Department of Neurophysiology, Paul Flechsig Institute of Brain Research, University of Leipzig, D-04109 Leipzig, Germany
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ABSTRACT |
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Bringmann, A.,
S. Schopf, and
A. Reichenbach.
Developmental Regulation of Calcium Channel-Mediated Currents in
Retinal Glial (Müller) Cells.
J. Neurophysiol. 84: 2975-2983, 2000.
Whole cell voltage-clamp
recordings of freshly isolated cells were used to study changes in the
currents through voltage-gated Ca2+ channels
during the postnatal development of immature radial glial cells into
Müller cells of the rabbit retina. Using
Ba2+ or Ca2+ ions as charge
carriers, currents through transient low-voltage-activated (LVA)
Ca2+ channels were recorded in cells from early
postnatal stages, with an activation threshold at 60 mV and a peak
current at
25 mV. To increase the amplitude of currents through
Ca2+ channels, Na+ ions
were used as the main charge carriers, and currents were recorded in
divalent cation-free bath solutions. Currents through transient LVA
Ca2+ channels were found in all radial glial
cells from retinae between postnatal days 2 and 37. The currents
activated at potentials positive to
80 mV and displayed a maximum at
40 mV. The amplitude of LVA currents increased during the first
postnatal week; after postnatal day 6, the amplitude remained virtually
constant. The density of LVA currents was highest at early postnatal
days (days 2-5: 13 pA/pF) and decreased to a stable, moderate level
within the first three postnatal weeks (3 pA/pF). A significant
expression of currents through sustained, high-voltage-activated
Ca2+ channels was found after the third postnatal
week in ~25% of the investigated cells. The early and sole
expression of transient currents at high-density may suggest that LVA
Ca2+ channels are involved in early developmental
processes of rabbit Müller cells.
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INTRODUCTION |
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Müller (radial
glial) cells are the principal type of glial cells in the mammalian
retina. One of the main functions of Müller cells is the spatial
buffering of the extracellular K+ concentration
(Newman and Reichenbach 1996; Newman et al.
1984
). This function is mediated by the prominent
K+ permeability of Müller cell membranes,
in particular, by inwardly rectifying K+ channels
(Brew et al. 1986
; Newman 1993
). In
addition to the dominant expression of K+
channels, Müller cells may express other ion channels in their membranes; among them are voltage-gated Na+
(Chao et al. 1994
; Francke et al. 1996
)
and Ca2+ channels (Newman 1985
).
In cultured human Müller cells, the presence of distinct types of
voltage-gated Ca2+ channels has been described
(Puro and Mano 1991
; Puro et al. 1996
).
The functional roles of Na+ and
Ca2+ channels in Müller cells are still
unclear, although sparse previous data indicated their involvement in
transdifferentiation and proliferative processes in cases of retinal
pathobiology. Voltage-gated Na+ currents, for
example, were found to be increased in their amplitude in human
Müller cells obtained from patients with various eye diseases
(Francke et al. 1996
). L-type Ca2+
channels, on the other hand, have been implicated in the regulation of
the proliferative activity of cultured Müller cells (Puro and Mano 1991
; Uchihori and Puro 1991
).
There is convincing evidence that, generally, glial cells may express
voltage-gated Ca2+ channels of different distinct
types (Sontheimer 1994; Steinhäuser 1993
). Whereas the postnatal development of
Ca2+ channel expression was extensively studied
in neurons (for example, McCobb et al. 1989
;
Pirchio et al. 1990
; Thompson and Wong
1989
; Yaari et al. 1987
), only a few data are
available about developmental changes of voltage-gated
Ca2+ channels in glial cells. Cultured glial
precursor cells of the oligodendrocyte lineage express two types of
Ca2+ currents that can be distinguished by the
voltage dependence of activation: low- and high-voltage-activated
Ca2+ currents (LVA and HVA currents)
(Verkhratsky et al. 1990
). Glioblasts and
oligodendrocytes of the murine corpus callosum express
Na+ and Ca2+ currents
during early postnatal stages but not in the differentiated stage
(Berger et al. 1992
). In neurons, T-type LVA
Ca2+ channels are predominantly expressed at
early embryonic or neonatal stages when they generate spontaneous
Ca2+ transients necessary for morphogenesis
(Gu and Spitzer 1993
); thereafter, their disappearance
is accompanied by an increase in HVA Ca2+
currents (Carbone and Lux 1984
; Kostyuk et al.
1993
; McCobb et al. 1989
; Tarasenko et
al. 1998
). In certain thalamic and hypothalamic neurons,
however, T-type LVA Ca2+ channels play an
important role in the organization of slow rhythmic activity in adult
animals (Akaike et al. 1989
; Huguenard and Prince 1992
).
The first aim of the present study was to detect voltage-gated
Ca2+ currents in freshly isolated Müller
cells (and/or their precursors) of the rabbit. Using
Ba2+ or Ca2+ ions as charge
carriers, transient (T-type) LVA currents were found in cells from
young postnatal animals. The second aim was to determine whether the
expression level of Ca2+ channels in Müller
cells is developmentally regulated. If the Ca2+
channels are implicated in the proliferative activity of Müller cells (Puro and Mano 1991; Uchihori and Puro
1991
), one may assume that the activity of
Ca2+ channels decreases during the postnatal
differentiation of mitotically active precursor cells, via immature
radial glial cells, into mature Müller cells. Since the
amplitudes of the Ba2+ and
Ca2+ currents were very small, it was
advantageous to maximize the currents through voltage-gated
Ca2+ channels. For this purpose, their
developmental regulation was investigated by recording currents of
monovalent cations. As previously described (Almers and
McCleskey 1984
; Hess and Tsien 1984
;
Kostyuk et al. 1983
; Lux et al. 1989
,
1990
), voltage-gated Ca2+ channels
mediate currents of monovalent ions when divalent cations are largely
absent in the extracellular solution.
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METHODS |
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Cell isolation and identification
Rabbits were kept in the animal center of the Leipzig University Medical School. Animals were deeply anesthetized by urethan (2.0 g/kg) before decapitation and enucleation of the eye balls. Isolated retinae were incubated at 37°C for 30 min in Ca2+- and Mg2+-free phosphate-buffered saline (PBS), pH 7.4, which contained papain (15 U/ml; Boehringer, Mannheim, Germany) and desoxyribonuclease I (200 U/ml; Sigma, Deisenhofen, Germany). After washing in PBS, the tissue was titrated by a wide-pore pipette. The resulting suspensions of dissociated singular cells were stored at 4-8°C in serum free modified Eagle's medium (Sigma) until use within 3 h after isolation.
Müller cells were identified by their unique bipolar morphology
and size (>100 µm length). While this discriminated them unequivocally from bipolar neurons and photoreceptor cells (which may
roughly resemble immature Müller cells in shape but are much shorter), it was impossible to distinguish immature Müller cells from late progenitor cells that are still present during the first few
postnatal days. The latter, producing both glial (Müller) and
neuronal (bipolar and photoreceptor cells) by their last division, share the same set of antigenic molecules with the Müller cells (Reichenbach 1993 and references therein) and cannot be
distinctly labeled by any known antibody or marker. Thus both (a
majority of) immature Müller cells and (less and less) late
progenitor cells may have been grouped together between postnatal days
(P) 2 and 6. However, as there seems to be a gradual rather than a clear-cut transition between the two cell types (cf. Bringmann et al. 1999
), this was not considered as a major obstacle of
our study.
Electrophysiological recordings
Whole cell voltage-clamp currents (Hamill et al.
1981) were measured using a List EPC-7 amplifier (List
Electronics, Darmstadt, Germany) and the TIDA 5.72 computer program
(HEKA elektronik, Lambrecht, Germany). High frequencies >4 kHz were
cutoff. The series resistance (13-16 M
) was compensated by
30-50%. Records were made at room temperature (22-25°C). Patch
electrodes of 4-7 M
resistance were pulled from borosilicate glass
(GB150F8P, Biologic, Science Products, Frankfurt/M., Germany). The
Ca2+ channel-mediated currents were evoked by a
standard step protocols (Vh,
80 mV;
depolarizing voltage steps from a 500-ms prepulse). For LVA current
activation, depolarizing voltage steps were applied to voltages between
100 and +20 mV with an increment of 10 mV, after a prepulse to
120
mV. For HVA current activation, voltage steps were applied to voltages
between
100 and +20 mV, after a prepulse to
60 mV. To assess LVA
current inactivation, voltage steps to
50 mV were applied after
prepulses to different potentials (between
120 and 0 mV; increment,
10 mV). For HVA current inactivation, voltage steps to
20 mV were
applied after prepulses to potentials between
80 and +40 mV
(increment, 10 mV). The traces were not leak subtracted. Leak currents
were subtracted when I-V curves were calculated. Data were
not corrected for liquid junction potentials since they did not exceed
3 mV. The membrane capacitance of the cells was measured by integrating
the uncompensated capacitive artifact evoked by a hyperpolarizing
voltage step from
80 to
90 mV. For recording the capacitive
artifact, the sampling rate was 30 kHz, and the frequencies 10 kHz
were cutoff. Measurements of time-dependent changes of the
Ca2+ channel-mediated currents indicated that,
after disruption of the membrane, dialysis of the cell interior was
completed within 3 min, and, thereafter, current amplitudes remained
stable for ~10 min. Thus the "run-down" of the
Ca2+ channel-mediated currents was considered to
be negligible within this period of time.
Solutions
The pipette solution consisted of (in mM) 10 NaCl, 130 CsCl, 1 CaCl2, 1 MgCl2, 10 ethyleneglycolbis(aminoethyl)-(ether)tetra-acetate (EGTA), and 10 N-2-hydroxyethyl-piperazine-N'2-ethanesulphonic acid (HEPES). The pH of 7.2 was adjusted with Tris-base. The
Ba2+ currents were recorded with a bath solution
containing (in mM) 20 BaCl2, 130 CsCl, 5 HEPES,
and 10 glucose (pH 7.4 adjusted with Tris-base). The
Ca2+ currents were recorded using a bath solution
that consisted of (in mM) 105 NaCl, 10 CaCl2, 1 MgCl2, 10 HEPES, and 11 glucose (pH 7.4). To
increase the amplitude of the currents through
Ca2+ channels, currents of monovalent cations in
divalent cation-free bath solutions were recorded. The bath solution
consisted of (in mM) 113 NaCl, 10 HEPES, 11 glucose, and 1 EGTA (pH
7.4). To obtain Na+-free solution,
Na+ was equimolarly replaced by
choline+. To investigate the
Ca2+ dependence of the Na+
currents, the control data were obtained in Ca2+-
and Mg2+-free solution containing 1 mM EGTA. The
solutions with 0.1, 1, 5, and 10 µM free Ca2+
were made by adding 0.625, 0.944, 0.993, and 1.004 mM
CaCl2, respectively, to the control solution. The
effect of other divalent cations was tested in solutions without EGTA.
Substances
Nimodipine and flunarizine were from Calbiochem (Bad Soden, Germany). Tetrodotoxin was obtained from Alomone Labs (Jerusalem, Israel). All other substances were from Sigma. Lipophilic drugs were dissolved in dimethylsulphoxide. Vehicle controls were prepared as above without addition of the drug. Drugs were applied by changing the perfusate within the recording chamber.
Data presentation
Single-channel currents depicted as negative (downward deflections) represent cation fluxes from the extra- into the intracellular compartment. Amplitude histograms of single-channel currents were established by means of the TIDA 5.72 computer program. Single-channel currents were evaluated from the current steps between peaks of the amplitude histograms. Statistical analysis (regression analysis, Bonferroni corrected P values using ANOVA) and curve fits were made using the Prism program (Graphpad Software, San Diego, CA). Data are expressed as means ± SD.
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RESULTS |
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Ba2+ and Ca2+ currents
To isolate Ba2+ currents through
Ca2+ channels in Müller cells of young
rabbits [postnatal days (P) 10-11], voltage step protocols were
applied using 20 mM extracellular Ba2+ as the
charge carrier, in K+-free bathing and pipette
solutions. Under these conditions, a depolarization-activated inward
current was found to be present in the whole cell records (Fig.
1A). The mean peak amplitude
of this current was small, with 10.6 ± 4.5 pA (n = 14). The current activated at potentials positive to 60 mV; the
peak was at
22.9 ± 7.5 mV (n = 32; Fig.
1B). The inward current was completely blocked by exposure
to extracellular Cd2+ (1 mM; n = 9) and is, therefore assumed to be mediated by
Ca2+ channels. The Ba2+
currents showed transient time-dependent activation kinetics (Fig.
1A) similar to the T-type currents previously described in
cultured human Müller cells (Puro and Mano 1991
).
Moreover the relatively low activation threshold is indicative of
currents through LVA Ca2+ channels. Transient
inwardly directed currents were also observed when
Ca2+ ions (10 mM) were used as the charge carrier
(Fig. 1C). The Ca2+ currents activated
at potentials positive to
60 mV, peaked at
28.1 ± 8.8 mV
(n = 5) and were blocked by external
Cd2+ ions (1 mM). Again, the peak amplitude of
the Ca2+ currents was small, with 8.2 ± 3.2 pA (n = 5).
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Since the amplitudes of the Ca2+ channel-mediated currents were very small when Ba2+ or Ca2+ ions were used in the extracellular solution, we used Na+ ions as the main charge carrier in all further experiments to increase the amplitude of the currents flowing through Ca2+ channels. Therefore bath solutions without divalent cations were used.
Na+ currents through Ca2+ channels
Using Ca2+-, Mg2+-, and K+-free bath solutions, large inwardly directed currents were evoked by depolarizing voltage steps. Figure 2A shows records in a cell from a P24 rabbit. The voltage-gated, inwardly directed currents were strongly diminished in their amplitudes when Ca2+ and Mg2+ ions were present in the bath solution at physiological concentrations (2 and 1 mM, respectively; middle). The amplitude of the inwardly directed currents recovered after a washout of the divalent cations (right). The inwardly directed currents recorded in divalent cation-free bath solution were mediated by external Na+ ions because they were absent when the bath solution was Na+ free (Fig. 2B).
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Because the activation kinetics of the inwardly directed currents were transient (Fig. 2), we assumed that these currents represent Na+ fluxes through transient (T-type) Ca2+ channels. Therefore we tested whether blockers of voltage-gated Ca2+ channels may affect the currents. As shown in Fig. 2C, the T-type channel blocker flunarizine (at a concentration of 5 µM) almost completely blocked the inwardly directed currents. The dose-dependence of the peak current inhibition revealed that 0.6 µM flunarizine inhibited 50% of the transient currents (Fig. 2D). Nimodipine also blocked this current, although at significant higher concentrations (IC50 = 3.6 µM). A blocking effect of nimodipine (10 µM) was also observed on the Ba2+ currents through LVA Ca2+ channels (not shown). Tetrodotoxin, a blocker of fast Na+ channels, did not inhibit the transient Na+ currents (n = 9; Fig. 2E). As indicated by both the sensitivity to Ca2+ channel blockers and the insensitivity to tetrodotoxin, the transient Na+ currents are mediated by T-type Ca2+ channels.
The Na+ currents through Ca2+ channels were blocked by extracellular divalent cations (Fig. 2A). Ca2+ ions decreased the amplitude of the transient currents in the low micromolar range, in a dose-dependent manner (Fig. 3A). The current inhibition by external Ca2+ ions was prominent over the entire voltage range with slightly greater inhibiting effects at more depolarized voltages (Fig. 3B). The dose dependences of the LVA-current block by various divalent cations are illustrated in Fig. 3C. The peak current was reduced to 50% by 0.6 µM Cd2+, by 0.7 µM Ca2+, by 1.7 µM Ni2+, by 3.0 µM Cu2+, by 31.4 µM Ba2+, and by 35.9 µM Mg2+.
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Although the vast majority of cells expressed only currents through
transient LVA channels (623 out of 653 investigated cells), cells from
rabbits older than two weeks also showed currents through sustained,
long-lasting HVA channels. In three of five cells from adult animals,
both LVA and HVA currents were found. However, the amplitudes of the
HVA currents were regularly very small, as compared with the amplitudes
of the LVA currents. An example of records in a cell expressing both
types of currents is shown in Fig.
4A. The two components of
currents were separated by a variation of the prepulses from 120 to
60 mV. Depolarizing voltage steps after prepulses to
120 mV evoked
both current components, while the transient component was inactivated
after prepulses to
60 mV. The difference between both records
represents the transient component. Figure 4B shows the mean
peak current-voltage curves for the sum (prepulse to
120 mV), for the
HVA (prepulse to
60 mV), and for the LVA currents (difference) in
four cells. The transient component had an activation threshold of
82.0 ± 5.4 mV and peaked at
43.5 ± 3.9 mV, while the
noninactivating, sustained current component activated at
52.7 ± 6.4 mV and showed a maximum at
12.0 ± 5.1 mV. According to
their voltage dependence, these two types of currents could be
attributed to LVA and to HVA Ca2+ currents,
respectively. The LVA currents gated ~20 mV more negative in divalent
cation-free solutions when compared with Ba2+- or
to Ca2+-containing solutions (Fig. 1,
B and C) due to alterations of the membrane
surface charges. The currents of monovalent cations through
Ca2+ channels reversed between +25 and +30 mV
(Fig. 4B). The true reversal potential of these currents was
not determined since omitting the divalent cations from the bath
solution increased the "leak" conductance of the records, possibly
reflecting the opening of unspecific cation channels that were normally
blocked by divalent cations (not shown). The flow of cations through
these channels would shift the reversal potential of the
Ca2+ channel-mediated currents toward more
negative values. When the Cs+ ions within the
pipette solution were equimolarly replaced by N-methyl-D-glucamine, no shift of the reversal
potential was observed, indicating that the reversal potential was not
influenced by a possible flow of Cs+ ions through
K+ channels.
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The voltage dependences of the steady-state inactivation and activation
of both components of monovalent cation currents are illustrated in
Fig. 4C. For establishing activation curves, the current
amplitudes at different potentials were transformed into conductances
(g) using a reversal potential of the currents of +25 mV
based on the data shown in Fig. 4B. Conductances were then normalized to the maximum conductance and fitted by a Boltzmann relation. The voltages at which half-maximal activation occurred were
63.3 ± 6.3 mV for the transient LVA current and
29.7 ± 5.2 mV for the sustained HVA current. The slopes characterizing the
voltage sensitivity of the channels were comparable for the activation
of the LVA current (4.4 ± 1.8 mV/e-fold change) and of
the HVA current (5.6 ± 1.6 mV/e-fold change). To
record the steady-state inactivation, the peak currents at different
prepotentials (V) were normalized to the peak currents
recorded from V =
120 mV (LVA) and from
V =
80 mV (HVA), respectively (Fig. 4C,
). The fractional currents, normalized to the maximum currents, were fitted with a Boltzmann equation. The voltages at which half-maximal inactivation occurred were
88.4 ± 6.1 mV (LVA) and
39.2 ± 3.5 mV (HVA). The slopes of the inactivation curves were
5.4 ± 1.5 mV for the LVA current and
8.3 ± 3.2 mV for the HVA
current. The "window currents," representing the overlap of the
activation and inactivation curves, were significantly different and
ranged from about
90 to
60 mV in the case of the transient LVA
currents and from about
60 to
20 mV in the case of the sustained
HVA currents. Despite the difficulty to determine the reversal
potential of the currents of monovalent cations through voltage-gated
Ca2+ channels (see preceding text), the voltage
range of LVA current activation and inactivation was well within the
range previously reported for these currents in other cell types
(Lux et al. 1989
, 1990
).
Cells from early postnatal stages had a rather small cell membrane area
and the amplitude of the Ca2+ channel-mediated
currents was small (see following text), indicating that they may
express only few Ca2+ channels in their membranes
assuming that no developmental alterations of the channel conductance
occurred. In some of these cells, it was possible to record
single-channel activity in the whole cell mode. Examples of such
records in cells from P3 and P4 rabbits are shown in Fig.
5A, at different depolarizing
steps from a prepulse to 120 mV. Channel openings as well as
time-dependent inactivation increased at stronger depolarizations.
Figure 5B shows the mean current-voltage relation of
single-channel currents in six cells. The mean slope conductance was
16.2 pS; the extrapolated reversal potential was at about +30 mV.
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Developmental changes of Ca2+ channel-mediated currents
In the developing rabbit retina, proliferative activity is present
up to about P7 (Germer et al. 1997; Reichenbach
et al. 1991
). Thus within the first postnatal week, postmitotic
radial glial cells develop from mitotically active progenitors. This was reflected by an increasing number of cells in the cell suspensions that displayed a unique bipolar morphology, involving a so-called endfoot at the end of one stem process, and a length of >100 µM (Bringmann et al. 1999
). Between P6 and P20, radial
glial cells differentiate into Müller cells as indicated by the
enhanced expression of inwardly rectifying K+
currents, the main membrane current expressed by adult cells (Bringmann et al. 1999
). Figure
6A (left)
illustrates whole-cell records of two typical cells that were found in
the cell suspension derived from a retina of a P2 rabbit. The records
were made in Ca2+-, Mg2+-,
and K+-free bath solution; only
depolarization-induced inwardly directed Na+
currents are shown. While the ganglion cell displayed three different Na+ currents (through fast
Na+ channels and through both transient LVA and
sustained HVA channels), the radial glial cell displayed only currents
through transient LVA Ca2+ channels. The
expression of fast Na+ currents and different
types of Ca2+ currents in early postnatal
ganglion cells was previously described for the rat retina
(Schmid and Guenther 1996
). In the further course of
development, the amplitude of the transient LVA currents in radial
glial cells increased, as shown by the example of the record in a cell
from a P29 rabbit (Fig. 6A, right).
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During the maturation of Müller cells, the cell membrane
capacitance increased (Fig. 6B) (Bringmann et al.
1999). The mean membrane capacitance of the radial glial cells
was small between days 2 and 4 (5.1 ± 2.3 pF, n = 38) and increased to 47.4 ± 12.4 pF after day 30 (n = 36; P < 0.001, ANOVA).
All investigated radial glial/Müller cells from rabbit retinae
between P2 and P37 (n = 647) expressed currents through
transient LVA Ca2+ channels. The amplitude of the
LVA currents was found to increase during the postnatal development.
Figure 6C illustrates the mean amplitudes of the peak LVA
currents in dependence on the postnatal age. Between P2 and P7, the
peak amplitude of the LVA currents increased from 45.3 ± 22.7 pA
at day 2 to 145.9 ± 99.6 pA at day 7 in correlation to both the
postnatal age (r = 0.44, n = 111 cells,
P < 0.001) and to the cell membrane capacitance
(r = 0.64, P < 0.001). After day 6, the peak amplitude of the LVA current remained largely constant
(130.5 ± 86.7 pA, n = 552 cells from P7 to P37),
indicating that the number of LVA Ca2+ channels
per radial glial cell did not significantly change after the first
postnatal week when the channel conductance remained unaltered during
development. On the other hand, the peak density of the LVA currents
was found to be maximal at early postnatal stages (Fig. 6D).
Between days 2 and 5, the mean density was 12.9 ± 7.6 pA/pF
(n = 61); thereafter, the density decreased to a mean of 2.6 ± 1.7 pA/pF after day 18 (n = 163;
P < 0.001). Between days 3 and 19, the peak current
density decreased in correlation to both the postnatal age
(r = 0.55, n = 492, P < 0.001) and the cell membrane capacitance (r =
0.46, P < 0.001). After day 18, the density remained
constant; no correlations were found to the postnatal age or to the
membrane capacitance. In cells from adult animals, the LVA currents
displayed a peak density of 2.0 ± 0.7 pA/pF (n = 6). The postnatal decrease of the LVA current density was not
accompanied by significant changes of the steady-state current
inactivation and activation (not shown) nor by changes of the current
density-voltage curves (Fig. 6E).
Currents through HVA channels were scarcely observed in cells from the first three postnatal weeks. With the exception of two cells from a P12 and a P14 rabbit, all other investigated cells from rabbits up to P23 (n = 524) expressed only LVA currents. After the first three postnatal weeks, a subpopulation of the cells displayed both LVA and HVA currents. In cells that displayed both current types, the density of the HVA currents was always smaller than that of the LVA currents (Fig. 4B). In 118 investigated cells from P24 to P37 rabbits, 25 cells (21%) expressed both current types. In these cells, the peak HVA current displayed a mean density of 0.6 ± 0.5 pA/pF while the peak LVA current had a mean density of 3.3 ± 3.1 pA/pF. An age-dependent increase of the incidence of cells that expressed both current types was not observed between P24 (28%) and P37 (20%).
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DISCUSSION |
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In cultured human Müller cells, using
Ba2+ ions as the charge carrier, the presence of
transient LVA and of sustained HVA currents was previously described
(Puro and Mano 1991; Puro et al. 1996
). Here, we demonstrate that freshly isolated Müller cells of the rabbit retina may express both types of Ca2+
channel-mediated currents with significantly different expression patterns during the postnatal development. The inwardly directed Ba2+ and Ca2+ currents in
cells from young postnatal age activated at potentials positive to
60
mV, peaked at about
25 mV and displayed a transient time-dependent
activation kinetics (Fig. 1). These features are consistent with the
expression of T-type Ca2+ channels. However,
since the peak amplitudes of the Ba2+ and
Ca2+ currents were small, the postnatal
development of Ca2+ channel expression was
investigated using Na+ ions as the main charge
carrier in divalent cation-free external solution. The increased
amplitude of Na+ currents through T-type
Ca2+ channels may be partly explained by the
greater single-channel conductance (16 pS) as compared with
Ba2+ or to Ca2+ currents
through the same channels (5-9 pS) (for a review, see Ertel et
al. 1997
). The omission of the divalent cations from the bath
solution caused a negative shift of the activation kinetics of the
T-type channels by ~20 mV when compared with
Ba2+ currents via alteration of the membrane
surface charges. The transient Na+ currents were
blocked in the low micromolar range by various divalent cations (Fig.
3). The values of half-maximal inactivation of the
Na+ currents through Ca2+
channels by Ca2+ (0.7 µM) and
Mg2+ ions (35.9 µM) are in the range previously
described for neuronal Ca2+ channels (0.7 and 39 µM, respectively) (Carbone et al. 1997
).
The transient Na+ currents described in this
paper were not mediated by fast transient Na+
channels, for the following reasons: although Müller cells from different mammalian species, including man, express fast transient Na+ channels (Chao et al. 1994;
Francke et al. 1996
), rabbit Müller cells were
never shown to express this channel type; currents mediated by fast
transient Na+ channels in human cells displayed a
significantly faster activation and inactivation kinetics as compared
with the transient Na+ currents described in the
present paper; and the Na+ channel-mediated
currents in human cells were largely reduced in their amplitudes by
tetrodotoxin (10 µM) (Francke et al. 1996
) while the
transient Na+ currents in rabbit Müller
cells were insensitive to tetrodotoxin. The inwardly directed transient
Na+ currents in young postnatal rabbit
Müller cells were blocked by low micromolar concentrations of the
T-type channel blocker flunarizine while the L-type channel blocker
nimodipine depressed the current amplitude with an
IC50 value significantly larger than flunarizine
(Fig. 2D). Dihydropyridine-sensitive LVA
Ca2+ channels were previously described to be
present in various neuronal and glial cell preparations (Akaike
et al. 1989
; Akopian et al. 1996
; Koike
et al. 1993
; Takahashi and Akaike 1991
).
From the above-mentioned data, it can be concluded that rabbit retinal
Müller (glial) cells may express both LVA and HVA type
Ca2+ channels. These two types of
Ca2+ channel-mediated currents displayed a
significantly different developmental regulation during the postnatal
differentiation of immature precursor cells into mature Müller
cells, i.e., during the first three postnatal weeks (Bringmann
et al. 1999). While the transient currents were early expressed
at high densities in developing cells, their density fell to a stable
moderate level in the course of further differentiation into mature
Müller cells (Fig. 6D). When the channel conductance
did not change during development, the data indicate that the number of
LVA channels per cell increases within the first postnatal week but
remains constant thereafter (Fig. 6C). As a result, the
density of the LVA currents decreases along with the increasing cell
membrane area after the first postnatal week. While all investigated
rabbit cells from all developmental stages displayed transient LVA
currents, HVA currents were expressed only at later developmental
stages. The different expression patterns probably indicate that the
two currents have different functional roles in Müller cells.
There are two possible functions of the early postnatal expression of
LVA currents at high-density. First, the activity of LVA channels may
be necessary for the precursor proliferation that occurs in the rabbit
retina up to postnatal day 7 (Germer et al. 1997).
According to this idea, the postnatal decrease of the density would
reflect the cessation of retinal proliferation. This assumption is
supported by the fact that proliferation of cultured Müller cells
is dependent on the activity of voltage-gated Ca2+ channels. In cultured Müller cells of
the guinea pig, the proliferation induced by epidermal growth factor
was found to be blocked by flunarizine and by nimodipine, with
flunarizine being the more effective substance (own unpublished
results). Second, LVA channels may have functional roles in early
differentiation processes of retinal radial glial cells. In neuronal
development, T-type Ca2+ channels were implicated
in the regulation of the morphogenesis and of the circuit specification
via mediating spontaneous Ca2+ transients
(Gu and Spitzer 1993
) and autorhythmic oscillatory activity, respectively (Bertolino and Llinas 1992
).
Waves of synchronous bursting activity were described to occur in the
immature retina (Meister et al. 1991
; Wong et al.
1992
; Zhou 1998
), and it may be possible that
immature radial glial cells are involved in this oscillatory electrical
activity, which may enhance the activity of voltage-gated
Ca2+ channels in these cells. LVA channels may
mediate the Ca2+ entry into developing radial
glial cells, an event that, in turn, may trigger intracellular events
necessary for cell differentiation such as for the outgrowth of glial
side branches. Further investigations, however, are necessary to
determine whether the activity of LVA channels contribute to the
morphogenesis of Müller cells. On the other hand, the expression
of both LVA and HVA currents in cells from adult animals indicates that
voltage-gated Ca2+ channels may also have, yet
unknown, functions in mature Müller cells.
The Ca2+ channel-mediated currents develop in
relation to various types of K+ currents in
Müller cells. Figure 7 summarizes
the densities and activities, respectively, of distinct membrane
conductances during the postnatal development from late retinal
progenitor cells into radial glial, and into immature and mature
Müller cells, respectively (for staging of the postnatal radial
glia development, see Bringmann et al. 1999).
Developmental alterations of distinct radial glia membrane conductances
occurred partly in relation to different markers of the retinal
activity; for example, the amplitude of the LVA currents increased up
to about day 6, then, the density of the inwardly rectifying
K+ currents begins to increase, presumably along
with the light-induced ganglion cell activity. The main indicator of
differentiation of immature radial glial cells into mature Müller
cells is the strong up-regulation of the density of inwardly rectifying
K+ currents that occurs between postnatal days 6 and 20 that may mainly underlie the hyperpolarization of the
Müller cell membrane observed during postnatal maturation
(Bringmann et al. 1999
). The hyperpolarization of the
cell membrane may cause a decrease of the opening probability of
depolarization-activated channels. Indeed a developmental decrease of
the activity of Ca2+-activated
K+ channels of big conductance (BK) was
previously described (Bringmann et al. 1999
). It was
suggested that both BK and voltage-gated Ca2+
channels may work together to enhance the Ca2+
entry from the extracellular space after receptor activation, for
example, when both channel types would be co-localized in Müller
cell membranes (Bringmann et al. 2000
). The
stabilization of the membrane potential at hyperpolarized values should
also decrease the activity of LVA Ca2+ channels;
a decreased LVA channel-mediated Ca2+ influx may
contribute to the observed decrease in the activity of BK channels.
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ACKNOWLEDGMENTS |
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This study was supported by grants from the Bundesministerium für Bildung, Forschung und Technologie (BMBF), Interdisciplinary Center for Clinical Research at the University of Leipzig (01KS9504, Project C5) and from the Deutsche Forschungsgemeinschaft (Bonn, Germany; Re 849/8-1).
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FOOTNOTES |
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Address for reprint requests: A. Bringmann, University of Leipzig, Paul Flechsig Institute of Brain Research, Dept. of Neurophysiology, Jahnallee 59, D-04109 Leipzig, Germany (E-mail: bria{at}server3.medizin.uni-leipzig.de).
Received 27 April 2000; accepted in final form 6 September 2000.
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REFERENCES |
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