Slow conformational changes of the voltage sensor during the mode shift in hyperpolarization-activated cyclic-nucleotide-gated channels

Abstract

Hyperpolarization-activated cyclic-nucleotide-gated (HCN) channels are activated by hyperpolarizations that cause inward movements of the positive charges in the fourth transmembrane domain (S4), which triggers channel opening. If HCN channels are held open for prolonged times (>50 ms), HCN channels undergo a mode shift, which in sea urchin (spHCN) channels induces a >50 mV shift in the midpoint of activation. The mechanism underlying the mode shift is unknown. The mode shift could be attributable to conformational changes in the pore domain that stabilize the open state of the channel, which would indirectly shift the voltage dependence of the channel, or attributable to conformational changes in the voltage-sensing domain that stabilize the inward position of S4, thereby directly shifting the voltage dependence of the channel. We used voltage-clamp fluorometry to detect S4 movements and to correlate S4 movements to the different activation steps in spHCN channels. We here show that fluorophores attached to S4 report on fluorescence changes during the mode shift, demonstrating that the mode shift is not simply attributable to a stabilization of the pore domain but that S4 undergoes conformational changes during the mode shift. We propose a model in which the mode shift is attributable to a slow, lateral movement in S4 that is triggered by the initial S4 gating-charge movement and channel opening. The mode shift gives rise to a short-term, activity-dependent memory in HCN channels, which has been shown previously to be important for the stable rhythmic firing of pacemaking neurons and could significantly affect synaptic integration.

Figure 1.

Different models for generating shifts in the voltage dependence of HCN channels. Experimental shifts in the midpoints of the Q(V) (A) and G(V) (B) accompanying the mode shift in spHCN channels (Männikkö et al., 2005), induced by voltage steps to −80 mV for 100 ms. A, Q(V) measured from a holding potential of −10 mV (●) and −80 mV (○) for nonconducting P435Y spHCN channels. B, G(V) measured as isochronal tail currents at +50 mV after equilibration at the indicated voltage for 80 ms with (♢) or without (□) a 100 ms prepulse to −80 mV. Holding potential was 0 mV. The Q(V) and G(V) were well fit by Boltzmann curves, i.e., G(V) = 1/(1 + exp(q(V − V_1/2)/kT)). C, Increasing lengths of the prepulse to −80 mV increased the size of the G(V) shift (■). The G(V) shift is measured as the shift in the midpoint of activation, V_1/2, as determined in B. The time course for the G(V) shift was slower than the onset of the ionic current during the prepulse (dotted line), showing that the mode shift is slower than channel opening. D–F, A change in V_1/2 may be attributable to a change in the free energy of opening of the activation gate ΔW_co (D), a change in the free energy of S4 movement ΔW_cc (E), or a change in both (F) (for energy calculations, see Materials and Methods, Modeling). D, A putative conformational change in the pore domain (shown here as an increased electrostatic interaction between S5 and S6) that stabilizes the open conformation would shift the voltage dependence by changing ΔW_co. E, A putative conformational change in the voltage sensor domain (shown here as an increased electrostatic interaction between S2 and S4) that stabilizes the open conformation would shift the voltage dependence by changing ΔW_cc. F, A putative global conformational change of whole channel protein that stabilizes the open conformation would shift the voltage dependence by changing both ΔW_co and ΔW_cc.

Figure 2.

Fluorescence from 326C is slower than ionic current activation. A, Sequence alignment of spHCN and HCN1 S4 domains. Marked residues exhibit state-dependent modification by intracellular (*), extracellular (#), or both intracellular and extracellular reagents (&), or state-independent modification by either intracellular or extracellular reagents (o) (Männikkö et al., 2002; Bell et al., 2004; Vemana et al., 2004). B, C, Representative current (top) and fluorescence (bottom) records from Alexa-488 C5-maleimide-exposed wt spHCN channels (B) and 326C channels (C). Channels were held at 0 mV and then stepped to negative potentials (0 to −140 mV), followed by a step to +50 mV. Note that 326C channel currents precede the fluorescence change and that wt channels do not show a fluorescence change. D, E, Steady-state (D) and kinetic (E) analysis of the voltage dependence of the conductance (■) and fluorescence (red circles) from C. The fluorescence signal was slower than ionic current activation across a range of voltages. Average time constants (milliseconds) for fluorescence and ionic current were, respectively, as follows: 132.1 ± 11.6 and 75.6 ± 15.1 (−80 mV); 98.5 ± 5.1 and 32.7 ± 6.3 (−100 mV); 75.4 ± 4.3 and 22.3 ± 3.8 (−120 mV); and 61.1 ± 4.2 and 15.6 ± 2.7 (−140 mV); p < 0.01 at each voltage. F, G, Scaled and overlaid current (black line) and fluorescence (red line) in response to a voltage step to −120 mV for 326C (F) and 332C (G) channels. Note that the fluorescence signal has been inverted for a direct kinetic comparison with the current traces. The positive direction of current (I) and fluorescence (F) is indicated by black arrows.

Figure 3.

Fluorescence from 326C channels correlates with the mode shift but not inactivation. A, B, Currents in response to voltage steps from 0 to −120 mV, in 20 mV increments from Alexa-488-labeled 326C channels that do not show any apparent cAMP-dependent inactivation (A) and from Alexa-488-labeled 326C/620G channels that do exhibit cAMP-dependent inactivation (B). C, D, Scaled and overlaid current (black line) and fluorescence (red line) in response to a voltage step to −120 mV for non-inactivating 326C channels (C) and inactivating 326C/620G channels (D) (inset, the inactivation component of the current trace scaled and overlaid with the fluorescence change). The time constants for the currents were 28 ms for activation and 360 ms for inactivation, whereas the time constant for the fluorescence change was 105 ms. E, Development of a delay in the tail currents at +50 mV after increasing lengths of a −100 mV prepulse for Alexa-488-labeled 326C channels. Inset, Voltage protocol used. F, Time course of the development of the tail-current delay (from E), measured as the time t_1/2 (■) for the tail-current amplitude to decay 50%. The ionic current (dotted line) and the fluorescence (red line) during the prepulse are shown for comparison. Note that the fluorescence change overlays the change in tail-current shape, a change that has been shown previously to correlate with the mode shift (Männikkö et al., 2005). Inset, Enlargement of the current during the step to +50 mV, from E. Note that in C, D, and F, the fluorescence signal has been inverted for a direct kinetic comparison with the current traces.

Figure 4.

Fluorescence from consecutive N-terminal S4 residues report on the mode shift. Left, Representative current (top) and fluorescence (bottom) records for Alexa-488 C5-maleimide-labeled residues 324C–326C. Channels were held at 0 mV and then stepped to negative potentials (0 to −140 mV), followed by a step to +50 mV. x- and y-axis calibration bars indicate time (seconds) and current (microamperes), respectively. Note the different timescale for 325C. Center, Normalized current (black) and fluorescence (red) traces during steps to −120 mV. Black squares indicate the time course of the mode shift for each Alexa-488-labeled mutant, measured as the time to half-decay of the tail currents as in Figure 3F. Right, Average time constants for channel activation (I), fluorescence (F), and the mode shift (Shift). *p < 0.01, Student’s t test.

Figure 5.

Mode shift in mammalian HCN channels affects response to trains of IPSPs. A, A train of negative voltage pulses (10 Hz, ΔV of −20 mV), simulating a train of IPSPs, applied to an oocyte expressing HCN1 channels with deleted CNBD (see Materials and Methods). The oocyte was held at −60 mV before the train of IPSPs. After one IPSP, the tail currents were faster than after a train of five IPSPs. Tail potential of −64 mV. Bottom, Normalized tail currents showing a kinetic change in the decay of the HCN1 currents after longer trains of IPSPs. B, C, Computer simulations of model HCN channels (similar to model 1; Materials and Methods). Top, Simulated currents in response to one simulated IPSP (black line) or a train of five simulated IPSPs (red line) (10 Hz, ΔV of −20 mV) from model HCN channels with mode shift (B) and without mode shift (C) (i.e., channels always in mode I). The y-axis is an arbitrary current scale. Inset, Voltage protocol used. Bottom, Normalized tail currents showing a kinetic change in the decay of the HCN currents after longer trains of IPSPs for the model with mode shift, suggesting that the change in the HCN1 tail current kinetics is attributable to the mode shift.

Figure 6.

The mode shift induces an integration effect on the ADP after IPSPs. Simulations of the voltage response to 1 or 10 IPSPs (10 Hz, 20 pA) in a simple passive cell (i.e., below threshold; see Materials and Methods) with HCN channels with a mode shift (A) or HCN channels without any mode shift (B). The HCN channels create a sag in the IPSP response and an ADP after the IPSPs. Enlargement of the ADPs after the IPSP(s) in the model with the mode shift (C; from A) and in the model without the mode shift (D; from B). In the model with a mode shift, the size of the ADP is dependent on the number of IPSP preceding the ADP (C). In the model without a mode shift, the ADP is not very dependent on the number of IPSPs (D).

Figure 7.

Model for pore and voltage-sensor movement during mode shift in HCN channels. The transmembrane domains of HCN were placed after homology modeling with the crystal structure of Kv1.2 channel (Long et al., 2005). Only two of the four pore domains and voltage-sensing domains are shown. We propose the following: (1) S4 undergoes a conformational change during the mode shift that brings the S4-positive charges closer to one or both of the two S2-negative charges, thereby stabilizing the inward position of S4. In addition, a second (2) conformational change occurs in the deeper regions of the pore that allosterically influence the conformational changes in S4. The conformational change of the pore is modulated by K⁺ that binds deep in the selectivity filter and thereby prevents the conformational change. External Cs⁺ can prevent K⁺ binding by binding to a more external site in the pore.