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1Division of Behavioral Development, Department of Developmental Physiology, 2Division of Cerebral Structure, Department of Cerebral Research, and 3Section of Mammalian Transgenesis, Center for Genetic Analysis of Behavior, National Institute for Physiological Sciences; and 4The Graduate University for Advanced Studies, Okazaki, Japan
Submitted 18 May 2007; accepted in final form 22 January 2008
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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; Isa and Sasaki 2002; Sparks 1986; Wurtz and Albano 1980). The wide field vertical (WFV) cells are a class of projection neurons distributed in the deep portion of the superficial SC (sSC) and are distinguished from other cell types by their expansive dendritic arborizations (Langer and Lund 1974) (Fig. 1A1), which cover a relatively large area of the topographic visual map of the sSC. WFV cells receive retinal inputs on the distal part of the dendrites (Fukuda et al. 1978; Nagata and Hayashi 1979), where they have characteristic "bottlebrush" dendritic endings (Major et al. 2000). Previous evidence indicated that WFV cells have large receptive fields and are selectively sensitive to moving visual stimuli (Fukuda and Iwama 1978; Humphrey 1968; Mooney et al. 1984, 1988).
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; Lo et al. 1998; Lopez-Barneo and Llinas 1988; Saito and Isa 1999), although its properties and functions have not been examined in detail. The channels carrying the Ih current, hyperpolarization-activated cyclic nucleotide-gated channels (HCN), are widely distributed in the brain and contribute to various neuronal functions, such as setting resting membrane potentials, pacemaking of firing activities, and dendritic integration (Pape 1996; Robinson and Siegelbaum 2003). In the sSC, all known HCN channel isoforms (HCN1–4) have been detected in previous in situ hybridization (Monteggia et al. 2000; Santoro et al. 2000) and immunohistochemical (Notomi and Shigemoto 2004) studies. The HCN channel subunits expressed in WFV cells are unknown. However, the expression of HCN1 at the dendrites of WFV cells has been assumed from these studies. Therefore there is a possibility that Ih plays important roles in dendritic processing in WFV cells as has been shown in the dendrites of neocortical and hippocampal pyramidal cells (Berger et al. 2001; Magee 1998, 1999; Williams and Stuart 2000). In the present study, we first characterized the properties of Ih in WFV cells in the rat sSC and corroborated the dendritic localization of HCN1. Second, we demonstrated that the distal dendrites of WFV cells express voltage-dependent Na+ channels so that forward propagating dendritic spikes are initiated at the distal dendrites in response to optic fiber inputs. Finally, we examined the effect of blocking Ih on spike initiation in the dendrites. The results indicate that the dendritic Ih together with a highly excitable dendritic membrane enable robustly timed, reliable responses of WFV cells to synaptic inputs onto their distal dendrites and undoubtedly play a critical role in the processing of moving visual stimuli.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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Sixteen to 22-day-old Long Evans and Wistar rats were used in electrophysiological experiments and immunohistochemical staining for HCN1. An adult Thy1-green fluorescent protein (GFP) transgenic rat was produced by pronuclear DNA microinjection of a Thy1-GFP construct (Feng et al. 2000) into Wistar rats (for details, see Takahashi et al. 1999) and used for immunohistochemical staining. All experiments were approved by the Animal Research Committee of the Okazaki National Research Institutes, and all efforts were made to minimize both the suffering and the number of animals used in this study.
Slice preparation and electrophysiological recording
Rats were deeply anesthetized with ether and decapitated. Depth of anesthesia was carefully confirmed by absence of reflexes to toe pinches. Frontal or sagittal SC slices (250–400 µm thick) were cut in modified Ringer solution, which contained (mM) 200 sucrose, 2.5 KCl, 1.25 NaH2PO4, 10 MgSO4, 0.5 CaCl2, 26 NaHCO3, and 11 glucose (pH 7.4, bubbled with 95% O2-5% CO2). Slices were incubated in standard Ringer solution at room temperature for >1 h before recording. The standard Ringer solution contained (mM) 125 NaCl, 2.5 KCl, 2 CaCl2, 1 MgCl2, 26 NaHCO3, 1.25 NaH2PO4, and 25 glucose (pH 7.4, bubbled with 95% O2-5% CO2).
Whole cell patch-clamp recordings were obtained by visual control of patch pipettes. Slices were mounted in a recording chamber on an upright microscope (BX61WI, OLYMPUS, Tokyo, Japan) and continuously superfused with standard Ringer solution. Patch pipettes were prepared from borosilicate glass capillaries and were filled with the internal solution containing (in mM) 150 K-gluconate, 10 KCl, 2 MgCl2, 2 Na2ATP, 0.2 EGTA, 10 HEPES, and 0.1 spermine, adjusted to pH 7.3 with KOH. To stain recorded neurons, biocytin (5 mg/ml; Sigma, St. Louis, MO) or Lucifer yellow (1 mg/ml; Sigma) was dissolved in the solution. Cells were classified according to the morphological criteria of Langer and Lund (1974). Some of the WFV cells were identified based only on the presence of the voltage sag. The resistance of the electrodes was 3–7 M
in Ringer solution. The actual membrane potential was corrected by a liquid junction potential of –10 mV. In voltage-clamp recordings, series resistance was compensated 
70%. To stimulate optic fibers, cathodal square-wave current pulses with a duration of 200 µs were applied through a concentric bipolar electrode placed at the rostral (sagittal slices) or lateral end (frontal slices) of the stratum opticum in sSC. The stimulation was applied repeatedly at 0.05–0.2 Hz. Glass micropipettes identical to the patch pipette were used for the local application of glutamate (1 mM; Sigma) and tetrodotoxin (TTX, 10 µM; Sankyo, Tokyo, Japan). Glutamate was ejected with an air pressure pulse (5–20 psi, 10- to 50-ms duration) using a pneumatic picopump (PV820, World Precision Instruments, Sarasota, FL) at 0.033–0.1 Hz. TTX was ejected with a manually applied air pressure from an injection syringe. The application of TTX was started 10 s before and stopped a few seconds before or after the glutamate applications. Alexa Fluor 568 (10 mM; Molecular Probes, Eugene, OR) was included in these drug solutions to visualize the solution spread. ZD7288 (30 or 100 µM; AstraZeneca, London, UK), 6-cyano-7-nitroquinoxaline-2, 3-dione (CNQX, 10 µM), and D-2-amino-5-phosphonovaleric acid (APV, 50 µM; both from Sigma) were bath applied. Most of the recordings were performed at room temperature, but some were at physiological temperature (35 –36°C). Data were recorded with an EPC9 patch-clamp amplifier and PULSE software (Heka, Lambrecht, Germany).
After recording, the slices were fixed with 4% paraformaldehyde in 0.12 M phosphate buffer (pH 7.4) for >1 day at 4°C. After fixation, biocytin-filled neurons were visualized by the Vectastain ABC method (Vector, Burlingame, CA). Details are described elsewhere (Isa et al. 1998).
Data analysis
To analyze the activation kinetics of Ih, the amplitude and time constants of the current responses were estimated from single- or double-exponential functions that fit the current responses to voltage steps from –55 mV. The total current responses to the voltage step were fitted with offset current plus exponential functions. The ratio of the fast exponential component of the response to that of the entire response was calculated as Af/(Af + As), where Af and As are the amplitudes of the fast and the slow components, respectively. Steady-state activation curves were determined from the tail current amplitudes measured at –75 mV after cells were kept at holding potentials of –55 to –125 mV for 2–3 s. The current-voltage relationships were then fitted with the Boltzmann function: I = Ioff + Imax/{1 + exp[(V – V1/2)/s]}, where Ioff is the offset current, Imax is the maximum tail current, V is the test holding potential, V1/2 is the half-activation voltage, and s is the slope of the Boltzmann curve.
The onset of action potentials and excitatory postsynaptic potentials (EPSPs) were determined as the points where the slopes exceeded threshold. The threshold was manually set in each cell at 5–11 V/s for action potentials and 0.2–0.5 V/s for EPSPs. The initial rising slope of an EPSP was defined as the slope between the onset of the EPSP and the point 2 ms after the onset.
All values are given as means ± SE. Unpaired and paired t-tests and Wilcoxon signed-rank tests were used for assessing statistical significance. Differences were considered to be significant at P < 0.05.
Immunohistochemistry
Rats were stereotaxically injected with recombinant Sindbis virus vector (Furuta et al. 2001) under deep pentobarbital anesthesia (20 mg/kg ip). A part of the cerebral cortex overlying the SC was sucked out, and the vector solution (1 µl) was pressure-injected through a glass micropipette (tip diameter of 50–100 µm) inserted into the deep SC. The incisions were sutured, and the animals were allowed to survive for 24 h.
For immunohistochemical processing, one Thy1-GFP transgenic rat and two wild-type rats injected with Sindbis virus vector were deeply anesthetized with pentobarbital (50 mg/kg ip), perfused transcardially with PBS for 1 min, followed by 4% paraformaldehyde, 0.05% glutaraldehyde, and 15% saturated picric acid in 0.1 M PB, pH 7.4 for 12 min. Coronal sections (50 µm thick) were cut into 25 mM PBS with a vibrating microslicer (VT-1000, Leica, Wetzlar, Germany). Fluorescence signals were observed in the sSC region by a fluorescence microscope; GFP-expressing WFV cells were identified based on their morphology and photographed. After fluorescence observation, sections were washed three times in 0.1 M PB and cryoprotected in 25% sucrose and 20% glycerol in 0.02 M PB for 3 h. After freeze-thawing in liquid nitrogen, sections were washed in 50 mM TBS three times, and incubated for 1 h at room temperature in TBS containing 20% normal goat serum to block nonspecific binding. After blocking, the sections were incubated with guinea pig anti-HCN1 antibody (1 µg/ml) (Notomi and Shigemoto 2004) and rabbit anti-GFP antibody (0.05 µg/ml) (Tamamaki et al. 2000) in TBS containing 1% normal goat serum for 24 h at 4°C. After washing in TBS, sections were incubated with 1.4-nm gold-labeled anti-guinea pig IgG (Nanoprobes, Stony Brook, NY) and biotinylated anti-rabbit IgG (Vector, Burlingame, CA) secondary antibodies at 4°C overnight and then treated with an HQ Silver kit (Nanoprobes). Finally, the sections were incubated with ABC-Elite (Vector) and then 0.02% diaminobenzidine-4HCl solution in 0.003% H2O2. After osmification, the immunostained sections were block-stained with uranyl acetate, dehydrated, and flat-embedded in Epon (Durcupan, Fluka). Ultrathin sections containing GFP-expressing WFV cells were prepared and examined for HCN1 signals with a Tecnai 10 electron microscope.
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RESULTS |
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. Some WFV cells were identified based only on the presence of the voltage sag (see following text). Comparison of Ih in various morphologically identified sSC neurons
In response to hyperpolarizing current pulses, WFV cells showed a prominent voltage sag caused by Ih as shown previously (Fig. 1A) (Isa et al. 1998; Lo et al. 1998; Lopez-Barneo and Llinas 1988; Saito and Isa 1999). However, other neurons in the sSC also expressed Ih currents (Fig. 1B) (Endo et al. 2003), although they appeared to be much less conspicuous than those seen in WFV cells. To compare the Ih of different cell types, we analyzed the activation time constants and current amplitudes, which were estimated by fitting current responses to a voltage step from –55 to –105 mV with single-exponential functions. The total current responses to the voltage step were fitted with offset currents plus exponential functions. In morphologically identified WFV cells (n = 63), the Ih activation time constant was 242 ± 8 (SE) ms and the amplitude was 647 ± 36 pA. The current density, calculated as the Ih amplitude normalized by membrane capacitance, was 14.0 ± 0.7 A/F. Among 23 neurons classified as cell types other than WFV cells, 21 cells showed slowly developing inward currents that were successfully fitted with single-exponential functions. In some of these cells, it appears that the slow inward currents contributed little or nothing to membrane properties (Fig. 1C). The time constant of the Ih in these cells was 1,720 ± 500 ms, the amplitude was 26.8 ± 4.8 pA, and the current density was 1.25 ± 0.17 F/A; all these values are significantly different from those in the WFV cells (P < 0.0001, unpaired t-test). Thus WFV cells have Ih with faster activations and larger amplitudes and are clearly distinguished from other cell types based on these properties (Fig. 1, D and E).
WFV cells express HCN1 in distal dendrites
The voltage range of Ih activation in WFV cells was then estimated from the amplitudes of the tail currents measured at –75 mV after various test potentials for 3 s (n = 30, Fig. 2A). The current-voltage relationship was fitted with the Boltzmann function (Fig. 2B). The fitted Ih had a half-activation voltage of –90 ± 0.7 mV and a slope of 9.6 ± 0.2 mV. From this we estimated that 9% of Ih channels are activated at the resting membrane potential of these cells (-68.4 ± 0.9 mV, measured immediately after patch membrane was broken), implying that Ih contributes to the resting membrane properties of WFV cells. We obtained similar results at physiological temperature (35–36°C, n = 7). The half-activation voltage and the slope were –89 ± 1.7 and 12 ± 1.1 mV, respectively. In this case, 13% of Ih channels are considered to be active at the resting membrane potential (–66.6 ± 1.4 mV).
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; Santoro et al. 2000). Given that Ih in WFV cells shows distinctly faster activation than it does in other cell types and that HCN1 subunits have the fastest activation kinetics among the four known subunits of HCN channels, it seems conceivable that WFV cells express HCN1. Because it has been shown that the kinetics of native Ih reflect the expressed subunit composition and are comparable to that of the corresponding HCN subunits measured in heterologous expression system (Chen et al. 2001; Santoro et al. 2000), we compared the activation kinetics of Ih in WFV cells to available data for mouse HCN channels expressed in Xenopus oocytes (Chen et al. 2001; Santoro et al. 2000; Ulens and Tytgat 2001). As exemplified in Fig. 2C, although the WFV Ih could be fitted to a single-exponential function, the fit to a double-exponential function was much better. The time constants of the fast component decreased from 160 to 70 ms as cells were hyperpolarized. Likewise for the slower time constants, from 1,400 to 320 ms (n = 68, Fig. 2D). These values are 1.5- to fourfold larger than those of the mHCN1, however, still 1/3 to 1/7 of those for mHCN2, and comparable to neocortical and hippocampal pyramidal cells that express high level of HCN1 and somewhat lower level of HCN2 (Chen et al. 2001; Franz et al. 2000; Santoro et al. 2000). The amplitude ratio of the fast component of Ih in WFV cells predominated over the entire voltage range (74.4 ± 0.6% at –105 mV) and showed little voltage dependency (Fig. 2E). These properties also resemble those of HCN1 and contrast with HCN2 that shows steep voltage dependency (Chen et al. 2001). Considering the activation kinetics of HCN3 and -4 is even slower than HCN2, these results are consistent with the assumption that WFV cells express HCN1 subunits as the dominant contributor to the Ih current, although we cannot exclude coexpression or heteromerization with other subunits.
Our recent immunohistochemical study described a gradient of dense neuropil staining of the HCN1 subunit in rat sSC, which was weak in the deep portion of the sSC and steadily increased in intensity toward the dorsal surface of the SC (Notomi and Shigemoto 2004). These observations suggest that WFV cells express HCN1 in distal dendrites rather than in the soma. To clarify dendritic expression of HCN1 in WFV cells, we examined the immunohistochemical localization of HCN1 at the electron microscopic level. Figure 3 shows results obtained from a transgenic rat (Thy1-GFP rat) in which the GFP-expressing WFV cells were sparsely distributed in the sSC (Fig. 3A). Immunoreactivity was examined in a single morphologically identified WFV cell (Fig. 3, A and B), and we found immunopositive signals for HCN1 in the distal dendritic branch of this cell (Fig. 3E). On the other hand, no labeling was found in the soma (Fig. 3C) or proximal part of the dendrite (Fig. 3D). In two other rats, WFV cells were identified by the injection of a recombinant Sindbis virus vector which labeled infected cells with GFP (Furuta et al. 2001). We also found immunopositive signals for HCN1 in the dendrites of these WFV cells (data not shown).
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HCN1 channels located in the dendrites should have a strong influence on dendritic processing in WFV cells. WFV cells respond to afferent excitation with spikes that rise from a very hyperpolarized onset potential (Fig. 4A) and that are assumed to be initiated at distal dendrites (Isa et al. 1998). In response to optic fiber stimulation, we typically observed that the apparent threshold of the first action potential was only a few millivolts positive to the baseline membrane potential and was more negative as the baseline potential was set at more hyperpolarized levels (Fig. 4, A2 and B). These spikes were easily induced by threshold level stimuli (Fig. 4A2). The second and later spikes usually showed more positive onset potentials (data not shown), but we did not analyze them in this study. We concluded that the low onset spikes were induced via synaptic transmission and not by direct activation of axons or dendrites because they were abolished when EPSPs were inhibited by ionotropic glutamate receptor antagonists (50 µM APV and 10 µM CNQX, n = 3; (Fig. 4C). Occasionally, full-sized spikes failed to occur when baseline membrane potentials were set at relatively hyperpolarized levels, and spikes with smaller amplitudes (15–20 mV) were observed (Fig. 4D, *). A notch on the rising phase of full-sized spikes, indicating a decrease in the slope, was observed more often (Fig. 4D2, arrowhead). In contrast to afferent activations, depolarizing current injections from the recording pipette, which should have activated primarily voltage-dependent channels distributed on the soma or axon initial segment, induced action potentials with higher and more constant onset membrane potentials of around –50 mV, irrespective of baseline potentials (Fig. 4, A1 and B). These results indicate that when WFV cells receive optic fiber inputs, action potentials can be initiated at sites that are electrotonically distant from the soma, most likely at distal dendrites.
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We examined the effect of the Ih blocker, ZD7288 (100 µM) on the response of WFV cells to optic fiber stimulation. The stimulation strength was set so that EPSPs were induced without failure and action potentials were induced with a probability of 0.44–1.0. All neurons examined (n = 13) displayed low onset spikes before bath application of ZD7288. When we applied ZD7288, WFV cells were hyperpolarized by 2–27 mV from the set membrane potential of –65 to –76 mV (Fig. 6A, ZD7288). The input resistance significantly increased in the presence of ZD7288 (85.7 ± 9.4 M in control vs. 211.3 ± 26.0 M in the presence of ZD7288, n = 8 cells, P < 0.005, paired t-test). The probability of low onset spikes decreased, or in some cases, completely disappeared even when the baseline membrane potentials were maintained at the same level as before ZD7288 application (Fig. 6A, ZD7288 + current). Instead of the low onset spikes, WFV cells often displayed spikes with more depolarized onsets, which had longer and fluctuating latencies (Fig. 6, A and B). The onset membrane potential of these spikes was less dependent on the baseline potential. The higher onset spikes were often preceded by a hump around the peak of the EPSPs (Fig. 6A2, 
) and a gently rising slope (
). Therefore the higher onset spikes seemed to be generated via voltage-dependent, regenerative processes overriding the EPSPs and were clearly different from the lower onset spikes, which were immediate responses to synaptic inputs.
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–0.5 mV relative to control) with DC current injection. When we took all the spikes into consideration, the probability increased in 5 of 13 cells and decreased in 8 cells, however, did not change significantly as a whole (n = 13, P > 0.05, Wilcoxon signed-rank test; Fig. 7C). In contrast, when only low onset spikes were considered, the probability was reduced in all but one cell, reaching statistical significance (n = 13, P < 0.01, Wilcoxon signed-rank test; Fig. 7C). Mean latency of spike responses in individual WFV cells was 3.0 ± 0.3 ms in control (n = 12) and 53.3 ± 29.1 ms in the presence of ZD7288 (n = 12) when all spikes were taken into consideration. The latency of the low onset spikes was very constant. The SD of the latency from the EPSP onset to the spike initiation in individual WFV cells was 0.5 ± 0.1 ms in control (n = 13 cells) and 0.4 ± 0.1 ms in the presence of ZD7288 (n = 8). In contrast, the latency of the high onset spikes in ZD7288 fluctuated (SD = 18.7 ± 3.1 ms, n = 7). When all spikes were took into consideration, the fluctuation of timing of spike responses became larger; the SD value increased from 1.2 ± 0.4 ms in control (n = 13) to 20.1 ± 7.8 ms in the presence of ZD7288 (n = 9). The initial slopes of EPSPs elicited by optic fiber stimulations were measured from sweeps that lacked low onset spikes. Mean initial slope did not change significantly (91.2 ± 3.7% of control, n = 11 cells, P > 0.05, paired t-test), suggesting that monosynaptic optic fiber inputs were not influenced presynaptically by ZD7288.
Finally, we tested the effect of more specific concentration of ZD7288 (30 µM) on the responses to optic fiber stimulations at physiological temperature (35–36°C). We could reproduce parallel results to those obtained at room temperature and with 100 µM ZD7288 in five neurons (Fig. 7, 
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DISCUSSION |
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In the sSC, all known HCN channel isoforms (HCN1–4) have been detected in previous histological studies (Monteggia et al. 2000; Notomi and Shigemoto 2004; Santoro et al. 2000). The present electrophysiological recordings show that WFV cells have more HCN channels with faster activation kinetics compared with other cell types in the sSC. Moreover, we found that the activation kinetics of Ih in WFV cells resemble those of HCN1 channels expressed in Xenopus oocytes (Chen et al. 2001; Santoro et al. 2000; Ulens and Tytgat 2001), suggesting that HCN1 is the dominant contributor to the Ih in WFV cells. The present immuno-electron microscopic observations showed the presence of HCN1 in the WFV dendrites. In addition, previous studies showed that dense expression of HCN1 mRNAs in the SO where the somata of WFV cells are located (Monteggia et al. 2000; Santoro et al. 2000), whereas dense neuropile staining for HCN1 antibody in the dorsal sSC rather than in the SO (Notomi and Shigemoto 2004). Taken together, it is likely that HCN1 subunits are abundantly expressed in the dendrites of WFV cells, similar to hippocampal, subicular and neocortical layer 5 pyramidal cells, where HCN1 subunits are expressed mainly in the distal apical dendrites (Lörincz et al. 2002). However, we do not exclude the possibility that WFV cells may have a mixed population of various homo- or heteromeric channels. Some central neurons have been shown to coexpress multiple isoforms of HCN channels (Franz et al. 2000; Notomi and Shigemoto 2004). HCN1 and -2 subunits can form functional heteromeric channels with intermediate activation kinetics between those of HCN1 and -2 homomers (Chen et al. 2001; Ulens and Tytgat 2001). Such coexpression may explain the slightly slower activation of Ih in WFV cells than that in homomeric HCN1 channels. We conclude that WFV dendrites have abundant HCN1 channels, but the possible expression of other HCN subunits and their precise subcellular localization are the subject of future investigations.
Spike initiation in WFV cell dendrites
Previous studies have reported that WFV cells and their chicken homologues display spikes with very hyperpolarized onset potentials in response to optic fiber inputs (Isa et al. 1998; Luksch et al. 2001). These spikes are thought to be initiated at distal dendrites. In the present study, we were able to induce TTX-sensitive dendritic spikes by local application of glutamate to the distal dendrites of WFV cells (Fig. 5). We could induce dendritic spikes by glutamate applications at two different dendritic sites. These results indicate that each dendrite of WFV cell expresses voltage-dependent Na+ channels and are excitable sufficiently to generate forward propagating Na+-dependent dendritic spikes. These results also suggest that activation of a restricted number of dendritic branches is sufficient to induce spike responses without integration of synaptic inputs onto widely distributed dendrites at the WFV cell soma. In addition, these spikes were evoked in a Ca2+-free extracellular solution, indicating that the dendritic spikes do not need an activation of voltage-dependent Ca2+ channels.
The apical dendrites of neocortical and hippocampal pyramidal cells have active voltage-dependent channels, and forward propagating Na+-dependent dendiritic spikes readily occur when they receive strong synchronized synapic inputs; however, action potentials are initiated first in the soma-axonic region when the inputs are relatively weak (Gasparini et al. 2004; Golding and Spruston 1998; Stuart and Sakmann 1994). In WFV cells, because we could easily induce dendritic spikes at threshold stimulus intensities, it seemed that action potentials are triggered primarily in the dendrites in response to optic fiber inputs. These results suggest that optic fiber synapses exert powerful influences on the distal dendritic branches of WFV cells. This might be due to very strong nature of optic fiber inputs. Another possibility is that WFV cells may have dense voltage-dependent Na+ channels at the site of optic fiber inputs. In this regard, globus pallidus neurons were also reported to initiate action potentials first in distal dendrites, where they have specific clustering of voltage-dependent Na+ channels at sites of excitatory synaptic inputs (Hanson et al. 2004).
HCN channels facilitate dendritic spike responses
Because Ih is continuously active at resting membrane potentials, it is an important determinant of membrane properties such as input resistance, membrane time constant, and resting membrane potential (Pape 1996; Robinson and Siegelbaum 2003). In the present study, we found that blocking Ih reduced the probability of low onset spikes occurring and led instead to fluctuating long-latency spike responses to optic fiber stimulations. These effects must be due to the blockade of Ih in postsynaptic WFV cells. As the initial slopes of EPSPs were not changed, monosynaptic inputs from optic fibers were probably not affected by ZD7288. Given the very short latency of the low onset spikes, it is unlikely that the suppression was caused by a blockade of HCN channels that might be expressed in interposed neurons. We estimated that a part of Ih channels in WFV cells were active at resting membrane potential levels, and consistent with this, WFV cells became hyperpolarized when Ih was blocked. Although we do not know the actual membrane potential of the dendrites, Ih could act as a continuous depolarizing drive for the dendritic membrane. Therefore the simplest explanation of the present results is that under normal conditions, Ih fixes dendritic membrane potentials at a depolarized level, which is close to the spike threshold, and thereby facilitates the generation and/or the propagation of Na+ action potentials in the dendrites of WFV cells independently from somatic membrane potentials.
The functions of Ih in the apical dendrite of hippocampal and neocortical pyramidal cells have been elucidated. Dendritic Ih increases the electrotonic distance along the apical dendrite, resulting in an increased attenuation of EPSPs (Berger et al. 2001; Magee 1998). In addition, dendritic Ih shortens and normalizes the time course of synaptic potentials and enables distance-independent temporal summation of EPSPs in pyramidal cells (Magee 1999; Williams and Stuart 2000). We believe that the same is the case in WFV cells for subthreshold synaptic inputs, and this explains the generation of fluctuating, long-latency somatic spikes when the dendritic spikes failed to occur in the presence of the Ih blocker. However, unlike the apical dendrites of hippocampal and neocortical pyramidal cells, in which Ih fundamentally acts to limit signal propagation along the apical dendrites by decreasing dendritic membrane resistance, the present results indicate that Ih apparently enhances signal transmission along the dendrites of WFV cells. Thus the present results suggest dual functions of the dendritic Ih in WFV cells: ensuring high-fidelity dendritic spike responses by setting the dendritic membrane potential close to the spike threshold and suppressing long latency fluctuating spike responses by decreasing the dendritic membrane resistance.
Functional significances
Previous evidence indicated that WFV cells have large receptive fields, with diameters extending 
90° in visual angle, and are selectively sensitive to moving visual stimuli (Fukuda and Iwama 1978; Humphrey 1968; Mooney et al. 1984, 1988). WFV cells are thought to receive retinal inputs to the distal parts of their expansive dendritic trees (Fukuda et al. 1978; Nagata and Hayashi 1979). These morphological configurations should enable WFV cells to integrate inputs from a large area of the visual space. The dendritic properties elucidated in the present study may be necessary to overcome disadvantages associated with the long electrotonic distances from the site of synaptic inputs to the soma-axonic region and enable precisely timed spike responses of WFV cells.
A mechanism for motion processing in WFV cells and their avian homologue has been proposed (Luksch et al. 2001, 2004; Major et al. 2000) in which each dendritic ending of a WFV cell is assumed to receive inputs from a small area in the whole visual receptive field of the cell. Moving visual stimuli pass over these subfields and in turn induce dendritic responses in sequence. Critical cellular properties for discriminating moving and stationary visual stimuli are phasic retinotectal signal transfer and binary dendritic responses that interact in a mutually exclusive manner in the postsynaptic neuron (Luksch et al. 2004). Dendritic HCN channels may be an important substrate for this processing mechanism because the dual functions discussed in the preceding text would enhance high-probability spike responses, which are spatiotemporally coupled tightly with retinal inputs onto each dendritic ending. Thus the present study has revealed important cellular properties that may be implicated in processing the motion of visual stimuli.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Present address and address for reprint requests and other correspondence: T. Endo, Mammalian Locomotor Laboratory, Department of Neuroscience, Karolinska Institutet, Retzius Väg 8, 17177 Stockholm, Sweden (E-mail: toshiaki.endo{at}ki.se)
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, the activation phase of Ih. ZD7288 (100 µM) completely inhibited Ih (A3, bottom). Wave forms of the current and voltage pulses are shown at the top of A2 and A3, respectively. B: an example of horizontal cells. B1: a photograph of a horizontal cell filled with Lucifer yellow. B2 and B3: the responses to hyperpolarizing current (up to –120 pA with –40-pA step; B2) and voltage pulses (from –55 to –125 mV with –10-mV step; B3). Arrangement of figures is the same as in A. C: an example of narrow field vertical (NFV) cells. Arrangement of figures is the same as in A. This cell showed very few slowly developing inward currents in response to hyperpolarizing voltage pulses. D and E: Ih amplitude (D) and current density (E) are plotted against activation time constant. H-M, horizontal-multipolar cells; stellate, stellate cells; non-WFV, cells identified as other than WFVs but that could not be specifically classified due to insufficient staining. Scale bars in A1–C1 = 100 µm.
, solid line) and at physiological temperature (35–36°C, 
in A was reconstructed from 2 consecutive sections (50 µm thick each). The locations of the electron micrographs shown in C–E are indicated (
, the onset of EPSPs and action potentials, respectively. Spike responses in the control situation displayed short and fixed latencies from the onset of EPSPs, whereas spikes in the presence of ZD7288 showed longer and fluctuating latencies.