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MOLECULAR AND GENOMIC |
1 Department of Biomedicine and Surgery, Division of Cell Biology, Linköpings Universitet, Linköping, Sweden
2 Department of Neuroscience, The Nobel Institute for Neurophysiology, Karolinska Institutet, Stockholm, Sweden
3 Neurological Sciences Institute, Oregon Health and Science University, Beaverton, OR, USA
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Abstract |
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 Top Abstract IntroductionMethodsResultsDiscussionAppendixReferences |
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(Received 17 May 2006;
accepted after revision 15 June 2006;
first published online 15 June 2006)
Corresponding authors F. Elinder: Department of Biomedicine and Surgery, Division of Cell Biology, Linköpings Universitet, SE-581 85 Linköping, Sweden. Email: fredrik.elinder{at}ibk.liu.se; H. P. Larsson: Neurological Sciences Institute, Oregon Health and Science University, 505 NW 185th Avenue, Beaverton, OR 97006, USA. Email: larssonp{at}ohsu.edu
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Introduction |
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Even though there is a high degree of sequence homology among HCN channels, the different mammalian HCN channels have very different activation kinetics. HCN1 channels activate with a time constant of around 100 ms, HCN2 channels with a time constant of many hundreds of milliseconds, and HCN4 channels with a time constant of many seconds. A number of non-mammalian HCN channels have also been cloned, such as the spHCN channel from the sea urchin. The spHCN channel opens faster than the mammalian HCN channels, and activates with a time constant of around 20 ms.
We found earlier that the spHCN channel undergoes a mode shift in its voltage gating during long voltage steps (> 100 ms) (Männikkö et al. 2005), leading to voltage shifts in the gating charge versus voltage curve, Q(V), and in the conductance versus voltage curve, G(V), of > 50 mV. In addition, the kinetics of the activation and the tail currents are altered in parallel to these voltage shifts. The voltage shifts lead to an apparent voltage hysteresis in the ionic current of the spHCN channel. These mode shifts in the spHCN channel are similar to the voltage shifts of the Q(V) curve in the depolarization-activated ion channels during slow inactivation (Olcese et al. 1997). However, the mode shifts in spHCN occur about 100 times faster than the Q(V) shifts in Shaker K channels, and furthermore, they do not inactivate spHCN.
We hypothesized that the mode shifts in spHCN are caused by a stabilization of the voltage sensor in either the extruded or retracted position (Männikkö et al. 2005). We also found evidence that the mammalian HCN1 channel undergoes a similar hysteresis in its voltage dependence during physiological pacemaker activity (Männikkö et al. 2005). Computer simulations of a SA model cell suggested that this mode shift in voltage gating in the mammalian HCN1 channels prevents arrhythmic behaviour of pacemaker cells (Männikkö et al. 2005). In addition to HCN1 channels, HCN2 and HCN4 are also expressed in the SA node (Santoro & Tibbs, 1999; Shi et al. 1999; Moroni et al. 2001). Elucidation of whether HCN2 and HCN4 also undergo a mode shift is critical for an understanding of the role that HCN channels play as pacemaker channels.
Azene et al. (2005) studied HCN1, HCN2 and HCN4 channels under non-equilibrium conditions and introduced a concept they called I(V) hysteresis, which is slightly different from our mode shift-caused voltage hysteresis. They recorded HCN currents using dynamic voltage clamp simulating action potentials in pacemaker cells and defined I(V) hysteresis as the separation of the current traces during the ascending and the descending voltage (Azene et al. 2005). They concluded that the HCN4 channel does not display any I(V) voltage hysteresis. This result could mean that the slower HCN channels do not undergo the mode shifts observed in the faster HCN channels and do not display the anti-arrhythmic properties of the faster HCN channels.
Here, we have studied the slower cardiac HCN2 and HCN4 channels to investigate whether the mode shifts and voltage hysteresis are preserved in the slower HCN channels. We found that some of the characteristics of the mode shift in the faster HCN channels, such as a speeding up of the activation kinetics in response to longer prepulses, were not present in the ionic currents from HCN2 and HCN4 channels. However, the tail currents from HCN2 and HCN4 channels displayed qualitatively the same characteristics as the tail currents from the faster-activating spHCN and HCN1 channels, indicating that HCN2 and HCN4 channels also undergo mode shifts.
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Methods |
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We performed experiments on hyperpolarized-activated HCN channels expressed in Xenopus oocytes. The mouse HCN2 and human HCN4 channels were used (Ludwig et al. 1999; Chen et al. 2000). Site-directed mutagenesis, cRNA synthesis, and cRNA injection into Xenopus laevis oocytes were performed as previously described (Larsson & Elinder, 2000). Prior to oocyte extraction, the Xenopus laevis were anaesthesized by immersion in a 0.3% solution of tricaine. The effectiveness of anaesthesia on the animal was assessed by the loss of response to a skin pinch, and loss of its ability to right itself after being placed on its back. After oocyte extraction, the incision was closed by the sequential suturing (with absorbable, monofilament thread) of the abdominal musculature and skin. The animals were finally killed using an overdose of tricaine (3% for 30 min), followed by decapitation and pithing of the brain and spinal cord. All experiments were carried out according to the guidelines laid down by our institutions' animal welfare committees.
Electrophysiology and solutions
We recorded the currents using a two-electrode voltage-clamp technique as previously described (Männikkö et al. 2002), with the CA-1B amplifier (Dagan Corp., Minneapolis, MN, USA). We used a ‘100K’-bath solution (mM): 89 KCl, 15 Hepes, 0.4 CaCl2 and 0.8 MgCl2. In some experiments, we used a ‘1K’-bath solution in which 88 mM KCl was changed to NaCl. KOH (or NaOH for low K+ solutions) was added to adjust the pH to 7.4, yielding a final K+ (Na+) concentration of about 100 mM. All experiments were performed at room temperature (20–23°C).
Computer modelling
The equations for the computer modelling used to simulate the ion currents in Fig. 5 are described in the Appendix. The equations and procedures to simulate the action potentials in the SA node (Figs 8 and 9) follow the description in Männikkö et al. 2005), with the values given in the legend to Fig. 8, which in turn is based on an earlier model (Zhang et al. 2000).
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Results |
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We have earlier shown that the tail currents in spHCN and HCN1 channels are clearly dependent on the length of the activating voltage pulse (Männikkö et al. 2005). This unusual feature is in sharp contrast to other non-inactivating, voltage-activated ion channels, which have closing kinetics that are independent of the length of the activating voltage pulse (Hahin, 1988; Zagotta et al. 1994). The prepulse-dependent closing kinetics were not predicted by conventional kinetic models, including a recently developed model for cloned mammalian HCN channels (Altomare et al. 2001; Männikkö et al. 2005). In the spHCN channel, the change in the tail current had a similar time course to the Q(V) shift underlying the voltage hysteresis (Männikkö et al. 2005), suggesting that the change in the tail kinetics and the Q(V) shift were caused by the same mechanism.
In the research reported here, we found that the mammalian HCN2 channel also had prepulse-dependent tail currents. After a brief negative pulse, the tail currents were roughly single exponential, while after longer negative pulses, the tail currents displayed a delay followed by an exponential decay (Fig. 1A and B). The tail currents after long prepulses (e.g. Fig. 1C) were fitted to the following equation:
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| (1) |
is the time constant, and w is an exponent determining the sigmoidicity of the tail. The average values were
= 75 ± 7 ms and w
= 4.3 ± 1.0 at +50 mV (n
= 3). We interpret the change in the tail kinetics to mean that the HCN2 channel has at least four open states and that it enters the additional open states only after the longer, activating pulses. The changes in the tail currents from the mammalian HCN2 channel were similar to the changes in tail currents from the spHCN and HCN1 channels (Männikkö
et al. 2005).
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In the mammalian HCN1 channel, the development of the tail delay was much faster at low external K+ concentrations than at high external K+ concentrations, suggesting that K+ binding slows the hysteresis conformational changes (Männikkö
et al. 2005). HCN2 channels showed a similar dependence on the external K+ concentration. In a 1 mM external K+ concentration, the tail delay developed with a time constant of
= 97 ± 33 ms at –160 mV (n
= 3; Fig. 1D). In a 100 mM external K+ concentration, the tail delay developed with a time constant of
= 377 ± 110 ms at –160 mV (n
= 3; Fig. 1D). The addition of 1 mM Cs+ to the 100 mM K+ solution blocked 91% of the inward current in the HCN2 channel at –160 mV, most likely by an open-channel block by Cs+ (Ludwig et al. 1998; Santoro et al. 1998). The addition of 1 mM Cs+ to the 100 mM K+ solution also prevented external K+ from slowing the development of the tail delay. In a 100 mM K+ solution with 1 mM Cs+, the tail delay developed with a time constant of
= 96 ± 49 ms at –160 mV (n
= 3; Fig. 1D). This finding suggests that external K+ modulates the development of the tail delay through a K+-binding site in the pore.
The development of the tail delay was slightly dependent on the prepulse potential (
= 0.39 ± 0.04, n
= 2; Fig. 2A and B). However, in a sequential model where the mode shift occurs preferentially from the open state, most of the voltage dependence can be attributed to the voltage dependence of opening: for prepulses to more negative potentials; the channels will spend more time in the open state than for prepulses of the same duration to a less negative potential. In Fig. 2C, we have corrected the lengths of the prepulse steps to the relative time the channel actual spends open. With this correction, the voltage dependence is reduced to
= 0.17 ± 0.005 (Fig. 2D; n
= 2), suggesting that the mode shift by itself is not very voltage dependent in HCN2 channels.
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cAMP does not affect the development of the tail delay
Wang et al. (2002) showed that cAMP binding to the HCN2 channel alters the voltage dependence of the HCN2 channel and slows channel closing. Therefore, we investigated a cAMP-insensitive mutant of HCN2 (R591E; Wang et al. 2002) to determine whether the changes in the tail currents in the HCN2 channel are due to cAMP binding. We found that the R591E channel also displays a similar delay in the tail currents after longer hyperpolarizing prepulses (Fig. 3A and B), suggesting that the effect was cAMP independent. Therefore, we conclude that the development of the tail delay in the mammalian HCN2 channel is independent of cAMP binding.
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We earlier showed that the activation kinetics of the spHCN channel were dependent on the length of a hyperpolarizing prepulse (Männikkö
et al. 2005). We interpreted the increasing speed of the activation kinetics in the spHCN channel to be caused by the mode shift and the shift of the Q(V). Therefore, we investigated whether the HCN2 channel also displays a similar prepulse-dependent reactivation. Figure 4A shows the reactivation time course of HCN2 channels at –140 mV after prepulses of different duration, followed by a tail current of 400 ms to +80 mV to allow for the complete closure of the channel. In contrast to the large changes in the reactivation time course previously found in HCN1 and spHCN channels, in HCN2 channels we found either no clear changes or very small changes (< 15%) in the reactivation time course. The activation time constant for HCN2 channels is plotted in Fig. 4B (
); data for HCN1 (
) are shown for comparison. Not even for tail steps of shorter duration (durations that did not completely close all channels) did we find any significant changes in the reactivation time constant (Fig. 4C–E). To investigate possible reasons for this apparent lack of prepulse-dependent reactivation kinetics in the HCN2 channel, we performed computer simulations of a four-state model developed previously (Männikkö
et al. 2005).
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We found earlier that a simple, four-state model could better reproduce the effects of voltage hysteresis in HCN channels (Männikkö et al. 2005) than could other previously proposed HCN-channel models with more states, such as the linear Hodgkin-Huxley-type model with five states (Hodgkin & Huxley, 1952) and an allosteric HCN model with 10 states (Altomare et al. 2001).
In the present investigation, we used our four-state model to examine the possible mode-shift effects on channels with different kinetics (Model 1, see Appendix). In this model, the channel can be in two modes. In mode I, the gating-charge movement and the channel opening occur at very negative potentials, while in mode II, they are shifted to more depolarized potentials. The I 
II transition is favoured in the open state, while the II I transition is favoured in the closed state. We have used the minimal number of parameters necessary to simulate the four-state model. In this model, there are only five independent parameters: VI, VII, z, k and 
. VI is the voltage where 
I and ßI are equal, VII is the voltage where II and ßII are equal, z is the gating charge that moves between the closed and open states C and O (assumed to be equal for mode I and mode II), k is the rate constant of opening () and closing (ß) when the rates are equal (at VI for mode I and at VII for mode II), and is the voltage-independent rate constant between the modes (
=
O
=
C, see Appendix for details).
To explore Model 1, we simulated ion currents with different rates of mode shift . We used the following parameter values: VI
=
–120 mV, VII
=
–60 mV, z
= 2, k
= 10 s–1, and
= 0.1–1000 s–1. Thus, the mode shift is 
Vmode
=
VII
–
VI
= 60 mV (compatible with experimental data; Männikkö
et al. 2005). Figure 5A shows open probabilities for a prepulse-activation step to –130 mV, followed by a tail step to 0 mV and a subsequent reactivation step to –130 mV. The length of the prepulse step varied between 50 and 450 ms, in 50-ms increments. In the four examples shown in Fig. 5A, the currents during the prepulse steps were similar in size and time course, but the tail currents and the reactivation currents differed markedly. Figure 5B shows the tail currents in detail, after normalization and change in time scale. For
= 1 s–1 and 10 s–1, there was a clear difference in the tail currents after the different prepulses, while the tails hardly changed for
= 0.1 and
= 100 s–1. Figure 5C shows the reactivation currents after different lengths of the prepulses at greater magnification. Again, the prepulse effects are clearest for
= 1 and
= 10.
To quantify the change in tail currents, we measured the amplitudes at 10 ms after the onset of the tail current and plotted them versus the prepulse length (Fig. 6A; values for eight different are plotted). The largest changes were seen for
= 1–10 s–1. We also plotted the ratio of the current amplitude after a 50 ms prepulse, divided by the current amplitude after a 500 ms prepulse versus
values (Fig. 6B). The reactivation was analysed in a similar manner. The current after 100 ms reactivation was plotted versus the prepulse length in Fig. 6C, which shows how reactivation varied with . The relative reactivation change was plotted versus
values in Fig. 6D. The largest effect of the mode shifts was again seen for
= 3 s–1, but it was much smaller than for the tail currents. To compare the effects of the mode shifts on the tail currents and the reactivation currents, the relative current changes were normalized and plotted versus the quotient k/ (Fig. 6E). As shown in Fig. 6E, both peaked around k/
= 3, when the activation rate was three times faster than the mode shift rate. However, they differed markedly in that the curve for reactivation was much sharper. For example, at k/
= 0.33, large tail-current effects were seen, while hardly any reactivation effects were seen. This result is probably due to the necessity for the channels to be closed in order to measure the reactivation rate, which allows some portion of the channel population to recover to mode I during the tail pulse.
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= 1–10 s–1). From our data, we calculated the k/ value for each of the HCN channels: spHCN k/
3, HCN1 k/
1, and HCN2 k/
0.3 (see Table 1). Thus, for spHCN and HCN1 channels, the k/ was in the range of values for which the tail current changes were as large as possible (Fig. 6B). For HCN2 channels, which have a smaller k/ ratio, the simulations predict a clear but small change in tail current but essentially no change for the reactivation (Fig. 6D) – predictions that were similar to our findings for the HCN2 channel (Fig. 4). However, for a channel such as the HCN4 channel, with 10 times slower activation kinetics (i.e. k/
= 0.03), no tail change is predicted.
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We tested whether the slowest member of the mammalian HCN channel family, HCN4, also undergoes mode shifts. In Fig. 7A and B, we show that the HCN4 channel also has prepulse-dependent tail currents, as do the other HCN channels that we tested. After a brief negative pulse, the tail currents of HCN4 were roughly single exponential, while after longer negative pulses, they displayed a delay followed by an exponential decay (Fig. 7B). The development of this delay had a time constant of
= 533 ± 188 ms at –160 mV (n
= 2; Fig. 7C) in 100 mM K+. We interpret this finding to mean that the HCN4 channel has more than one open state and that the channel enters additional open states only after longer activating pulses.
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mode shift is too fast compared to the activation rate in the HCN4 channel. In 100 mM K+, the HCN4 channel had a ratio of k/
= 0.78 at –160 mV (opening
= 687 ± 258 ms, mode shift
= 533 ± 188 ms), which was at the lower range of where tail changes are detectable in the computer simulations (Fig. 6E). In more physiological solutions (low external K+ solutions, which probably speed up the mode-shift kinetics) and more physiological voltages (less-negative activation voltages, which slow the activation kinetics), we expect k/ to be even smaller (see Table 1). Thus, the changes in the tail currents from the mammalian HCN4 channel were similar to the changes we found in the tail currents from spHCN, HCN1 and HCN2 channels (Männikkö
et al. 2005; Fig. 1), suggesting that all HCN channels may undergo mode shifts. Simulation of pacemaker activity in SA cells
In a previous investigation, we showed with computer simulations that a two-state HCN1 channel produced arrhythmic firing in a sino-atrial model cell. The arrhythmic firing was prevented by the introduction of a mode shift of 60 mV in the HCN channel model, making it a two-mode, four-state HCN channel model (Männikkö
et al. 2005). To test the effect of the mode shift in slower HCN channels, we performed simulations of a SA node cell (Zhang et al. 2000) with HCN channels, with different k/ relations. As seen in Fig. 8A, there was a clear arrhythmia in the SA model with a two-state HCN channel that had no mode shift. To quantify this arrhythmia, we measured the time between the peaks of the action potentials for a period of about 16 s and calculated the mean and the root-mean-square (r.m.s.) deviation for the time between peaks in simulations with k values between 1 and 1000 s–1 (Fig. 8C). For slow- and fast-activating channels (i.e. for small and large k values) the r.m.s. deviation was almost zero, but for a broad range of k values (k
= 10–1000), the time between peaks greatly varied, indicating arrhythmia in the firing rate.
Introducing a mode shift of 60 mV in the HCN channels completely eliminated this arrhythmia (Fig. 8B). This introduction of a mode shift had an anti-arrhythmic effect for all k values (Fig. 8C, dashed line, r.m.s. = 0). In Fig. 8C, we indicate the activation kinetics for the four HCN channels in which we studied the mode shift (Table 1). We found HCN1 and spHCN in the arrhythmic region, while HCN2 was just outside this region. HCN4 was clearly outside the arrhythmic region. However, we note that our experiments were performed at room temperature, not at body temperature. Assuming a Q10 = 3 (i.e. the kinetics of opening increased by a factor of 3 when the temperature increased by 10°C; (Halliwell & Adams, 1982; Tokimasa & Akasu, 1990; Magee, 1998), we expect that the opening kinetics are roughly five times faster at 37°C (dotted arrows in Fig. 8C) than at room temperature. Thus, at 37°C, HCN2 is clearly in the range where a two-state HCN channel could cause arrhythmia, and where a mode shift could prevent the arrhythmia. Even at 37°C, the HCN4 channel was outside the arrhythmic range.
Why are there fast HCN channels in pacemaker cells, if they potentially can cause arrhythmia? We propose that fast HCN channels are critical to achieving a wider frequency range for the autonomic regulation of the heart rate through changes in the intracellular cAMP concentration. Sympathetic stimulation increases the intracellular cAMP concentration, and a parasympathetic stimulation decreases the cAMP concentration (DiFrancesco, 1993; Robinson & Siegelbaum, 2003; Baruscotti et al. 2005). cAMP modulates HCN channels by shifting the G(V) curve in a positive direction along the voltage axis and by speeding up the activation kinetics (DiFrancesco, 1993; Robinson & Siegelbaum, 2003; Baruscotti et al. 2005). We investigated the frequency versus cAMP relation by simulating the cell activity of the SA node using HCN channels with different midpoints of the G(V) curve. For an SA cell without HCN channels, the automatic rhythm was 3.0 Hz, with no dependence on cAMP (dashed line in Fig. 8D). Introducing a slow HCN channel (i.e. HCN4 channels) without a mode shift (i.e. a two-state model) increased the heart rate, but the dependence of the firing rate on cAMP was low – a G(V) shift of 30 mV was required to increase the heart rate from 4 to 5 Hz (• in Fig. 8D). An increase in the opening kinetics led to a higher cAMP-to-frequency dependence, but this increase introduced arrhythmia in the firing rate (see Fig. 8A and C). However, the addition of a mode shift in combination with the faster opening kinetics increased the cAMP-to-frequency dependence without causing arrhythmia (Fig. 8D). For example, the addition of a mode shift increased the cAMP-to-frequency dependence by a factor of 2.5 (
) compared to the cAMP-to-frequency dependence for the two-state model without a mode shift.
Why is the pacemaker model cell firing arrhythmic (Fig. 8A) in some cases and rhythmic (Fig. 8B) in other cases? In Fig. 9, we show the membrane voltage, the major ionic currents, and the HCN channel open probability in a rhythmic simulation compared to an arrhythmic simulation. All parameters are identical between the two simulations (number of channels, activation kinetics, steady-state activation voltage, etc.). The only difference is that in the arrhythmic case we used a one-mode HCN channel with V
=
–75 mV, while in the rhythmic case we used our two-mode HCN channel with VI
=
–105 mV and VII
=
–45 mV (i.e. V
=
–75 ± 30 mV). In the rhythmic simulation (Fig. 9A), the initial hyperpolarization, in addition to closing the L-type Ca2+ channels (red line), causes an increased current through the HERG channel (green line), due to fast recovery from inactivation and slow closing in HERG channels. Further hyperpolarization closes HERG channels and activates HCN channels. The inward current through HCN channels (blue line) slowly depolarizes the cell, which activates the L-type Ca2+ channels. This in turns triggers a new action potential, which closes HCN channels, activates and quickly inactivates HERG channels, and activates other K+ channels (not shown for clarity). The current through the K+ channels leads to a slow hyperpolarization, which again recovers the HERG channels and closes the Ca2+ channels. In the arrhythmic simulation (Fig. 9B), the initial hyperpolarizing phase is similar to the rhythmic case (compare continuous and dashed lines in Fig. 9C). However, the hyperpolarization activates more HCN channels, which leads to a faster depolarization and a curtailed hyperpolarization. The shorter time spent at hyperpolarized potentials leads to fewer HERG channel having time to close. This increased outward current through open HERG channels (at t
= 4.1 s) prevents the Ca2+ channels from causing a new action potential. It is not until enough HERG channels have closed (at t
= 4.3 s) that the Ca2+ channels can initiate a new delayed action potential. The larger HCN current in the arrhythmic simulation is due to the fact that these HCN channels open at a less hyperpolarized potential than in the arrhythmic case, where the HCN channels have to first open through mode I which has a more hyperpolarized activation voltage. Eventually, the channels will reach the same open probability in both cases (at t
= 4.15 s), since the two-mode channels will open to mode II by mass action. The increased open probability at hyperpolarized potentials occurs for all HCN channels with kinetics that are roughly faster than the frequency of the action potentials (f
= 5 s–1). For slower HCN channels (k < 5 s–1), the channels will not have time to significantly increase their open probability during the hyperpolarization, and therefore the HCN channels cannot speed up the depolarization phase and prevent HERG channel closing. This explains why slower two-state HCN channels will not introduce arrhythmia in the pacemaker model (Fig. 8C).
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Discussion |
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of activation kinetics (k) to mode shift kinetics (). Only in the middle range of k/ ratios (0.3–30) are the effects from the mode shifts easily observable. However, the range was slightly different for the different voltage protocols. For example, the changes in activation kinetics were only seen for a narrower range of k/ ratios (1–30). We propose that all HCN channels undergo mode-shift conformational changes and that these changes underlie the voltage hysteresis, but that in the slower HCN channels, the kinetic effects from the mode shifts are not readily observable. The mechanism for the mode shift
What underlies the mode shifts or voltage hysteresis in HCN channels? We hypothesized earlier that once an HCN channel opens, a conformational change occurs at the interface between S4 and the pore domain that stabilizes the open state (Männikkö et al. 2005). This conformational change gives rise to a depolarizing shift in the voltage dependence of activation, thereby creating the voltage hysteresis. The rate of the mode shift is similar across the different HCN channels: 70–140 ms in 1 mM K+ and 200–600 ms in 100 mM K+ (see Table 1). These rates are similar despite the very different activation time constants for the different HCN channels: the fast spHCN channel opens with a time constant of 20 ms, while the slow HCN4 channel opens with a time constant of 4000 ms. In all mammalian HCN channels that we tested (i.e. HCN1, HCN2, and HCN4), the rate of mode shift depended on the external K+ concentrations; the mode shifts significantly slowed when we increased the K+ concentrations, suggesting that a conserved process in all HCN channels causes the mode shift. The slowing of the mode shift in HCN channels by elevating the external K+ concentrations could be caused by a foot-in-the-door effect by K+ (Lopez-Barneo et al. 1993) – that is, K+ binding in the pore slows the mode shift transition. At low K+ concentrations, or with a high concentration of an external blocker (e.g. Cs+) that prevents K+ access to the pore, the binding site is empty, leading to a fast mode-shift transition.
A possible mechanism for the transient increase in tail current
In contrast to the tail currents in spHCN and HCN1 channels, the tail currents in HCN2 channels displayed a small rising phase before the decline – a hook (< 4% in total amplitude). The reason for this hook is not clear. The four-state model, in which we have assumed that the two open states have the same single-channel conductance, does not reproduce the hook in the tail currents. One possible explanation for the hook is that different open states in HCN2 channels have slightly different single-channel conductances. The sigmoidicity of the tail currents (w = 4.3) suggests that there are more than four open states (Fig. 1), and we have earlier suggested that a two-mode 20-state model would better reproduce the kinetics details of the ion currents in the HCN channels (Männikkö et al. 2005). In simulations of two-mode HCN-channel models that had more than one open state in each mode, the rises in the tail currents were reproduced when some of the open states in mode II had a single-channel conductance that was 80–90% of the conductance in the other open states (data not shown). Another possible explanation for the hook is that some of the HCN2 channels may undergo some type of inactivation during the hyperpolarization pulse, in which case the hook would represent the recovery from this inactivation during the tail currents. HCN2 channels have been proposed to undergo a type of inactivation that is holding potential dependent (Shin et al. 2004). However, this inactivation mechanism, in its present form, does not explain the hook, because the recovery from inactivation in that model is through the closed state, not through the open state (Shin et al. 2004). Therefore, this inactivation model does not generate any significant hook in the tail currents.
The role of the mode shift for pacemaker activity
Azene et al. (2005) cite the slow gating of the HCN4 channel as the reason for the absence of I(V) hysteresis in their recordings of the HCN4 channel. However, we propose that the HCN4 channel undergoes mode shifts, similar to those that give rise to voltage hysteresis in the spHCN channel. Here we have shown that it is very hard to detect mode shifts in channels that have activation kinetics that are slower than the mode-shift kinetics, which is probably the case for the HCN4 channel in low (physiological) external K+ concentrations (Fig. 4E). However, we propose that the slower HCN4 channel does undergo mode shifts in physiological conditions, but that these mode shifts do not give rise to any apparent voltage hysteresis in the ionic currents due to the slow activation kinetics.
Why do HCN channels undergo mode shifts and voltage hysteresis? In computer simulations, we have earlier shown that voltage hysteresis in the faster HCN1 channel is important for stabilizing the rhythmic firing in a pacemaker cell model (Männikkö et al. 2005). Here, we have also shown that HCN2 channels, at 37°C, are important for stabilizing the firing rate in a pacemaker cell model. In addition, we show that faster HCN channels without mode shifts cause a shorter and less negative hyperpolarization after an action potential, thereby reducing the number of HERG channels that close during the hyperpolarization. This leads to an outward K+ current that prevents and delays the initiation of the next action potential, thereby causing arrhythmic firing. The slower HCN4 channels are not fast enough to change their open probability substantially during the rhythmic firing of, for example, the SA node cells; therefore, HCN4 channels do not induce arrhythmic firing in a model of the SA node cell. Therefore, the mode shifts in the HCN4 channel most likely do not play a role in stabilizing the rhythmic behaviour of pacemaker cells. However, the mode shifts may be important for shifting the open probability of these slower HCN channels to a more depolarized, physiological voltage range. Furthermore, HCN1, HCN2 and HCN4 subunits are all expressed in the SA node, and it has been shown that different HCN subunits can combine to form heterotetrameric channels with intermediate opening kinetics (Chen et al. 2001; Ulens & Tytgat, 2001; Altomare et al. 2003). These heteromeric channels have opening kinetics (Chen et al. 2001; Ulens & Tytgat, 2001; Altomare et al. 2003) in a range that would cause arrhythmia in our model of the SA node cell, if these channels were simple two-state channels. However, these heteromeric HCN channels probably undergo a mode shift that prevents arrhythmia in pacemaker cells.
The simulations in this study were done with only one subtype of HCN channels at a time. A more physiological model would incorporate all three subtypes of HCN-channel subunits (and potentially heteromeric HCN channels). However, the actual channel subtypes underlying the f current in SA node cells and the relative contribution of the different subtypes to the f currents are not known. For example, the f currents are faster that the currents from homomeric HCN4 channels (Altomare et al. 2003). Even currents from a heteromeric channel with 50% HCN1 subunits and 50% HCN4 subunits are slower than the f currents (Altomare et al. 2003). This suggests that a high percentage of the f currents is generated by channels constituted of faster HCN-channel subunits or that some unknown cofactor speeds up the HCN channel currents in SA node cells. Our simulations show that faster HCN channels have a propensity to cause arrhythmia in SA node cells, and that the mode shift removes the arrhythmia. We have not tried to mix different subtypes of HCN channels in our model since the relative contributions of the HCN-channel subunits underlying the f currents have not yet been experimentally determined.
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Appendix |
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TopAbstractIntroductionMethodsResultsDiscussionAppendixReferences |
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| (Model 1) |
II transition is favoured in the open states, while the II I transition is favoured in the closed states. The voltage-dependent rate constants i and ßi are described by
|
| (A1) |
|
| (A2) |
i
=
ßi), zi and zßi are the valencies of the transitions (total gating charge zi
=
zi
+
zßi), e0 is the elementary charge, V is the membrane voltage, kb is Boltzmann's constant, T is the absolute temperature, and i indicates mode I or II. We assumed all mode shift rates (vertical transitions) to be voltage independent (thus, zi
=
zii
=
z); furthermore, we assumed that O
=
C; C
=
O; k
=
kI
=
kII; and 
. All of these assumptions were compatible with experimental data (Männikkö
et al. 2005). The difference between VII and VI is the mode shift, denoted by Vmode. Thus:
|
| (A3) |
|
| (A4) |
|
| (A5) |
|
| (A6) |
To explore Model 1, we simulated ion currents with different rates of mode shift , where
=
O
=
C. In these computations, we used the following parameter values: VI
=
–120 mV, VII
=
–60 mV, z
= 2, k
= 10 s–1, and
= 0.1–1000 s–1. Thus, the mode shift is Vmode
=
VII
–
VI
= 60 mV (compatible with experimental data; Männikkö
et al. 2005).
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References |
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TopAbstractIntroductionMethodsResultsDiscussionAppendixReferences |
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Altomare
C, Terragni
B, Brioschi
C, Milanesi
R, Pagliuca
C, Viscomi
C, Moroni
A, Baruscotti
M
&
DiFrancesco
D (2003). Heteromeric HCN1-HCN4 channels: a comparison with native pacemaker channels from the rabbit sinoatrial node. J Physiol
549, 347–359.
Azene
EM, Xue
T, Marban
E, Tomaselli
GF
&
Li
RA (2005). Non-equilibrium behavior of HCN channels: insights into the role of HCN channels in native and engineered pacemakers. Cardiovasc Res
67, 263–273.
Baruscotti M, Bucchi A & Difrancesco D (2005). Physiology and pharmacology of the cardiac pacemaker (‘funny’) current. Pharmacol Ther 107, 59–79.[CrossRef][Medline]
Chen
J, Mitcheson
JS, Lin
M
&
Sanguinetti
MC (2000). Functional roles of charged residues in the putative voltage sensor of the HCN2 pacemaker channel. J Biol Chem
275, 36465–36471.
Chen
S, Wang
J
&
Siegelbaum
SA (2001). Properties of hyperpolarization-activated pacemaker current defined by coassembly of HCN1 and HCN2 subunits and basal modulation by cyclic nucleotide. J Gen Physiol
117, 491–504.
DiFrancesco D (1993). Pacemaker mechanisms in cardiac tissue. Annu Rev Physiol 55, 455–472.[CrossRef][Medline]
Hahin
R (1988). Removal of inactivation causes time-invariant sodium current decays. J Gen Physiol
92, 331–350.
Halliwell JV & Adams PR (1982). Voltage-clamp analysis of muscarinic excitation in hippocampal neurons. Brain Res 250, 71–92.[CrossRef][Medline]
Hodgkin
AL
&
Huxley
AF (1952). A quantitative description of membrane current and its application to conduction and excitation in nerve. J Physiol
117, 500–544.
Larsson HP & Elinder F (2000). A conserved glutamate is important for slow inactivation in K+ channels. Neuron 27, 573–583.[CrossRef][Medline]
Lopez-Barneo J, Hoshi T, Heinemann SH & Aldrich RW (1993). Effects of external cations and mutations in the pore region on C-type inactivation of Shaker potassium channels. Receptors Channels 1, 61–71.[Medline]
Ludwig A, Budde T, Stieber J, Moosmang S, Wahl C, Holthoff K, Langebartels A, Wotjak C, Munsch T, Zong X, Feil S, Feil R, Lancel M, Chien KR, Konnerth A, Pape HC, Biel M & Hofmann F (2003). Absence epilepsy and sinus dysrhythmia in mice lacking the pacemaker channel HCN2. EMBO J 22, 216–224.[CrossRef][Medline]
Ludwig A, Zong X, Hofmann F & Biel M (1999). Structure and function of cardiac pacemaker channels. Cell Physiol Biochem 9, 179–186.[CrossRef][Medline]
Ludwig A, Zong X, Jeglitsch M, Hofmann F & Biel M (1998). A family of hyperpolarization-activated mammalian cation channels. Nature 393, 587–591.
Magee
JC (1998). Dendritic hyperpolarization-activated currents modify the integrative properties of hippocampal CA1 pyramidal neurons. J Neurosci
18, 7613–7624.
Männikkö R, Elinder F & Larsson HP (2002). Voltage-sensing mechanism is conserved among ion channels gated by opposite voltages. Nature 419, 837–841.
Männikkö
R, Pandey
S, Larsson
HP
&
Elinder
F (2005). Hysteresis in the voltage dependence of HCN channels: conversion between two modes affects pacemaker properties. J Gen Physiol
125, 305–326.
Moroni
A,
Gorza
L, Beltrame
M, Gravante
B, Vaccari
T, Bianchi
ME, Altomare
C, Longhi
R, Heurteaux
C, Vitadello
M, Malgaroli
A
&
DiFrancesco
D (2001). Hyperpolarization-activated cyclic nucleotide-gated channel 1 is a molecular determinant of the cardiac pacemaker current If. J Biol Chem
276, 29233–29241.
Nolan MF, Malleret G, Lee KH, Gibbs E, Dudman JT, Santoro B, Yin D, Thompson RF, Siegelbaum SA, Kandel ER & Morozov A (2003). The hyperpolarization-activated HCN1 channel is important for motor learning and neuronal integration by cerebellar Purkinje cells. Cell 115, 551–564.[CrossRef][Medline]
Olcese
R, Latorre
R, Toro
L, Bezanilla
F
&
Stefani
E (1997). Correlation between charge movement and ionic current during slow inactivation in Shaker K+ channels. J Gen Physiol
110, 579–589.
Pape HC (1996). Queer current and pacemaker: the hyperpolarization-activated cation current in neurons. Annu Rev Physiol 58, 299–327.[CrossRef][Medline]
Robinson RB & Siegelbaum SA (2003). Hyperpolarization-activated cation currents: from molecules to physiological function. Annu Rev Physiol 65, 453–480.[CrossRef][Medline]
Santoro B, Liu DT, Yao H, Bartsch D, Kandel ER, Siegelbaum SA & Tibbs GR (1998). Identification of a gene encoding a hyperpolarization-activated pacemaker channel of brain. Cell 93, 717–729.[CrossRef][Medline]
Santoro B & Tibbs GR (1999). The HCN gene family: molecular basis of the hyperpolarization-activated pacemaker channels. Ann N Y Acad Sci 868, 741–764.[CrossRef][Medline]
Schulze-Bahr E, Neu A, Friederich P, Kaupp UB, Breithardt G, Pongs O & Isbrandt D (2003). Pacemaker channel dysfunction in a patient with sinus node disease. J Clin Invest 111, 1537–1545.[CrossRef][Medline]
Shi W, Wymore R, Yu H, Wu J, Wymore RT, Pan Z, Robinson RB, Dixon JE, McKinnon D & Cohen IS (1999). Distribution and prevalence of hyperpolarization-activated cation channel (HCN) mRNA expression in cardiac tissues. Circ Res 85, e1–6.
Shin KS, Maertens C, Proenza C, Rothberg BS & Yellen G (2004). Inactivation in HCN channels results from reclosure of the activation gate: desensitization to voltage. Neuron 41, 737–744.[CrossRef][Medline]
Tokimasa
T
&
Akasu
T (1990). Cyclic AMP regulates an inward rectifying sodium-potassium current in dissociated bull-frog sympathetic neurones. J Physiol
420, 409–429.
) solutions. In this experiment, the time constants were 59 ms, 580 ms and 35 ms, respectively. Holding potential (Vhold) = 0 mV in all recordings.
), measured as the tail current amplitude 50 ms after the onset of the tail potential, fitted with a single exponential. The x-axis was corrected for the relative time that the channel spent open during the prepulse, i.e. instead of plotting tail current versus prepulse length (as in 
