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MOLECULAR AND GENOMIC |
1 From the Department of Physiology, Faculty of Medicine, Kyoto University, Kyoto 606-8501, Japan
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Abstract |
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-helical structure of the S1 transmembrane segment. Tryptophan replacements of residues responsible for the different kinetics between HCN1 and HCN4 made the activation kinetics slower than the wild-type HCN1. Tryptophan mutations introduced in the middle of S1 (L139W and V143W) prevented normal channel closure. Furthermore, a negatively charged residue at position 139 (L139D) induced a positive voltage shift of activation by 125 mV. Thus, L139 and V143 probably face a mobile part of the S4 voltage sensor and may interact with it. These results suggest that the secondary structure of S1 is -helical and profoundly affects the motion of the voltage sensor.
(Received 5 November 2006;
accepted after revision 18 December 2006;
first published online 21 December 2006)
Corresponding author H. Ohmori: Department of Physiology, Faculty of Medicine, Kyoto University, Kyoto 606-8501, Japan. Email: ohmori{at}nbiol.med.kyoto-u.ac.jp
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Introduction |
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To investigate the structure and the orientation of S1 architecture, we adopted a tryptophan (Trp) perturbation mutagenesis strategy (Choe et al. 1995; Sharp et al. 1995). The premise of the approach is that replacing the native amino acid by Trp will disturb channel function by influencing nearby residues in other transmembrane segments, without affecting residues exposed to lipid. Nevertheless, the bulky hydrophobic side-chains of Trp residues often experience hydrophobic interactions and stabilize protein–protein interfaces (York & Nunberg, 2004), and therefore the results from Trp perturbation scans must be carefully interpreted. A Trp perturbation study and an Ala perturbation study for Kv channels each demonstrated that S1–S3 transmembrane regions are -helical structures (Monks et al. 1999; Hong & Miller, 2000; Li-Smerin et al. 2000). Since HCN channels share the basic transmembrane organization and topology with Kv channels (Santoro et al. 1998; Ludwig et al. 1998), and the S4 voltage sensors of HCN and Kv channels move in the same direction upon voltage changes (Männikkö et al. 2002), we expected to find similar results using Trp perturbation to probe HCN channels. However, HCN channels are decidedly different from Kv channels in that they are activated by membrane hyperpolarization, while depolarizing potentials activate Kv channels. In addition, Kv channels and HCN channels are different in the local S4 environment; the NH2-terminal half of S4 in HCN channels is static (Bell et al. 2004; Vemana et al. 2004), while it is mobile for Kv channels upon voltage gating (Larsson et al. 1996). Moreover, the primary amino acid sequences of the S1 segments from HCN and Kv channels are quite different (Santoro et al. 1998; Ludwig et al. 1998). Based upon these considerations, a Trp perturbation scan of HCN1 was undertaken. We found that HCN1 channel function was disrupted periodically by S1 Trp substitutions, suggesting an -helical structure, that some mutants unexpectedly prevented normal channel closure, and that two residues are implicated in affecting the S4 voltage sensor.
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Methods |
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The HCN1 channel was cloned from mouse brain by PCR as previously described (Ishii et al. 2001). To facilitate subcloning, 289 amino acid residues in the C-terminal region of HCN1 were deleted. In HCN1, C-terminal deletion did not affect gating kinetics or voltage dependence (Ishii et al. 2001). In this study, HCN1 without the C-terminal region is referred to as ‘WT’. All mutations were introduced into WT channels by overlap PCR as previously described (Ishii et al. 2001). The nucleotide sequences of all mutant channels were verified using dideoxy chain termination sequencing (BigDye Terminator Cycle Sequencing, Applied Biosystems, Inc., Foster City, CA, USA). The T7 promoter sequence was introduced into an oocyte expression vector, pBF (Fakler et al. 1994). The WT channel and all mutants cloned in pBF were linearized by appropriate restriction endonucleases and were transcribed in vitro with T7 RNA polymerase (Ambion, Austin, TX, USA) in the presence of 2.5 mM m7G(5')ppp(5')G Cap Analog (Ambion). To determine expression levels, some mutants were subcloned into a mammalian expression vector, pCI (Promega, Madison, WI, USA), and were transfected in COS7 cells or HEK 293 cells using Lipofectamine 2000 (Invitrogen, Carlsbad, CA, USA). Transfected COS7 cells were subjected to a whole-cell patch clamp recording as previously described (Ishii et al. 2001).
Immunofluorescence
After transfection, HEK 293 cells on coverslips were incubated for 24 h with 100 µM cycloheximide. The cycloheximide-treated HEK 293 cells were fixed in ice-cold 4% paraformaldehyde in phosphate-buffered saline (PBS) (Invitrogen) for 20 min. The cells were then washed with PBS containing 1% bovine serum albumin (BSA), permeabilized with 0.1% Triton X-100 in PBS containing 1% BSA for 20 min, washed with PBS containing 1% BSA, and blocked with 1% normal donkey serum (NDS) in PBS for 1 h at room temperature. Subsequently, the cells were incubated with a guinea-pig polyclonal antibody to HCN1, a kind gift from Dr R. Shigemoto (NIPS, Okazaki, Japan; Notomi & Shigemoto, 2004), at a dilution of 1:2000 in PBS containing 1% NDS for 1 h at room temperature. The antibody was removed, and the cells were washed again with PBS containing 1% BSA. The cells were then incubated with a goat antiguinea-pig antibody conjugated with red fluorescent dye (Alexa Fluor-594 goat antiguinea-pig IgG; Molecular Probes, Eugene, OR, USA) at a dilution of 1:200 in PBS containing 1% NDS for 1 h at room temperature in the dark. The antibody was removed and the cells were washed with PBS containing 1% BSA. The coverslips were dried completely and mounted on slides using FluoroGuard Antifade Reagent (Bio-Rad, Hercules, CA, USA). The cells were observed using a confocal laser-scanning microscope (CSU10; Yokogawa, Tokyo, Japan).
Oocyte expression, electrophysiology and data analysis
Xenopus care and handling were in accordance with the guiding principles and regulations of Kyoto University. Frogs were anaesthetized by immersion in a 0.2% solution of tricaine. A segment of Xenopus ovary was treated with 2% collagenase (Worthington, Lakewood, NJ, USA) and then mature stage V and VI oocytes were defolliculated and isolated manually. The capped RNA was dissolved in sterile water at 1 mg ml–1. Fifty nanolitres of the RNA solution was microinjected into an oocyte using a microinjector (Nanoject, Drummond, Broomall, PA, USA). Injected oocytes were incubated at 18°C in modified Barth's medium (88 mM NaCl, 1 mM KCl, 2.4 mM NaHCO3, 0.3 mM CaNO3, 0.41 mM CaCl2 and 0.82 mM MgSO4) supplemented with 0.1 mg ml–1 gentamicin. Currents were recorded 2–3 days after injection using two-electrode voltage-clamp (Axoclamp2B, Axon Instruments). Data acquisition was performed using Digidata 1320A and AxoGraph 4.6 (Axon Instruments). Currents were filtered at 2 kHz and sampled at 10 kHz. The extracellular (bath) solution contained (mM): 96 KCl, 1.8 CaCl2, 1 MgCl2, 10 Hepes and pH was adjusted by 6 KOH (pH 7.6). Electrodes were filled with 3 M KCl. Electrodes had resistances of 0.3–1.5 M
. The holding potential was –20 mV. All experiments were performed at 25.0 ± 0.5°C. Tail-current amplitudes were measured 2–3 ms after the pulse at –120 or –140 mV. Normalized tail current amplitude was plotted versus test potential to obtain the voltage-dependent activation curve and fitted with a Boltzmann function:
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| (1) |
is the membrane potential for the half-maximal activation, and S is the slope factor. Activation and deactivation kinetics were compared among various mutants and WT. Activation kinetics, which were fitted well by a double exponential function (Santoro et al. 2000), were quantified by the time required for half-opening (t) at –120 mV when the hyperpolarizing pulse duration was 1200 ms. Figures 2A and 3A demonstrate traces using shorter pulses as representative records. The time constants for deactivation, 
d, were determined by fitting the tail current with a single exponential function without the initial delay. The ‘open-state stabilization energy’ is defined as:
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| (2) |
and S were calculated from the Boltzman function (Monks et al. 1999; Hong & Miller, 2000; Lesso & Li, 2003). The mutations made in the present study induced significant changes in side-chain volume. To compensate for the differences in side-chain volume changes, we calculated a weighted 
G0 (G0w) using the following equation as described by Li-Smerin et al. (2000) and Subbiah et al. (2005):
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| (3) |
Volave (44.9 Å3) is the average change in side-chain volume introduced by Trp in this study and Vol is the change in side-chain volume for the specific mutant. Large depolarizing voltage steps were applied in the case of L139 and V143 mutants. In these mutants, more than 50% of channels were still open even at extreme depolarization (+100 mV), and voltage activation curves were extremely positive shifted. Therefore, HCN currents were estimated as the Cs+-sensitive currents (2 mM Cs+) for L139. Normalized Cs+-sensitive tail current amplitude for L139 mutants was plotted and fitted with the following function:
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Results |
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-helical structure in S1
Tryptophan-scanning mutagenesis of non-Trp residues 130-Phe to 149-Ile except for 134-Trp of the S1 segment was carried out. The C-terminal deleted HCN1 (WT) and 19 Trp mutants were expressed in Xenopus oocytes and were examined by the two-electrode voltage-clamp technique. Four Trp mutants (M138W, M142W, L146W and I149W) failed to generate currents (Circled residues in Fig. 1A). Three of the four residues were conserved among all the HCN subtypes (Fig. 1A). The expression levels of these mutants were further examined in the mammalian expression system; these four mutants were subcloned into a mammalian expression vector, pCI, and were examined by whole-cell patch clamp in COS7 cells. Only I149W generated currents; however, the current level was unstable and was too small to determine the gating parameters. The current was at most 100 pA after a –100 mV hyperpolarizing voltage step from a holding potential of –20 mV. The other three mutants also failed to generate currents in the mammalian expression system. Using confocal microscopy we examined the localization of mutant channels labelled with an anti-HCN1 antibody. In transfected HEK293 cells, all four mutant channels appeared to be expressed in the plasma membrane, similar to WT channels, whereas cells expressing green fluorescent protein (GFP) had diffuse cytosolic fluorescence (Fig. 1B). Therefore, it is unlikely that the failure of these channels to express currents results from trafficking defects resulting from the Trp substitutions. Two or three residues separate each of the Trp-substituted positions (Fig. 1A). Assuming that the structure of S1 is an -helix and that one turn is 3.6 residues, these residues should cluster on one face of the -helical wheel (shaded diamonds in Fig. 6B).
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Out of the 19 Trp mutants, 11 generated robust currents with time-dependent activation kinetics in response to hyperpolarizing voltage steps. Representative current traces are shown in Fig. 2A for WT and four other mutants. Their gating parameters were estimated. The V for I148W was shifted most positively (Fig. 2D). This is consistent with the fact that G0w for I148W is the largest, meaning that among all the mutants the open probability of I148W is greatest at 0 mV (Fig. 2B). Half-activation time was measured from current responses to voltage steps to –120 mV for 1.2 s. The current activations of D135W, I137W and M141W were considerably slower than WT (Fig. 2E). In the previous study, which compared the kinetics between HCN1 and HCN4 (Ishii et al. 2001), we found that the residues at positions 137 and 141 are responsible for the difference of activation kinetics (Ishii et al. 2001). The introduction of corresponding residues of HCN4 into these two positions slowed the activation kinetics of HCN1. Deactivation time constants were measured at +20 mV after activation of the channels by voltage steps to –120 mV (Fig. 2F). Two mutants (I137W and M141W) showed similar deactivation kinetics to WT, and the other nine mutants showed significantly slower deactivation than WT. This may indicate that HCN1 channels are optimized for fast deactivation and mutations necessarily slow deactivation gating.
Gating parameters varied among the different mutants but the gating changes did not show clear -periodicity. However, a complete spectrum of mutants could not be studied because a large number of mutant channels, eight out of 19, did not generate sizeable currents that could be well quantified. In contrast, only two Kv Trp mutants failed to generate current (Hong & Miller, 2000). The large number of mutants that did not yield currents in this study does not permit conclusions about the periodicity of gating changes.
Mutations of the negatively charged residue in S1 slow activation kinetics
A single negatively charged residue, D135, resides in S1. Activation kinetics for D135W were the slowest among all of the Trp mutants (2.2-fold; P < 0.05; Fig. 2E) as described in the subsection above. Deactivation kinetics for D135W were also significantly slower than those for WT (2.7-fold; P < 0.05; Fig. 2F). A neutral residue (asparagine, N) or positive residues (histidine, H; arginine, R; and lysine, K) were introduced at position 135 to examine the effects of the charge at this position. The D135K mutant generated no currents. The rest of the mutations shifted the activation curve to the negative direction by 21.3 and 24.4 mV for D135N and D135H, respectively (Fig. 3C); the negative shift by D135R was too large to be analysed accurately. All three mutations slowed activation kinetics significantly compared with WT (2.5-fold, 4.8-fold and 5.9-fold for D135N, D135H and D135R, respectively; P < 0.05; Fig. 3D), and this was accompanied by decreasing G0 by more than 1.5 kcal mol–1 (Fig. 3B). Deactivation kinetics were slowed by the mutation D135H (1.4-fold at +20 mV; P < 0.05) and D135R (1.6-fold at +20 mV; P < 0.05), but not by D135N (Fig. 3E).
Two mutations disrupt channel closure
The introduction of Trp into position 139 or 143 did not generate typical HCN currents (Fig. 4Aa and Ba), although the currents were still larger at hyperpolarizing voltage steps than at depolarizing steps, especially for L139W. The elicited currents were relatively small in these two mutants. Consequently, the endogenous currents were not negligible, especially at depolarizing potentials. Therefore, HCN currents for L139W were estimated as Cs+-sensitive currents by subtraction after block by 2 mM Cs+ (Fig. 4Ac). There were two problems with using Cs+: (1) extent of block is dependent on membrane potentials (Woodhull, 1973; Moroni et al. 2000); and (2) a block by 2 mM Cs+ is not complete (80% at –100 mV in our system). Therefore, HCN currents by subtraction after block by 2 mM Cs+ were underestimated especially at depolarizing potentials. However, since the tail current was measured at a fixed voltage, the voltage dependence of the block did not affect the evaluation of the tail current. We also tried blocking the channels with the HCN channel blocker ZD7288, but this proved unsatisfactory (see Methods). It was confirmed that the endogenous currents were not affected by 2 mM Cs+ by assaying oocytes without RNA injection (data not shown). There were no outward currents in the subtracted traces (Fig. 4Ac) because outward HCN currents were not blocked by Cs+ and arithmetically removed by subtraction (Männikkö et al. 2005). The tail currents for L139W indicated that 52% of the channels were not closed even at +100 mV (Fig. 4C). The V and slope factors were –0.9 mV and +24.2 mV, respectively. The V143W mutant showed almost no voltage dependence, and appeared constitutively open. The currents were not significantly blocked at –100 mV in 5 min after application of ZD7288 (1 mM; Fig. 4Bb and D). In contrast, the currents generated by WT channels were blocked by about 80% in 5 min by ZD7288 (data not shown). Five millimolar Cs+ blocked the currents by 80% (P = 0.002) at –100 mV, although not significantly at +50 mV (Fig. 4Bc and D). This property of voltage-dependent block of V143W current by Cs+ was similar to the results found for native HCN channels (Mayer & Westbrook, 1983). The L139 and V143 residues are separated by three residues, indicating that these two residues probably line the same face of an -helical wheel in S1.
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Mutation L139W disrupted channel closure and shifted the voltage dependence greatly in the positive direction (V = –0.9 mV). This suggests that position 139 may directly, or indirectly via other transmembrane domains (S2 or S3), interact with the movement of the voltage sensor. The putative moving part of the voltage sensor is the S4 segment, which contains many positive charges. Therefore, we introduced a negatively charged side-chain (aspartate, D) or positively charged side-chain (lysine, K) into position 139 to determine the effect of the charge at this position. Like L139W, both charge mutants disrupted channel closure, whereas the introduction of a small side-chain (glycine, G, or alanine, A) or a polar side-chain (aspargine, N) into position 139 did not interfere with channel closure (Fig. 5A and B). Open probabilities for L139D and L139K were 79% at +140 mV and 72% at +40 mV, respectively (Fig. 5B). Mutation L139D markedly shifted the voltage dependence. The V for L139D was +39.2 mV, whereas values of V for WT and L139K were –85.6 and –71.6 mV, respectively. The negative charge at position 139 (139D) produced an extreme shift of the voltage dependence. This supports the idea that the residue at position 139 can interact strongly with the moving part of the voltage sensor. The slope factor for L139K (14.1 mV) was almost the same as that for WT (14.4 mV), but the slope factors for the rest of the L139 mutants (24.2 mV for L139W, 24.2 mV for L139D, 19.7 mV for L139G, 22.3 mV for L139A and 20.0 mV for L139N) were significantly larger than for WT, indicating that the charge movement of the voltage sensor might be restricted in these mutants (L139W, L139D, L139G, L139A and L139N).
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-helical wheel may be divided into three parts
The -helical wheel and a net diagram of the HCN1 S1 segment are displayed in Fig. 6. Two Trp mutants (F132W and N145W) generated only small time-dependent currents, which were too small (< 100 nA at –120 mV) to be analysed. We classified the effects of Trp substitutions into four categories as illustrated in Fig. 6. First (open circles in Fig. 6), 11 mutants displayed robust currents (> 1 µA at –120 mV) like WT; second (shaded diamonds in Fig. 6), four mutants generated no currents; third (shaded circles in Fig. 6), two mutants generated only small currents; and fourth (shaded squares in Fig. 6), two mutants showed incomplete channel closure. It is apparent that the residues of the first group (open circles in Fig. 6) are located close to each other and that these residues cover about half the surface of the helical wheel (I in Fig. 6B). There are two exceptions in the first group, and these two residues are located close to each other but are separated from the other first-group residues: D135W and R131W in cluster II. Residues of the second group (shaded diamonds in Fig. 6) comprise another cluster (II in Fig. 6B), and two residues of the fourth group (shaded squares, residues 139 and 143 in Fig. 6) were also clustered with each other (III in Fig. 6B). One of the two residues of the third group (shaded circles in Fig. 6) is located in cluster II (N145W, Fig. 6B), and the other residue in cluster III (F132W, Fig. 6B). Thus, the circumference of the -helical wheel can be divided into three parts of functionally different characters (I, II and III in Fig. 6B).
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Discussion |
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-helical structure. However, in contrast to S1 in Kv channels (Hong & Miller, 2000), as many as eight mutants out of 19 failed to generate sufficient currents for accurate analysis. Although we could not find a clear periodicity in the gating parameters (Fig. 2), the expression level of current suggested an -helical periodicity in HCN channels (Fig. 6). The gating parameters were altered by Trp mutations in Kv channels with an -helical periodicity (Hong & Miller, 2000). This discrepancy between HCN and Kv channels suggests that S1 might interact with other transmembrane regions more intimately in HCN channels than in the Kv channels, or that HCN channels themselves might be more vulnerable to slight changes in conformation than Kv channels.
As already described for Kv channels (Monks et al. 1999; Hong & Miller, 2000), variable residues among subtypes had a tendency to be resistant to Trp substitution, and are thought to be located in a lipid-exposed surface. This rule is almost true for HCN channels because there exist seven variable residues (filled squares in Fig. 6), which cluster on a single partition of the -helical wheel (cluster I in Fig. 6B; except for the one residue I149), and these six variable residues were resistant to the Trp mutation (Fig. 6). We previously demonstrated that two positions (137 and 141) out of seven variable residues were responsible for the difference in activation kinetics between HCN1 and HCN4. The introduction of Trp into these two positions also slowed the channel kinetics. The side-chains at these two positions seem to face lipid according to our results (Fig. 6B); a change of a side-chain at such a position is generally inefficient in affecting the channel gating. However, the side-chain replacements in these two positions of HCN1 significantly altered channel gating. The changes in the two positions may lead to an overall structural change in S1.
Mutations at four positions (shaded diamonds in Fig. 6; positions 138, 142, 146 and 149) failed to generate currents, and these residues cluster in another area of the -helical wheel (II in Fig. 6B). Considering that Trp possesses a large hydrophobic side-chain, together with the results in Kv (Monks et al. 1999; Hong & Miller, 2000) and Kir channels (Choe et al. 1995; Collins et al. 1997), these four residues are likely to be located in a protein–protein interface.
Two mutations resulted in channels that failed to close properly (L139W and V143W; shaded squares in Fig. 6). Interestingly, V143W did not show any voltage-dependent gating and the currents were not significantly blocked by 1 mM ZD7288. Shin et al. (2001) proposed two models of ZD7288 blockade, one with preferential closed state block and the other with two open states having different blocker affinities. The V143W mutant did not appear to have a closed state and appeared insensitive to ZD7288. Therefore, the preferential closed state block model seems more reasonable, based upon this single mutant channel. It is noteworthy that voltage-independent currents of HCN2 were blocked by ZD7288 (Proenza & Yellen, 2006). This result may show that V143W has no closed state because a closed state is necessary for block by ZD7288 in the preferential closed state block model.
The two positions L139 and V143 are separated by three residues and are located close to each other on the other area of the -helical wheel (III in Fig. 6B). In HCN channels, it was previously reported that mutations showing disrupted closure were found in S4, the S4–S5 linker, and in the C-linker (Chen et al. 2000, 2001; Decher et al. 2004). As previously described (Chen et al. 2001), there are three possibilities for disrupting channel closure. First, the voltage sensor could be fixed in an open position. Second, the channel gate could be stabilized in an open state. Third, the coupling between the voltage sensor and the channel gate could be disturbed. Since S1 is believed to be at the periphery of the channel, it is unlikely that these residues (L139 and V143) are the coupling between the voltage sensor and the pore gate or that these residues could be part of the gate itself. Therefore, it is likely that these two residues modify the movement of the voltage sensor. In addition, because of its insensitivity to block by ZD7288, V143W is expected to spend much less time in the closed state compared with the voltage-independent currents of HCN2, which may suggest that the gate is locked open by paralysing the voltage sensor instead of by slippage in the coupling between the gate and the voltage sensor.
A collapsing gating canal model was proposed by a cysteine-scanning study (Bell et al. 2004). In this model, an extensive aqueous crevice (the gating canal) collapses around and buries the C-terminal tail of S4 in the closed state. If these two residues (139 and 143) form a part of the gating canal, mutations at these positions may profoundly affect channel gating.
Two X-ray crystal structures of the Kv channel from Aeropyrum pernix (KvAP channel) are available (Jiang et al. 2003). One is of the whole channel and the other is of the isolated voltage-sensor domain (S1–S4). The full KvAP structure suggested that the entire S1 and S2 segments are in the interior of the membrane (Cohen et al. 2003). In contrast, the isolated S1–S4 structure showed salt bridges between S2 and S4 (also see Silverman et al. 2003). In addition, the results from Ala- and Trp-scanning studies were consistent with the result of the isolated S1–S4 structure (Cohen et al. 2003). The crystal structure of Kv1.2 also shows that the structure of voltage-sensor S1–S4 region is isolated from the pore domain of the channel (Long et al. 2005). If a similar arrangement occurs for HCN channels, and the variable residues (I in Fig. 6B) are located in a lipid-exposed surface of -helix, then: (1) the surface of the helical wheel, which generates no current by Trp mutation (II in Fig. 6B), might interact with S2; and (2) the other surface showing incomplete channel closure (III in Fig. 6B) might interact with S4. This model seems consistent with our results.
A recent report using site-directed spin labelling and electron paramagnetic resonance spectroscopy suggested that the S1 segment of KvAP is surrounded by other parts of the protein, and a hypothesis was proposed that S1 lies at the interface between the voltage-sensing domain and pore domain in KvAP (Cuello et al. 2004). This result is not compatible with our results or with other studies of Kv channels. If the hypothesis of Cuello et al. (2004) was true, then residues (139 and 143) showing disrupted channel closure may couple the voltage sensor and the gate. Their hypothesis, however, seems unlikely for HCN channels because the present results demonstrate that half of S1 is facing the protein–lipid interface and that mutations at positions 139 and 143 markedly shift voltage dependence, accompanied by the disruption of normal channel closure.
In conclusion, the S1 segment in HCN channels shows an -helical periodicity. Some mutations in S1 extremely alter the minimal open probability and voltage dependence, which suggests that S1 interacts with the moving part of the voltage sensor.
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References |
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