state at this potential whereas HCN1 and KAT1 channels are in the closed state. Thus, the orientation of the S4 and S5 segments is likely to reflect the state of the voltage sensor (which is a function of membrane potential) rather than the state of the channel gate. That HCN1 channels may not form the intersubunit Cd2+ coordination site seen in Kv channels at hyperpolarized potentials is consistent with the notion that the S4 segments in HCN1 channels show a more restricted motion during voltage-sensing compared to the S4 segments of Kv channels (5, 50).
The finding that S4 and S5 in the same subunit are in close proximity when the membrane is depolarized is difficult to reconcile with the X-ray crystal structure of the homologous mammalian Kv1.2 channel (31), where the S1-S4 voltage-sensing subdomain lies close to the S5 segment of a neighboring subunit but far from its own S5 segment. Although the structure is somewhat more compatible with the finding of intersubunit interactions between residues at the top of S4 and S5 in Shaker channels (8, 18, 25), even here a discrepancy exists as the Shaker channel S4-S5 intersubunit interactions occur preferentially in the resting state (18), whereas the X-ray structure is of the open channel. Moreover, the intersubunit distance between the top of S4 and S5 in the crystal structure is too great to explain high-affinity metal coordination (27).
To gain more insight as to whether the distance between V246C and C303 in HCN1 is compatible with the Kv1.2 structure, we modeled HCN1 based on the Kv1.2 structure using a multiple sequence alignment (Figure 6). As there is some ambiguity in the alignments, we explored structures based on four distinct possible alignments. The intrasubunit distances between V246C and C303 ranged from 33-43 Å in the four structures, all too large to support formation of disulfide bonds or coordinate Cd2+ binding. The intersubunit distances, ranging from 14-23 Å, are less than the intrasubunit distances but are still at the upper limit for disulfide bond formation and too large to form a high-affinity Cd2+ binding site. Thus, the structure of HCN1 may differ substantially from that of Kv1.2. Alternatively, the Kv1.2 structure may be partially distorted (27). Finally, the V246C-C303 intrasubunit disulfide bonds or Cd2+ coordination may occur during rare sojourns of the HCN1 channel into a distorted state that becomes trapped by covalent bond formation or Cd2+ binding. However, any such distortion cannot be too great as the 20 mV shift in voltage-gating represents a relatively modest functional change.
Given the apparent similarities in channel architecture identified here that give rise to intrasubunit interactions between S4 and S5 in depolarization activated Kv channels and the hyperpolarization activated HCN1 (and potentially also in KAT1) channels, our results argue that the opposite polarity of voltage sensitivity arises elsewhere in the HCN channel structure. This could occur at the intracellular side of the membrane where the S4-S5 linker is thought to couple voltage-sensor movement to channel gating (15, 16, 40). Alternatively, HCN1 voltage-gating polarity could be determined by conformational changes in the channel regions surrounding S4 as described previously (5, 50).
We thank Keri Fogle and Renee Saenger for technical assistance and Margaret Wood and the Department of Anesthesiology for continuing support. D.C.B. was supported by the Wellcome Trust (UK) and an American Heart Association grant (0325454T). S.A.S was supported by the Howard Hughes Medical Institute & grant R01NS36658 from the National Institutes of Health. G.R.T. was supported by The Whitehall Foundation (2003-05-02-REN)