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NEUROSCIENCE |
1 Physiologisches Institut der Universität Freiburg, Hermann-Herder-Str. 7, D-79104 Freiburg, Germany
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Abstract |
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(Received 20 December 2005;
accepted after revision 5 May 2006;
first published online 11 May 2006)
Corresponding author P. Jonas: Physiologisches Institut, Universität Freiburg, Hermann-Herder-Str. 7, D-79104 Freiburg, Germany. Email: peter.jonas{at}physiologie.uni-freiburg.de
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Introduction |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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BCs are fast signalling devices with highly specialized membrane properties (Jonas et al. 2004). BCs show fast excitatory postsynaptic potentials and currents, mediated by rapidly gated AMPA-type glutamate receptors (Geiger et al. 1997). BCs generate APs at frequencies of several hundred Hertz under physiological conditions, as a result of the expression of voltage-gated Na+ channels with rapid recovery from inactivation (Martina & Jonas, 1997) and voltage-gated K+ channels with high activation threshold and rapid deactivation (Martina et al. 1998). Finally, BCs show a relatively positive resting potential, a low input resistance and a fast apparent membrane time constant (Geiger et al. 1997; Mott et al. 1997; Lübke et al. 1998; Jonas et al. 2004). While the positive resting potential sets the BCs into a ready-to-fire mode near the AP threshold from which excitatory inputs can efficiently trigger APs (Fricker et al. 1999), the fast apparent membrane time constant leads to a brief duration of excitatory postsynaptic potentials (EPSPs), allowing BCs to selectively respond to coincident principal neuron activity (Geiger et al. 1997).
Although the passive membrane properties are key determinants of synaptic integration and the input–output relation of fast-spiking BCs, the molecular basis is poorly understood. In the simplest possible scenario, resting potential and input resistance of a cell are determined by a single conductance. Primary candidates for such a conductance would be inwardly rectifying K+ channels or two-pore domain K+ channels (Goldstein et al. 2001). However, both types of channels are highly K+-selective and thus cannot explain the combination of depolarized resting potential and low input resistance. Alternatively, resting potential and input resistance could be determined by the opposing actions of K+-selective conductances and the non-selective cation conductance Ih, as recently proposed for cortical pyramidal neurons (Day et al. 2005; reviewed by Pape, 1996; Robinson & Siegelbaum, 2003). Ih channels are assembled from four subunits (HCN1–HCN4), which are abundantly expressed in the central nervous system (Pape, 1996; Gauss et al. 1998; Ludwig et al. 1998; Santoro et al. 1998, 2000; Magee, 1999; Williams & Stuart, 2000; Vasilyev & Barish, 2002). Functional studies indicate that Ih is expressed in hippocampal dendrite-inhibitory interneurons (Maccaferri & McBain, 1996; Lupica et al. 2001). Furthermore, immunocytochemical analysis revealed that HCN1–4 subunits are present in axons and presynaptic terminals of GABAergic interneurons in the hippocampus (Notomi & Shigemoto, 2004). However, unlike dendrite-inhibitory interneurons (Maccaferri & McBain, 1996; Lupica et al. 2001), BCs do not show an obvious ‘sag’ during hyperpolarizing current injection (Lübke et al. 1998), which argues against the expression of Ih. Thus, whether fast-spiking BCs express Ih has remained unknown.
In addition to setting resting potential and input resistance, several alternative functions of Ih have been suggested. First, Ih could contribute to the afterpotential following a single AP or a train of APs (Maccaferri et al. 1993). Second, Ih may participate in the regulation of transmitter release, as proposed at the crayfish neuromuscular junction (Beaumont & Zucker, 2000) and at hippocampal mossy fibre terminals (Mellor et al. 2002). Finally, Ih is thought to be involved in rhythmogenesis. For example, Ih contributes to slow rhythmic activity in thalamocortical neurons (McCormick & Pape, 1990). By analogy, if Ih was expressed in fast-spiking GABAergic interneurons, it could have major influence on the AP phenotype, the properties of GABA release at interneuron output synapses, and the rhythmic properties of interneuron networks (Bartos et al. 2002; Vida et al. 2006). However, none of these hypotheses has been tested experimentally.
In this paper, we attempted to answer three questions. First, do fast-spiking BCs in the hippocampus express HCN channels? Second, if so, what is the subunit composition of these channels? Third, are HCN channels also expressed in axons or presynaptic terminals of fast-spiking GABAergic interneurons? We found that HCN channels are expressed in BCs of the dentate gyrus at both somatodendritic and axonal locations and that they shape both the input and the output properties of these cells.
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Methods |
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Transverse hippocampal slices (thickness, 300 µm) were cut from the brains of 17- to 21-day-old Wistar rats using a vibratome (Dosaka, Kyoto, Japan). Animals were killed by rapid decapitation without anaesthesias in accordence with national and institutional guidelines. Experiments were approved by the Animal Care Committe Freiburg according to 
15 of the Tierschutzgesetz (registry T-04/10). Experiments were performed under visual control using infrared differential interference contrast videomicroscopy. BCs in the dentate gyrus were identified by the location of their soma in the granule cell layer near the hilar border and their fast-spiking AP phenotype (AP frequency for 800 ms, 1 nA pulses > 50 Hz; Martina et al. 1998). Current- and voltage-clamp recordings were made with a Multiclamp 700A amplifier (Axon Instruments, Union City, CA, USA). Patch pipettes were pulled from thick-walled borosilicate glass tubing. When filled with internal solution, the resistance was 1.5–3 M
. Series resistance was 6–15 M; it was compensated in current-clamp recordings, and carefully monitored but not compensated during voltage-clamp recordings. Signals were low-pass filtered at 2, 4 or 10 kHz (4-pole low-pass Bessel), and sampled at 5, 10 or 40 kHz. Pulse generation and data acquisition were performed using a 1401plus interface (CED, Cambridge, UK)/PC system with FPulse (home-made) running under Igor (version 5.01, Lake Oswego, OR, USA). In current-clamp experiments, the resting membrane potential was measured directly after obtaining access to the cell interior (–61.1 ± 0.3 mV; range, –65 to –54 mV; n
= 73) and then set to –70 mV by injecting a constant negative holding current (
150 pA). In voltage-clamp recordings, the holding potential was set to –50 mV. For antidromic AP propagation experiments, a stimulus electrode (glass pipette filled with Hepes-buffered Na+-rich solution; resistance 
1 M) was placed in the granule cell layer at a distance of 200–1000 µm from the soma of the recorded BC. The stimulus intensity was 3–16 V and the stimulus duration was 0.2 ms. The location of the stimulation pipette tip was monitored carefully during the experiment, and the recording was terminated if a shift was observed. Effects of both ZD 7288 and Cs+ were assessed between 10 and 30 min after onset of the application of the substance. The recording temperature was 21–24°C.
Recording of miniature IPSCs (mIPSCs) and GABA-activated currents in granule cells
mIPSCs in dentate gyrus granule cells were recorded in the whole-cell configuration in the presence of 1 µM TTX, 10 µM 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX) and 20 µM D-2-amino-5-phosphonopentanoic acid (D-AP5). The internal solution contained 140 mM KCl, and the holding potential was set to –70 mV. Currents activated by 10-ms pulses of 1 mM GABA were measured in outside-out and nucleated patches isolated from granule cells using fast-application techniques (Jonas, 1995). The exchange time (20–80%), measured with an open patch pipette during a change between 100% and 10% Hepes-buffered Na+-rich external solution, was 200–300 µs for nucleated patch recordings (perfusion rate, 50–70 µl min–1) and 50–150 µs for outside-out patches (perfusion rate, 200 µl min–1). Pulses of GABA were applied every 5 or 8 s. Patches were held at –50 mV throughout the recordings.
Analysis
In current-clamp experiments, input resistance was determined from the voltage at the end of 800-ms hyperpolarizing current pulses. Apparent membrane time constants were measured by plotting voltage differences logarithmically in the range of 100% to 5% of the maximal amplitude, and fitting the final 20-ms epoch of each trace by linear regression. AP amplitude and half-duration were measured from the baseline preceding the current pulse. The maximal rate of rise was determined from the first derivative of the AP waveform. The fast afterhyperpolarization (AHP) amplitude was measured as the minimum voltage directly after the AP, the slow AHP was determined as the mean voltage after the fast AHP (18 ms after spike onset). AP frequency was determined as the inverse of the mean interspike interval. In voltage-clamp experiments, quantitative analysis of Ih was performed by digital subtraction of traces in the absence and presence of ZD 728 or Cs+ (see Figs 3 and 4). Rise and decay of current traces were fitted by exponential functions. Conductance–voltage data were fitted with a Boltzmann function:
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= 1/(
+
ß), with (V) =
aexp[V/k1] and ß(V) =
bexp[–V/k2], where a and b are rate constants at 0 mV and k1 and k2 are slope factors. Conductance was calculated as G
=
I/(V
–
Vrev), where I is current and Vrev is reversal potential. For display purposes, currents between 0 and 1 ms after voltage jumps were blanked. Fitting was made using the non-linear least-squares algorithms of Mathematica 4.1 (Wolfram Research, Champaign, IL, USA). PNa/PK was determined from Vrev according to the Goldman equation:
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The physiological external solution contained (mM): NaCl 125, NaHCO3 25, KCl 2.5, NaH2PO4 1.25, CaCl2 2, MgCl2 1 and glucose 25. The Hepes-buffered Na+-rich external solution contained (mM): NaCl 135, KCl 5.4, CaCl2 1.8, MgCl2 1 and Hepes 5; pH adjusted to 7.2 with NaOH. The internal solution used in electrophysiological recordings contained (mM): KCl 140 or potassium gluconate 120 and KCl 20, EGTA 10, MgCl2 2, Na2ATP 2 and Hepes 10; pH adjusted to 7.3 with KOH. The internal solution used for reverse transcription followed by quantitative polymerase chain reaction (RT-qPCR) experiments contained (mM): KCl 140, EGTA 5, MgCl2 3 and Hepes 5; pH adjusted to 7.3 with KOH. Chemicals were as follows: 4-ethylphenylamino-1,2-dimethyl-6-methylaminopyrim-idinium chloride (ZD 7288, Tocris), 6-cyano-7-nitro-quinoxaline-2,3-dione (CNQX, Tocris), D-2-amino-5-phosphonopentanoic acid (D-AP5, Tocris), bicuculline methiodide (BIC, Tocris), tetrodotoxin (TTX, Alomone), 4-aminopyridine (4-AP, Sigma) and 
-aminobutyric acid (GABA, Sigma).
Single-cell RT-qPCR
Single-cell expression analysis using RT-qPCR was performed as previously described (Monyer & Jonas, 1995; Franz et al. 2000; Lien et al. 2002; Liss, 2002). Patch-clamp capillaries were heated for 4 h at 220°C prior to use. After determining the AP phenotype in the whole-cell configuration, the cytoplasm of a recorded fast-spiking BC was harvested into the recording pipette. Experiments in which the gigaseal was disrupted or debris was attached to the outer surface of the pipette were discarded. A 0.5-ml reaction tube was filled with 6 µl reverse transcription (RT) reaction buffer containing 1.4 x first strand buffer, 24 mM dithiothreitol (DTT), deoxynucleotidetriphosphates (dNTPs, 1.2 mM each), 1.7 x hexanucleotide mix (Hoffmann-La Roche, Basel, Switzerland) and 3 pmol Oligo(dT) primer. After expelling the contents of the patch pipette (8 µl), the RT reaction was initiated by adding 100 U Superscript II (Invitrogen, Karlsruhe, Germany) and 20 U RNasin (Promega, Mannheim, Germany). After incubation for > 2 h at 37°C, cDNA was ethanol-precipitated in the presence of 1 µg glycogen (Ambion, Huntingdon, Cambridgeshire, UK), 250 ng poly(C) RNA (Amersham Biosciences, Freiburg, Germany) and 250 ng poly(dC) DNA (Amersham) with 0.1 vol 3 M Na acetate (pH 4.8) and 3.5 vol ethanol at –20°C overnight and followed by centrifugation (4°C, 20 000 g, 50 min). The supernatant was replaced by 100 µl 75% ethanol and then centrifuged again (4°C, 20 000 g, 40 min). After removal of the supernatant, the cDNA pellet was dried at 45°C, and resuspended in 8 µl sterile water (Sigma) and 27 µl of 2 x TaqMan MasterMix (Applied Biosystems, Darmstadt, Germany).
The cDNA solution of a single cell was split into two aliquots (16.5 µl), each used for the amplification of a single cDNA. Real-time qPCR was performed in a total volume of 25 µl with 880 nM primer (each) and 200 nM TaqMan probe in an ABI Prism 7000 sequence detection system (Applied Biosystems) using a default temperature cycle protocol (50°C for 2 min, 95°C for 10 min followed by 48 cycles with 95°C for 15 s and 60°C for 1 min). Primers and TaqMan probes were designed with PrimerExpress software (version 2.0; Applied Biosystems) and selected for maximal specificity and intron-overspanning amplicons. Sequences of primers and Taqman probes were as follows (F, forward; R, reverse; T, TaqMan probe; referring to published sequences in the GenBank of the National Center for Biotechnology Information, http://www.ncbi.nlm.nih.gov, HCN1, NM_053375; HCN2, NM_053684; HCN3, NM_053685; HCN4, NM_021658; neurofilament 3 (NF3), NM_017029; glial fibrillary acidic protein (GFAP), NM_017009):
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The abundance of HCN1–4 transcripts was quantified on the basis of the interpolated cycle number in which fluorescence reached an arbitrary threshold of 0.2 fluorescence units (Ct value), using background fluorescence in cycles 7–29 as baseline. The relative abundance of HCN2 was calculated as 2–
Ct, where Ct
=
Ct(HCN2) –
Ct(HCN1), leading to normalization to the HCN1 expression level.
Several control experiments were performed to validate the results of RT-qPCR analysis. Amplification efficiency for HCN1–4 was assessed using cerebellar cDNA (mRNA extraction with DYNABeads, DYNAL, Oslo, Norway; cDNA synthesis as described for single-cell RT, 10-fold dilution); efficiency per cycle was 1.94 for HCN1, 1.97 for HCN2, 2.00 for HCN3 and 1.95 for HCN4. Possible differences in the amplification efficiencies for HCN1 and HCN2 were also tested using a plasmid containing both HCN1 and HCN2 fragments (2–200 plasmid copies per qPCR); Ct values under these conditions were almost identical (Ct = 34.8 ± 0.2 and 34.9 ± 0.01 for 20 plasmid copies, n = 3). The molecular weight of the HCN1 and HCN2 amplicons from single-cell RT-qPCR was examined on ethidium bromide-stained agarose-TAE gels; sizes were in close agreement with the expected length. In some cases, amplicons were verified by cloning into a pBluescript SK– vector (Stratagene, La Jolla, CA, USA) followed by sequencing. To exclude the possibility of contaminations, sterile water controls were run in parallel to every qPCR, and the reverse transcriptase was omitted in a subset of cells. No amplification signal was detected after 48 qPCR cycles under these conditions.
Statistical analysis
Data are reported as mean ± S.E.M.; error bars in the Figures also represent S.E.M. and are shown only if they exceeded the size of the symbol. Statistical significance was assessed using a two-sided Wilcoxon signed rank test for paired samples at a given significance level (P).
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Results |
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We examined the contribution of Ih to the resting membrane properties of fast-spiking BCs in the dentate gyrus under current-clamp conditions (Fig. 1). To minimize cell-to-cell variability, the initial resting potential of recorded neurons was set to –70 mV by injection of hyperpolarizing current. Although a sag during hyperpolarizing current pulses was not apparent under these conditions (Fig. 1A and B; Maccaferri & McBain, 1996; Lupica et al. 2001), bath application of the Ih channel blocker ZD 7288 at a concentration of 30 µM led to a hyperpolarization of the membrane potential by 5.7 ± 1.5 mV (P < 0.05; n
= 6 BCs). Correlated with the hyperpolarization, the somatic input resistance, measured from the voltage deflection at the end of a hyperpolarizing 100-pA, 800-ms current pulse, increased from 64.3 ± 8.6 to 106.6 ± 15.0 M (P < 0.05; Fig. 1C–F). Furthermore, the apparent membrane time constant measured by logarithmic fitting of the responses to hyperpolarizing current pulses increased from 12.5 ± 0.8 to 71.6 ± 8.0 ms (P < 0.05; Fig. 1B and G). As reported previously (Harris & Constanti, 1995; Beaumont & Zucker, 2000; Chevaleyre & Castillo, 2002), the effects of ZD 7288 were largely irreversible after washout (Fig. 1C and D). Thus, the Ih channel blocker ZD 7288 hyperpolarized fast-spiking BCs, increased their input resistance and prolonged their apparent membrane time constant, consistent with a contribution of Ih to resting membrane characteristics.
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We next examined Ih expressed in BCs under voltage-clamp conditions (Fig. 3). Test pulses from a holding potential of –50 mV to a potential of –120 mV activated an inward current with time-dependent onset (Fig. 3A). ZD 7288 at a concentration of 1–100 µM blocked this current in a concentration-dependent manner. Plotting the amplitude of the ZD 7288-sensitive current against ZD 7288 concentration revealed a half-maximal inhibitory concentration of 1.4 µM and a Hill coefficient of 1.3 (n
= 9 BCs; Fig. 3B). Thus, ZD 7288 blocked hyperpolarization-activated currents in fast-spiking BCs with micromolar affinity. Both recombinant and native Ih channels are also blocked by external Cs+ (Santoro et al. 1998; Chevaleyre & Castillo, 2002). We therefore tested whether external Cs+ was able to mimic the effects of ZD 7288 in fast-spiking BCs (Fig. 3C). As with ZD 7288, 30–3000 µM Cs+ blocked the hyperpolarization-activated inward current in a concentration-dependent manner. The half-maximal inhibitory concentration was 37 µM, and the Hill coefficient was 1.1 (n
= 5 BCs; Fig. 3D). These results show that fast-spiking BCs express a hyperpolarization-activated conductance with pharmacological properties characteristic of Ih. Subtraction of currents under control conditions and in the presence of saturating concentrations of either ZD 7288 or Cs+, revealed both an instantaneous and a time-dependent component of Ih during test pulses (Fig. 3E). Both components were blocked with comparable concentration dependence and time course (Fig. 3F). For Ih isolated by subtracting currents 20 min after application of 30 µM ZD 7288 from those under control conditions, the relative contribution of the instantaneous component was 34 ± 8% (n
= 8 BCs), suggesting that a fraction of Ih channels is open at the holding potential of –50 mV (Macri & Accili, 2004; Rodrigues & Oertel, 2006).
We next determined the ion selectivity and the gating properties of Ih expressed in fast-spiking BCs (Fig. 4). To examine the ion selectivity, a prepulse to –120 mV was applied from a holding potential of –50 mV to maximally activate the channels, followed by a test pulse to potentials between –110 and –60 mV to determine the current reversal potential (Fig. 4A). Ih was isolated pharmacologically by subtracting currents in the presence of 30 µM ZD 7288 from those under control conditions. The current at the beginning of the test pulse was plotted against test-pulse voltage, and analysed by linear regression. Under the ionic conditions used, the extrapolated reversal potential was –27.4 mV (n = 8 BCs; Fig. 4B), corresponding to PNa/PK of 0.36. Thus, Ih channels expressed in fast-spiking BCs are non-selective cation channels with moderate selectivity for K+ over Na+.
To determine the voltage dependence of activation of Ih, test pulses to potentials between –60 and –120 mV were applied from a holding potential of –50 mV (Fig. 4C). The activation curve was obtained from the total current at the end of the test pulse, which was converted into conductance using the current reversal potential determined previously in the same cell. Fitting the data with a Boltzmann function gave a midpoint potential of –83.9 mV and a slope factor of 13.1 mV, with a small voltage-independent component of 0.08 (n = 8 BCs; see Methods; Fig. 4D). Activation and deactivation time courses were further analysed by fitting the rise and decay phase of the current at different test-pulse amplitude with exponential functions (Fig. 4E). The activation time constant measured at –120 mV was 190.4 ± 23.1 ms, while the deactivation time constant at –50 mV was 133.8 ± 37.9 ms. Plotting activation and deactivation time constants against voltage revealed a bell-shaped relationship with a maximal value at approximately –90 mV (n = 8 BCs; Fig. 4F), comparable to the midpoint potential of the activation curve.
Ih channels expressed in fast-spiking BCs are likely assembled from HCN1 and HCN2 subunits
To examine the putative subunit composition of the native Ih channels, we analysed the expression of HCN1–4 subunit mRNA in fast-spiking BCs by single-cell RT-qPCR (see Methods; Fig. 5). The harvested cytoplasm from each single cell was reverse transcribed, and the resulting cDNA was split into two aliquots. The first aliquot was amplified with primers for one of the HCN subunits, whereas the second aliquot was amplified with primers for another HCN subunit, NF3 or GFAP. Single-cell RT-qPCR analysis revealed that fast-spiking BCs expressed HCN1 and HCN2 (Fig. 5A), whereas HCN3 and HCN4 were not detectable (Fig. 5B and C). In 5 of 5 BCs tested with primers for HCN1 and HCN2, both HCN subunits were coexpressed in the same cell, suggesting the formation of heteromeric channels (Chen et al. 2001). The HCN2/HCN1 cDNA ratio, quantified using the difference in Ct values in the qPCR (Fig. 5A, inset), was 0.61 ± 0.14 (n = 5 BCs; Fig. 5D). Cells tested were positive for NF3, but negative for GFAP, as expected for selective harvesting from neurons (n = 3 BCs). To summarize, single-cell RT-qPCR revealed that fast-spiking BCs coexpress HCN1 and HCN2 transcripts with comparable relative abundance, whereas HCN3 and HCN4 transcripts were not detectable.
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Previous studies suggested that Ih is present in both peripheral and central axons (Baker et al. 1987; Soleng et al. 2003). We therefore probed the possible axonal localization of Ih in fast-spiking BCs (Fig. 6). The BC axon was stimulated extracellularly, and antidromic APs were recorded at the soma under current-clamp conditions while the resting potential was held at –70 mV by continuous adjustment of the holding current (Fig. 6A). Under control conditions, suprathreshold stimuli evoked APs with 100% reliability (Fig. 6B). With the same stimulus intensity, bath application of 30 µM ZD 7288 reduced the reliability of AP initiation and finally abolished antidromic spikes (Fig. 6C). To distinguish between stimulation and conduction failures, the stimulus intensity was increased. AP initiation was restored under these conditions, arguing for stimulation rather than conduction failures (Fig. 6D). To quantify this observation, the probability of evoking an AP was plotted against stimulus intensity (Fig. 6E). ZD 7288 (30 µM) increased the current threshold (defined as the current leading to 50% successes) to 155 ± 8% of the control value (P < 0.05; n = 6 BCs; Fig. 6F). As a block of Ih would be expected to hyperpolarize the axon, which in turn would increase the threshold of AP initiation, these results suggest axonal localization of Ih in fast-spiking BCs.
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Discussion |
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Functional properties and subunit composition of Ih channels in BCs
Previous studies suggested that ZD 7288, in addition to blocking Ih channels, affects AMPA-type glutamate receptors (Chen, 2004). However, several lines of evidence suggest that the block of Ih in fast-spiking BCs is relatively specific. First, Ih was blocked by both ZD 7288 and external Cs+. Second, ZD 7288 had no effects on the rate of rise and the duration of the AP in fast-spiking interneurons, suggesting that it does not affect voltage-gated Na+ and K+ channels. Finally, ZD 7288 did not change the amplitude of currents activated by fast application of GABA, showing that it does not affect GABAA receptors. Therefore, the use of ZD 7288 to isolate pharmacologically Ih in voltage-clamp experiments appears to be justified.
How do the functional properties of Ih channels expressed in fast-spiking BCs compare to those expressed in other types of central neurons? Ih channels in BCs are non-selective cation channels with a slight preference for K+ over Na+ ions. PNa/PK is 0.36, similar to Ih reported in other cell types (e.g. 0.36 in photoreceptors; Wollmuth & Hille, 1992; see also DiFrancesco, 1981). Furthermore, the midpoint potential of the activation curve of the time-dependent component of Ih is –83.9 mV. This is also comparable to values found in other cell types (–90 to –83 mV; Franz et al. 2000). Thus, Ih expressed in fast-spiking BCs shows an ion selectivity and a voltage range of activation similar to other types of neurons.
The activation time constant of Ih channels expressed in fast-spiking BCs is 190 ms at –120 mV and 21–24°C. This is slower than Ih in hippocampal CA1 pyramidal neurons (64 ms) and neocortical layer 5 pyramidal neurons (84 ms), but faster than in substantia nigra dopaminergic neurons (482 ms) and thalamocortical neurons (602 ms; 22–24°C; Franz et al. 2000). When recombinantly expressed in host cells, HCN1 shows the fastest gating, whereas HCN4 shows the slowest gating (time constants, HCN1 < HCN2 HCN3 < HCN4; Santoro et al. 2000; Chen et al. 2001). Thus, the intermediate gating kinetics of Ih in BCs is consistent with the idea that the native channel is an HCN1/HCN2 heteromer, as suggested by RT-qPCR analysis.
Both ZD 7288- and Cs+-sensitive currents evoked by hyperpolarizing test pulses from a holding potential of –50 mV showed an instantaneous and a time-dependent current component (Fig. 3E). In ZD 7288-sensitive currents, the relative contribution of the instantaneous component was 34%. As the instantaneous component was evident in both ZD 7288- and Cs+-sensitive currents, it seems unlikely that it is mediated by a channel other than Ih. Despite considerable variability across cells, our results suggest two contributions to this instantaneous component. First, the activation curve given by the fitted Boltzmann function is relatively shallow (slope factor, 13.1 mV), implying that Ih is not completely deactivated at –50 mV. Second, the activation curve shows a voltage-independent component (Fig. 4D). These results are consistent with the hypothesis that Ih channels expressed in fast-spiking BCs are heterogeneous, with one subpopulation activating and deactivating in a time- and voltage-dependent manner and another subpopulation constitutively open. Instantaneous components in the ZD 7288-sensitive current have been recently reported for both recombinant (Macri & Accili, 2004; Proenza & Yellen, 2006) and native HCN channels (Day et al. 2005; Rodrigues & Oertel, 2006). However, in recombinant channels the instantaneous component appears to be sensitive to ZD 7288, but not Cs+ (Macri & Accili, 2004).
HCN channels shape integrative properties of fast-spiking interneurons
Our results indicate that Ih channels contribute to the low input resistance and the fast apparent membrane time constant of dentate gyrus BCs. As the decay of EPSPs is shaped by the membrane time constant, HCN channels will limit the duration of EPSPs, and thereby sharpen the temporal window for coincidence detection of synaptic inputs (Geiger et al. 1997; Jonas et al. 2004; Yamada et al. 2005). Thus, the expression of Ih will contribute to the repertoire of fast signalling mechanisms in fast-spiking BCs (Jonas et al. 2004).
Our results further suggest that Ih channels shape the input–output relation of fast-spiking BCs. As Ih channels are non-selective cation channels, they will shift the membrane potential away from the resting potential into the depolarizing direction. However, their activation by hyperpolarization also implements a negative feedback loop. If depolarization is too large, Ih channels will deactivate, and the membrane potential will return to the original resting value. Thus, the interneurons are clamped at a subthreshold potential, setting them into a ready-to-fire mode from which APs can be triggered readily. In this scenario, despite the low input resistance of BCs, activation of a small number of excitatory synaptic inputs is sufficient to trigger an AP (Geiger et al. 1997).
The presence of a sag during hyperpolarizing current injection is thought to be the hallmark of neurons expressing Ih (Pape, 1996; Robinson & Siegelbaum, 2003). However, although Ih channels are expressed in BCs, a pronounced sag under current-clamp conditions is not evident. Thus, our results show that the correlation between sag and Ih expression is not absolute. Two factors may explain this apparent discrepancy. First, the activation kinetics of the time-dependent component of Ih in BCs temporally overlaps with a slow component of the membrane time constant uncovered by ZD 7288 application (Fig. 1A and B). Second, the relatively shallow voltage dependence of the activation curve (Fig. 4D) implies relatively small changes in open probability of Ih during hyperpolarizing current pulses.
HCN channels are present in axons and presynaptic terminals of fast-spiking interneurons
In experiments in which APs were evoked by extracellular axonal stimulation, ZD 7288 increased the proportion of AP failures. Antidromic spikes were restored by increasing stimulus intensity, suggesting failures of initiation rather than failures of conduction. The simplest interpretation of these findings is that block of Ih channels hyperpolarizes the axon, and thereby increases the threshold for AP initiation by brief extracellular stimuli. The physiological significance of the presence of HCN channels in axons is not known. One possibility is that Ih increases the reliability of AP propagation in fine axonal branches (reviewed by Debanne, 2004). This may be relevant for the propagation of APs in BC axons, which arborize very extensively (Freund & Buzsáki, 1996). In Schaffer collateral axons, it was proposed that Ih compensates for the hyperpolarization induced by the activity of the electrogenic Na+–K+-ATPase (Soleng et al. 2003). Considering the high AP frequency and metabolic rate of BCs, Ih may have a similar function in these neurons.
Our results further suggest that Ih is present in inhibitory presynaptic terminals. ZD 7288 reduced the frequency of mIPSCs in dentate gyrus granule cells. Although these neurons receive inhibitory input from several types of interneurons (Freund & Buzsáki, 1996), previous studies suggested that mIPSCs in granule cells mainly originate from synapses located close to the soma (Soltesz et al. 1995). Thus, it is likely that the effects of ZD 7288 on mIPSCs in granule cells are generated mainly via block of Ih channels, located either in BC terminals or electrotonically close to these terminals. The present data differ from results in both the hippocampal CA1 region and the cerebellum, where ZD 7288 changes the frequency of spontaneous AP-dependent, but not AP-independent IPSCs (Southan et al. 2000; Lupica et al. 2001). This difference may be explained by differential location of Ih channels, which may be close to release sites in BCs, but at remote locations in other interneuron types.
The simplest interpretation of the reduction in mIPSC frequency is that presynaptic Ih exerts a tonic depolarizing influence in inhibitory terminals, and that block of Ih inhibits transmitter release via hyperpolarization. This hypothesis is further supported by the observation that increasing the external K+ concentration from 2.5 to 5 mM reverses the effect of ZD 7288 on mIPSCs (Y. Aponte, unpublished observations). How changes in membrane potential are converted into changes in frequency of mIPSCs is unknown. One possibility is that depolarization by HCN channels activates a small fraction of high-threshold Ca2+ channels in presynaptic terminals. In the calyx of Held, subthreshold depolarization activates a proportion of presynaptic P/Q-type Ca2+ channels sufficient to facilitate transmitter release (Awatramani et al. 2005). As synaptic transmission at BC–granule cell synapses is also mediated by P/Q-type Ca2+ channels (Hefft & Jonas, 2005), a similar mechanism may operate at BC output synapses. Alternatively, depolarization by HCN channels could activate low-threshold (i.e. T-type) Ca2+ channels, or change the driving force for electrogenic Na+–Ca2+ exchanges. However, we cannot exclude the possibility that Ih is directly coupled to the release machinery, as proposed at the crayfish neuromuscular junction (Beaumont & Zucker, 2000).
Role of HCN channels for rhythmogenesis in interneuron networks
Ih channels are thought to be involved in rhythmogenesis (Pape, 1996). In particular, their role in the generation of thalamic spindle oscillations is well established (Wang & Rinzel, 1993; Pape, 1996). Fast-spiking BCs in the hippocampus in vivo also generate APs rhythmically, firing gamma-frequency bursts of spikes in a theta-modulated manner (Penttonen et al. 1998). Can Ih promote the generation of nested theta–gamma activity in networks of hippocampal BCs? The maximal activation time constant of Ih in fast-spiking BCs is 330 ms, which after consideration of temperature differences would roughly correspond to the upper theta-frequency band. Thus, Ih channels could amplify theta oscillations driven by external septal and entorhinal inputs (Hu et al. 2002; Rotstein et al. 2005). Furthermore, Ih would be a source of tonic excitatory drive necessary for the emergence of gamma oscillations in interneuron networks (Wang & Buzsáki, 1996). Unlike tonic excitatory drive generated by metabotropic or kainate-type glutamate receptors, the drive mediated by Ih has self-regulatory properties. The activation level of Ih will be higher in interneurons that receive a small drive via other sources than in cells that receive a large drive. Thus, Ih will have a homogenizing effect on the firing frequency of interneurons, which could contribute the stability of coherent gamma oscillations in heterogeneous interneuron networks (Vida et al. 2006).
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). Time constants were determined by fitting traces with single exponential functions. Red curve, fitted 
