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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
J Neurosci. Author manuscript; available in PMC 2010 May 24.
Published in final edited form as:
PMCID: PMC2875068

Astrocytic Dysfunction in Epileptogenesis: Consequences of Altered Potassium and Glutamate Homeostasis?


Focal epilepsy often develops following traumatic, ischemic or infectious brain injury. While the electrical activity of the epileptic brain is well characterized, the mechanisms underlying epileptogenesis are poorly understood. We have recently shown that in the rat neocortex, long-lasting breakdown of the blood-brain barrier (BBB) or direct exposure of the neocortex to serum-derived albumin leads to rapid up-regulation of the astrocytic marker, glial fibrillary acidic protein (GFAP), followed by delayed (within 4–7 days) development of an epileptic focus. We investigated the role of astrocytes in epileptogenesis in the BBB-breakdown and albumin models of epileptogenesis. We found similar, robust changes in astrocytic gene expression in the neocortex within hours following treatment with deoxycholic acid (BBB breakdown) or albumin. These changes predict reduced clearance capacity for both extracellular glutamate and potassium. Electrophysiological recordings in-vitro confirmed the reduced clearance of activity-dependent accumulation of both potassium and glutamate 24 h following exposure to albumin. We used a NEURON model to simulate the consequences of reduced astrocytic uptake of potassium and glutamate on excitatory postsynaptic potentials (EPSPs). The model predicted that the accumulation of glutamate is associated with frequency-dependent (>100 Hz) decreased facilitation of EPSPs, while potassium accumulation leads to frequency-dependant (10–50 Hz) and N-methyl-D-aspartic acid (NMDA)-dependent synaptic facilitation. In-vitro electrophysiological recordings during epileptogenesis confirmed frequency-dependant synaptic facilitation leading to seizure-like activity. Our data indicate a transcription-mediated astrocytic transformation early during epileptogenesis. We suggest that the resulting reduction in the clearance of extracellular potassium underlies frequency-dependent neuronal hyper-excitability and network synchronization.

Keywords: Albumin, Astrocytes, Blood-brain barrier, Glutamate, Post-synaptic potentials, Potassium


Epilepsy is one of the most common neurological disorders, affecting 0.5–2% of the population worldwide. While the characteristic electrical activity in the epileptic cortex has been studied extensively, the mechanisms underlying epileptogenesis are poorly understood. Focal neocortical epilepsy often develops following traumatic, ischemic or infectious brain injury. Under these conditions, vascular damage is common and includes local breakdown of the blood-brain barrier (BBB; Abbott et al., 2006; Neuwelt, 2004; Tomkins et al., 2001). The BBB has long been recognized as crucial for maintenance of the brain’s micro-environment, but it was only recently documented that disruption of the blood brain barrier plays an important role in the pathogenesis of epilepsy (Seiffert et al., 2004; van Vliet et al., 2007; Marchi et al., 2007). It was found, for example, that in the rat neocortex, long-lasting BBB disruption leads to gradual development (within 4–7 days) of an epileptic focus that persists for weeks (Seiffert et al., 2004; Ivens et al., 2007; Tomkins et al., 2007). Our experiments further indicated that during BBB breakdown, serum-derived albumin diffuses into the brain’s extracellular space and is rapidly transported into astrocytes via a specific receptor-mediated mechanism. Albumin uptake by astrocytes was followed by a rapid (within hours) up-regulation of the astrocytic marker glial fibrillary acidic protein (GFAP; Seiffert et al., 2004; Ivens et al., 2007), suggesting that astrocytic dysfunction plays a role in injury-induced epileptogenesis.

Data accumulating from human and animal studies supports the notion that glial cells make an important contribution to the control of neuronal function under both normal and pathological conditions (for reviews see Araque et al., 2001; Seifert et al., 2006; Wetherington et al., 2008). Studies on epileptic tissue show significant alterations in the expression of astrocytic proteins, including increased expression of GFAP (Bordey and Sontheimer, 1998) and reduced expression of proteins involved in the regulation of extracellular potassium ([K+]o) and glutamate (Hinterkeuser et al., 2000; Schroder et al., 2000). Despite the vast body of data confirming changes in the morphology and function of astrocytes in epileptic tissue, the direct role of these cells in the development of the epileptic network (i.e., in epileptogenesis) remains unclear. In this study, we combined molecular, electrophysiological and computer modeling approaches to investigate the potential role of vascular-injury-induced and albumin-induced early transformation of astrocytes in altered neuronal excitability and epileptogenesis.

Materials and Methods

Animals were housed and handled according to the directives of the internationally accredited Animal Care and Use Committees (IACUC) at Charité University Medicine, Berlin, and Ben-Gurion University of the Negev, Beer-Sheva. All experimental procedures were approved by the ethical committees supervising experiments on animals at Charité University Medicine (in-vivo approval no.: G0104/05, in-vitro: T0228/04) and Ben-Gurion University of the Negev (approval no.: BGU-R-71-2006).

In-vivo experiments

The in-vivo experiments were performed as previously described in Seiffert et al. (2004). In brief, adult male Wistar rats (120–250 g) were anesthetized using ketamine and xylazine and placed in a stereotactic frame. A 4-mm diameter bone window was drilled over the somatosensory cortex, the dura was opened and the underlying cortex was perfused with artificial cerebrospinal fluid (ACSF). For the “treated” rats group, the BBB-disrupting agent deoxycholic acid sodium salt (DOC, 2 mM, Sigma-Aldrich, Steinheim, Germany) or bovine serum albumin (0.1 mM, >98% in agarose cell electrophoresis; catalogue no. A7906, Sigma Aldrich, Steinheim, Germany) was added to the ACSF. Albumin concentrations corresponded to 25% of the normal serum concentration [determined to be 0.4 mM for 10 rats, see also Geursen and Grigor (1987); final osmolarity of 303–305 mOsmol/l]. For the sham-operated control group, the cortex was perfused with ACSF. The composition of the ACSF was (in mM): 129 NaCl, 21 NaHCO3, 1.25 NaH2PO4, 1.8 MgSO4, 1.6 CaCl2, 3 KCl, and 10 glucose. Rats were sacrificed at 7–8, 24, or 48 h following treatment, before the onset of epileptiform activity (>4 days, see Seiffert et al., 2004).


Total RNA from animals treated with DOC or with albumin was isolated from the somatosensory cortex, directly under the craniotomy area, using the TRIzol® reagent (Invitrogen, Carlsbad, CA), and prepared using the Affymetrix GeneChip one-cycle target labeling kit (Affymetrix, Santa Clara, CA). Biotinylated cRNA was then fragmented and hybridized to the GeneChip Rat Genome 230 2.0 Array according to manufacturer’s protocols (Affymetrix Technical Manual). The array data was normalized by using GCRMA (GC Robust Multi-Array Average) or RMA (Robust Multi-Array Average) analysis. One array was run for each treatment (DOC and albumin) and for every contralateral hemisphere for the following time points: 7/8, 24, and 48 h. The data from a sham-treated animal (24 h) was used to normalize the other arrays. To identify genes involved in astrocytic functions, we used GeneCards (, querying for “astrocyte”. For comparison of the relative changes in the expression of astrocytic vs. neuronal genes, we used gene sets published by Cahoy and colleagues (2008) of astrocytic and neuronal enriched genes (expressed by S100β+ and S100β−/PDGFRα−/MOG-cells, respectively). Cluster analysis was performed with MATLAB by assessing the expression relationship as the Euclidean distance in N-dimensional space between measurements (N denotes number of gene transcripts). Arrays were then clustered according to distance data, by using the Unweighted Pair Group Method with Arithmetic mean method (UPGMA, Gronau and Moran, 2007).

In-vitro astrocytic and neuronal culture preparations

Primary neuronal cortical cultures were prepared from embryonic day 18 rats as reported previously (Kaufer et al., 2004). Briefly, cells were dissociated with a papain solution for 20 min at 37°C. After the removal of the papain solution, the tissue was resuspended in growth medium [MEM with Earle’s salts containing 2.5% B27 supplement, 0.1% mito serum extender, 5% fetal bovine serum (FBS), 20 mM glucose, and 5 mM L-glutamine] and dissociated by mechanical trituration. The cells were plated, and after 4 h in vitro the cell culture medium was replaced with neurobasal medium supplemented with 2% B27 supplement and 0.5 mM GlutaMAX. The cells were maintained in 5% CO2 at 37°C. After 7 days in vitro, cytosine arabinofuranoside (AraC) (10 μM) was added to the cultures. After 10 days in vitro, the cells were incubated with 0.4 mM albumin for 24 h at 37°C. For astrocytic cultures, astrocytes were isolated from the cerebral cortices of P0 rat pups. Cells were dissociated with papain and mechanical trituration. The cells were cultured in high-glucose Dulbecco’s modified eagle medium supplemented with 10% FBS and 1% penicillin/streptomycin at 37°C and in 5% CO2 (medium was replaced every 3–4 days). After 10 days in vitro, the culture medium was replaced with serum-free high-glucose DMEM (containing 1% penicillin/streptomycin) for 18 h. The cells were then incubated in serum-free medium containing 0.4 mM albumin for 24 h at 37°C. For immunostainings cells were washed with phosphate buffered saline (PBS) and fixed in 4% paraformaldehyde for 15 min. The cells were permeabilized with 0.2% Triton X-100 in PBS for 5 min and washed in PBS. They were then incubated with 5% normal donkey serum in PBS for one hour at room temperature followed by overnight incubation at 4°C with either mouse anti-NeuN (1:1000; Chemicon, Temecula, CA) or mouse anti-GFAP (1:1000; Cell Signaling Technology, Beverly, MA). The cells were washed in PBS, incubated with donkey anti-mouse Cy3 (1:1000; Jackson ImmunoResearch, West Grove, PA) for 1 hour at room temperature, and then counterstained with DAPI.

Real-time polymerase chain reaction

Total RNA was isolated from the somatosensory cortices of animals treated with DOC or albumin (24 h treatment; n = 3) or from primary cultures (astrocytic and neuronal, n = 3 independent experiments). Expression levels were determined by real-time reverse transcriptase-PCR (RT-PCR) with an iQ5 detection system (Bio-Rad, Hercules, CA) using gene-specific primer pairs. RT-PCR data were analyzed using the PCR Miner program (Zhao and Fernald, 2005), and fold changes in gene expression were represented relative to sham-operated controls (in-vivo samples) or serum-deprived controls (in-vitro samples). Ribosomal 18S RNA (18S rRNA) was used as an internal control for variations in sample preparation. For samples from in-vivo treatments, RT-PCR was performed with the iScript one-step RT-PCR kit (Bio-Rad). Control RT-PCR reactions were performed without reverse transcriptase to verify amplification of genomic DNA. For in-vitro samples, DNase treatment was applied, followed by first-strand cDNA synthesis (iScript cDNA Synthesis kit, Bio-Rad). PCR reactions were carried out with iQ SYBR Green Supermix (Bio-Rad). Primer specificity was verified by melt curve analysis. The amplification cycles for 18S, Gja1, GS, SLC1A2, SLC1A3 (GLAST) and Kcnj10 consisted of 40 cyclesof 10 s at 95°C, 30 s at 55°C, and 30 s at 72°C. The amplification cycles for Gjb2 and Gjb6 consisted of 40 cyclesof 10 s at 95°C, 30 s at 60°C, and 30 s at 72°C.

Primer sequences (forward, reverse) were as follows: 18S rRNA (GenBank accession number M11188.1, 5″-CCATCCAATCGGTAGTAGCG-3″, 5″ GTAACCCGTTGAACCCCATT-3″); SLC1A3 (GenBank accession number NM_019225.1; 5″-GAGGCCATGGAGACTCTGAC- 3″, 5″-CGAAGCACATGGAGAAGACA-3″); GS (GenBank accession number NM_017073.3; 5″-AGCGACATGTACCTCCATCC-3″, 5″ TACAGCTGTGCCTCAGGTTG-3″); Kcnj10 (GenBank accession number X83585.1; 5″-GAGACGACGCAGACAGAGAG-3″, 5″CCACTGCATGTCAATGAAGG-3″); Gjb2 (GenBank accession number NM_001004099.1; 5″-GGTTTGTGATGTGAGCATGG-3″, 5″-CTCAGCACACCAAGGATGAA-3″); Gjb6 (GenBank accession number NM_053388.1; 5″-GCCAAGATGAGTCACAGCAA- 3″, 5″-TCAGAGCTGGATCACAATCG-3″); Gja1 (GenBank accession number NM_012567.2; 5″-TCCTTGGTGTCTCTCGCTTT-3″, 5″-TTTGGAGATCCGCAGTCTTT-3″); SLC1A2 (GenBank accession number NM_017215.2; 5″-GGTCAATGTAGTGGGCGATT-3″, 5″-GGACTGCGTCTTGGTCATTT-3″).

In-vitro electrophysiological recordings

For electrophysiological experiments, rats were deeply anesthetized with isoflurane and then decapitated. Brains were quickly removed, and transverse hippocampal-cortical slices (400 μm thick) were prepared using a vibratome (Campden Instruments, Loughborough, UK). Slices were maintained in a humidified, carbogenated (5% CO2 and 95% O2) gas atmosphere at 36 ± 1°C and perfused with ACSF in a standard interface chamber (Seiffert et al., 2004; Ivens et al., 2007). To mimic the altered ionic environment during BBB disruption, recordings were acquired in a serum-adapted electrolyte solution (sACSF; see Seiffert et al. 2004). sACSF was similar in composition to the ACSF except for different concentrations of MgSO4 (0.8 mM), CaCl2 (1.3 mM), KCl (5.7mM) and glutamine (1 mM). “Treated” slices were incubated with sACSF containing 0.1 mM bovine serum albumin for 2 h before transfer to the perfusion chamber.

Electrophysiological recordings were obtained 6–10 h following perfusion with sACSF. Control slices were treated similarly, using sACSF without albumin. For extracellular recordings, glass microelectrodes (~3 MΩ, 154 mM NaCl) were positioned in layer 4 of the neocortex. Slices were stimulated with brief (100 μs) pulses, by using bipolar stimulation electrodes placed at the border between white and gray matter in the same cortical column. Trains of 50 stimuli were applied at 2, 5, 10, 20, 50 and 100 Hz, at 2.5× threshold stimulation intensity. Signals were amplified (SEC-10L; NPI Electronics, Tamm, Germany), filtered at 2 kHz, displayed on an oscilloscope, digitized on-line (CED-1401micro; Cambridge Electronics Design, Cambridge, UK) and stored for off-line analysis. Extracellular potassium concentrations ([K+]o) were measured with ion-sensitive microelectrodes (ISMEs; Lux and Neher, 1973; Jauch et al., 2002).

In vitro intracellular recordings were obtained from pyramidal neurons (layer 2–3) 23–28 h following the in vivo treatment with albumin or from control rats. Currents were recorded using the whole cell patch configuration, as described previously (Pavlovsky et al., 2003). In brief, glass pipettes were pulled from capillaries using a vertical puller (Narashige, Greenvale, NY) and filled with a solution comprising (in mM): 150 CsCl, 1 MgCl2, 10 HEPES, 4 Na2ATP, 0.1 CaCl2, and 1.1 mM EGTA, pH adjusted to 7.2 with a final osmolarity of 290–310 mOsm. Cells were visualized using infrared differential interference phase contrast video-microscopy. Recordings were performed using AxoPatch 700B (Axon Instruments, Foster City, CA), digitized at 10 kHz and recorded using pClamp 9.2 (Axon Instruments, Foster City, CA). Patch pipette’s resistance was 4–5 MΩ. Series resistance was not electronically compensated; however, cells in which series resistance varied by more than 25% were excluded from the analysis. Stimulation protocols were started at least 5 min following impalement to allow intracellular dialysis with the pipette solution. Excitatory post-synaptic currents (EPSCs) were evoked – using a bipolar stimulating electrode positioned <200 μm from the recorded cell – at 75% of the intensity producing maximal EPSCs. N-methyl-D-aspartic acid (NMDA) currents were recorded in the presence of blockers of α-amino-3-hydroxy-5-methyl-4-isoxazole-propionic acid (AMPA)/kainate (i.e., 30 μM CNQX) and of gamma-aminobutyric acid A (GABAA) receptors (i.e., 10 μM bicuculine methiodide). Cells were voltage clamped to +40 mV to alleviate NMDA receptor blockade and inactivate fast Na+ currents. In some experiments, dihydrokainic acid (DHK, Tocris, Bristol, UK), 100 μM, was added to the extracellular solution to selectively block, the astrocyte specific glutamate transporter, SLC1A2 (see Arriza et al., 1994).

Computer simulations

A computer model was implemented using the NEURON modeling environment (Hines and Carnevale, 1997) with 20-μstime steps. The model consisted of a multi-compartment isolated cell, simulating a layer 2/3 cortical neuron, using only passive membrane properties. Geometric parameters and spatial relationships of the 74 compartments were modeled after Traub and colleagues (2003). Resting membrane potential was set at −65 mV (determined by Na+ and K+ conductance); membrane capacitance Cm was 0.9 μF/cm2; and the cytoplasmic resistance was set at 250 Ω/cm2. Simulated excitatory inputs consisted of eight synapses on apical dendrites (located 1368 μm from the soma), contributing currents with AMPA and NMDA kinetics modeled after Saftenku (2005) and Kampa et al. (2004), respectively. AMPA to NMDA maximal current ratios were set at 1 (Myme et al., 2003). Synaptic currents were triggered by a surge of ‘glutamate’, decaying with first-order kinetics (baseline time constant = 1.2 ms). Down-regulation of uptake mechanisms was simulated by changing the time constant of the decay function, similar to the effect of the application of DL-threo-β-benzyloxyaspartate (DL-TBOA, Diamond, 2005). To investigate the effects of altered [K+]o, each compartment was enveloped by a fixed space in which potassium was allowed to accumulate. [K+]o diffused’ either into the bathing solution or into astrocytes with KIR kinetics. Since KIR channel conductance is proportional to [K+]o (Sakmann and Trube, 1984), K+ influx into ‘astrocytes’ was determined by the local potassium gradient ([K+]o – [K+]bath) modulated by KIR conductance (log[K+]o; adapted from Ciani et al., 1978).


where Ik - momentary K+ flux (nA/cm2), Ikrest - resting K+ flux (nA/cm2),τECS - time constant for potassium diffusion into the extracellular space, τastrocytic - time constant for potassium diffusion into astrocytes, F - Faraday constant, C - ratio of astrocytic K+ uptake relative to extracellular diffusion, and V - radius of enveloping extracellular space, set at 20 nm (Egelman and Montague, 1999; Savtchenko et al., 2000). The ionic flux equation describes first-order potassium clearance by both free diffusion and ‘astrocytic’ uptake (see Kager et al., 2000). Lateral diffusion of K+ ions was not taken into account. To simulate a decrease in astrocytic potassium clearance, τastrocytic was increased to mimic a reduction in astrocytic KIR channels. “Resting” ion concentrations were set at (in [mM]): [Na+]o, 145; [Na+]i, 12; [K+]o, 3.5; [K+]i, 140.

Statistical analysis

Data are expressed as means ± SEM. Differences between treated and control slices were determined by the Mann-Whitney U test for two independent samples. Statistical tests were performed using SPSS 13.0 for Windows. The level of statistical significance was set at p < 0.05, unless otherwise stated.


Astrocytic transcriptional changes following BBB opening or exposure to albumin

To explore changes in astrocytic gene expression during epileptogenesis, we analyzed gene-array data from DOC- and albumin-treated brains (n = 3 from each treatment) during the first 48 h after treatment and prior to the development of epileptiform activity (Ivens et al., 2007; Seiffert et al., 2004). When compared to sham-operated controls, the two treatments, at each time point, resulted in similar changes in expression of astrocytic-enriched genes with a correlation coefficients between the different treatments (see Methods) of r2 = 0.69, 0.82, 0.85 for 8, 24 and 48 h following treatment, respectively, p<0.0001, Fig. 1a). Unsupervised hierarchical cluster analysis revealed further similarities between changes in transcripts levels in treated cortices (which cluster according to time after treatment) (Fig. 1b) while transcripts changes in the contralateral, untreated, hemispheres are relatively dissimilar and cluster together.

Figure 1
Transcriptional changes in astrocytes following exposure to albumin or BBB disruption

In a recent study, Cahoy and colleagues (2008) created a transcriptome database reflecting cell type-specific, comprehensive mRNA expression levels in astrocytes, neurons and oligodendrocytes. We used these gene lists to classify genes into “astrocytic” or “neuronal” categories. When compared with the “neuronal” category at all examined time points, the “astrocytic” category included a higher average number of genes that underwent a change in expression of more than ±150% (Fig. 1c). Comparison between the results at 8 and 48 h after treatment showed an increase in the average number of genes that reached 150% change in both groups (34 vs. 40 for astrocytes and 21 vs. 28 for neurons, 8 and 48 h after treatment, respectively). We also examined the expression levels of genes reported as over-expressed in reactive astrocytes (Ridet et al., 1997) and found a large overlap with over-expressed genes 8, 24 and 48 h following both treatments (Fig. 1d). These results are consistent with the hypothesis that an early and prominent change in astrocytic gene expression is an important early feature of BBB-breakdown or albumin-induced epileptogenesis (see Discussion).

Altered expression of astrocytic potassium and glutamate regulating genes

In their pioneering study, Kuffler and Potter (1964) established that astrocytes are crucial for the control of the brain’s extracellular environment. Specifically, these cells limit the accumulation of [K+]o and glutamate (Oliet et al., 2001; Newman et al., 2004), thus potentially contributing to the regulation of neuronal excitability. We therefore searched our gene array results for changes in the level of expression of several potassium and glutamate homeostasis-related genes. We found that transcripts coding for the predominantly astrocytic (Kcnj10), but not neuronal (e.g. Kcnj2 or Kcnc1, see Butt and Kalsi, 2006), inward-rectifying K+ channel (KIR) were down-regulated. In addition, the mRNA coding for the astrocytic glutamate transporters of the solute carrier family 1, subfamily A members SLC1A2 and SLC1A3 (see Su et al., 2003; Chaudhry et al., 1995), but not for SLC1A4, was down-regulated. In contrast, SLC1A1 (preferentially expressed in neurons; see Rothstein et al., 1994) did not show significant changes in expression levels. Glutaminase (Gls, Gls2) and glutamine synthetase (GS), both of which are predominantly expressed in astrocytes (Derouiche and Frotscher, 1991) and are responsible for regulating glutamate levels, were also down-regulated (Fig. 2a). Furthermore, our gene arrays showed that at most time points there was a significant down-regulation of gap junction proteins (Gja1, Gjb2, Gjb6) 24 h following treatment, a finding that indicates reduced spatial buffering capacity (see Wallraff et al, 2006). Real-time RT-PCR confirmed the main observations obtained from the gene arrays, i.e., significant up-regulation of GFAP, and down-regulation of KCNJ10 (KIR 4.1, data not shown and see Fig. 5 in Ivens et al. 2007) as well as KCNJ3, SLC1A2 and SLC1A3, Gja1, Gjb2 and Gjb6 at all time points (connexins 43, 26 and 30, respectively, Fig. 2b). In contrast, glutamine synthetase did not show significant down-regulation (Fig. 2b).

Figure 2
Alterations in astrocytic potassium and glutamate regulating genes

The microarray results hinted at a rapid and robust change in astrocytic gene expression in vivo following BBB breakdown or brain exposure to serum albumin. To further validate the specificity of the astrocytic response to albumin, we exposed cell cultures enriched with either astrocytes or neurons (see Methods) to albumin for 24 h. Significantly, the astrocytic cultures responded with significant down-regulation of the same transcripts found to respond in vivo to albumin (SLC1A3, GS, Gja1, Gjb2 and Gjb6, Fig. 2c). No significant differences in expression levels of the same transcripts were found in the neuronal-enriched culture (except for downregulation of GS and upregulation of Gjb6 mRNA levels, Fig. 2d), supporting the notion that the changes observed in-vivo do indeed reflect an astrocytic response.

Does epileptogenesis involve reduced glutamate and potassium clearance?

To confirm that the transcriptional changes induced by albumin were associated with altered cellular functions, we investigated the clearance of extracellular glutamate and potassium in cortical slices 24 hours following albumin treatment in vivo. To measure synaptic glutamate levels during neuronal activation, we recorded the slowly inactivating (Lester et al., 1990) NMDA currents in cortical neurons by using the whole-cell patch configuration (in the presence of non-NMDA glutamate and GABA receptor blockers, see Methods). Cells were clamped at +40 mV to prevent a potential confounding effect of post-synaptic depolarization due to the accumulation of synaptic [K+]o. Mean single EPSC rise-time and amplitude were similar in both control and albumin-treated groups [14.5±0.5 vs. 13.2±0.7 ms and 505±100 vs. 492±140 pA, for rise-time (not shown) and amplitude, respectively in treated vs. controls, Fig. 3a, inset], suggesting that no changes in post-synaptic NMDA receptor density or properties at this time point (data not shown). We measured synaptic glutamate elicited by 50 extracellular stimulations at 2, 5, 10, 20, 50 and 100 Hz before and after adding the astrocytic SLC1A2 specific inhibitor, DHK. In neurons from control animals, DHK had no effect on single EPSCs or EPSCs elicited at low stimulation frequencies (<20 Hz). In contrast, stimulation frequencies > 20 Hz resulted in increased NMDA currents (or reduced depression when normalized to the first stimulus, Fig 3b–c, left), suggesting that astrocytic glutamate transporters efficiently reduce synaptic glutamate levels at high frequencies of neuronal activation. The same experiments were then repeated 24 h following cortical application of albumin (i.e. during epileptogenesis). In contrast to the control experiments, DHK had no effect on EPSC amplitude in treated slices (Fig. 3b–c, right), supporting reduced expression of the astrocytic transporter SLC1A2. Repetitive stimulation, however, resulted in a stronger depression of EPSC amplitude in treated slices as compared to controls (see Discussion).

Figure 3
Electrophysiological evidence for reduced glutamate and potassium buffering during epileptogenesis

To study K+ clearance from the extracellular space, we recorded from control and treated slices (24 h following treatment with albumin) by using ISMEs. We previously reported slower decay kinetics of [K+]o in response to pressure application in BBB-treated animals (Ivens et al., 2007). Here we tested for [K+]o accumulation during neuronal activation at different frequencies of stimulation. In slices from control animals, the increase in [K+]o was limited to 25% of baseline levels (<3.75mM) at all stimulation frequencies with the employed stimulation intensities and number of stimuli. In contrast, in treated slices [K+]o accumulation was significantly higher at frequencies ≥ 10 Hz, reaching 6.7 mM (Fig. 3d–e).

Modeling reduced K+ clearance results in frequency-dependent facilitation of excitatory post-synaptic potentials

To elucidate the possible contribution of astrocytic dysfunction to neuronal excitability, we developed a NEURON-based model of a post-synaptic neuron and an astrocyte. To evaluate the role of increased [K+]o accumulation and glutamate accumulation, we focused on examining changes to excitatory synaptic currents in the post-synaptic neuron (see Methods). Excitatory synaptic input was simulated by simultaneous application of glutamate at all 8 distal dendritic processes (Fig. 4a). In light of our experimental data, we simulated the reduction in K+ clearance by manipulating a [K+]o-regulated potassium removal mechanism (IKIR), while keeping the diffusion component constant. In the absence of neuronal activity, reducing KIR-mediated potassium clearance had no effect on resting [K+]o and thus had a negligible effect on the rising phase and maximal amplitude of a single excitatory post-synaptic potential (EPSP) (Fig. 4b). Reducing potassium buffering and consequent increased K+ accumulation during repetitive stimulation resulted in enhanced EPSP duration due to slower repolarization (due to a reduced driving force for K+ and a slight increase in NMDA-mediated current, see below and Fig. 4b). During repetitive activation, the accumulation of [K+]o near the dendritic compartment reached a maximum of 8.7 (and 16 mM) for reduction to 50% (and 10%) of astrocytic [K+]o buffering capacity, respectively (Fig. 4c). [K+]o accumulation during repetitive stimulation had a differential effect on AMPA- and NMDA-mediated currents: while the AMPA current showed frequency-dependent depression due to receptor desensitization, the NMDA component was strongly facilitated due to membrane depolarization (Fig. 4b). Reducing astrocytic potassium uptake from the extracellular space to 10% of control values resulted in an increase in total charge transfer mediated by the NMDA component of 44, 344 and 84% at 10, 20 and 100 Hz, respectively, while the AMPA charge transfer decreased by 5, 24, and 15%, respectively (Fig. 4d). Overall, there was a frequency-dependent increase in EPSP amplitude (Fig. 4e) associated with longer decay time (Fig. 4f). Repeated simulations with no NMDA conductance (GNMDA = 0, with concomitant increased AMPA conductance, to achieve similar depolarization for a single stimulus) resulted in a much smaller facilitation (compare Fig. 4g and h).

Figure 4
Application of NEURON-based model to determine the effects of [K+]o accumulation

Modeling reduced glutamate clearance results in frequency-dependent depression of excitatory post-synaptic potentials

We next used the NEURON model and simulation paradigms described above to test the expected effect of reduced glutamate uptake. We simulated the reduction in glutamate uptake by slowing the transmitter’s synaptic decay function. A twofold increase in the glutamate decay time constant resulted in a 48% increase in EPSP amplitude (from 25 to 37 mV) at a single post-synaptic dendrite and a 60% increase in the amplitude of the summated somatic EPSP (Fig. 5a). While for a single stimulation both AMPA and NMDA-components were increased, with repetitive activation, a marked decrease in the AMPA current (due to receptor desensitization, see Otis et al., 1996) and a strong facilitation of the NMDA current (due to post-synaptic depolarization, see Mayer et al., 1984 and Fig. 5b-c) were measured. Somatic EPSP facilitation (ratio of 5th to 1st EPSP amplitude, Fig. 5c) was maximal at 100 Hz with our initial conditions for glutamate clearance. Inhibiting glutamate clearance did not affect EPSP facilitation at low stimulation frequencies (<20 Hz) but reduced it at high stimulation frequencies (>80 Hz). The decreased facilitation was due to a reduced AMPA current through the desensitized receptors, thus keeping the membrane potential below the threshold for NMDA receptor activation. In simulations performed in the absence of NMDA conductance, EPSP facilitation was reduced at most stimulation frequencies, with only a small (<150%) residual facilitation measured at high stimulation frequencies (>100 Hz, Fig. 5d).

Figure 5
Application of NEURON-based model to determine the effects of glutamate accumulation

Modeling the concerted effect of reduced potassium and glutamate clearance

The simulations showed that while synaptic glutamate levels mainly affected the 1st EPSP in the train, an activity-dependent increase in [K+]o mainly enhanced EPSP facilitation in a frequency-dependent manner. Since our molecular data indicated a decrease in both potassiumand glutamate buffering mechanisms, we simulated their joint effect on synaptic transmission. Decreasing the clearance of [K+]o led to maximal EPSP facilitation when stimulating at 20 Hz, while a concurrent twofold reduction in glutamate uptake shifted the optimal frequency for maximal facilitation to 10 Hz (Fig. 6a). Concurrent reductions in glutamate and [K+]o clearance led to increases in the duration of the 1st EPSP, which in turn elicited increased and longer NMDA receptor activation per stimulus. The longer EPSPs allowed for a larger charge transfer with longer inter-stimulus intervals (i.e., reduced frequency, Fig. 6b) thus lowering the optimal stimulation frequency. To assess the sensitivity of the synaptic response during repetitive stimulation (at 20 Hz), we used several glutamate decay time constants and varying levels of [K+]o uptake. We plotted the maximal EPSP amplitude as a function of [K+]o; Figure 6c demonstrates that increasing synaptic glutamate led to small increases in the maximal EPSP amplitudes for all levels of [K+]o. However, synaptic facilitation was decreased with reduced glutamate uptake: thus, synaptic [K+]o accumulation to 10 mM was associated with 40% EPSP facilitation (upon the 5th stimulation) under baseline glutamate clearance, but with only 22% facilitation when glutamate decay time was doubled (Fig. 6d).

Figure 6
Modeling the concerted effect of reduced potassium and glutamate clearance

Electrophysiological evidence for frequency-dependent synaptic facilitation during epileptogenesis

Our simulation data predicted maximal EPSP facilitation at 20 Hz when [K+]o clearance is reduced and decreased facilitation (at 50–100 Hz) when the only change induced is glutamate accumulation in the synaptic cleft. We therefore measured field potentials in response to stimulation at various frequencies in brain slices during “epileptogenesis” (exposure to albumin in sACSF) compared to controls (sACSF alone). Comparison of the field potential amplitude and absolute integral during the first five stimuli revealed a significant reduction in both measures only under 100 Hz stimulation [amplitude: 1.13 ± 0.12 vs. 0.46 ± 0.03 mV, 1.44 ± 0.36 vs. 0.47 ± 0.09 mV and area: 2.4 ± 0.2 vs. 0.9 ± 0.02 V*s, 4.4 ± 1.1 vs. 0.5 ± 0.7 V*s, 1st vs. 5th stimulus, control (n = 5) and treated (n = 4), respectively, p<0.05]. Comparing field potential duration (measured at 1/3 maximal amplitude) for the 1st vs. the 5th stimulus among different frequencies did not reveal any changes in control slices. In contrast, in treated slices the field potential was significantly prolonged at 10 and 20 Hz (10 Hz: 7.5 ± 0.4 vs. 9.4 ± 5.5 ms, 6.5 ± 0.7 vs. 13.1 ± 2.6 ms, 20 Hz: 6.0 ± 0.9 vs. 6.8 ± 0.8 ms, 6.6 ± 0.8 vs. 12.9 ± 2.9 ms for 1st vs. 5th stimulus in control and treated, respectively, p<0.05, Fig. 7c). Interestingly, in the “treated” group, stimulation-induced frequency-dependent, long-lasting epileptiform discharges occurred most reliably during 10-Hz stimulation (4 of 4 slices, n = 3 animals), and sometimes at 20 Hz (3 of 4 slices) and 5 Hz (2 of 4 slices), but never at higher frequencies (Fig. 7d). Epileptiform discharges were observed in one control slice without any apparent frequency dependence (5 to 50 Hz, 1 of 5 slices, n = 3 animals, Fig. 7d).

Figure 7
Recording in vitro shows frequency-dependent increased neuronal excitability and hyper-synchronous network activity during albumin-mediated epileptogenesis

Taken together, our experiments show that exposure to albumin in-vitro induces changes in neuronal excitability and that evoked network activity facilitates, and often turns into, robust epileptiform discharges upon repetitive stimulation. We found that 10–20 Hz is the most reliable frequency, as was also predicted by our K+ recording data (Fig. 3d) and by our model in the case of reduction in [K+]o clearance with or without glutamate accumulation (see Fig. 4g and Discussion).


The primary goal of the present study was to study the role of astrocytes in epileptogenesis. With the BBB disruption and albumin-induced models of epileptogenesis, the following findings were obtained early during epileptogenesis: (1) Similar significant changes in astrocytic gene expression occurred following the two treatments. (2) Transcriptional data predicted disturbed homeostasis of extracellular potassium and glutamate. (3) Intracellular recordings confirmed reduced astrocytic glutamate uptake, which did not, however, seem to account for increased neuronal excitability. (4) Recordings with ISMEs confirmed activity-dependent impaired [K+]o buffering. (5) A NEURON-based model predicted that reduced [K+]o buffering leads to frequency- and NMDA-R dependent facilitation of EPSPs, with maximal facilitation around 20Hz, while reduced clearance of glutamate results in a modest increase in the amplitude and duration of a single EPSP with decreased facilitation during repetitive stimulation. (6) Finally, extracellular recordings in cortical slices confirmed the frequency-dependent facilitation of synaptic activity predicted by the model and showed epileptiform discharges at preferred stimulation frequencies of 10–20 Hz. Overall, the present study suggests a key role for reduced astrocytic-mediated clearance of [K+]o in activity-dependent facilitation of synaptic activity during epileptogenesis.

The working hypothesis for the present study is based on our previous experimental results, which demonstrated that BBB breakdown induces epileptogenesis (Seiffert et al., 2004; Ivens et al., 2007; Tomkins et al., 2007) and increases the expression of the astrocytic marker, GFAP, within hours following the epileptogenic treatment. This rapid astrocytic response was observed before the emergence of epileptiform activity, leading to the hypothesis that astrocytes play a role in the epileptogenic process (see also Tian et al., 2005; Ding et al., 2007). Indeed, changes in the structure and function of astrocytes are found in a wide variety of brain insults, including epilepsy, in both animals and man (Bordey et al., 2001; Kivi et al., 2000; Herman, 2002; Jauch et al., 2002; Schroder et al., 1999). However, the role of astrocytic dysfunction in disease progression and neuronal dysfunction is not well understood (for reviews see Heinemann et al., 1999, Seifert et al., 2006 and Schwarcz, 2008). Since BBB breakdown causes albumin extravasation from brain vessels and its specific uptake by astrocytes (Ivens et al., 2007), we put forward the hypothesis that under BBB breakdown astrocytic gene expression is directly modulated by albumin. A role for serum albumin in epileptogenesis is also supported by experiments showing that albumin induced focal epileptiform activity in a dose-dependent manner (Seiffert et al., 2004). Importantly, albumin-induced epileptogenesis is observed only after a window period of several hours (in vitro) or days (in vivo), with no apparent effect on neuronal membrane characteristics, firing properties or amplitude and duration of single EPSCs (Seiffert 2004, Fig. 3 and data not shown). Furthermore, the specific uptake of albumin by astrocytes, together with the rapid changes in the level of astrocyte-specific proteins (Ivens et al., 2007), led us to hypothesize that a concerted astrocytic transcriptional response may underlie epileptogenesis under these conditions. Using microarray technology we confirmed that a large number of astrocytic genes do show significant changes in expression levels as early as 8 h following an epileptogenic event, several days before epileptic activity emerges (Seiffert et al., 2004; Ivens et al., 2007). However, it is plausible that under pathological conditions, different cell populations may express previously unexpressed transcripts, thus confounding our results. Nevertheless, the cell-specific mRNA changes in response to albumin exposure that were evident in our astrocytic- and neuronal-enriched cultures constitute further support for our conclusion that the observed changes do occur preferentially in astrocytes (Fig. 2). The cluster analysis and the strikingly high correlation between expression profiles of both BBB breakdown and albumin treatments at the various time points support the notion that the extravasation of the most abundant serum protein, albumin, through the injured vessels plays a role in the transcriptional modulation of astrocytic genes. An alternative hypothesis – that albumin itself is disruptive to the BBB, leading to the extravasation of some other blood-derived mediator – is less likely, since local application of albumin did not increase BBB permeability to large molecules, as previously measured using systemic injection of Evans-blue (Seiffert et al., 2004).

Since astrocytes are known to be key contributors to [K+]o buffering, we searched our microarrays for expression levels of astrocytic K+ channels (Barres et al., 1990). Indeed, Kir 4.1 (but not other K+ channels, see Fig. 2a), previously shown to be expressed in neocortical astrocytes (Hibino et al., 2004; Higashi et al., 2001), was down-regulated, leading to activity- dependent accumulation of [K+]o in treated cortical slices (Fig. 3 and see Ivens et al., 2007). Reduced [K+]o clearance has been reported in the injured brain (D’Ambrosio et al., 1999), and a loss of IKIR has been found in reactive astrocytes around freeze lesions (Bordey et al., 2001), after ischemic insults (Koller et al., 2000) and direct injuries (Schroder et al., 1999), and in epileptic Tsc1 knock-out mice (Jansen et al., 2005) as well as in human subjects with temporal lobe epilepsy (TLE, Bordey and Sontheimer, 1998; Hinterkeuser et al., 2000; Kivi et al., 2000; Jauch et al., 2002). It is noteworthy that KIR 4.1 knock-out mice display seizure activity very early in life consistent with the idea that down regulation of KIR 4.1 channels may contribute to epileptogenesis (Djukic et al., 2007). The hypothesis that elevated [K+]o could lead to seizure initiation (Fertziger and Ranck, 1970) is not a new one; however, in this study we show that a selective down-regulation of the KIR 4.1 channel occurs prior to the emergence of epileptic activity, thus highlighting the potential role of potassium accumulation in epileptogenesis. To what extent KIR channels contribute to the spatial buffering of [K+]o is not entirely known and may differ between brain regions. In hippocampal slices, low concentrations of Ba2+ augmented stimulus-induced K+ by 147% (to more than 9mM, Gabriel et al., 1998), while in the neocortex, Ba2+ slowed down the clearance of iontophoretically applied K+ by only 70% (Ivens et al., 2007). Notably, in both preparations, epileptogenesis was associated with a reduced effect of Ba+2 indicating reduced IKIR. The relatively high [K+]o found during stimulation in the neocrotex in our study may reflect altered extracellular space volume or impairments in additional buffering mechanisms such as reduced expression of leak K+ channels (not supported by our microarray but see Pasler et al., 2007) and/or gap junction proteins (which were found to be down regulated in our molecular experiments, see figure 2). However, in connexin knockout mice (which lack gap junctions in the hippocampus), potassium clearance capacity seems almost conserved (Wallraff et al., 2006). In addition, reduced [K+]o clearance is expected to enhance potassium accumulation due to delayed neuronal repolarization and facilitated synaptic potentials (Fig. 4). Moreover, the rectification properties of KIR channels predict that their contribution to K+ clearance becomes critical when local [K+]o are high, thus further augmenting their importance during high frequency stimulation (Fig. 3d and see Chen and Nicholson, 2000; Newman, 1993).

Due to the lack of pharmacological tools that specifically block astrocytic KIR channels, it is difficult to experimentally its role in controlling neuronal excitability. We therefore used computer simulations that predicted that synaptic accumulation of potassiumwill lead to frequency-dependent facilitation of EPSPs. Importantly, [K+]o accumulation under these conditions did not have an effect on a single EPSC, consistent with the observation of normal field potentials during epileptogenesis (Seiffert et al., 2004). In contrast, EPSCs were strongly facilitated at stimulation frequencies between 10–50 Hz (maximum around 20 Hz) due to membrane depolarization and increased NMDA conductance. Under these conditions, facilitation (>300%) was associated with synaptic [K+]o levels reaching ~8 mM. This value may seem high considering that during normal neuronal activity (Heinemann et al., 1990) or that recorded in the present study (Fig. 3). Yet these levels were measured using ISMEs at a distance from the narrow synaptic cleft (<200 Å, Egelman and Montague, 1999; Savtchenko et al., 2000). In fact, the model predicts that during neuronal activity, [K+]o levels in the extracellular space near the soma are three times lower (due to diffusion and astrocytic uptake) than levels in the synaptic cleft; in line with the results of Somjen and colleagues (2008).

In addition to the “potassium hypothesis”, our molecular experiments point to reduced expression of astrocytic glutamate transporters in both albumin- and DOC-treated rats. In the normal brain, passive diffusion and transport clears released glutamate into neurons (via members of the solute carrier protein family - SLC1A) and astrocytes (specifically SLC1A2, SLC1A3 and SLC1A4). The relative contribution of each of these mechanisms in the neocortex is unknown. Studies in the hippocampus indicated that astrocytes play a central role in the removal of synaptic glutamate (Bergles and Jahr, 1998) whereas neuronal transport plays a negligible role (Sarantis et al., 1993); leading to the hypothesis that down-regulation of astrocytic glutamate transporters entails glutamate accumulation in the synaptic cleft and increased excitability. A reduction in glutamate uptake mechanisms has previously been reported in various neurodegenerative diseases frequently associated with BBB breakdown (e.g., Zlokovic, 2008; Rothstein et al., 1992). However, evidence for the role of glutamate uptake mechanisms in epilepsy remains inconclusive (Eid et al., 2008). Although Tanaka et al. (1997) demonstrated spontaneous seizures in SLC1A2 knock-out mice, studies of tissue from human TLE patients have failed to provide conclusive evidence for reduced glutamate uptake (Proper et al., 2002; Tessler et al., 1999). Moreover, the consequences of reduced glutamate uptake are still controversial, with considerable heterogeneity between various brain regions and preparations (Hestrin et al., 1990; Sarantis et al., 1993; Turecek and Trussell, 2000; Arnth-Jensen et al., 2002). To determine the functional consequences of the observed transcriptional response on glutamate homeostatic mechanisms, we recorded whole-cell glutamatergic currents before and after application of DHK, a selective inhibitor of the astrocytic glutamate transporter SLC1A2 (Arriza et al., 1994; Rothstein et al., 1994) in slices from control and treated rats. In agreement with previous studies, we found no effect for DHK on single EPSCs or EPSCs evoked by low-frequency stimulation (Hestrin et al., 1990; Diamond and Jahr, 2000; Takayasu et al., 2004). We did, however, find increased glutamatergic currents under DHK with stimulation frequencies higher than 20 Hz in control slices, similar to the effect of TBOA in the hippocampus (Arnth-Jensen et al., 2002). In contrast, in slices from albumin-treated animals, DHK had no effect on EPSCs, supporting the observed down-regulation of SLC1A2. Yet, despite the reduced DHK effect, we did not observe an increase in EPSCs 24 h following the epileptogenic insult. This finding may be explained by activation of compensatory mechanisms (e.g., up-regulation of neuronal glutamate uptake proteins, see Fig. 2a), excessive activation of pre-synaptic mGluR2 (Scanziani et al., 1997; Iserhot et al., 2004) or altered expression of NMDA receptor subunits.

Our computer model predicted that reduced glutamate uptake alone will result in reduced synaptic facilitation due to desensitization of AMPA receptors. These predictions are in agreement with previous studies in brainstem slices showing a frequency-dependent decrease in AMPA currents after incubation with the glutamate uptake blockers, THA and DHK (Turecek and Trussell, 2000), but stand in contrast to recordings in hippocampal neurons showing increased NMDA-mediated EPSPs under similar conditions (Arnth-Jensen et al., 2002). The difference may be because the latter study was conducted with holding voltages of +40 mV, resulting in a higher open probability of NMDA-R operated ion channels. While our molecular and electrophysiological experiments do indicate a reduction in astrocytic glutamate uptake during the early stages of epileptogenesis, both the electrophysiological data and computer simulations suggest that only at high stimulation frequencies (>100 Hz), does sufficient transmitter accumulate to facilitate EPSCs (see below).

Changes in astrocytic membrane potential were not implemented in our model. Extracellular accumulation of potassiumresults in astrocytic depolarization, which inhibits glutamate uptake (or even causes a reversal of the uptake mechanism; see Szatkowski et al., 1990). Simulating a concurrent reduction in both uptake mechanisms, resulted in a reduction of the stimulation frequency at which maximal facilitation occurs (from ~ 20 to 10 Hz). The simulations predicted that when both [K+]o and glutamate accumulate in the synapse, more NMDA receptors are activated at any given [K+]o (Fig. 6). In addition, we expect such depolarization to further reduce the cationic-coupled glutamatergic transport, thus enhancing voltage-dependent NMDA currents. The optimal frequency for synaptic facilitation recorded in brain slices exposed to albumin in sACSF was around 20 Hz, which as predicted by the model, was mainly due to reduced clearance of [K+]o. This stimulation frequency – 20 Hz – also proved to be the minimal stimulation frequency at which we measured [K+]o accumulation during epileptogenesis. It is striking that repetitive stimulation under these conditions often resulted in seconds-long, seizure-like activity, highlighting the potential role for BBB breakdown and brain exposure to serum albumin in neuronal hypersynchronicity, enhanced excitability and the generation of seizures. A plausible hypothesis would be that the increased, repeated activation of NMDA receptors leads to non-specific synaptic plasticity, thus strengthening excitatory synapses and causing persistent hyperexcitability (Li and Prince, 2002; Shao and Dudek, 2004). This premise also provides a satisfactory explanation for the efficacy of NMDA-R antagonists in improving cortical function in animal studies of brain injury and stroke – conditions in which the BBB is frequently impaired (Hickenbottom and Grotta, 1998; Sonkusare et al., 2005) and seizures are often observed. While further studies are needed to confirm this hypothesis, we propose astrocytic reaction in the injured cortex, and specifically impaired buffering of extracellular K+, as novel targets for the prevention and treatment of injury-related neocortical epilepsies.


This study was supported by the Sonderforschungsbereich TR3 (AF and UH), the German-Israel Foundation for Scientific Research and Development (AF), the Israel Science Foundation (566/07, AF), Minerva Stiftung (YD), the Israel-USA Binational Science foundation (AF and DK), and the CURE foundation (DK and AF).

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