Department of Neurobiology, University of Alabama at Birmingham, Birmingham, Alabama 35294
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ABSTRACT |
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Bordey, A., S. A. Lyons, J. J. Hablitz, and H. Sontheimer. Electrophysiological Characteristics of Reactive Astrocytes in Experimental Cortical Dysplasia. J. Neurophysiol. 85: 1719-1731, 2001. Neocortical freeze lesions have been widely used to study neuronal mechanisms underlying hyperexcitability in dysplastic cortex. Comparatively little attention has been given to biophysical changes in the surrounding astrocytes that show profound morphological and biochemical alterations, often referred to as reactive gliosis. Astrocytes are thought to aid normal neuronal function by buffering extracellular K+. Compromised astrocytic K+ buffering has been proposed to contribute to neuronal dysfunction. Astrocytic K+ buffering is mediated, partially, by the activity of inwardly rectifying K+ channels (KIR) and may involve intracellular redistribution of K+ through gap-junctions. We characterized K+ channel expression and gap-junction coupling between astrocytes in freeze-lesion-induced dysplastic neocortex. Whole cell patch-clamp recordings were obtained from astrocytes in slices from postnatal day (P) 16-P24 rats that had received a freeze-lesion on P1. A marked increase in glial fibrillary acidic protein immunoreactivity was observed along the entire length of the freeze lesion. Clusters of proliferative (bromo-deoxyuridine nuclear staining, BrdU+) astrocytes were seen near the depth of the microsulcus. Astrocytes in cortical layer I surrounding the lesion were characterized by a significant reduction in KIR. BrdU-positive astrocytes near the depth of the microsulcus showed essentially no expression of KIR channels but markedly enhanced expression of delayed rectifier K+ (KDR) channels. These proliferative cells showed virtually no dye coupling, whereas astrocytes in the hyperexcitable zone adjacent to the microsulcus displayed prominent dye-coupling as well as large KIR and outward K+ currents. These findings suggest that reactive gliosis is accompanied by a loss of KIR currents and reduced gap junction coupling, which in turn suggests a compromised K+ buffering capacity.
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INTRODUCTION |
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One of the
functions attributed to astrocytes is the buffering of extracellular
[K+], which appears to be of critical
importance to assure normal neuronal excitability (D'Ambrosio
et al. 1999; Janigro et al. 1997
; Newman
1995
). It has been demonstrated that astrocytes are liberally
endowed with voltage-activated K+ channels
(Barres et al. 1990
; Bevan et al. 1985
;
Sontheimer 1994
), which mediate the diffusional uptake
of K+. Inwardly rectifying
K+ (KIR)
channels in particular play an important role in
K+ buffering by astrocytes (D'Ambrosio et
al. 1999
; Ransom and Sontheimer 1995
) and
retinal glial cells (Newman 1993
; Newman et al.
1984
). It is likely that intracellular redistribution of
K+ (spatial buffering) is further facilitated by
intracellular coupling of astrocytes through gap junctions
(Giaume et al. 1991
; Mobbs et al. 1988
;
Yamamoto et al. 1990
). The expression of
voltage-activated K+ channels changes during
early postnatal development (Bordey and Sontheimer 1997
;
Steinhauser et al. 1992
) in a way that suggests that
K+ buffering is not fully established at birth
but develops in the first 3-4 wk of postnatal life. During this
developmental time period, neuronal activity causes much larger
K+ fluctuations than in the adult (Ransom
et al. 1986
; Sykova 1983
; Sykova et al.
1992
), again suggesting that K+ buffering
by astrocytes develops as these cells mature. Interestingly, in
cultured astrocytes, this maturation, where expression of
KIR channels is a hallmark, can be
reversed by injury (MacFarlane and Sontheimer 1997
).
Mechanical injury to astrocytes leads to a dedifferentiation and
proliferation of astrocytes at injury sites. This process is
accompanied by a loss of KIR channels.
The astrocytic response to injury of the CNS is often referred to as
"reactive gliosis." Although not fully understood, this stereotypic, mainly astrocytic, response to injury is characterized by
an initial dedifferentiation of astrocytes and enhanced cell proliferation, ultimately leading to a dense astrocytic scar. Astrocytes hypertrophy and show marked upregulation of the intermediate filament protein glial fibrillary acidic protein (GFAP) (Eng
1985; Eng and Ghirnikar 1994
). Reactive gliosis
is often found in association with diseases and is a prominent feature
in stroke and epilepsy. It has been shown that freeze-induced
neocortical malformations are intrinsically hyperexcitable
(Hablitz and Defazio 1998
; Jacobs et al.
1996
; Luhmann and Raabe 1996
). Experimental
microgyria is also associated with increased immunocytochemical
staining for glial markers (Hablitz and Defazio 1998
).
While the biochemical and antigenic changes that occur in conjunction
with reactive gliosis are well studied (Ridet et al. 1997), the changes in biophysical properties of reactive
astrocytes are just being revealed. A decrease in
KIR current amplitudes has been
reported after injury of cultured astrocytes (MacFarlane and
Sontheimer 1997
) and after a posttraumatic injury in vivo (D'Ambrosio et al. 1999
; Schroder et al.
1999
), leading to an impaired K+
homeostasis (D'Ambrosio et al. 1999
). However, these
findings are still controversial in light of conflicting previous
studies. In 1970, Pollen and Trachtenberg suggested that
a decisive factor in the development of posttraumatic focal epilepsy
was a degradation of glial function, especially with respect to the
buffering of K+. A loss of
KIR channels was reported in human
reactive astrocytes associated with epileptic seizure foci
(Bordey and Sontheimer 1998
) and in rat hippocampal
slices rendered seizure prone following a fluid percussion injury
(D'Ambrosio et al. 1999
). However,
Glötzner (1973)
, using alumina hydroxide
gel-induced epileptogenic lesions in cats, found that neuroglial
K+ buffering was even more efficient in
transporting K+ away from the epileptic sites
than under normal conditions. This finding is not consistent with a
loss in KIR currents that would reduce
their buffering capacity but could result from an increase in
intercellular coupling among astrocytes as has been reported for human
epileptic astrocytes (Lee et al. 1995
; Naus et
al. 1991
). Adding even more complexity to the issue is the fact
that a subpopulation of reactive astrocytes proliferates
(Garcia-Estrada et al. 1993
; Guénard et al.
1996
; Krushel et al. 1995
; Niquet et al.
1994
; Ritchie and Rogart 1977
). Proliferative
astrocytes differ significantly in their biophysical properties from
differentiated astrocytes. Most notably, they show reduced
KIR channel activity
(MacFarlane and Sontheimer 1997
).
These conflicting findings in different models of injury could result
from examination of different populations of reactive astrocytes.
Indeed, the above-mentioned in vivo or in situ studies thus far have
focused on elucidating properties of astrocytes associated with
epileptic seizure foci. In an effort to characterize properties of
reactive astrocytes associated with glial scars, we used a cortical
freeze-lesion model of brain injury that is characterized by the
development of abnormal microgyria. This maldevelopment is associated
with neuronal loss and a hyperexcitable zone in adjacent cortex
(Hablitz and Defazio 1998; Jacobs et al. 1996
; Luhmann and Raabe 1996
). This model is
thus extremely useful for the comparison of reactive astrocytes in and
around the lesion with those in the hyperexcitable region. We used
whole cell patch-clamp recordings to characterize the electrical
profile of reactive astrocytes in different regions of freeze-lesioned
neocortex. BrdU, which intercalates into the nuclear DNA, was detected
by an antibody and was used to distinguish proliferating astrocytes in
these slices. We found that proliferative astrocytes were concentrated at the base of the microsulcus and lacked expression of
KIR channels and were not coupled by
gap-junctions as judged from dye injections. By contrast, astrocytes in
the hyperexcitable zone were not proliferating and displayed increased
intercellular coupling associated with prominent expression of
KIR channels. If indeed,
KIR channels and gap junctions each
serve a role in K+ buffering, these results
suggest that astrocytes directly adjacent to the lesion would be
compromised in their ability to buffer K+ through
diffusional uptake by KIR channels and
redistribution through gap junctions, whereas the ones in the
hyperexcitable zone might be more proficient.
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METHODS |
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Freeze lesion
Freeze lesions were performed as previously reported
(Hablitz and Defazio 1998). Briefly, timed-pregnant
Sprague-Dawley dams were maintained in our animal facility until
parturition. On P1, rat pups were anesthetized via hypothermia. After a
midline scalp incision, the skin was retracted, and a 5-mm brass probe
cooled to
60°C was applied for 3 s to the skull 1.5 mm lateral
to the midline. After suturing, animals were allowed to recover under a
heating lamp for 30 min. They were then returned to their cage and
allowed to survive for 16-24 days prior to experimentation. The
institutional review board approved this procedure.
Slice preparation
Methods used for preparation of thin cortical slices were
described previously (Bordey and Sontheimer 2000;
Hablitz and Defazio 1998
). Briefly, rats were
anesthetized using pentobarbital (50 mg/kg) and decapitated. The brain
was quickly removed and placed in ice-cold (4°C) artificial
cerebrospinal fluid (ACSF) containing (in mM) 125 NaCl, 2.5 KCl, 2 CaCl2, 1.5 MgCl2, 1.25 NaH2PO4, 25 NaHCO3, 10 glucose, and 5 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES) and was continuously oxygenated with 95%
O2-5% CO2. The brain was
hemisected, and a block of cortex containing the microsulcus was glued
(cyanoacrylate glue) to the stage of a Vibratome. Slices (250- to
300-µm thick) were cut in cold oxygenated ACSF and transferred to a
chamber filled with ACSF at room temperature. After a recovery period
of 2 h in ACSF, slices were placed in a flow-through chamber
continuously perfused with oxygenated ACSF at room temperature. The
chamber was mounted on the stage of an upright microscope (Nikon
Optiphot2) equipped with a ×40 (2-mm working distance),
water-immersion objective and Nomarski optics.
Whole cell recordings and data analysis
Whole cell patch-clamp recordings were obtained as previously
described (Bordey and Sontheimer 2000; Hamill et
al. 1981
). Patch pipettes were pulled from thin-walled
borosilicate glass (OD, 1.55 mm; ID, 1.2 mm; WPI, TW150F-40) on a PP-83
puller (Narishige, Japan). Pipettes had resistances of 4-7 M
when
filled with the following solution (in mM): 140 KCl, 0.2 CaCl2, 1.0 MgCl2, 10 ethylene glycol-bis(aminoethyl ether)-N,N,N',N'-tetraacetic
acid (EGTA), 4 Na2ATP, and 10 HEPES, pH adjusted
to 7.2 with tris (hydroxymethyl)aminomethane (Tris). To label cells for
later morphological identification, 0.1-0.2% Lucifer yellow (LY,
dilithium salt) was added to the pipette solution. Voltage-clamp
recordings were performed using an Axopatch-200A amplifier (Axon
Instruments). Current signals were low-pass filtered at 5 kHz and
digitized on-line at 25-100 kHz using a Digidata 1200 digitizing board
(Axon Instruments) interfaced with an IBM-compatible computer system.
Data acquisition, storage and analysis were done using pClamp version 7 (Axon Instruments). For all measurements, capacitance compensation and
series resistance compensation (40-80%) were used to minimize voltage
errors. Settings were determined by compensating the transients of a
small (5 mV) 10 ms hyperpolarizing voltage step; the capacitance
reading of the amplifier was used as value for the whole cell
capacitance. After compensation, series resistances ranged from 7 to 12 M
.
Capacitive and leak subtraction was done off-line using Clampfit 7 (Axon Instruments). Since voltage- or time-dependent currents were
activated at almost every step potentials in cells with a "complex"
electrical profile, we used a modified tail protocol to calculate
membrane input resistance (Rm, Fig.
1). This protocol has been commonly used
to detect KIR currents. The cell was
stepped to 0 mV and then hyperpolarized from 0 to 160 mV. Between
0 and
60 mV, long-lasting outward K+ currents
were inactivating, and at potentials more hyperpolarized than
80 mV,
KIR currents were activated (Fig.
1A). There was thus a small window of membrane potentials
(
90 and
60 mV at the beginning and the end of the pulse,
respectively) without any voltage-activated currents. This can be seen
on the traces and in the current-voltage relationship (Fig.
1B). Membrane resistance was determined from the inverse
slope of a linear fit to the current-voltage relationship at the
membrane potentials not showing voltage-activated currents (Fig.
1C). In the cell illustrated, the membrane resistance was
280 M
(coefficient of correlation, r = 0.999). After
off-line leak subtraction using Rm
(Fig. 1A, right), the maximum current measured at about
150 mV is indicative of the true KIR
current amplitude. For cells displaying a "passive" electrical
profile, we measured the membrane input resistance, called
R*m by applying a 10-mV
hyperpolarizing pulse from a holding potential of
80 mV.
R*m values are contaminated by
the activation of inward currents present at
80 mV and are thus
underestimated. To activate transient and long-lasting outward
K+ currents, depolarizing pulses were applied
from
70 to +80 mV following a prepulse to
50 and
110 mV,
respectively. After off-line leak subtraction using
Rm,
KDR and
KA current amplitudes were measured at
+50 mV at the end of the protocol and at the peak value, respectively.
For cells displaying a "passive" electrical profile, inward
currents were measured at
160 mV on nonleak subtracted protocols.
These cells were stepped from
160 to 80 mV from a holding potential
of
80 mV. The corresponding conductance, G, of each
K+ and Na+ currents was
calculated using the following equation: G = I/(V
Vi)
where V is the membrane command potential,
Vi is the equilibrium (Nernst)
potential for the ion under consideration. Statistical values
(means ± SE, with n being the number of cells tested)
were evaluated with a statistical graphing and curve-fitting program (Origin, MicroCal). All statistical analysis was performed using GraphPAD (InStat) software. Student's two-tailed, unpaired
t-test was used to compare pairs of data sets that followed
normal standard deviation distribution. ANOVA was used for multiple
comparisons or for data that did not have normal standard deviation
distributions.
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Proliferation assay
Slices from freeze-lesioned and age-matched control animals were
incubated for 2 h in ACSF containing 10 µmol/l BrdU (Boehringer Mannheim). After cell recordings, slices were fixed in 4%
paraformaldehyde at 4°C for 3-6 h. Slices were rinsed thoroughly
with PBS and prepared for BrdU staining according to the adapted
protocol of Zupanc (1998)
. Briefly, slices were
permeabilized for 5 min in a PBS buffer containing 0.5% triton and
0.1% BSA and rinsed. DNA was denatured by 2 M HCl for 30 min at 37°C
and neutralized twice with
Na2B4O7
rinses (5 min) followed by three PBS rinses (5 min). After a 20-min
blocking step in a PBS buffer containing 0.1% Triton, 10% normal goat
serum, and 1% BSA, the slices were incubated with 1:100 dilution of
mouse anti-BrdU FITC-conjugated antibody (DAKO Corporation). This 2-h
incubation in PBS buffer was followed by three rinses (5 min) in PBS.
Slices were then mounted on glass coverslips with fluorescent mounting
medium (Vector). In some cases, the LY-filled cell was preserved, and
BrdU staining was determined for that cell. BrdU immunostaining
controls were performed by incubating slices without BrdU and using
only the fluorescently tagged antibody to BrdU. Note that we may have
underestimated the true extend of cell proliferation as only those
cells that were in S phase during slice incubation would have been BrDU labeled.
GFAP staining
Cells chosen visually for recordings were archived using a CCD camera (Watec Instruments) in combination with a video printer (Sony) for later (off-line) comparison to LY fills. After recordings, slices were transferred to a fixation medium containing 4% paraformaldehyde in PBS. Slices were washed three times in PBS for 1 h and incubated for 10 min with 0.2% Triton X-100 and 1% normal goat serum (Vector) in PBS. Slices were then incubated for 24 h at 4°C with the primary polyclonal antibody to GFAP (IncStar; rabbit anti-mouse GFAP, dilution 1:100) in PBS in the presence of 1% normal goat serum and 0.2% Triton. Slices were washed three times with PBS for 5 min and incubated with the secondary antibody (goat-anti-rabbit IgG), conjugated to rhodamine (dilution 1:100, Atlantic Antibodies) for 2 h at room temperature. Slices were mounted in Gel Mounting Media (Fisher) on glass coverslips. LY labeling and GFAP staining were visualized on a Leica DM microscope at ×20, ×40, and ×100 magnifications. Images were captured with an Optronics DEI-750 integrating camera and printed on an Epson color printer. No bleed-through was observed between the rhodamine and FITC fluorescence channels in control experiments.
Chemicals were purchased from Sigma, unless otherwise noted.
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RESULTS |
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Cortical freeze lesions contain reactive astrocytes and a population of proliferative astrocytes
A representative photomicrograph of a cerebral cortex section from
a P16 rat that received a freeze lesion on P1 is shown in Fig.
2. The lesion induced an abnormal
microsulcus. Layer I of the cortex followed the entire extent of the
microsulcus. At the base of this malformation, the cytoarchitectonic
organization of the cortical layers was disrupted as has been
previously described in this model (Dvorak et al. 1978;
Hablitz and Defazio 1998
; Humphreys et al.
1991
; Jacobs et al. 1996
). This base region of
the microsulcus showed extensive neuronal cell loss. By contrast,
normal cortical lamination was observed adjacent to the microsulcus. At
a distance of ~400 µm from the microsulcus, neurons display
epileptiform responses following electrical stimulation, and thus this
region has been referred to as an "hyperexcitable" region
(Hablitz and Defazio 1998
; Jacobs et al.
1996
; Luhmann and Raabe 1996
).
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Immunohistochemical staining for the astrocyte-specific intermediate
filament protein, GFAP, showed profound changes in GFAP immunoreactivity in postlesion cortical slices (Fig.
3A). These changes are
consistent with widespread reactive gliosis. An increase in GFAP
immunostaining was particularly evident along and at the base of the
abnormal microsulcus. GFAP immunostaining was more pronounced around
the microsulcus and in the region immediately adjacent to the
microsulcus (<200 µm). At distances >200 µm from the microsulcus
including the hyperexcitable region, staining was similar to that
observed in the contralateral, unlesioned cortex. Astrocytes appeared
to be hypertrophic with thickened processes when examined at higher
magnification (Fig. 3B). Animals that were sham-operated did
not show these morphological features typically referred to as reactive
astrocytes (data not shown). Reactive astrocytes have been shown to be
a heterogeneous cell population containing both hypertrophic
differentiated cells and proliferating astrocytes. This has also been
demonstrated in the hippocampus of kainic acid-injected rats that show
both proliferative and nonproliferative cells (Niquet et al.
1994). Similarly, in freeze-lesioned animals, Hablitz
and DeFazio (1998)
showed that some of the reactive astrocytes
were positively stained for vimentin, a marker believed to be specific
for immature, dividing glial cells (Eng and Ghirnikar
1994
). To assess this heterogeneity further, we incubated
sections containing the microsulcus with BrdU, a DNA marker that is
selectively incorporated by dividing cells during S phase of the cell
cycle. Slices were incubated for a minimum of 2 h and then stained
with a monoclonal antibody against BrdU. Figure
4 shows a representative example of the
BrdU immunoreactivity in a section from a freeze lesion compared with
an equivalent unlesioned section from the contralateral hemisphere
(Fig. 4, A and B, respectively). BrdU-positive
cells show bright red cell nuclei. Nuclear BrDU staining was prominent
on the lesioned side of the cortex. The contralateral, unlesioned side
showed few if any BrDU-positive cells, with primarily unspecific,
nonnuclear background labeling. Although proliferative cells were found
throughout the lesioned cortex, there was a tendency to cluster at the
base of the microsulcus. Microglial cells also proliferate in response to CNS injury. However, BrdU-positive cells were localized to regions
of high astrogliosis (intense GFAP staining, see preceding text) and
most likely resembled astrocytes. We have established the time course
for microglial and astrocytic proliferation and have found that
microglial proliferation predominates at 24-72 h after injury and is
essentially absent 1 wk after lesioning, while astrocyte proliferation
continues for many weeks (F. Love and H. Sontheimer,
unpublished data). This was also demonstrated in an axotomy model of
injury (Boucsein et al. 2000
). These data suggest that
the freeze-lesion gives rise to reactive gliosis composed of both
proliferative (BrdU-positive) astrocytes concentrated near the base of
the microsulcus and nonproliferative hypertropic reactive astrocytes
concentrated in adjacent areas. These phenotypes are reminiscent of
gliotic lesions in human biopsy specimens from patients with
mesio-temporal lobe sclerosis (Kallioinen et al. 1987
;
McNamara 1994
).
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Electrophysiological characterization of astrocytes in unlesioned neocortex
The electrical properties of cortical astrocytes were studied in
lesioned and unlesioned brains using whole cell patch-clamp recordings
from acutely isolated cortical slices obtained from 16- to 24-day-old
rats. Astrocytes were visually identified in different cortical layers
as cells with somata of <10 µm diameter. These cells did not fire
action potentials during seal formation nor on current injection in
current-clamp recordings. Identification of these glial cells as
astrocytes was confirmed by filling the cells with LY for subsequent
morphological analysis. Astrocytes all exhibited stellate morphology
and often extended endfeet onto blood vessels (see Fig. 7A,
control). Whole cell patch-clamp recordings were obtained from 50 control cells presumed to be astrocytes by the preceding criteria.
Astrocytes in cortical layer I had a typical complex pattern of ion
channel expression (Bordey and Sontheimer 2000).
Briefly, they expressed a composite outward potassium current that
included transient (KA) and
long-lasting delayed rectifier (KDR)
K+ currents (Fig.
5E). Inward sodium currents
and inwardly rectifying K+ currents
(KIR) were typically expressed (Fig.
5F) as previously described in our studies on hippocampal
and neocortical slices (Bordey and Sontheimer 1997
,
2000
; D'Ambrosio et al. 1998
;
Steinhauser et al. 1992
). Astrocytes in deeper cortical
layers either expressed a complex pattern of ion channels as described
for cortical layer I astrocytes (Fig. 5) or displayed a more
"passive" current pattern (Fig. 8A). This "passive"
electrical profile was, however, simply due to extensive gap-junctional
coupling between these cells (Fig. 7) preventing proper voltage-clamp
and hence activation of voltage-gated channels (more in the following
text). Dye coupling could only be observed in cells that were located
50 µm from the surface of the slice. Astrocytes showing no
cell-to-cell coupling as judged by LY injection displayed a complex
electrical profile and were located <50 µm deep into the
slice. In thin 100-µm-thick slices, LY coupling between astrocytes
was consistently absent and all recorded cells showed a complex pattern
of ion channel expression. This observation may account for some of the
differing results in previous studies (D'Ambrosio et al.
1998
; Steinhauser et al. 1992
). We did not
further explore the reason for this difference in cell coupling but
took care to monitor for each cell the exact location within the slice.
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Passive electrophysiological properties of astrocytes in freeze lesions
The electrophysiological changes of posttraumatic astrocytes were
studied in acute cortical slices from P16 to 24 freeze-lesioned animals. Reactive glial cells in sections containing the microsulcus are not likely to be reactive microglial cells since, as previously mentioned, reactive microglial cells regain a complete normal immunological and electrophysiological phenotype after 14 days (Boucsein et al. 2000). Moreover, microglia have not
been shown to extend endfeet onto blood vessels. Whole cell patch-clamp
recordings were obtained from 81 cells in the different layers of
slices containing a microsulcus and compared with 50 cells in control slices. Representative recordings of both are illustrated in Fig. 5.
The mean resting membrane potential
(Vr) was
81.6 ± 5.8 mV (mean ± SD, n = 50) for control astrocytes and
79.8 ± 8.6 mV (n = 81) for postlesion
astrocytes. The difference in Vr was
not significant. When calculated with a "tail" protocol allowing
leak current measurement in complex cells (see METHODS),
Rm was significantly different
(P = 0.0001) with values of 239.5 ± 133.6 and
636.0 ± 509.4 M
in astrocytes from control (n = 36) and freeze-lesioned animals (n = 54),
respectively. Mean membrane capacitances were not significantly
different between control and lesion groups with values of 42.1 ± 21.5 pF (n = 50) and 38.1 ± 23.1 pF
(n = 81), respectively. To interpret these results, we
further analyzed our data by comparing changes between different
regions, specifically layer I adjacent to the lesion and deeper layers.
In slices containing a microsulcus, the deeper layers can be subdivided
into two regions: the proliferative zone (seen by staining with BrdU)
and the hyperexcitable zone including dye-coupled and non-dye-coupled
cells (Fig. 2).
Mean passive membrane properties are summarized in Table
1. In layer I, while mean
Vr values were not significantly
different between lesioned and unlesioned animals (81 vs.
78.5 mV),
Rm (597 vs. 218 M
), and
Cm (27 vs. 41 pF) values were
significantly different in cells from lesioned cortex as compared with
controls. In the hyperexcitable zone, astrocytes that displayed a
"passive" current profile had a significantly more hyperpolarized
Vr in postlesion slices (
87 vs.
82 mV in unlesioned slices). Mean Cm values were not significantly
different. The difference in mean
R*m values were marginally
significant. Rm values could not be
obtained in these cells since there appeared essentially passive. In
the same region, astrocytes with a "complex" current profile
display no significant changes in their mean passive membrane
properties. In the proliferative zone, the mean values of all the
passive parameters were significantly different from the ones from
complex cells in either control or lesion-containing slices. These
cells had a more depolarized Vr (
70
mV), a higher input resistance, Rm
(788 M
), and a smaller cell capacitance (21 pF).
|
Layer I astrocytes near the microsulcus display reduced inward potassium current amplitudes
Differentiated astrocytes express inwardly rectifying
K+ currents that have been suggested to be
involved in buffering of extracellular K+
(D'Ambrosio et al. 1999; Newman 1993
;
Newman et al. 1984
; Ransom and Sontheimer
1995
). These currents have also been shown to be developmentally regulated (Bordey and Sontheimer 1997
;
Steinhauser et al. 1992
) and to decrease in amplitude in
reactive astrocytes in culture (MacFarlane and Sontheimer
1997
). Changes in KIR current amplitude in layer I astrocytes adjacent to the lesion were therefore evaluated. There was a 48% decrease in the relative
KIR current expression in these
astrocytes (Fig. 5, C and F). This was determined from conductance densities for KIR
currents in which the conductance of
KIR currents was divided by the
membrane capacitance to normalize for cell size
(gKIR) (446 pS/pF in unlesioned and
232 pS/pF in postlesion; Table 2). This
difference was statistically significant (P < 0.001).
There was also a small decrease in the number of cells displaying
KIR currents. The density of
conductance for KDR
(gKDR) and Na+
channels (gNa) was, however, not
significantly different between control and postlesion cell groups
(Table 2). On the other hand, KA mean
conductance density (gKA) was 39%
larger (Table 2) in cells near the microsulcus than in control cells.
This difference was statistically significant (P < 0.05). To describe the relative changes in the contribution of outward
and inward K+ conductances, the ratio of total
outward K+ conductance
(gKout) divided by
gKIR was computed. This ratio is a
direct indicator of the relative contribution of
KIR current to the total current and,
assuming that inward K+ currents are indeed
responsible for diffusion-mediated K+ buffering
by astrocytes, a larger ratio reflects improved
K+ cell buffering capacity. As can be seen in
Table 2, there is an apparent decrease in the buffering capacity of
layer I astrocytes adjacent to the lesion with
gKout/gKIR
ratios larger than 7.
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Proliferative astrocytes lose KIR currents after a freeze lesion
De novo proliferation of astrocytes has been shown to be
accompanied by a change in K+ current amplitudes
in vitro (MacFarlane and Sontheimer 1997), with a
parallel decrease and increase in KIR
and KDR current amplitudes, respectively. We thus recorded reactive astrocytes at the base of the
microsulcus where proliferative astrocytes were prominent as indicated
by BrdU staining (Fig. 4). After an incubation period of
2 h in BrdU,
astrocytes were recorded and filled with LY. BrdU-immunostaining of
LY-filled cells was then performed to allow identification of ion
channel expression in single proliferative astrocytes. Figure
6A shows two representative
BrdU-positive astrocytes filled with LY. The cell on the
left shows a confocal image in which the LY fill (green,
smaller image middle top) and BrDu labeling (red, small
image middle bottom) were merged showing the nuclear localization of the BrDU label. The example on the right shows a
fluorescent micrograph in which several cell nuclei were BrDU positive
(red) including the LY-filled cell. None of the BrdU-positive cells
showed dye coupling. Of 26 recorded and LY-filled astrocytes in the
proliferative zone near the microsulcus, only 5 cells were recovered
that still had an intact cell body allowing for unequivocal identification using BrDU antibodies. All five were BrDU positive. The
other cells had their cell body removed on withdrawal of the patch
pipette. However, these cells showed identical current profiles as the
BrdU-positive cells and for this reason were presumed to have also been
proliferating cells. These cells were included in the pooled data to
evaluate changes in K+ current amplitudes (Table
2). Specifically, proliferating cells had large outward
K+ currents but lacked
KIR currents as shown for a
representative example in Fig. 6B. A representative
"complex cell" recording from the hyperexcitable zone and the
corresponding I-V curve are shown in Fig. 6, C
and D. There was a significant 72% decrease in
gKIR between control complex cells in
layer II-VI and proliferative cells (460 and 127 pS/pF, respectively,
see Table 2). This change was also accompanied by a notable decrease in
the number of cells expressing KIR
channels (100% in control vs. 31% in postlesion groups). This
apparent loss of KIR currents was
consistent with the more depolarized
Vr from proliferative astrocytes (
71
mV in lesion vs.
84 mV in control). There was a significant 114% increase in gKDR in cells near the
microsulcus, whereas gKA did not
change significantly. These changes resulted in a marked increase in
the
gKout/gKIR
ratio (7.7 from proliferative cells compared with 2.3 from control
complex cells), suggesting a loss of the K+
buffering capacity in these proliferative astrocytes.
|
Sodium currents have been reported to be increased in reactive
astrocytes in situ (Bordey and Sontheimer 1998) and in
proliferative astrocytes in vitro (MacFarlane and Sontheimer
1997
). Astrocytes near the microsulcus did not show a
statistically significant increase in
gNa. MacFarlane and Sontheimer
(1997)
also reported a switch from TTX-sensitive to
TTX-resistant Na+ currents in reactive spinal
cord astrocytes in vitro. TTX sensitivity was tested in five
proliferative astrocytes (2 were recovered for BrdU staining and were
positive). In each cell tested, Na+ currents were
completely and reversibly blocked by TTX 100 nM (data not shown). The
injury-mediated loss of TTX sensitivity may thus be a phenomenon
typical of spinal cord injury.
Astrocytes in the hyperexcitable zone display increased dye coupling
Lee et al. (1995) reported that gap-junction
coupling was more pronounced in cells isolated and cultured from
hyperexcitable tissue surrounding human epileptic seizure foci than
from uninvolved comparison tissues. By filling the cells with LY, we
attempted to assess the extent of cell-to-cell coupling between
astrocytes. We counted the number of LY-filled cells surrounding one
single recorded astrocyte in the hyperexcitable zone. To include cells in multiple planes, we performed confocal reconstruction of coupled cells, as shown in Fig. 7, A
and B. The number of LY-coupled cells was significantly
higher in postlesion section as compared with control sections
[13.6 ± 4.99 cells (n = 6) in control and
46.5 ± 15.19 cells (n = 5) in postlesion
(P < 0.001), Fig. 7B]. This increase in
intercellular coupling correlated with a more negative Vr of postlesion astrocytes and with a
decrease in the input resistance. There was, however, no significant
increase in Cm (8% increase in
postlesion sections). This discrepancy between an increase in the
extent of intercellular coupling and
Cm could be explained by the limited
capacity of the patch-clamp to detect distal membrane capacitances.
LY-coupled astrocytes expressed a passive current profile. The
conductance density for inward currents was identical in cells from
control and freeze-lesioned cortex (1,115 ± 471 and 1,307 ± 518 pS/pF, respectively). The inward current was sensitive to barium (1 mM), indicating that these inward currents were due to
K+ channel activation (Fig.
8, A and B). Figure
8C shows the barium-sensitive inward current and its
I-V curve. The ratios of outward versus inward conductances
were 0.79 ± 0.23 and 0.64 ± 0.12 in control and postlesion
slices, respectively. Overall, these results suggest an increased
K+ conductance in astrocytes of freeze-lesioned
neocortex and an increase in interconnectivity in this region.
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DISCUSSION |
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This study demonstrates significant electrophysiological changes in reactive astrocytes in a model of cortical maldevelopment associated with epileptiform activity. By examining different populations of reactive astrocytes, our results begin to reconcile previous contrasting reports on the properties of astrocytes in epileptogenic tissue. We show that a subset of reactive astrocytes near the freeze-induced microsulcus is proliferative. It appears likely that these cells may have lost their capacity to buffer extracellular K+ since they no longer express KIR channels, presumed to mediate this function. However, astrocytes surrounding the microsulcus, a region that is considered "hyperexcitable," display increased K+ conductances and enhanced intercellular coupling, features suggesting enhanced K+ buffer capacity.
Some reactive astrocytes surrounding the microsulcus are proliferative
Our results show that the abnormal microsulcus is surrounded by
reactive astrocytes as indicated by increased immunostaining for GFAP.
In addition, some of these reactive astrocytes are BrdU-positive, indicating that proliferative cells are located where neuronal loss has
occurred. The emergence of proliferative astrocytes is commonly
observed after brain injury and often accompanies neuronal loss. As an
example, a convulsant dose of kainic acid injected intracerebroventricularly into rats causes degeneration of hippocampal neuronal cells (Ben-Ari 1985). This degeneration is
accompanied by astrocytic proliferation (Murabe et al.
1981
; Niquet et al. 1994
). Although reactive
gliosis has been extensively observed in different brain injury models,
the pathways leading to the activation and the proliferation of
reactive astrocytes are still unknown. Astrocyte proliferation has been
shown to be modulated by various factors including neurotransmitters
and growth factors (Abbracchio et al. 1994
;
Gómez-Pinilla et al. 1995
; Hodges-Savola et
al. 1996
; Huff et al. 1990
; Sawada et al.
1993
; Scherer and Schnitzer 1994
; Selmaj
et al. 1990
; Stachowiak et al. 1997
). In the
case of the freeze-lesion model, it is believed that the freezing probe
induces focal hypoxia resulting in cell death and subsequent abnormal
lamination (Dvorak and Feit 1977
; Humphreys et
al. 1991
). A cascade of phenomena might thus occur leading to
increased extracellular K+ and hyperexcitability
and increased neurotransmitter release, as well as release of various
growth factors and cytokines.
Two populations of reactive astrocytes coexist in the freeze lesion model and may differentially affect neuronal development and function. We focused on the electrical properties of these cells, most notably in their K+ conductances, as they are likely to affect the cell's ability to buffer K+. We recorded astrocytes surrounding the lesion (layer I) and at the base of the lesion focusing on BrdU-positive cells. Although we did not observe a significant increase in GFAP immunostaining in the hyperexcitable region known to be present 2 wk after the initial in vivo lesion, we recorded astrocytes in this region and compared their properties to normal astrocytes.
Proliferative astrocytes lack KIR currents and expressed large KDR currents
A functional correlation between K+ channel
expression and cell proliferation has been commonly observed
(Chiu and Wilson 1989; MacFarlane and Sontheimer
1997
; Nilius and Wohlrab 1992
; Pappone and Ortizmiranda 1993
; Puro et al. 1989
;
Woodfork et al. 1995
). A recent study of reactive,
proliferative astrocytes in vitro showed changes in
K+ channel expression similar to the one
described in our study (MacFarlane and Sontheimer 1997
).
In comparison with control astrocytes in all cortical layers except
layer I, 70% of the presumably proliferative cells lack
KIR channel expression, whereas they
all expressed KDR channels. In the
cells expressing KIR channels, there
was a significant decrease in KIR
current amplitude accompanied by a significant increase in
KDR current amplitude. These cells
also had a significantly more depolarized resting membrane potential and smaller membrane capacitance reflecting their smaller cell size.
These changes in ion channel expression mirror those associated with
development (Bordey and Sontheimer 1997
;
Steinhauser et al. 1992
), suggesting that these
reactive, proliferative astrocytes regain an immature phenotype. In the
freeze lesion model, N-methyl-D-aspartate and
GABA channels also express subunits more typical of immature neurons
(DeFazio and Hablitz 1999
, 2000
).
Similarly, reactive astrocytes in the portion of layer I overlying the lesion displayed significant changes in their electrical profile. A decrease in KIR current amplitudes and an increase in KA current amplitudes mainly characterized these changes. There was also a decrease in cell-to-cell coupling. As previously mentioned, such changes inversely mirror changes accompanying gliogenesis and might reflect a delayed maturation of these astrocytes.
Astrocytes in the hyperexcitable zone express increased intercellular coupling
Conflicting hypotheses exist concerning the buffering capacity of
glial cells in epileptic foci (Bordey and Sontheimer
1998; Glötzner 1973
; Pollen and
Trachtenberg 1970
). Walz and Wuttke (1999)
showed that reactive astrocytes in slices containing gliotic hippocampal CA1 still have the capacity to limit the increases in
extracellular K+ that are produced by hyperactive
neurons following kainic acid injection. Astrocytes from the epileptic
foci also have increased gap junction coupling (Lee et al.
1995
). This could facilitate the redistribution of
K+ from the epileptic site to sites with lower
K+ levels. Increased levels of neurotransmitter
such as glutamate exist at epileptic foci (Carlson et al.
1992
; During and Spencer 1993
; Meldrum
1994
; Ronne-Engstrom et al. 1992
). Since glia
actively take up glutamate, enhanced intercellular coupling would help the sequestration and redistribution of glutamate through the glial
syncytium. Increased coupling could be seen as an adaptive response of
reactive astrocytes to neuronal hyperexcitability. Consistent with this
idea of increased gap junctional coupling, propagation of paroxysmal
activities is accompanied by either changes in the membrane surface
area and/or the interglial communication via gap junction
(Amzica and Neckelmann 1999
). Our finding of a 240%
increase in intercellular coupling demonstrates that astrocytes surrounding hyperexcitable neurons potentially have an increased capacity to allow ionic diffusion between cells and this would likely
include K+. These cells, by effectively expanding
their communication compartment, could also provide neuroprotective
effects following brain injury. Increased intercellular coupling could
however have epileptogenic effects by increasing the synchronization of
discharges (Naus et al. 1991
). Thus taken the loss of
coupling at the lesion site and the enhanced coupling in astrocytes
together, one might envision a scenario where at the lesion,
extracellular ion such as glutamate and K+ may
accumulate. If these changes induce an astrocytic response, such as a
spreading Ca2+ wave (Finkbeiner
1992
), its propagation may be facilitated by the enhanced
coupling of cells in the surrounding hyperexcitable tissue.
If one equates expression of KIR channels with an enhanced ability of cells to take up K+, our results suggest that, in the core of the freeze-induced malformation and in adjacent areas, there may be a decreased capacity to buffer K+. This would in turn lead to neuronal dysfunction and eventually neuronal death. Similarly, if one equates the expression of gap junctions with an enhanced ability to spatially redistribute K+ through the glial syncytium, our results suggest that in the hyperexcitable zone there is an increase capacity to spatially buffer K+ accompanying neuronal activity, while spatial buffering is lost in the lesion proper. Clearly, further studies are needed to functionally link these changes in astrocytic properties to changes in the neuronal microenvironment at the lesion sites.
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ACKNOWLEDGMENTS |
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This work was supported by National Institute of Neurological Disorders and Stroke Grants R01-NS-31234 and NS-22373.
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FOOTNOTES |
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Address for reprint requests: H. Sontheimer, Dept. of Neurobiology, The University of Alabama at Birmingham, 1719 6th Ave. S., CIRC Rm. 545, Birmingham, AL 35294 (E-mail: hws{at}nrc.uab.edu).
Received 4 October 2000; accepted in final form 8 December 2000.
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REFERENCES |
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