| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
1 Department of Neurosurgery,, University of Wisconsin Medical School, Madison, Wisconsin 53792; 2 Department of Physiology, University of Wisconsin Medical School, Madison, Wisconsin 53792; 3 Division of Biomedical Science, University of California-Riverside, California 92521
Submitted 30 December 2002; accepted in final form 31 March 2003
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
|---|
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
|---|
). In immature
neurons, NKCC1 maintains an outwardly directed electrochemical gradient for
Cl– that permits GABA-induced membrane depolarization
(Alvarez-Leefmans 2001;
Alvarez-Leefmans et al. 1988;
Schomberg 2003). The excitatory action of GABA is important for the
development of nervous system (Ben Ari et
al. 1997; Yuste and Katz
1991).
In various types of immature neurons, activation of 
-aminobutyric
acid-A (GABAA) receptors initiates a depolarizing efflux of
Cl– that triggers an influx of Ca2+
(Ben Ari et al. 1989). This
Ca2+ then activates processes of maturation and
differentiation (Ben Ari et al.
1997; Marty et al.
1996). GABA also triggers a depolarizing efflux of
Cl– in cultured oligodendrocytes
(Kettenmann et al. 1991).
Re-accumulation of intracellular Cl– after depletion is
blocked by removal of extracellular Na+ or inhibition of NKCC1
activity (Hoppe and Kettenmann
1989a,b).
Thus NKCC1 may be important in maintaining the high
[Cl–]i in oligodendrocytes. While the trophic
effect of GABA on neurons is now well established
(Fukuda et al. 1998;
Ikeda et al. 1997;
Marty et al. 1996), the effect
of GABA on oligodendrocyte survival and growth remains uncertain.
Here we report that NKCC1 is abundantly expressed in oligodendrocytes and that its activity is regulated by GABAA receptors. GABA-mediated oligodendrocyte survival depends on NKCC1 activity.
|
METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
|---|
Eagle's balanced salt solution (EBSS), Leibovitz's L-15 medium (L-15 medium), and Dulbecco's modified Eagle's medium (DMEM) were from Gibco (Grand Island, NY). Deoxyribonuclease I (DNase), poly-L-ornithine, platelet-derived growth factor (PDGF), and N1 medium were from Sigma (St. Louis, MO). Mouse anti-2',3'-cyclic nucleotide 3'-phosphodiesterase monoclonal antibody (CNPase) was from Chemi-Con International (Temecula, CA). N-glycanase was from Glyko (Novato, CA). Fura-2-acetoxymethyl ester (AM) was from Molecular Probes (Eugene, OR). Rip monoclonal antibody was from Developmental Studies Hybridoma Bank (Iowa City, IA).
Enriched oligodendrocyte culture
Spinal cords were removed from 7- to 10-day-old rat pups (Sprague-Dawley)
and transferred to a 60-mm dish containing ice-cold L-15 medium. Under a
dissecting microscope, the meninges and nerve roots were removed. The spinal
cords were cut into 1-mm3 pieces and incubated in EBSS containing
0.25 mg/ml trypsin and 50 µg/ml DNase at 37°C for 30 min with shaking.
The enzymatic activity of trypsin was stopped by adding fetal bovine serum
(FBS). After centrifugation, the tissues were resuspended with L-15 medium
(10% FBS and 50 µg/ml DNase) and triturated. Cell suspension was
centrifuged at 30,000 g at 4°C in 29% Percoll as described before
(Grever et al. 1999). The cell
pellet was suspended in DMEM supplemented with N1 medium, 10 ng/ml biotin,
0.5% FBS, and 10 ng/ml PDGF. In each well, 2 x 104 (96-well
plate), 6 x 104 (24-well plate), and 12 x
104 (6-well plate) cells were plated. The plates were coated with
poly-L-ornithine (0.1 mg/ml). Cultures were maintained in 5%
CO2 atmosphere at 37°C and refed every 2 days with DMEM
containing freshly prepared PDGF. More than ninety percent of cells in culture
were oligodendrocytes, analyzed with immuno-flow cytometry. All animal
procedures were conducted in strict compliance with the NIH Guide for the Care
and Use of Laboratory Animals and approved by the University of Wisconsin
Center for Health Sciences Research Animal Care Committee.
Immunofluorescence staining
Cultured cells grown on poly-L-ornithine-coated chamber slides
were rinsed with PBS (pH 7.4) and fixed with 4% paraformaldehyde in PBS for 40
min at room temperature. After rinsing, cells were incubated with blocking
solution (10% goat serum, 0.4% Triton X-100, and 1% bovine serum albumin in
PBS) for 1 h. Cells were incubated with a primary antibody in blocking
solution overnight at 4°C. After rinsing, cells were incubated with
fluorescein isothiocyanate (FITC)-conjugated or Texas red conjugated secondary
antibodies for 1 h. The images were captured by a laser-scanning confocal
microscope (Bio-Rad MRC 1000, Bio-Rad, Hercules, CA), as described before
(Su et al. 2000).
Gel electrophoresis and immunoblotting
Cultured oligodendrocytes on six-well plates were washed with ice-cold PBS
(pH 7.4) containing 2 mM EDTA and protease inhibitors as described previously
(Sun and Murali 1999). Cells
were scraped from the plate and lysed in PBS by 30 s of sonication at 4°C
by an ultrasonic processor (Sonics and Materials, Danbury, CT). Rat spinal
cords at different ages were isolated and tissue homogenate was prepared as
described before (Yan et al.
2001). Samples were denatured in SDS reducing buffer (1:2 by
volume, Bio-Rad) and heated at 37°C for 15 min before gel electrophoresis.
The sample and prestained molecular mass markers (Bio-Rad) were separated
electrophoretically on 10% gel (Laemmli
1970) or gradient gels (4–15%, Bio-Rad). The resolved
proteins were transferred electrophoretically to a polyvinylidene difluoride
(PVDF) membrane (0.45 µm; Millipore, Bedford, MA). The blots were incubated
in 7.5% nonfat dry milk in Tris-buffered saline (TBS) for 2 h at room
temperature and incubated overnight with a primary antibody. The blots were
then rinsed five times with TBS and incubated with horseradish peroxidase
conjugated secondary IgG for 1 h. After washing, bound antibody was visualized
using the enhanced chemiluminescence assay (ECL, Amersham Pharmacia Biotech,
Piscataway, NJ). NKCC1 was detected with monoclonal antibody T4
(Lytle et al. 1995).
Anti-myelin basic protein (MBP) and anti-
III tubulin monoclonal
antibodies were used to detect MBP and III tubulin, respectively. To
obtain quantitative analysis of expression of the proteins, the blots on a
film after ECL reaction were scanned using a Hewlett-Packard Scanjet (4c/t)
scanner. The intensity of each band was measured by UN-SCAN-It gel software
(Silk Scientific, Orem, UT). Western blot analysis using T4 antibody was
carried out with 10- to 15-µg protein loads and ECL detection (20-s
exposure time). We determined previously
(Yan et al. 2001) that the T4
signal is proportional to protein with loads between 15 and 45 µg.
For deglycosylation studies, crude cell lysate proteins (50 µg) were solubilized with 0.1% SDS, heated at 100°C for 5 min, incubated with 0.5 units of N-glycanase F for 2 h at 37°C, and separated by SDS-PAGE as described above.
Assay for NKCC1 activity
NKCC1 activity was measured as bumetanide-sensitive 86Rb influx,
using 86Rb as a tracer of K+
(Sun and Murali 1999).
Cultured oligodendrocytes on 4–5 days in vitro (DIV 4–5) were
equilibrated for 10–30 min at room temperature with an isotonic
HEPES-buffered MEM (290 mOsm). The concentrations of components in HEPES-MEM
were described before (Su et al.
2000). Cells were incubated for 10 min in HEPES-MEM with or
without 10 µM bumetanide. The cells were exposed to 2 µCi/ml of
86Rb in HEPES-MEM for 3 min at room temperature, in either the
presence or the absence of 10 µM bumetanide. 86Rb influx was
terminated by rinsing cells with ice-cold 0.1 M MgCl2. Cells were
extracted in 1% SDS, and radioactivity in cell lysates was analyzed by liquid
scintillation counting (1900CA, Packard, Downers Grove, IL). 86Rb
influx rate was calculated as the slope of 86Rb uptake over time
and expressed in nanomoles of 86Rb per milligram of protein per
minute. Six determinations were obtained in each experiment throughout the
study, and protein content was measured in each sample by the bicinchoninic
acid method (Smith et al.
1985).
Intracellular Cl– content measurement
Cells on 96-well plates were preincubated in HEPES-MEM containing
36Cl (0.4 µCi/ml) for 30 min at room temperature. The cells were
then incubated in HEPES-MEM containing 36Cl (0.4 µCi/ml) in the
presence of either bumetanide (10 µM), muscimol (10 µM), muscimol plus
bicuculline (10 µM), or muscimol plus bumetanide for 20 min. Intracellular
36Cl content measurement was terminated by two washes with a
washing buffer, as described previously
(Su et al. 2002).
Radioactivity of cellular extract in 1% SDS was analyzed by liquid
scintillation counting. In each experiment, specific activities
(counts/µmol/min) of 36Cl were determined for each assay
condition and used to calculate intracellular Cl– content
(µmol/mg protein).
Measurement of relative cell volume changes in a single cell
Relative cell volume changes were estimated using video-enhanced
differential interference contrast (DIC) microscopy, as described in our
previous study (Su et al.
2002). Oligodendrocytes cultured on
poly-L-ornithine-coated coverslips were placed in an open-bath
imaging chamber (Warner Instruments, Hamen, CT; bath volume 40 µl) on the
stage of a Nikon TE 300 inverted epifluorescence microscope. Cells were
equilibrated with an isotonic HEPES-MEM (312 mOsm) for 15 min at room
temperature (Su et al. 2002).
Cells were exposed sequentially to isotonic HEPES-MEM, HEPES-MEM plus muscimol
(10 µM), muscimol plus bicuculline (10 µM), and hypertonic HEPES-MEM
(480 mOsm). Between each treatment, cells were reequilibrated with isotonic
HEPES-MEM. Cells were visualized using a Nikon 60x Plan Apo oil
immersion objective lens and cell images were recorded every minute as
described previously (Su et al.
2002). The mean cross-sectional area (CSA) was calculated after
tracing the perimeter of the cell body with MetaMorph image-processing
software (Universal Imaging, Downingtown, PA).
The control CSA values were obtained when cells were exposed to isotonic HEPES-MEM only. Relative changes of mean cross-sectional area (CSAr) were calculated as experimental CSA/control CSA. Hypertonic buffer of 480 mOsm was prepared by holding the salt concentrations constant and adding 165 mM sucrose.
This approach to estimate cell volume changes can be criticized because it does not consider the cell volume changes in the z-axis. Thus we may underestimate the actual cell volume changes.
Cell survival assay
Calcein-AM and propidium iodide were used to determine survival of
oligodendrocytes. Cells were grown in DMEM with supplements (N1, Biotin, PDGF)
for 2–3 days. Cells were then cultured for 18 h in either DMEM plus
supplements, DMEM without supplements, DMEM plus 30 µM muscimol, DMEM plus
30 µM muscimol and 10 µM bicuculline, or DMEM plus 30 µM muscimol and
10 µM bumetanide, respectively. To assay the effect of 25 mM
[K+]o on survival of oligodendrocytes, cells were
cultured in DMEM plus 25 mM KCl, DMEM plus 25 mM KCl, and 1 µM nifedipine
for 18 h. Nifedipine is stable in culture medium for 
30 h
(Franklin et al. 1995).
Muscimol and bicuculline are stable in cultures for 6 days
(Marty et al. 1996). Therefore
media containing the drugs were not changed during the 18 h incubation. At the
end of 18 h incubation, cells were incubated in 1 µg/ml calcein-AM and 10
µg/ml propidium iodide in DMEM at 37°C for 35 min. After rinsing, cells
were counted using the inverted fluorescent microscope (Nikon TE 300). More
than 1000 cells were counted in each condition in a blind manner. Cell
mortality was calculated as the ratio of propidium iodide positive cells to
the sum of calcein-AM- and propidium iodide positive cells.
Measurement of changes in intracellular Ca2+
Cultured oligodendrocytes grown on polyLornithine-coated
cover-slips were loaded with 10 µM fura-2-AM in HEPES-MEM at room
temperature for 30 min. The coverslips were placed in the open bath imaging
chamber containing HEPES-MEM at room temperature. The chamber was mounted on
the stage of the Nikon TE 300 inverted epifluorescence microscope.
Oligodendrocytes were excited every 10 s at 340 and 380 nm and the emission
fluorescence at 510 nm was recorded. The ratio of 340/380 was used to indicate
the [Ca 2+]i as described before
(Grynkiewicz et al. 1985;
Su et al. 2000).
Images were collected and analyzed with MetaFluor image-processing
software.
Statistics
Throughout the study, statistical significance was determined by analysis of variance (ANOVA; Bonferroni/Dunn) at a confidence of 95% (P < 0.05).
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
|---|
As shown in Fig. 1, A and
D, oligodendrocytes in primary cultures were identified
by immunocytochemical detection of the oligodendrocyte-specific proteins Rip
and CNP (Jhaveri et al. 1992).
Oligodendrocytes in DIV 2–3 culture exhibited a "spider's
web"-like morphology with many slender and branched processes
(Fig. 1, A–G,
H). NKCC1 was detected in the cell body and processed with
polyclonal antibody NT antiserum against the N-terminus of human NKCC1
(Fig. 1, B and
E) and monoclonal antibody T4
(Fig. 1G),
respectively. Colocalization of Rip and NKCC1 or CNP and NKCC1 was found in
oligodendrocytes (Fig. 1, C and
F).
|
Development-dependent expression of NKCC1 in oligodendrocytes from rat spinal cord
Expression of NKCC1 in spinal cords and cultured oligodendrocytes was also
evaluated by immunoblotting. As shown in
Fig. 2A, expression of
NKCC1 in rat spinal cord varied over the course of postnatal development. In
the rat spinal cord, NKCC1 was low at P1 yet progressively increased from P6
through adulthood (Fig. 2, A and
C). A development-dependent pattern of expression was
found for MBP. In contrast, levels of -tubulin did not change
significantly from P1 to adulthood (Fig. 2,
A and C). NKCC1 was also detected in cultured
oligodendrocytes (Fig.
2B). Deglycosylation of the native 146-kDa NKCC1 protein
in oligodendrocytes with N-glycanase resulted in a core protein of
approximately 135 kDa.
|
GABAA receptor activation stimulates NKCC1 activity
In cultured oligodendrocytes (DIV 4–5) under control conditions, NKCC1 activity (bumetanide-sensitive 86Rb influx rate) was 9.1 ± 1.8 nmol/mg protein/min (Fig. 3A). This component constituted 39.3% of the total 86Rb influx and required the simultaneous presence of extracellular Na+ (Fig. 3A) or Cl– (data not shown). Activation of the GABAA receptor with muscimol stimulated NKCC1 activity approximately 50% (Fig. 3B). This stimulation was blocked by the GABAA receptor antagonist bicuculline.
|
GABAA receptor activation modulates intracellular 36Cl content in oligodendrocytes
As shown in Fig. 4A, cells were preequilibrated in HEPES-MEM with 36Cl (0.4 µCi/ml) for 0–50 min. A steady-state level of intracellular 36Cl was obtained by a 30-min incubation and maintained during the 50-min equilibration. Thus in the rest of the study, a 30-min preincubation was performed. After a 30-min equilibration with 36Cl, changes of intracellular Cl– content were measured when cells were exposed to HEPES-MEM containing 36Cl (0.4µCi/ml) with or without 30 µM muscimol for 20 min. Activation of the GABAA receptor with muscimol reduced intracellular Cl– content from 0.70 ± 0.05 to 0.48 ± 0.03 µmol/mg protein (P < 0.05, Fig. 4B). This response was blocked by bicuculline (Fig. 4B). Inhibition of NKCC1 activity with bumetanide had no significant effect on resting levels of Cl– content or on loss of Cl– evoked by muscimol (Fig. 4B).
|
Activation of GABAA receptors leads to oligodendrocyte shrinkage
We determined whether activation of the GABAA receptor causes shrinkage in oligodendrocytes. Figure 5, A and B, illustrates that muscimol caused CSAr to decrease in oligodendrocytes gradually over 10 min (0.95 ± 0.01; P < 0.05). This decrease is reversible and CSAr recovered to the basal levels in isotonic HEPES-MEM. In contrast, in the presence of both muscimol and the GABAA receptor inhibitor bicuculline, no significant changes in CSAr were found (1.00 ± 0.01, P > 0.05). The same oligodendrocytes remained responsive to a subsequent hypertonic challenge (0.91 ± 0.01, P < 0.05) and then returned to the basal cell volume levels when they were returned to isotonic HEPES-MEM (1.00 ± 0.01).
|
GABAA receptor activation modulates oligodendrocyte [Ca2+]i
Intracellular free Ca2+ ([Ca2+]i) was monitored using the Ca2+-sensitive dye fura-2. Muscimol evoked a transient increase in [Ca2+]i (Fig. 6A). The increases in intracellular Ca2+ in the first and second exposures of 100 µM muscimol were similar (0.106 ± 0.006 and 0.100 ± 0.003, respectively, Fig. 6A). This muscimol-induced response was blocked not only by the GABAA receptor antagonist bicuculline, but also by the NKCC1 inhibitor bumetanide (Fig. 6, B and C). The average change of intracellular Ca2+ induced by 100 µM muscimol was 0.106 ± 0.006. In the presence of bicuculline or bumetanide, the average change was 0.008 ± 0.005 or 0.006 ± 0.008, respectively (Fig. 6D, P < 0.05).
|
GABAA receptors affect oligodendrocyte survival
Activation of GABAA receptors in immature neurons leads to
Ca2+ influx and this is thought to play an important
role in cell maturation during development
(Ben Ari et al. 1997;
LoTurco et al. 1995). Whether
GABA-mediated Ca2+ influx has a similar trophic effect
on oligodendrocytes remains unknown. Survival of oligodendrocytes in vitro
depends on the presence of growth factors
(Noble et al. 1988;
Raff et al. 1988). Consistent
with these early reports, we found that removal of PDGF and other supplements
led to cell death (Fig. 7). In
contrast, activation of GABAA receptors with muscimol significantly
reduced mortality following removal of growth factors. Whereas bumetanide did
not affect basal cell death (22.8 ± 1.3%) or cell death induced by
removal of PDGF and other supplements (Fig.
7), it prevented the effect of muscimol. Thus the improved
survival of oligodendrocytes with the GABAA receptor activation
appears to require NKCC1 activation.
|
High [K+]o-mediated cell survival
In neurons, the mechanism by which GABAA receptor activation
improves survival is believed to involve Ca2+ entry
through depolarization-activated Ca2+ channels. Membrane
depolarization produced by raising extracellular potassium
([K+]o) is known to have a trophic effect in neurons
(Franklin et al. 1995). This
maneuver had a similar effect in oligodendrocytes
(Fig. 8). Raising
[K+]o to 25 mM prevented cell death induced by removal
of PDGF and other supplements, but not in the presence of the L-type
Ca2+ channel antagonist nifedipine (1 µM). Neither
nifedipine nor bicuculline had any effects on the basal levels of cell
mortality in DMEM-containing supplements
(Fig. 8). Cell mortality was
not affected when NKCC1 activity was blocked by bumetanide in the presence of
25 mM [K+]o (Fig.
8). Measurements of intracellular Ca2+
revealed that 25 mM [K+]o increased
[Ca2+]i through a mechanism blocked by
nifedipine but not by bumetanide (Fig. 9,
A–C). Taken together, our data suggest that a rise
of intracellular Ca2+ mediated by either muscimol or
elevated [K+]o plays an important role in
oligodendrocyte survival.
|
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
|---|
), GABA may boost
NKCC1 activity following Cl– efflux, a reduction in
[Cl–]i, and subsequent cell shrinkage. In accord
with this hypothesis, activating GABAA receptors brought about a
decrease in oligodendrocyte Cl– content and cell
shrinkage. However, concurrent inhibition of NKCC1-mediated Cl– influx and activation of GABA-induced Cl– efflux did not cause additive effects on a decrease in [Cl–]i. The mechanisms underlying this observation are not clear. This may suggest that other Cl– transport mechanisms (such as Na+-dependent and independent Cl–/HCO3– transporters and/or Cl– channels) may conduct Cl– influx and prevent a further loss of intracellular Cl–.
GABAergic neurons develop earlier than glutamatergic neurons and GABA
induces depolarization in immature neurons
(Kaila 1994). This
GABA-induced depolarization elevates [Ca2+]i
via activation of voltage-dependent Ca2+ channels and
contributes to several aspects of the CNS development, including gene
expression (Bading et al. 1993;
Vaccarino et al. 1992) and
neuronal growth and differentiation (Ben
Ari et al. 1997; LoTurco et
al. 1995). Our study suggests that GABA-mediated
Ca2+-influx likewise exerts a trophic effect on
oligodendrocytes. We found that activation of GABAA receptors
triggered a rise in intracellular Ca2+ through L-type
voltage-gated Ca2+ channels and promoted oligodendrocyte
survival following withdrawal of PDGF and other supplements. Inhibition of
NKCC1 blocked this trophic effect. These data further support the view that
NKCC1 activity is required for GABAA receptor function in
oligodendrocytes.
A functional link between the GABAA receptor and L-type
voltage-gated Ca2+ channels has been suggested in
oligodendrocytes (Kirchhoff and Kettenmann
1992). We found that activating these channels by membrane
depolarization (high [K+]o) also promotes survival of
oligodendrocytes. These findings are consistent with high
[K+]o-mediated trophic effects in developing neurons
(Collins and Lile 1989;
Franklin et al. 1995;
Koike et al. 1989).
In the current study, 30–100 µM muscimol was used. We believe that
the effects mediated by 30–100 µM muscimol are physiologically
relevant. The in vivo ambient concentration of GABA in the extracellular space
is 0.8–2.9 µM, which is sufficient to activate GABAA
receptors (Lerma et al. 1986).
GABA in the GABAergic synaptic cleft can reach 0.5–1.0 mM
(Maconochie et al. 1994). GABA
released from neuronal growth cone or neurites could increase the local GABA
concentration above the ambient concentration range and mediate the
coordination of neuron and glial interaction in vivo. It has been hypothesized
that activation of nearby glial GABA receptors could be the means for
oligodendrocyte precursors to detect migrating neurites. Thus GABA may play a
role in oligodendrocyte maturation
(Kettenmann et al. 1991). Our
findings imply that GABA release from the nearby neurons can exert trophic
effects on oligodendrocytes by activating GABAA receptor-mediated
Cl– efflux and triggering opening of voltage-dependent
Ca2+ channels. However, these actions require the
function of NKCC1, which maintains the intracellular Cl–
above the Cl– equilibrium potential, and enable GABA to cause
membrane depolarization. Therefore NKCC1 may play an important role in
oligodendrocyte development.
The GABAA receptor is a heteropentamer drawn from a repertoire
of 
1–6, 1–3, 1–3, 
, 
, 
,
, and 
1–3 subunits in the CNS
(Lambert et al. 2001). Various
subunit combinations influence the physiological and pharmacological
properties of the receptor. Currently, the subunit combination of
GABAA receptor in oligodendrocytes is not clear. The
pharmacological studies of the GABAA receptor in oligodendrocytes
indicate that GABA activates a Cl– conductance in a
dose-dependent manner (Kettenmann et al.
1991). The GABA response in oligodendrocyte precursor cells is
blocked by picrotoxin and bicuculline. Pentobarbital and flunitrazepam
increase the GABA-induced currents but -carbolines act as inverse
benzodiazepine agonists (Kettenmann et al.
1991).
In summary, our results demonstrate that NKCC1 is expressed at relatively high levels in oligodendrocytes where it helps maintain [Cl–]i above electrochemical equilibrium in opposition to GABA-induced Cl– efflux. Our data also suggest that NKCC1 activity may be important in oligodendrocyte development in association with GABAA receptor function.
|
DISCLOSURES |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
|---|
|
FOOTNOTES |
|---|
Address for reprint requests: D. Sun, Department of Neurological Surgery, University of Wisconsin Medical School, H4/332 Clinical Science Center, 600 Highland Ave., Madison, WI 53792 (E-mail: sun{at}neurosurg.wisc.edu).
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
|---|
Alvarez-Leefmans FJ, Gamino SM, Giraldez F, and Nogueron I.
Intracellular chloride regulation in amphibian dorsal root ganglion neurones
studied with ion-selective microelectrodes. J Physiol
406: 225–246,
1988.
Bading H, Ginty
DD, and Greenberg ME. Regulation of gene expression in hippocampal neurons
by distinct calcium signaling pathways. Science
260: 181–186,
1993.
Ben Ari Y, Cherubini E, Aniksztejn L, Roisin MP, and Charriaut-Marlangue C. Mechanism of induction of long term potentiation by the mast cell degranulating peptide. Pharmacopsychiatry 22(Suppl. 2): 107–110, 1989.
Ben Ari Y, Khazipov R, Leinekugel X, Caillard O, and Gaiarsa JL. GABAA, NMDA and AMPA receptors: a developmentally regulated `menage a trois'. Trends Neurosci 20: 523–529, 1997.[ISI][Medline]
Collins F and Lile JD. The role of dihydropyridine-sensitive voltage-gated calcium channels in potassium-mediated neuronal survival. Brain Res 502: 99–108, 1989.[ISI][Medline]
Franklin JL, Sanz-Rodriguez C, Juhasz A, Deckwerth TL, and Johnson EM Jr. Chronic depolarization prevents programmed death of sympathetic neurons in vitro but does not support growth: requirement for Ca2+ influx but not Trk activation. J Neurosci 15: 643–664, 1995.[Abstract]
Fukuda A,
Muramatsu K, Okabe A, Shimano Y, Hida H, Fujimoto I, and Nishino H.
Changes in intracellular Ca2+ induced by
GABAA receptor activation and reduction in Cl–
gradient in neonatal rat neocortex. J Neurophysiol
79: 439–446,
1998.
Grever WE, Zhang S, Ge B, and Duncan ID. Fractionation and enrichment of oligodendrocytes from developing human brain. J Neurosci Res 57: 304–314, 1999.[ISI][Medline]
Grynkiewicz G,
Poenie M, and Tsien RY. A new generation of Ca2+
indicators with greatly improved fluorescence properties. J Biol
Chem 260:
3440–3450, 1985.
Hoppe D and Kettenmann H. Carrier-mediated Cl– transport in cultured mouse oligodendrocytes. J Neurosci Res 23: 467–475, 1989a.[ISI][Medline]
Hoppe D and Kettenmann H. GABA triggers a Cl– efflux from cultured mouse oligodendrocytes. Neurosci Lett 97: 334–339, 1989b.[ISI][Medline]
Ikeda Y, Nishiyama N, Saito H, and Katsuki H. Furosemide-sensitive calcium rise induced by GABAA-receptor stimulation in cultures of embryonic rat striatal neurons. Jpn J Pharmacol 74: 165–169, 1997.[Medline]
Jhaveri S, Erzurumlu RS, Friedman B, and Schneider GE. Oligodendrocytes and myelin formation along the optic tract of the developing hamster: an immunohistochemical study using the Rip antibody. Glia 6: 138–148, 1992.[ISI][Medline]
Kaila K. Ionic basis of GABAA receptor channel function in the nervous system. Prog Neurobiol 42: 489–537, 1994.[ISI][Medline]
Kettenmann H, Blankenfeld GV, and Trotter J. Physiological properties of oligodendrocytes during development. Ann NY Acad Sci 633: 64–77, 1991.[Abstract]
Kirchhoff F and Kettenmann H. GABA triggers a [Ca2+]i increase in murine precursor cells of the oligodendrocyte lineage. Eur J Neurosci 4: 1049–1058, 1992.[ISI][Medline]
Koike T, Martin
DP, and Johnson EM Jr. Role of Ca2+ channels in the
ability of membrane depolarization to prevent neuronal death induced by
trophic-factor deprivation: evidence that levels of internal
Ca2+ determine nerve growth factor dependence of
sympathetic ganglion cells. Proc Natl Acad Sci USA
86: 6421–6425,
1989.
Laemmli UK. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227: 680–685, 1970.[Medline]
Lambert JJ, Belelli D, Harney SC, Peters JA, and Frenguelli BG. Modulation of native and recombinant GABAA receptors by endogenous and synthetic neuroactive steroids. Brain Res Rev 37: 68–80, 2001.[Medline]
Lerma J, Herranz AS, Herreras O, Abraira V, and Martin R. In vivo determination of extracellular concentration of amino acids in the rat hippocampus. A method based on brain dialysis and computerized analysis. Brain Res 384: 145–155, 1986.[ISI][Medline]
LoTurco JJ, Owens DF, Heath MJ, Davis MB, and Kriegstein AR. GABA and glutamate depolarize cortical progenitor cells and inhibit DNA synthesis. Neuron 15: 1287–1298, 1995.[ISI][Medline]
Lytle C.
Activation of the avian erythrocyte Na-K-Cl cotransporter protein by cell
shrinkage, cAMP, fluoride, and calyculin-A involves phosphorylation at common
sites. J Biol Chem 272:
15069–15077, 1997.
Lytle C, Xu JC, Biemesderfer D, and Forbush B III. Distribution and diversity of Na-K-Cl cotransport proteins: a study with monoclonal antibodies. Am J Physiol 269: C1496–C1505, 1995.
Maconochie DJ, Zempel JM, and Steinbach JH. How quickly can GABAA receptors open? Neuron 12: 61–71, 1994.[ISI][Medline]
Marty S, Berninger B, Carroll P, and Thoenen H. GABAergic stimulation regulates the phenotype of hippocampal interneurons through the regulation of brain-derived neurotrophic factor. Neuron 16: 565–570, 1996.[ISI][Medline]
Noble M, Murray K, Stroobant P, Waterfield MD, and Riddle P. Platelet-derived growth factor promotes division and motility and inhibits premature differentiation of the oligodendrocyte/type-2 astrocyte progenitor cell. Nature 333: 560–562, 1988.[Medline]
Raff MC, Lillien LE, Richardson WD, Burne JF, and Noble MD. Platelet-derived growth factor from astrocytes drives the clock that times oligodendrocyte development in culture. Nature 333: 562–565, 1988.[Medline]
Russell JM.
Sodium-potassium-chloride cotransport. Physiol Rev
80: 211–276,
2000.
Schomberg SL,
Bauer J, Kintner DB, Su G, Flemmer A, Forbush B, and Sun D.
Cross-talk between the GABAA receptor and the
Na+-K+-Cl– cotransporter is mediated by
intracellular Cl–. J Neurophysiol
89: 159–167,
2003.
Smith PK, Krohn RI, Hermanson GT, Mallia AK, Gartner FH, Provenzano MD, Fujimoto EK, Goeke NM, Olson BJ, and Klenk DC. Measurement of protein using bicinchoninic acid. Anal Biochem 150: 76–85, 1985.[ISI][Medline]
Su G, Haworth
RA, Dempsey RJ, and Sun D. Regulation of
Na+-K+-Cl– cotransporter in primary
astrocytes by dibutyryl cAMP and high [K+]o.
Am J Physiol Cell Physiol 279:
C1710–C1721, 2000.
Su G, Kintner D, Flagella M, Shull GE, and Sun D. Cortical astrocytes from NA-K-Cl cotransporter null mice exhibit an absence of high [K+]o-induced swelling and a decrease in EAA release. Am J Physiol 282: C1147–C1160, 2002.
Sun D and
Murali SG. Na+ -K+-2Cl+ cotransporter in
immature cortical neurons: a role in intracellular Cl+ regulation.
J Neurophysiol 81:
1939–1948, 1999.
Vaccarino FM, Hayward MD, Nestler EJ, Duman RS, and Tallman JF. Differential induction of immediate early genes by excitatory amino acid receptor types in primary cultures of cortical and striatal neurons. Brain Res Mol Brain Res 12: 233–241, 1992.[Medline]
Yan YP, Dempsey RJ, and Sun D. Expression of Na+-K+-Cl+ cotransporter in rat brain during development and its localization in mature astrocytes. Brain Res 911: 43–55, 2001.[ISI][Medline]
Yuste R and Katz LC. Control of postsynaptic Ca2+ influx in developing neocortex by excitatory and inhibitory neurotransmitters. Neuron 6: 333–344, 1991.[ISI][Medline]
This article has been cited by other articles:
|
|
 |
|
  H. I. Rocha-Gonzalez, S. Mao, and F. J. Alvarez-Leefmans Na+,K+,2Cl- Cotransport and Intracellular Chloride Regulation in Rat Primary Sensory Neurons: Thermodynamic and Kinetic Aspects J Neurophysiol, July 1, 2008; 100(1): 169 - 184. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S. F. Pedersen, M. E. O'Donnell, S. E. Anderson, and P. M. Cala Physiology and pathophysiology of Na+/H+ exchange and Na+-K+-2Cl- cotransport in the heart, brain, and blood Am J Physiol Regulatory Integrative Comp Physiol, July 1, 2006; 291(1): R1 - R25. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Wouters, A. D. Laet, L. V. Donck, E. Delpire, P.-P. van Bogaert, J.-P. Timmermans, A. de Kerchove d'Exaerde, K. Smans, and J.-M. Vanderwinden Subtractive hybridization unravels a role for the ion cotransporter NKCC1 in the murine intestinal pacemaker Am J Physiol Gastrointest Liver Physiol, June 1, 2006; 290(6): G1219 - G1227. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 B. Lenart, D. B. Kintner, G. E. Shull, and D. Sun Na-K-Cl Cotransporter-Mediated Intracellular Na+ Accumulation Affects Ca2+ Signaling in Astrocytes in an In Vitro Ischemic Model J. Neurosci., October 27, 2004; 24(43): 9585 - 9597. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Visit Other APS Journals Online |
