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Molecular Endocrinology, doi:10.1210/me.2009-0132
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Molecular Endocrinology 23 (9): 1494-1504
Copyright © 2009 by The Endocrine Society

Thyroid Hormone (T3)-Induced Up-Regulation of Voltage-Activated Sodium Current in Cultured Postnatal Hippocampal Neurons Requires Secretion of Soluble Factors from Glial Cells

Vanessa Niederkinkhaus, Romy Marx, Gerd Hoffmann and Irmgard D. Dietzel

Department of Molecular Neurobiochemistry (V.N., R.M., G.H., I.D.D.) and International Graduate School of Neuroscience (V.N., I.D.D.), Ruhr-University Bochum, NC7/170, D-44780 Bochum, Germany

Address all correspondence and requests for reprints to: Irmgard D. Dietzel, Department of Molecular Neurobiochemistry, Ruhr-University Bochum, NC7-170, Universitätsstrasse 150, D-44780 Bochum, Germany. E-mail: Irmgard.D.Dietzel-Meyer{at}ruhr-uni-bochum.de.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 Results
 Discussion
 Materials and Methods
 REFERENCES
 
We have previously shown that treatment with the thyroid hormone T3 increases the voltage-gated Na+current density (Nav-D) in hippocampal neurons from postnatal rats, leading to accelerated action potential upstrokes and increased firing frequencies. Here we show that the Na+ current regulation depends on the presence of glial cells, which secrete a heat-instable soluble factor upon stimulation with T3. The effect of conditioned medium from T3-treated glial cells was mimicked by basic fibroblast growth factor (bFGF), known to be released from cerebellar glial cells after T3 treatment. Neutralization assays of astrocyte-conditioned media with anti-bFGF antibody inhibited the regulation of the Nav-D by T3. This suggests that the up-regulation of the neuronal sodium current density by T3 is not a direct effect but involves bFGF release and satellite cells. Thus glial cells can modulate neuronal excitability via secretion of paracrinely acting factors.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
Results
 Discussion
 Materials and Methods
 REFERENCES
 
Thyroid hormone deficiency in the developing brain leads to irreversible mental retardation accompanied by a mental slowing together with increased latencies of evoked potentials (1, 2, 3). Concomitantly, in hypothyroidism a decrease in electroencephalogram amplitudes and frequencies (4, 5), a reduced cortical excitability, as indicated by transcranial magnetic stimulation (6), as well as a slowing of peripheral conduction velocities (7, 8) have been reported. Hyperthyroidism, in turn, leads to nervousness, restlessness, and tremor (9) accompanied by increased frequencies of electroencephalogram waves (10) and can, in some cases, cause epileptic seizures (11). As a possible underlying mechanism we have previously shown that a lack of T3 down-regulates the voltage-gated Na+ current density (Nav-D) in hippocampal neurons from postnatal rats (12), leading to slowed action potential upstrokes and decreased firing frequencies (13). It remained unresolved, however, whether T3 acts directly on neurons or via secondary factors.

The role of glial cells as a target for T3-action has recently been recognized. Astrocytes (14, 15, 16, 17) as well as oligodendrocytes (18, 19) express thyroid hormone receptors. Cerebellar astrocytes secrete growth factors such as acidic and basic fibroblast growth factor (aFGF and bFGF), epidermal growth factor (EGF), TNFβ, and Il-3 after T3-stimulation which, in turn, exert mitogenic effects on cerebellar granule cells (20, 21, 22).

Information concerning the influence of growth factors on long-term regulation of voltage-gated Na+ channel expression in neurons is predominantly available from cell lines, such as human neuroblastoma (NB69) or PC12 cells. In these cells it has been observed that Na+ currents are up-regulated by astroglial-conditioned media (23), by nerve growth factor (24, 25), platelet-derived growth factor, and EGF as well as bFGF (24, 26, 27, 28). In neurons from the peripheral nervous system nerve growth factor has been shown to regulate Na+ channel expression, thus influencing pain sensitivity by increasing the expression of the Na+ channel {alpha}-subunit Nav1.8 and decreasing Nav1.3 (29).

Here we report that the up-regulation of the Nav-D in rat hippocampal neurons is mediated by a soluble factor, most likely bFGF, released from satellite cells after treatment with T3. Thus glial cells from the central nervous system are not only able to modulate synaptic transmission in the hippocampus (30, 31) but can also influence the electrical excitability of neurons in a T3-dependent manner.


   Results
TOP
 ABSTRACT
 INTRODUCTION
 Results
Discussion
 Materials and Methods
 REFERENCES
 
Effects of T3 on isolated Na+ and K+ currents
The aim of this study was to investigate whether the thyroid hormone T3 regulates hippocampal Na+ currents directly or via intermediate steps. To enable recordings under culture conditions favoring the survival of neurons, the culture medium was changed from DMEM, as used in previous investigations, to a neurobasal (NB)-based medium. We now tested whether under these culture conditions Na+ currents are selectively regulated by T3 as observed in our previous study (12). Cells were prepared from P4 rats and preincubated for 3 d in the presence of fetal calf serum (FCS). Then FCS was exchanged against B18 supplement containing none or 50 nM T3. After an incubation of an additional 4 d, isolated Na+ currents were recorded in solutions blocking K+ and Ca2+ currents using whole-cell patch clamp recording. As shown in Fig. 1Go, a preincubation with 50 nM T3 resulted in an increase of the Nav-D normalized to the cell capacitance (Cm, determined as described in Materials and Methods) from 53.1 ± 4.7 pA/pF (n = 22) to 71.8 ± 6.4 pA/pF (n = 27, P < 0.05).


Figure 1
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  		Fig. 1. Effect of 4 d preincubation with T3 on amplitudes of isolated Na+ currents. A, Original recordings from cells prepared from P4 rats, grown in cultures pretreated with FCS, and maintained for the last 4 d in the absence of T3 (top) or the presence of 50 nM T3 (bottom). B, Current to voltage relationships of Na+ currents from control and T3-treated cells. Error bars indicate ±SE. ms, Milliseconds.

 
As shown in Fig. 2Go, a preincubation with T3 did not influence voltage-activated K+ currents (Fig. 2Go, Aa and Ab). No significant differences in the average current to voltage relationships for peak and persistent current components were observed (Fig. 2EGo). Thyroid hormone preincubation neither influenced the peak A current amplitude nor the amplitude of the persistent KDR current recorded after 200 msec depolarization steps (Fig. 2Go, B and C). Likewise, no changes in the rise time of the A current (measured as time required for the current to increase from 20–80% of maximal level) as well as the time constant of inactivation of the A current component were observed (Fig. 2DGo). Thus, in accordance with our previous results and in contrast to observations in heart ventricle cells (32, 33) T3 treatment did not influence K+ currents in hippocampal neurons cultured under the present conditions. Because T3 preincubation had no effect on K+ currents we conducted further Na+ current investigations in the presence of unblocked K+ currents.


Figure 2
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  		Fig. 2. Effect of 4 d preincubation with T3 on isolated potassium currents. A, Original recordings from cells prepared from P4 rats and grown in cultures pretreated with FCS and maintained for the last 4 d in the absence or presence of T3. Families of K+ currents recorded from control (Aa) and T3-pretreated cells (Ab) using voltage steps of increasing depolarization as shown in inset. B, Separation of KDR and A current components. A current isolated by subtracting KDR from the total K+ current using voltage protocols as shown in inset. Original current traces recorded from a control cell (Ba) and a cell from a sister culture treated with 50 nM T3 for the 4 last days (Bb); traces to the right: KDR activated after a prepulse to –15 mV, traces to the left: A current calculated by subtracting KDR from the total current evoked after a prepulse to –100 mV. C, Average peak A currents (KA) and delayed rectifying currents recorded after a test pulse duration of 200 msec (KDR), in control and T3-treated cells for test pulse steps to +30 mV. D, Comparison of average 20–80% rise time and decay time constants of A currents in control and T3-treated cells. E, Average current to voltage relationships of peak and sustained current components, recorded after test pulse durations of 200 msec for control and T3-treated cells stimulated using the voltage protocol shown in the inset of panel A. Error bars indicate ±SE. ms, Milliseconds.

 
Acute effects of T3
T3 has been shown to directly induce bursting of Na+ channels as well as to increase Na+ currents via a G protein dependent mechanism in ventricle cells from the heart (34). To study whether T3 regulates Na+ currents directly in postnatal primary neurons, we tested the effects of 30 nM T3, which has been shown to up-regulate Na+ currents after 2–4 d in culture (13). After 15 min of superfusion, all 12 cells successfully recorded from showed no increase in the Na+ current recorded using near-physiological solutions (data not shown). The Na+ current varied between 252.3 ± 41.1 pA before, 224.7 ± 45.0 pA during, and 204.6 ± 35.8 pA after T3 application (n = 12, mean ± SE). Likewise, no change in the current-voltage relationship was observed.

Influence of T3 in neuron-enriched cultures
To investigate whether T3 acts directly on neurons after an extended incubation period, neuron-enriched cultures were prepared by incubating cells in NB medium replacing FCS by modified B18 supplement and additionally treating them with 4 µM cytosine β-D-arabinofuranoside (araC) to inhibit glial cell proliferation (Fig. 3AGo). Furthermore, mixed neuronal/glial cultures were grown in NB medium supplemented with FCS for the first 3 d in vitro (div), and no incubation with araC was performed (Fig. 3BGo). The first procedure led to neuron-enriched cultures, containing glial fibrillary acidic protein (GFAP)-positive astrocyte to neuron ratios varying between 0.9 ± 0.1 [postnatal d 2(P2)], 2.2 ± 0.1 [postnatal d 3 (P3)], and 1.3 ± 0.1 [postnatal d 4 (P4)] (three different preparations each) and mixed cultures containing an astrocyte to neuron ratio of 5.0 ± 0.4 (P2), 5.4 ± 0.7 (P3) up to 15.3±3.0 (P4) (at least four different preparations each; Fig. 3DGo). Before recording cells were either treated with none or 50 nM T3 for the last 4 div. No change of the cell capacitances was observed by T3 treatment; however, FCS-treated cells displayed a significant decrease of the cell capacitance compared with neuron-enriched cultures (Fig. 3CGo). A pretreatment with thyroid hormone increased the Na+ currents normalized to membrane capacitance only in mixed cultures originating from 4-d-old pups (43.1 ± 2.8 pA/pF control cells, n = 30 vs. 60.5 ± 3.9 pA/pF 50 nM T3, n = 31, cells recorded from five different preparations; Fig. 3HGo), which display a high glia to neuron ratio (see Fig. 3DGo). Normalized to membrane capacitance the Na+ current density observed in nonregulated neuron-enriched and mixed cultures varied from 35.3 ± 3.2 pA/pF to 53.2 ± 4.3 pA/pF (Fig. 3Go, G and H) (at least three preparations).


Figure 3
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  		Fig. 3. Influence of culture conditions on the regulation of voltage-gated Na+ currents by T3. Photomicrographs of cells immunostained with anti-βIII tubulin (neurons; white) and anti-GFAP antibodies (astrocytes; gray) of neuron-enriched (A) and mixed cultures grown in the presence of FCS (B). Nuclei visualized using Hoechst 33342. C, Mean capacitances from P2–P4 neuron-enriched and mixed cultures recorded after 3 d of preincubation and 4 d of treatment by T3 or vehicle. N, Neuron-enriched cultures treated with araC to reduce the number of glial cells; M, mixed cultures treated with FCS to promote proliferation of astrocytes. Each bar originates from at least 20 cells. D, Astrocyte to neuron ratio from neuron-enriched and mixed cultures. Significance evaluated with ANOVA followed by Dunnet’s post hoc test, comparing the T3-treated culture from P4 mixed cultures with all other culture types. Numbers below bars indicate numbers of preparations examined. E and F, Original current recordings from P4 hippocampal neurons in mixed cultures treated with 0 nM (E) or 50 nM T3 (F). G, Peak Nav-Ds in neuron-enriched cultures prepared from P2- to P4-old rats recorded after 3 d of preincubation and 4 d of treatment by T3 or vehicle. No regulation of the Nav-D by T3 treatment was observed. H, Na+ current densities in mixed cultures from P2–P4 rats. Note, that in neurons originating from P4 rats, containing the highest astrocyte to neuron ratio, an increase of the Nav-D by T3 treatment was observed. Peak currents (G and H) were determined at test potentials of –20 mV, starting from a holding potential of –85 mV and normalized to cell membrane capacitance. Numbers in bars indicate numbers of cells recorded. Error bars indicate SE (**, P < 0.01;***, P < 0.005).

 
Nav-D regulation by T3 requires the presence of satellite cells
The results illustrated in Fig. 3Go suggest that the presence of glial cells could be required for the action of T3 on the neuronal Nav-D. Alternatively, the presence of FCS in the mixed cultures could have promoted the T3 effect. To exclude a possible interference by FCS we cultured half of the neurons from 2-d-old pups directly on a 4-wk-old confluent astrocyte layer and incubated them with NB-medium supplemented with modified B18 to replace FCS. The second half of the neurons was used to prepare neuron-enriched control cultures in which the neurons were seeded directly on poly-L-lysine-coated cell culture dishes and treated with araC. The cultures were incubated with none or 30 nM T3 for 4 d after 3 div. Whereas in neuron-enriched control cultures the Nav-D was not influenced by T3 treatment (48.9 ± 4.9 pA/pF 0 nM T3, n = 33; 48.4 ± 5.1 pA/pF 30 nM T3, n = 33, six different preparations), neurons plated on a confluent astrocyte layer displayed an increase in the Nav-D from 34.7 ± 2.7 pA/pF (n = 32) in untreated cells to 56.4 ± 3.5 pA/pF (n = 38) in cells treated with 30 nM T3 (seven different preparations, Fig. 4AGo).


Figure 4
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  		Fig. 4. Influence of astrocytes on the regulation of the Na+ current density by T3. A, Peak Na+ current density of neurons from neuron-enriched cultures and from neurons plated on 4-wk-old astrocytes obtained from P2 rats incubated in the absence of FCS. Average peak Na+ currents measured at test potentials of –20 mV and normalized to cell membrane capacitance. Numbers in bar charts indicate numbers of cells investigated. B, Average current-voltage relationships for Na+ currents normalized to capacitance from neurons plated on astrocytes and cultured in the absence or presence of T3. C, Influence of T3 on steady-state inactivation of Na+ currents recorded from neurons cultured on astrocytes in the absence or presence of T3. Prepulse protocol: holding potential: –85 mV, incrementing prepulses in steps of 5 mV applied for 200 msec starting with hyperpolarizations to –105 mV followed by a depolarizing test pulse to –20 mV applied for 50 msec. Peak currents normalized to maximal peak currents. Solid lines represent fits to a modified Boltzmann equation. Error bars indicate SE (***, P < 0.005).

 
It has been observed that the voltage of inactivation of the Na+ current is shifted to more depolarized values after status epilepticus (35) and to the hyperpolarizing direction by the protein kinase C activators phorbol-12,13-diacetate (36) and 1-oleoyl-2-acetyl-sn-glycerol (37). To exclude a shift of the voltage of inactivation of the Na+ current to more depolarized values as a cause for the observed increase in the Nav-D, we investigated the steady state inactivation of Na+ currents in cultures on astrocyte layers treated with 0 nM or 30 nM T3. As previously demonstrated (13), no shift in inactivation was observed (IC50: –60.6 ± 0.8 mV 0 nM T3, n = 29; –60.0 ± 0.8 mV 30 nM T3, n = 26, seven different preparations; Fig. 4CGo). This is in line with the recent finding of an unaltered voltage dependence of inactivation of T3-up-regulated Na+ currents in fetal human neuroepithelial cells, but disagrees with observations on T3-up-regulated Na+ currents in human mesenchymal stem cells, in which a shift by 7 mV to more depolarized values was found (38). Treatment with T3 had no effect on cell capacitance, which amounted to 19.4 ± 1.0 pF (n = 33) for neuron-enriched control cells, 19.2 ± 1.0 pF (n = 33) for T3-treated neuron enriched cells, 18.4 ± 0.8 pF (n = 32) for neurons on astrocytes, and 19.3. ± 0.8 pF (n =38) for T3-treated neurons on an astrocyte layer. This indicates that the neuronal membrane surface area exposed to the whole-cell clamp did not vary significantly in sister cultures by T3 treatment and the presence of astrocytes.

These experiments demonstrate, that the presence of astrocytes, rather than exposure to FCS, is required for T3 to regulate the Nav-D. Furthermore, the experiments show that the regulation by T3 is not dependent on the age of the neurons at the day of preparation (P4) but is also found in neurons prepared at P2 if a sufficient number of astrocytes is present.

Glial cells influence the Nav-D via a heat-instable, soluble factor
We further tested whether direct contact of neurons to glial cells is necessary for T3 to regulate the Nav-D or whether glial cells secrete a putative soluble factor in response to T3 treatment. After 3 div neuron-enriched cultures from 2-d-old pups were cocultured for 4 d with cell culture inserts covered with a confluent glial cell layer (0.2-µm Anopore membrane; Nunc, Wiesbaden, Germany) either supplemented with none or 50 nM T3. Neuron-enriched cultures treated with araC and incubated in the presence or absence of 50 nM T3 for 4 d served as reference culture, which did not display an effect by T3.

The reference cultures did not exhibit a regulation of the Nav-D [41.4 ± 2.2 pA/pF 0 nM T3, (n = 41); 43.8 ± 2.7 pA/pF 50 nM T3 (n = 42), seven different preparations, Fig. 5Go]. In contrast, cultures containing astrocyte-coated inserts showed a 52% increase of the Nav-D [39.1 ± 2.3 pA/pF 0 nM T3 (n = 36); 59.6 ± 5.2 pA/pF 50 nM T3 (n = 33); six different preparations]. Next, we tested whether the direct presence of glial cells is essential during the whole T3 treatment period by incubating neuron-enriched cultures with astrocyte-conditioned medium (ACM) or ACM obtained from T3-treated astrocytes (T3-ACM). To obtain ACM and T3-ACM, confluent glial cell cultures were treated with none or 50 nM T3 for 2 d. Then neuron-enriched cultures were treated with ACM or T3-ACM for an additional 48 h. The Nav-D was significantly up-regulated by 31%, slightly less than observed in cocultures [39.9 ± 2.5 pA/pF ACM (n = 54); 52.1 ± 2.8 pA/pF T3-ACM (n = 50); 10 different preparations].


Figure 5
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  		Fig. 5. Effect of T3-induced release of soluble factors on average sodium current densities. Bars labeled "Neuron enriched" represent recordings from neurons cultured in NB medium and treated with araC to reduce growth of glial cells. Co-culture, Neuron-enriched cultures cocultured in the presence of astrocytes plated on cell culture inserts not touching the neurons. ACM, Neuron-enriched cultures after incubation with ACM (ACM/T3-ACM) for 48 h. Boiled ACM, Neuron-enriched cultures treated with boiled (5 min at 95 C) ACM/T3-ACM for 48 h. Numbers in bar charts indicate numbers of cells investigated. Error bars indicate SE (***, P < 0.005). All recordings from neuron-enriched cultures obtained from P2 rats treated for 4 d with either none or 50 nM T3.

 
To examine whether the Nav-D is regulated via a soluble protein, neuron-enriched cultures were incubated for 48 h with ACM or T3-ACM, each being preheated to 95 C for 5 min. Here, the effect of T3 was abolished [41.8 ± 3.5 pA/pF 0 nM T3 (n = 19); 41.8 ± 3.9 pA/pF 50 nM T3 (n = 16); four different preparations) suggesting that T3 treatment induces the release of a soluble protein from astrocytes.

Regulation of the Nav-D by bFGF
Because it has been reported that 0.6 nM bFGF increases the Nav-D in PC12 cells (26), we now investigated whether bFGF, known to be released from cerebellar glial cells after T3 treatment (20), regulates the Nav-D in neuron-enriched cultures lacking T3. After 3 div media were changed and cells were incubated with 6 nM bFGF for 1–6 d (d1–d6) to obtain clear effects. Basic FGF treatment induced characteristic morphological changes of astrocytes, which have been reported previously (Fig. 6Go, A and B) (39).


Figure 6
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  		Fig. 6. bFGF incubation in neuron-enriched cultures mimics T3 action in the presence of astrocytes. Immunohistochemical staining with GFAP of control cells (A) and bFGF-treated astrocytes (B); nuclei stained with Hoechst 33342. Note the characteristic change of astrocyte morphology due to treatment with 6 nM bFGF. Ca, Peak Na+current densities of neurons from neuron-enriched cultures lacking T3 treated with 6 nM bFGF for 1–6 d. Cb, Mean capacitances of the cells representing the source for the Nav-Ds shown in (Ca). Cc, Dose-response curve of Nav-Ds from neuron-enriched cultures prepared at P2 and recorded at d 7 after incubation with various concentrations of bFGF for 4 d. All data from at least nine different preparations. Numbers in bar charts indicate number of cells analyzed. D, Current to voltage relationship of peak Na+ currents. E, influence of bFGF on steady-state inactivation of Na+ currents of neurons treated with none or 6 nM bFGF for 4 d. For pulse protocol see legend of Fig. 4Go. Error bars indicate ±SE (*, P < 0.05; ***, P < 0.005).

 
In addition, the Nav-D was persistently increased after a single addition of bFGF for the following 6 d. The up-regulation varied between 48% (d6) and 116% (d3) (Fig. 6CGoa). As observed in T3-treated cells, a preincubation with bFGF changed the voltage dependence of activation by less than 5 mV (Fig. 6DGo) and did not influence the voltage dependence of inactivation (Fig. 6EGo). Cell capacitances were unaltered with the exception of d3, at which an increase by 35% occurred (Fig. 6CbGo). Changes in capacitance, however, were small compared with changes in Na+ current density, suggesting that bFGF regulates Na+ currents to a much larger extent than membrane surface.

A dose-response curve of bFGF effects, recorded 2–4 d after bFGF treatment, showed significant increases of the Nav-D starting at bFGF concentrations of 0.03 nM and 0.6 nM [38.4 ± 3.8 pA/pF (n = 154), control; 46.8 ± 2.3 pA/pF, 0.03 nM bFGF (n = 64)]. The largest increase by 80% was observed using a bFGF concentration of 6 nM [69.0 ± 3.6 pA/pF (n = 69), 12 different preparations; Fig. 6CcGo]. We did not test whether a saturation occurs using larger concentrations of bFGF because 6 nM bFGF induced morphological changes of astrocytes not seen with the T3 concentrations required to increase Nav-D (see Fig. 3BGo).

Neutralization assay
To test whether bFGF is an active component of T3-ACM, we performed a neutralization assay using monoclonal anti-bFGF antibodies (Upstate Biotechnology, Inc., Lake Placid, NY; catalog. no. 05-117). The working concentration of the antibody was tested by incubating neuron-enriched cultures either with 0.03 nM bFGF or with the antibody in addition to bFGF. This bFGF concentration was used because it already showed a regulation of the Nav-D but did not influence the morphology of the astrocytes, as observed in T3-treated cultures. As shown in Fig. 7AGo, the regulation of the Nav-D by 0.03 nM bFGF after 4 d of incubation was blocked by 8 µg/ml anti-bFGF.


Figure 7
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  		Fig. 7. Blockage of the T3-induced regulation of sodium current density by anti-bFGF antibodies. A, Reversal of the bFGF effect on Nav-D by anti-bFGF antibodies. Aa, Increase of average peak Na+ current density after 4 d of treatment with 0.03 nM bFGF. Ab, Neutralization of the effect of 0.03 nM bFGF by 8 µg/ml anti-bFGF. B, Effect of anti-bFGF on T3-induced Nav-D regulation. Ba, Increase in the Nav-D by T3-ACM treatment for 48 h, recorded using near-physiological solutions. Bb, Note the block of the increase of the Nav-D by coincubation of T3-ACM with 8 µg/ml anti-bFGF. Bc, Note the increase of the Nav-D by addition of 6 nM bFGF; however, no further significant difference between ACM + bFGF and T3-ACM + bFGF. Numbers in bar charts indicate number of cells investigated. Error bars indicate ± SE (***, P < 0.005).

 
Then, neuron-enriched cultures were incubated with ACM as well as T3-ACM both treated with anti-bFGF for 1 h at 37 C before the cells were incubated at 37 C for 48 h. ACM and T3-ACM treated cells served as control cultures. In accordance with the results shown in Fig. 5Go, the Nav-D in control cultures increased from 48.0 ± 2.7 pA/pF (ACM, n = 28) to 68.1 ± 5.8 pA/pF [T3-ACM (n = 24), six different preparations, Fig. 7BGoa]. Incubation with anti-bFGF blocked the effect of T3-ACM [ACM + anti-bFGF: 55.4 ± 4.2 pA/pF (n = 25); T3-ACM + anti-bFGF: 56.4 ± 4.6 pA/pF (n = 24); five different preparations].

To study whether T3 treatment would further increase the Nav-D in the presence of a high concentration of bFGF, neuron-enriched cultures were treated with ACM, respectively, T3-ACM each supplemented with 6 nM bFGF. The addition of 6 nM bFGF increased the Nav-D in ACM and T3-ACM [87.0 ± 10.4 pA/pF, ACM + bFGF (n = 13); 85.9 ± 10.7 pA/pF T3-ACM + bFGF (n =14), three different preparations, Fig. 7BcGo) by about the same values as observed for 6 nM bFGF treatment in neuron-enriched cultures (Fig. 6CGo). However, no significant difference was observed between ACM + bFGF and T3-ACM + bFGF cultures, suggesting that the effects of T3 and bFGF were not additive (Fig. 7BGo).


   Discussion
TOP
 ABSTRACT
 INTRODUCTION
 Results
 Discussion
Materials and Methods
 REFERENCES
 
This is, to our knowledge, the first study demonstrating a role of glial cells in regulating the excitability of neurons in the central nervous system and strengthens the view of glia as a paracrine tissue. This conclusion is based on the observation that thyroid hormone increased the Na+ current density in hippocampal neurons from postnatal rats only in the presence of a high astrocyte to neuron ratio. This regulation did not require direct physical contact between neurons and astrocytes. We further obtained evidence that astrocytes secrete a heat-instable factor that causes the regulation of the Nav-D in neurons. In addition, we demonstrated that bFGF, known to be released from cerebellar glial cells after treatment with T3 (20), increases the Nav-D in the absence of glial cells. Treatment of conditioned medium obtained from T3 treated astrocytes with anti-bFGF blocked the effect on the Nav-D. Our results suggest that bFGF is a major component in the cascade of T3 action and involved in the regulation of the Nav-D in hippocampal neurons.

Direct effects of T3
In cardiac myocytes acute exposure to T3 induces bursting of Na+ channels (34) and increases Na+ currents (40). This effect might be mediated by direct binding of thyroid hormone to plasma membrane receptors (41) such as the recently identified integrin {alpha}Vβ3 T4 receptors (42, 43). Such a membrane receptor could lead to Na+ channel regulation via activation of G proteins, including a shift in the steady-state inactivation in ventricular myocytes from adult guinea pigs after application of 1 nM T3 (44). The resulting increase in the membrane excitability may facilitate ventricular arrhythmias in hyperthyroid patients with hypertrophy, ischemia, and/or heart failure (45).

Using concentrations in the same order of magnitude as those used to elicit direct effects on heart cells, we did not observe direct effects of T3 within 15 min of application. Furthermore, we observed no shift of the voltage dependence of inactivation of the Na+ current to more depolarized values as characteristic for phosphorylation effects on Na+ channels (36, 37, 46), induced by activation of membrane receptors. Third, our own preliminary experiments using quantitative PCR (Ref. 47 ; and http://deposit.ddb.de/cgi-bin/dokserv?idn=987289209)present evidence that Nav 1.6 mRNA is regulated by T3 under the present conditions. This suggests that sodium currents in the heart, which are dominated by the Na+ channel {alpha}-subunit (Nav1.5) (48), are regulated by T3 in a different manner than Na+ currents in hippocampal neurons. However, in accordance with the observation of an increase in excitability induced by microelectrophoretically injected thyroid hormone on single neurons from cat (49), an additional, nongenomic pathway for T4-dependent modulation of Na+ currents has been reported recently in Rohon-Beard sensory neurons from zebrafish embryos (50). Here, an acute up-regulation of the Na+ current density by T4 (not T3) has been described to be mediated by T4 binding to {alpha}Vβ3-integrins. This suggests that multiple mechanisms leading to thyroid hormone-dependent Na+ current regulation at different time scales and/or developmental stages could have evolved.

T3 effects on glia
Glial cells express nuclear thyroid hormone receptors (14, 15, 17, 19, 51). Thyroid hormones induce phosphorylation of specific proteins (52) and cell proliferation and differentiation of primary astrocyte cultures, transforming flat, polygonal astrocytes into process-bearing cells (53, 54, 55). Most investigations concerning the involvement of T3 in neuron-glia interactions have been performed in the cerebellum. Here T3 induces the synthesis and secretion of growth factors by glial cells (21, 53), among others, bFGF, aFGF, TNF-β, and IL-3. In addition, a basal secretion of bFGF, aFGF, and TGF-β in the absence of T3 (20) has been reported. Gomes et al. (22) demonstrated that T3-treated cerebellar astrocytes secrete EGF, which induces external granular layer neurons to initiate outgrowth of neurites (56) and induces neuronal proliferation in combination with TNF-β (22). Because Trentin et al. (20) found bFGF to be the most prominent factor released by cerebellar astrocytes, we focused in our present study on the potential action of T3-induced release of bFGF from glial cells.

Interaction between glial cells and neurons
Glial cells have recently been shown to influence synapse formation, control synaptic strength, and participate in information processing by coordinating the activity among sets of neurons (57, 58, 59). They modulate synaptic transmission via release of glutamate (60), PGE2, and TNF-{alpha} (30, 61). Other known gliotransmitters include D-serine, aspartate, taurine, neuropepties, eicosanoids, steroids, growth factors (62, 63, 64), and ATP (65).

Despite the vast knowledge available concerning how glial cells influence synaptic transmission, little is known whether glia can influence neuronal function by regulating the expression pattern of voltage-gated ion currents. In the peripheral nervous system Schwann cells have been demonstrated to modulate Na+ channel expression in spinal sensory neurons in vitro (66); however, the molecule involved was not further characterized. Recently PGE2 was shown to increase Nav1.9 Na+ currents in small dorsal root ganglion neurons (67). Here we have elaborated, for the first time, a consecutive signal chain in which T3 leads to an up-regulation of Na+ currents via secretion of a soluble factor from glial cells involving bFGF.

The link between thyroid hormone and bFGF
Interestingly, in other tissues a similar interplay between two cell types mediated by T3-stimulated bFGF release has been described. Hence, T3 stimulates bone resorption in organ cultures, but studies of isolated osteoclasts have shown that this effect requires cocultured osteoblasts (68). The bone cells of the osteoblast and chondrocyte lineages are the major T3-responsive cells (69, 70). FGFR1 mRNA and protein are up-regulated in osteoblasts 6–48 h after T3 treatment (71), which could finally result in the bone loss of hyperthyroidism (72). As a second example, in a chick chorioallantoic membrane model, T4 was shown to be a proangiogenic factor, the effect mediated via released bFGF (73). Furthermore, it has been described that in T4-treated rat capillary endothelial cells, DNA synthesis occurs after a single injection of T4 coincident with an up-regulation in bFGF mRNA and an increase in bFGF protein concentration resulting in enhanced proliferation in the venular capillaries in the heart (74). Our present results extend these observations, indicating that bFGF release induced by T3-stimulated hippocampal astrocytes influences the sodium current density and thus the excitability of hippocampal neurons.


   Materials and Methods
TOP
 ABSTRACT
 INTRODUCTION
 Results
 Discussion
 Materials and Methods
REFERENCES
 
Preparation of cell cultures
Hippocampi were obtained from 2- to 4-d-old Wistar rats under sterile conditions and collected in ice-cold modified PBS containing [in millimolar concentration; chemicals when not stated otherwise were from Sigma (Schnelldorf, Germany): NaCl, 137 (J. T. Baker, Deventer, The Netherlands); KCl, 2.7 (Riedel-de-Haen, Seelze, Germany); Na2HPO4, 5 (J.T. Baker), KH2PO4, 0.89 (Riedel-de-Haen, Seelze, Germany); HEPES, 10 (Biomol, Hamburg, Germany); pyruvate, 1; glucose, 10 (J.T. Baker); L-alanyl glutamine, 1 (Invitrogen, Karlsruhe, Germany); and 1 mg/ml BSA, 25 U/ml penicillin, and 25 µg/ml streptomycin]. Additionally, 10 µg/ml deoxyribonuclease I and 5 µl/ml of a 2.5% trypsin (Invitrogen) solution were added. After incubation of the tissue under gentle agitation for 7 min at 37 C, cells were triturated 15 times with a plastic pipette tip. The dissociated cells were collected in 10 ml modified PBS and centrifuged at 145 x g at room temperature for 12 min. The pellet was resuspended in RPMI-1640 (PAA, Cölbe, Germany) containing 10% fetal calf serum (FCS, Invitrogen) and incubated at 37 C and 5% CO2 in humidified air for 1 h (B5060 incubator; Heraeus, Hanau, Germany). For coculture experiments, cells were incubated on cell culture inserts (0.2-µm Anopore membrane; Nunc, Wiesbaden, Germany). After preplating, the supernatant containing the neurons was collected in 10 ml RPMI-1640 and centrifuged at 145 x g at room temperature for 12 min. The pellet was resuspended in NB medium (Invitrogen) and either supplemented with modified B18 solution (75) to obtain neuron-enriched cultures or 10% FCS for mixed cultures. All cell culture media were supplemented with 10 U/ml penicillin, 10 µg/ml streptomycin, and 2 mM L-alanyl glutamine. End concentration of modified B18 medium contained in micromolar concentration: biotin, 20; L-carnitine hydrochloride, 600; corticosterone, 2.9; ethanolamine, 800; D-galactose, 4150; gluthatione red, 160; linoleic acid, 175, linolenic acid, 175; progesterone, 1; putrescine dihydrochloride, 9150; retinyl acetate, 10; sodium selenite, 1.9; DL-{alpha}-tocopherole, 115; DL-{alpha}-tocopherole acetate, 105; BSA, 1850; catalase, 0.5; insulin, 30; superoxide dismutase, 3.85; and transferrin, 3.1; in PBS..

Cells were seeded on 3.5-cm plastic or 12-mm cell culture dishes coated with poly-L-lysine at a density of 5–9 x 105 cells/cm2 and stored at 37 C and 5% CO2 in humidified air.

Neuron-enriched cultures were treated 2 d after preparation with 4 µM (araC) for 24 h.

Hippocampal glial cells were grown from the dishes used for preplating and kept in RPMI-1640 supplemented with 10% FCS at 37 C and 5% CO2 in humidified air. Media were changed once a week until cells reached confluence. One day before further use glial cells either in cell culture dishes or inserts were washed three times with serum-free NB medium containing modified B18 to remove FCS and stored at 37 C and 5% CO2 in humidified air.

For experiments investigating acute effects of T3 on sodium channel regulation, hippocampal neurons were prepared as described previously (13).

Cell treatment and preparation of astrocyte-conditioned medium
T3 (Sigma-Aldrich, Schnelldorf, Germany) was dissolved in 0.01 N NaOH solution to a concentration of 300 µM. A stock solution of 1 µl of this solution/ml NB medium containing modified B18 supplement was prepared yielding a concentration of 300 nM. Cell culture medium was changed 3 d after preparation, and cells were either incubated with 0 nM, 30 nM, or 50 nM T3 or varying concentrations of bFGF (R&D Systems, Wiesbaden-Nordenstadt, Germany). ACM, or T3-treated ACM (T3-ACM), respectively, were collected 2 d after incubation with T3, centrifuged at 2000 x g for 10 min, and either used directly or stored at –20 C for further use. For neutralization assays, anti-bFGF (8 µg/ml, Upstate Biotechnology, Inc., Lake Placid, NY) was added to ACM or T3-ACM, respectively, agitated at 37 C for 1 h and incubated with neuron-enriched cultures 5 d after preparation for 48 h at 37 C, 5% CO2 in humidified air.

Immunohistochemistry
Cultured cells were washed with PBS, fixed with 4% paraformaldehyde for 20 min at room temperature, and permeabilized with 0.05% Triton X-100 (Sigma-Aldrich, Schnelldorf, Germany) for 10 min at room temperature. Cells were blocked with 1% goat serum in PBS for 30 min and incubated for 45 min at room temperature with primary antibodies (mouse anti-β-tubulin III and rabbit antiglial fibrillary acidic protein; both Sigma-Aldrich, 1:350). Afterward cells were washed twice with PBS and incubated with secondary antibodies for 45 min at room temperature (Alexa Fluor 488 goat antimouse IgG, Alexa Fluor 594 goat antirabbit IgG; Invitrogen, 1:2000). Negative controls were performed by omitting the primary antibody during staining. No reactivity was observed in the absence of the primary antibody. Then cells were washed with PBS and incubated with Hoechst 33342 (1 µg/ml, Sigma-Aldrich) for 10 min at room temperature. Cell preparations were mounted on Immuno-Fluore (MP Biomedicals, Inc., Eschwege, Germany) and visualized by using an Olympus IX 51 microscope and analySIS B software (Olympus, Hamburg, Germany).

Patch-clamp recording conditions
Whole-cell recordings were performed with borosilicate pipettes (GB-150TF-8P; Science Products, Hofheim, Germany) pulled on a PP-830 puller (Narishige Europe, London, UK) yielding resistances between 4–5 M{Omega} after filling. Pipette solutions for recording whole-cell currents in near physiological composition contained in millimolar concentration: K+-gluconate, 100; EGTA, 1.1; CaCl2, 0.1; MgCl2, 5; NaCl, 5; HEPES, 10; Mg2+-ATP, 3. Bath solutions contained in millimolar concentration: NaCl, 110; CaCl2, 1.8; glucose, 10; HEPES, 10; KCl, 5.4; MgCl2, 0.8. Osmolarities of pipette and bath solutions were adjusted to the osmolarity of the NB medium. A liquid junction potential of this solution with respect to the bath solution of 15 mV was corrected offline. Peak Na+ currents were measured at a potential of –20 mV after correction of the liquid junction potential. Only Na+ currents showing an activation range exceeding 20 mV in the current to voltage relationship were included in the evaluation of the data. Because larger currents in T3-treated cells could lead to larger voltage errors compared with recordings from the control cells, voltage errors were calculated as products of series resistances, as measured after rupturing the membrane, and maximal Na+ current amplitudes. These were 4.4 ± 0.5 mV (n = 30) for recordings from control cells and 5.3 ± 0.6 mV (n = 31) for recordings from T3-treated cells in FCS containing cultures (for data shown in Fig. 3HGo, P4). The largest voltage errors occurred for the recordings of Na+ currents after treatment with 6 nM bFGF. Here, it amounted to 5.7 ± 0.5 mV (n = 77) in control cells and 9.3 ± 0.8 mV (n = 22) in bFGF-treated cells, explaining the 5-mV shift in the peak of the I/V relationship of the bFGF-treated cells with respect to the control cells (Fig. 6DGo).

Isolated Na+ currents were recorded using pipette solutions containing in millimolar concentration: CaCl2, 0.1; CsF, 100; EGTA, 1.1; HEPES, 10; MgCl2, 5; NaCl, 5; and bath solutions containing in millimolar concentration: 4-aminopyridine, 4; CaCl2, 1; CdCl2, 0.5; glucose, 10; HEPES, 10; MgCl2, 1; NaCl, 100; TEA-Cl, 10. The osmolarity of the bath solution was adjusted to 245 mosmol to match the osmolarity of the NB culture medium, and pH was adjusted to 7.35.

Pipette, bath, and cell culture solutions used in experiments investigating acute effects of T3 on hippocampal neurons were described elsewhere (13), and a superfusion system was used (76). Previously, after 15 min T3 treatment and 15 min after washout, voltage-activated whole-cell currents were measured and I/V-relationships registered every minute.

K+ currents were recorded using extracellular solutions containing in millimolar concentration: CaCl2, 1.8; CdCl2, 0.5; KCl, 5.4; MgCl2, 0.8; glucose, 10; N-methyl-D-glucamine, 110; HEPES, 10; TTX, 0.05 x 10–3. A liquid junction potential of this solution of –10 mV was corrected offline.

Total K+ currents were recorded using a series of voltage steps starting from a holding potential of –80 mV. To isolate the persistent KDR current, cells were predepolarized to a holding potential of –15 mV for 200 msec before stepping to a test potential of +30 mV (Fig. 2BGo, traces at the right). To recover the A current component from inactivation, cells were then prehyperpolarized to a prepulse potential of –100 mV for 200 msec and then pulsed to the same test potential of +30 mV (see inset in Fig. 2BGo). The A current component was then isolated by subtracting the KDR current from the total K+ current.

Recordings were performed at room temperature using a Patch-Clamp L/M-EPC7 amplifier (List Medical, Darmstadt, Germany). Series resistance errors were compensated by up to 30%. Signals were filtered using the EPC7 10-kHz lowpass filter and then digitized with a sampling rate of 2 kHz. Data were digitized with a DigiData 1200 board (Axon Instruments, Union City, CA) and stored on an IBM-compatible PC. Data evaluation was performed with the PClamp 6 software (Axon Instruments). Leakage and capacitive artifacts were subtracted using a P/4 protocol. Current densities were determined by normalizing the respective current to the cell capacitance calculated from the integral of the charging curve for a test potential step of 20 mV after compensation of the electrode capacitance. Cells with series resistances above 20 M{Omega} were discarded to minimize series resistance errors.

Student’s t test for statistical comparison of two different datasets was used; for multiple comparisons, e.g. the bFGF dose-response curve, ANOVA followed by Dunnett’s test was used. Errors are given as ± SE. Statistics were performed with the Origin 5.0 software (Microcal Software, Inc., Northhampton, MA) and the SSPS 15.0 software (SSPS GmbH Software, München, Germany).


   ACKNOWLEDGMENTS
 
We thank S. A. Mann, K. Chakrabarty, and R. Heumann (all at Ruhr-University Bochum at the time this work was performed) for helpful discussions, technical advice, and support. We thank H. Breuker-Siraj for help with the cell culture, M. Conrad for some of the recordings shown in Fig. 6Go, C and D, and D. Meyen for contributing to the recordings summarized in Fig. 6CcGo in the course of their undergraduate studies. The major part of the present publication is based on data collected in the PhD thesis of Vanessa Niederkinkhaus.


   FOOTNOTES
 
This work was supported by a fellowship from the International Graduate School of Neuroscience (to V.N.) and a fellowship of the country of North Rhine-Westphalia (to G.H.).

Present address for G.H.: PFM AG, Wankelstrasse 60, D-50996 Köln, Germany.

Disclosure Summary: The authors have nothing to disclose.

First Published Online May 21, 2009

Abbreviations: ACM, astrocyte-conditioned medium; aFGF, acidic fibroblast growth factor; araC, cytosine β-D-arabinofuranoside hydrochloride; bFGF, basic fibroblast growth factor; div, days in vitro; EGF, epidermal growth factor; FCS, fetal calf serum; GFAP, glial fibrillary acidic protein; Nav-D, voltage-gated Na+ current density; NB, neurobasal; P2, postnatal d 2.

Received for publication March 20, 2009. Accepted for publication May 15, 2009.


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 Discussion
 Materials and Methods
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Ligands:   Thyroid hormone




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