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. 1999 May 15;517(Pt 1):135–142. doi: 10.1111/j.1469-7793.1999.0135z.x

Elevation of intracellular Na+ induced by hyperpolarization at the dendrites of pyramidal neurones of mouse hippocampus

Hiroshi Tsubokawa *,†, Masami Miura *, Masanobu Kano *
PMCID: PMC2269327  PMID: 10226155

Abstract

  1. Whole-cell recordings were made from CA1 pyramidal cells in mouse hippocampal slices with patch pipettes containing the sodium indicator dye SBFI (sodium binding benzofuran isophthalate). Using a high-speed imaging system, we investigated changes in intracellular sodium concentration, [Na+]i, in response to hyperpolarizing pulses applied to the soma.

  2. In current-clamp recordings, we detected increases in [Na+]i during negative current injection. Hyperpolarization-induced [Na+]i elevation was more prominent in the middle apical dendrites than in the soma.

  3. In the voltage-clamp mode, hyperpolarization induced rapid increases in [Na+]i at the apical dendrites that were significantly faster than those at the soma. The signals were not affected by bath application of 1 μM TTX, but were reduced by 5 mM CsCl.

  4. Changes in membrane potential recorded from the apical dendrites in response to negative currents were significantly smaller than those recorded from the soma. In the presence of 5 mM CsCl, the I-V relationships measured at the soma and the dendrites became almost identical, indicating that CsCl-sensitive components are predominantly in the apical dendrites.

  5. These results suggest that hyperpolarization-induced [Na+]i elevations reflect Na+ influx through the non-selective cation channel (Ih channel), and that this channel is distributed predominantly in the apical dendrites. The non-uniform Na+ influx may contribute to integrative functions of the dendrites.

Neuronal excitability is precisely regulated by a combination of a number of voltage-gated channels, ligand-gated channels and other channels related to the intracellular metabotropic pathways. The h current (Ih) (also referred to as q current, Iq) is a voltage-gated non-selective cation conductance activated by membrane hyperpolarization. Although many electrophysiological and pharmacological studies have characterized this conductance in several preparations (Brown et al. 1990; Maccaferri et al. 1993; Mercuri et al. 1995; Perkins & Wong, 1995; Maccaferri & McBain, 1996; Magee, 1998), the physiological functions of Ih are not clear. Ih is reported to contribute pacemaker-like rhythmic activity in thalamic relay neurones (McCormick & Pape, 1990), and in hippocampal interneurones located in the stratum oriens-alveus (Maccaferri & McBain, 1996) and the hilus (Strata et al. 1997). However, in dopaminergic neurones in the midbrain, Ih is not a significant factor underlying the spontaneous pacemaker activity, although the presence of Ih is a typical feature of these neurones (Mercuri et al. 1995). Moreover, hippocampal and cortical pyramidal neurones have a large Ih (Brown et al. 1990; Maccaferri et al. 1993; Magee, 1998), but these neurones usually do not show pace-making activity.

To investigate Ih further, its ionic conductance should be isolated and analysed independently from other conductances. However, it is difficult to evaluate properties of an ionic conductance only by recording membrane potentials or currents, because concentration changes of one ion may affect the kinetics of other conductances (Wollmuth & Hille, 1992; Mercuri et al. 1995). To overcome this problem, we used optical recording of the Na+ influx through Ih channels combined with whole-cell recordings. Activity-induced changes of [Na+]i in single neurones in a slice preparation have previously been demonstrated in cerebellar Purkinje cells (Lasser-Ross & Ross, 1992) using the Na+-sensitive fluorescent dye SBFI (sodium binding benzofuran isophthalate). Using this method, we found a non-uniform distribution of [Na+]i changes during hyperpolarization in single CA1 pyramidal neurones in the mouse hippocampal slice. The hyperpolarization-induced Na+ signals were clearly larger in the apical dendrites than in the soma. Our results suggest that the Ih channel is distributed predominantly in the apical dendrites of mouse CA1 pyramidal cells.

METHODS

All experiments were carried out according to the guidelines laid down by the animal welfare committee of the Jichi Medical School. Three- to five-week-old mice (C57BL/6, purchased from Claire Japan Inc. or from Japan SLC Inc.) were deeply anaesthetized with ether and decapitated. The brains were quickly removed and hemisected on filter paper moistened with a cutting solution of the following composition (mM): 120 choline chloride, 3 KCl, 8 MgCl2, 1.25 NaH2PO4, 26 NaHCO3 and 20 glucose, equilibrated with 95 % O2-5 % CO2. Brain tissues containing the hippocampi on both sides were dissected out and put in a cutting chamber filled with ice-cold cutting solution. These two blocks were sliced into 300 μm sections transversely to their longitudinal axis by a microtome (VT1000S, Leica, Germany). The slices were immediately placed in a reservoir chamber filled with normal artificial cerebrospinal fluid (ACSF) solution, incubated at 35°C for about 30 min, and then maintained at room temperature. The normal ACSF solution was composed of (mM): 125 NaCl, 2.5 KCl, 2 CaCl2, 2 MgCl2, 1.25 NaHPO4, 26 NaHCO3 and 20 glucose, bubbled with a mixture of 95 % O2-5 % CO2, making the final pH 7.4. For recording, a single slice was transferred to a submerged chamber mounted on the stage of an upright microscope (Axioskop, Zeiss, Germany). The slice was superfused continuously with normal ACSF solution regulated at 33°C.

Electrical and optical recordings were made from CA1 pyramidal neurones in the slice using patch pipettes pulled from 1.5 mm o.d., thick-walled glass tubing (1511-M, Friedrich & Dimmock, USA). The pipette solution contained (mM): 115 potassium gluconate, 10 KCl, 10 NaCl, 10 Hepes, 2 Mg-ATP and 0.3 GTP, pH adjusted to 7.3 with KOH. For imaging experiments, 1 mM sodium binding benzofuran isophthalate (SBFI, Molecular Probes) was added. For perforated patch recording, the pipette contained 200 μg ml−1 nystatin, 0.4 % DMSO, and (mM): 120 potassium gluconate, 20 KCl, 2 MgCl2 and 10 Hepes, pH 7.2. A small amount of fura-2 was added to the pipette solution to check that the fluorescence was confined to the pipette, demonstrating that the membrane was not broken. Open resistance of the pipettes was 5-7 MΩ for somatic recordings and 7-11 MΩ for dendritic recordings. Tight seals (> 5 GΩ) were made on the soma or dendrite under visual control using a × 40 water-immersion lens. Pipette capacitance was compensated by using the automatic compensation function of the patch-clamp amplifier (EPC-9, HEKA, Germany). After allowing SBFI to diffuse into the cell, fluorescence images were recorded using a cooled CCD camera system (IMAGO, T.I.L.L. Photonics, Germany). The cell was excited every 32 ms at 350 ± 6 and 390 ± 6 nm using a monochrometer and the 350 nm/390 nm fluorescence ratio was measured at each location. Changes in [Na+]i are presented as the spatial average of Δ(F350/F390), where F350/F390 is the fluorescence ratio at control membrane potentials (corrected for background autofluorescence at each wavelength) and Δ indicates the time-dependent change in this ratio (corrected for bleaching). Each record was smoothed by a 5-9 point moving average to reduce noise. The main reason we used the ratio instead of direct measurement of fluorescence is that the ratio appears to be more sensitive for the detection of small changes in [Na+]i. Hyperpolarizing and/or depolarizing pulses were injected through the recording pipette in current-clamp or voltage-clamp mode. Tetrodotoxin (TTX, 1 μM), 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX, 10 μM), dl-2-amino-5-phosphonovaleric acid (APV, 50 μM), and bicuculline methiodide (BMI, 10 μM) were added to the superfusing solution in voltage-clamp and perforated patch recordings. Cells were identified as pyramidal neurones using both electrical and anatomical criteria.

BMI and CNQX were purchased from Tocris Cookson (Bristol, UK). TTX was from Wako Pure Chemical Industries Ltd (Osaka, Japan). All other compounds were obtained from Sigma.

RESULTS

Simultaneous electrical and optical recordings showed increases in the ratio of SBFI fluorescence at 350 nm and 390 nm excitation when the membrane was hyperpolarized. Figure 1 shows typical changes in the ratio of SBFI fluorescence in selected areas (see inset in Fig. 1A) of a CA1 pyramidal neurone. The resting membrane potential of this cell was -60 ± 1.4 mV (mean ±s.d.). Hyperpolarizing current injection (1 s, -400 pA) induced an increase in Δ(F350/F390) at the soma and the apical dendrites that was synchronized with the hyperpolarization (Fig. 1A and B). In the C57BL/6 mice used in the present study, the apical dendrites of CA1 pyramidal neurones reach the stratum lacunosum-moleculare, which is 270-280 μm from the stratum pyramidale. The increase in the somatic signal reached a plateau during hyperpolarization. However, in the middle of the apical dendrites, the fluorescence ratio continued to increase to the end of the pulse. In some trials changes in the SBFI signal at very proximal dendrites (10-50 μm from the soma) looked smaller than those of the soma, but the differences were not significant after averaging the signal in consecutive trials. A subsequent depolarizing pulse (1 s, 400 pA) induced a robust Na+ spike burst. Changes in SBFI fluorescence corresponding to this burst were observed in almost all regions of the cell. Figure 2 shows spatial differences of the SBFI signals in response to hyperpolarization. Data were obtained from 12 different neurones, and the representative responses of each cell were sampled for this analysis. Mean changes of fluorescence (Δ(F350/F390)) at the end of a 1 s hyperpolarizing pulse in the soma and in the regions of apical dendrites 20-80, 80-140 and 140-200 μm away from the soma were 1.10 ± 0.07, 2.56 ± 0.83, 4.99 ± 1.87 and 4.95 ± 1.57 (×10−3, means ±s.d.), respectively. Although absolute values of the fluorescence change varied from cell to cell, increases at the middle of the dendrites (80-200 μm from the soma) were significantly larger than those at the soma in all tested cells (P > 0.001, Student's paired t test).

Figure 1. Changes in SBFI signals in response to hyperpolarizing and depolarizing pulses.

Figure 1

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Somatic whole-cell current-clamp recording. A, inset, grey-scale picture of the fluorescence image of the recorded cell. Regions from which Δ(F350/F390) was measured are indicated by black (soma), red (40 μm from the soma), and blue (100 μm from the soma) squares. Middle and lower panels, membrane voltage response induced by current injection (-400 pA, 1 s, followed by 400 pA, 1 s) and average changes in the ratio of fluorescence (Δ(F350/F390)) of 4 consecutive trials. Total sampling time was 3450 ms. Black, red and blue lines represent Δ(F350/F390) measured in the corresponding regions in the inset. Vertical dashed and interrupted lines indicate periods of hyperpolarization and depolarization, respectively. B, the same graph as shown in A, with the ordinate enlarged to show changes during hyperpolarization.

Figure 2. Spatial distribution of the change in SBFI signals during hyperpolarization.

Figure 2

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Mean changes in the ratio of fluorescence (Δ(F350/F390)) at the soma and at three different regions in the apical dendrites (20-80, 80-140 and 140-200 μm from the soma). Each plot represents the mean ±s.d. obtained from 12 neurones. Vertical dashed lines indicate periods of hyperpolarization (-400 pA, 1 s). Asterisks indicate significant difference between the soma and the dendrites at the end of the 1 s hyperpolarizing pulse (P < 0.001, Student's paired t test).

Hyperpolarization-induced increases in SBFI signals were also detected when the membrane was pulsed in the voltage-clamp mode as exemplified in Fig. 3A-C. At least two components of inward current (fast activated and slow activated types) were observed in response to a hyperpolarizing pulse (4 s; holding potential, -60 mV) applied to the soma. The slow component was reversibly reduced by bath application of 5 mM CsCl (Fig. 3B), but was not affected by 200 μM BaCl2 (data not shown). Therefore, this slowly activated current was identified as Ih. During the hyperpolarizing step, increases in the SBFI signal were detected at the soma and the apical dendrites (Fig. 3C). When the somatic signal was compared with the dendritic signal in the same cell, the rise of the signal in the dendrites was always faster than the changes in the soma. However, the magnitude of changes in the dendrite at the end of the command step was smaller than that at the soma in many pairs. Pharmacological profiles of the SBFI signals at the soma and three different regions of the apical dendrites are summarized in Fig. 4. These SBFI signals were reversibly reduced by bath application of 5 mM CsCl but not affected by 200 μM BaCl2 even in the presence of 1 μM TTX, indicating that the changes correspond to Na+ influx induced by Ih. We quantified the CsCl-sensitive components of the signal by subtracting the traces recorded in CsCl from the control traces. The time courses of the mean increase in the CsCl-sensitive signals at the soma and at the dendrites are summarized in Fig. 5 (n= 6 cells). Each pair of columns indicates the mean increase ±s.d. in consecutive 500 ms periods during a 4 s hyperpolarization. During the initial 1.5 s, increases in dendritic signals were significantly larger than those in somatic signals (P < 0.01, Student's paired t test). Changes in the somatic signals did not exceed baseline fluctuations in the first column (within 500 ms). These results suggest that the Na+ influx by Ih occurred predominantly at the dendrites at the beginning of the hyperpolarization.

Figure 3. Changes in SBFI signals at the soma and the dendrites in response to hyperpolarization.

Figure 3

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Somatic whole-cell voltage-clamp recording. A, fluorescence image of the recorded cell. Regions of interest are shown by a black square (soma) and a red square (dendrite) on a grey-scale picture. B, current responses during a -60 mV hyperpolarizing step from -60 mV (holding potential). The control trace, the trace in the presence of 5 mM CsCl, and the trace during washing are superimposed. C, changes in Δ(F350/F390) in response to hyperpolarization from -60 mV to -120 mV. Black lines indicate the signal at the soma, and red lines indicate the signal at the dendrites. Mean of 8 control trials, mean of 8 trials during 5 mM CsCl application, and mean of 4 trials during washing (interrupted lines). Responses in each region are superimposed. Total sampling time was 8 s. Vertical dashed lines indicate the period of hyperpolarization.

Figure 4. Pharmacological properties of hyperpolarization-induced changes in SBFI signals.

Figure 4

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Averaged changes in SBFI signals were measured at 1 s after the beginning of hyperpolarization in normal bathing solution (control, ▪), in the presence of 200 μM BaCl2 (Inline graphic) and in the presence of 5 mM CsCl (□). Data were obtained from the soma and the apical dendrites (20-80, 80-140 and 140-200 μm from the soma) of six neurones. Asterisks indicate significant difference from the data in the normal solution (control) (P < 0.001, Student's paired t test).

Figure 5. Time course of mean changes in CsCl-sensitive components of SBFI signals.

Figure 5

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Somatic (▪) and dendritic (□) changes are shown. Each pair of columns indicates the mean increase ±s.d. in consecutive 500 ms periods during a 4 s hyperpolarization. Data were obtained from six cells. Regions of interest on the dendrites were 50-150 μm away from the soma. Asterisks indicate significant difference between the soma and the dendrites (P < 0.01, Student's paired t test).

To investigate whether there are differences between the dendritic Ih and the somatic Ih, changes in membrane potentials in response to negative current injection were examined with perforated patch recordings from the soma and the apical dendrites (Fig. 6). All the neurones tested (n= 10 for somatic recording, n= 11 for dendritic recording) had stable resting membrane potentials ranging from -58 to -65 mV. Current pulses of 500 ms were injected through the recording pipette. The pulse amplitude was incremented by 0.1 nA steps. In both somatic and dendritic recordings, current injection of less than -0.2 nA induced a negative shift of membrane potential with a clear ‘sag’. Voltage changes at the dendrites were smaller than those at the soma. CsCl (5 mM) increased the amplitude of voltage responses at both recording sites. Changes in membrane potential (ΔVm) during the sustained phase are plotted against the amplitude of injected current in Fig. 6B. ΔVm of each trace was measured at 400 ms after the beginning of the current pulse (see Fig. 6A). In the control condition, the slope of the current-voltage relationship at the dendrite was significantly smaller than that at the soma (P < 0.001, Student's paired t test). However, in the presence of CsCl, both slopes became steeper and overlapped. These results indicate that CsCl-sensitive conductances in the apical dendrites are larger than in the soma, and that the sum of the other conductances during sustained hyperpolarization is not significantly different between the soma and the dendrites.

Figure 6. Perforated patch recordings from the soma and dendrites.

Figure 6

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A, representative traces of membrane potential in response to negative current injections (-0.4 nA and -0.8 nA, 500 ms) at the soma and at the apical dendrite 100 μm away from the soma. Small regular waves on each trace are electrical noise due to the high access resistances of the perforated patch electrodes (30-60 MΩ). Upper traces are control, and lower traces are responses in the presence of 5 mM CsCl. B, changes in membrane potential (ΔVm) at the soma (○) and at the dendrite (•) plotted against amplitude of current commands, in control and in the presence of 5 mM CsCl. ΔVm of each response was obtained by subtracting the mean membrane potential 50-100 ms before current injection from the mean membrane potential 380-420 ms after the beginning of current injections. Data were obtained from 11 dendritic recordings (80-120 μm from the soma) and 10 somatic recordings. Asterisks indicate significant difference between the soma and the dendrites (P < 0.001, Student's paired t test).

DISCUSSION

Using SBFI and a cooled-CCD camera system, we found an uneven spatial distribution of [Na+]i elevations in response to hyperpolarization. The CsCl-sensitive components of these signals are thought to correspond to Na+ influx by Ih. The time course of the SBFI signal in the middle of the apical dendrites was faster than that at the soma. These results suggest that the Na+-permeable Ih channels are distributed predominantly in the apical dendrites.

Sensitivity of SBFI

Although the affinity of SBFI to Na+ is higher than that to other cations, the dissociation constant for Na+ is affected by K+ (Minta & Tsien, 1989). Since Ih is composed of both Na+ and K+ conductances, the SBFI signals we detected could reflect a combination of the increase in [Na+]i and the decrease in [K+]i. Some groups detected SBFI signals in response to single spikes (Jaffe et al. 1992) or trains of Na+ spikes (Callaway & Ross, 1997), but some did not (Mittmann et al. 1997). These experiments were done by using sharp microelectrodes, which might have made it difficult to control [K+]i. Therefore, this discrepancy might be caused by a difference in baseline [K+]i which could have affected the SBFI signals. However, Rose & Ransom (1997) reported that the effects of [K+]i on the SBFI signal were negligible in cultured hippocampal neurones. It is possible that other uncontrolled cytoplasmic components also affect the sensitivity of SBFI to Na+ (Harootunian et al. 1989). In the present study, we used patch pipettes containing a K+-based solution to keep the intracellular environment constant. We were able to detect changes in SBFI fluorescence in all neurones examined in response to hyperpolarization as well as depolarization.

Origin of increases in [Na+]i

Hyperpolarization-activated [Na+]i signals were not affected by 1 μM TTX, but were reversibly reduced by bath application of 5 mM CsCl. These pharmacological results indicate that the CsCl-sensitive component of the [Na+]i rise represents Na+ influx through Ih channels. There might be other components of the hyperpolarization-induced Na+ influx. If [Ca2+]i is high enough, a Ca2+-dependent route for Na+ entry, such as the Na+-Ca2+ exchanger (Kimura et al. 1987) and the Ca2+-activated non-selective cationic conductance (Kramer & Zucker, 1985), may be activated. These can cause an increase in [Na+]i because their reversal potentials are above the resting membrane potential. Although the spatial distribution of these possible routes is not clear, Jaffe et al. (1992) reported that [Na+]i increases which may be caused by activation of the Na+-Ca2+ exchanger were localized in the proximal apical dendrites of rat hippocampal neurones. Nevertheless, those transporter-mediated currents are thought to be Cs+ resistant. It is therefore likely that the [Na+]i signals we analysed are responsible for the activation of Ih channels.

Non-uniform distribution of the rise in [Na+]i

We examined the effects of hyperpolarization on [Na+]i in the voltage-clamp mode. Although the somatic hyperpolarization was not matched in the dendritic arbor due to a space clamp error, the rise in [Na+]i at the dendrites was always faster than that at the soma. Since the dendrites have a larger surface-to-volume ratio than the soma, dendritic changes in the fluorescence signal were usually larger. At 500 ms after the beginning of hyperpolarization, changes in the somatic signals did not exceed their baseline fluctuations. These results suggest three possible explanations for the spatial non-uniformity of the rise in [Na+]i. First, Ih channels may be predominantly distributed in the apical dendrites. Our perforated patch recordings suggest that CsCl-sensitive conductances are larger in the dendrites than in the soma (Fig. 6). Recently, non-uniformly distributed CsCl-sensitive conductances were reported in layer V cortical pyramidal neurones (Stuart & Spruston, 1998) and in hippocampal CA1 neurones (Magee, 1998) of rats. According to these data, CsCl-sensitive conductances are present at higher densities in the distal region of the apical dendrites and channel densities increase proportionally to the distance from the soma. Our imaging data strongly support this view. Therefore, a proportional increase of CsCl-sensitive conductances along the apical dendrites also appears to be the case in pyramidal neurones of mice. The second hypothesis is that Ih channels may be composed of two types of channel, one with a preference for Na+ and the other with a preference for K+. The Na+-preferring channels may distribute predominantly in the apical dendrites. Wollmuth (1995) reported the possibility that Ih channels contain at least two sites having a higher affinity for K+ over Na+ and another site with a higher affinity for Na+ over K+. However, Magee (1998) showed that electrophysiologically there were no significant differences in the reversal potentials between somatic Ih and dendritic Ih. Therefore, even if there are two types of Ih channel, the distribution ratio in the apical dendrites might be the same as that in the soma. Recently, two types of gene have been identified that are responsible for pacemaker currents (see Clapham, 1998). These genes may encode the Ih channel in the mammalian brain. The expressed proteins from these genes comprise ion channels that are approximately four times more permeant to K+ than to Na+. Since hippocampal pyramidal neurones have no pace-making activity, it is possible that other types of Ih channel are dominant in these neurones. The third hypothesis is that Ih channels may be localized in the apical dendrites, and not present in the soma. In this case, the Na+ signals seen in the soma are derived from Na+ diffusion from the apical dendrites. Since somatic whole-cell currents reflect summation of currents from all over the cell that are conducted to the soma, it is possible that the currents recorded in the soma reflect those occurring at the dendrites. Similarly, changes in voltage recorded with somatic patch electrodes reflect the events occurring in the cell as a whole. Negative current injected into the soma causes hyperpolarization of the dendrites as well as the soma, and it may activate the conductances localized at the dendrites (see Fig. 6). Na+ may diffuse from the Ih channel site in dendrites to the soma after Na+ influxes occur and, as a result, somatic [Na+]i may increase. The delay in the rise in [Na+]i between the dendrites and the soma may be explained by this mechanism. Taken together with our findings, it is likely that the three possibilities mentioned above contribute in a combinatorial manner to the uneven distribution of [Na+]i signals. Nevertheless, it is safe to conclude that Ih channels are predominantly distributed in the apical dendrites of CA1 pyramidal neurones.

Recent studies using dendritic patching and/or imaging techniques have revealed differences in functional properties between the soma and the apical dendrites in pyramidal neurones (Magee & Johnston, 1995; Miles et al. 1996; Tsubokawa & Ross, 1996, 1997; Hoffman et al. 1997; Meyer et al. 1997; Schiller et al. 1997; Schwindt & Crill, 1997; Stuart & Spruston, 1998), and in cerebellar Purkinje cells (Callaway et al. 1995; Callaway & Ross, 1997). The spatio-temporal non-uniformity of Na+ influx through Ih channels shown in this study may also contribute to characteristic properties of the dendrites such as voltage attenuation of synaptic potentials and backpropagating Na+ spikes.

Acknowledgments

We thank Professor William N. Ross, Dr Nechama Lasser-Ross and Dr Hiroyoshi Miyakawa for helpful discussions and comments on our manuscript. We also thank Professor Nobufumi Kawai for his continuous encouragement and support during the course of this study. This work was partly supported by grants from CREST (Core Research for Evolutional Science and Technology) of Japan Science and Technology Corporation (JST) (to M. K.), from Special Coordination Funds for promoting Science and Technology of the Science and Technology Agency of the Japanese Government (to M. K. and H. T.), from the Japanese Ministry of Education, Science, Sports and Culture (to M. K. and H. T.), and from the Novartis Foundation (Japan) for the promotion of science (to H. T.).

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