Brain Advance Access originally published online on June 15, 2005
Brain 2005 128(10):2396-2407; doi:10.1093/brain/awh568
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Metabolic dysfunction during neuronal activation in the ex vivo hippocampus from chronic epileptic rats and humans
1 Institute for Neurophysiology and 2 Department of Neurosurgery, Charité—Universitätsmedizin Berlin, Germany and 3 Department of Neurochemistry, Institute of Biomolecular Chemistry, Budapest, Hungary
Correspondence to: Dr. med. Oliver Kann, Institut für Neurophysiologie, Charité—Universitätsmedizin Berlin, Tucholskystrasse 2, 10117 Berlin, Germany E-mail: oliver.kann{at}charite.de
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Summary |
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Metabolic dysfunction has been implicated in the pathogenesis of temporal lobe epilepsy (TLE), but its manifestation during neuronal activation in the ex vivo hippocampus from TLE patients has not been shown. We characterized metabolic and mitochondrial functions in acute hippocampal slices from pilocarpine-treated, chronic epileptic rats and from pharmaco-resistant TLE patients. Recordings of NAD(P)H fluorescence indicated the status of cellular energy metabolism, and simultaneous monitoring of extracellular potassium concentration ([K+]o) allowed us to control the induction of neuronal activation. In control rats, electrical stimulation elicited biphasic NAD(P)H fluorescence transients that were characterized by a brief initial ‘drop’ and a subsequent prolonged ‘overshoot’ correlating to enhanced NAD(P)+ reduction. In chronic epileptic rats, overshoots were significantly smaller in area CA1, but not in the subiculum as compared to controls. In TLE patients, who were histopathologically classified in groups with and without Ammon's horn sclerosis (AHS, non-AHS), large drops and very small overshoots of NAD(P)H transients were observed in dentate gyrus, CA3, CA1 and subiculum. Nevertheless, monitoring mitochondrial membrane potential (
m) by mitochondria-specific, voltage-sensitive dye (rhodamine-123) revealed similar mitochondrial responses during neuronal activation with glutamate and protonophore application in area CA1 of control and chronic-epileptic rats. Applying confocal laser scanning microscopy, these findings were confirmed in individual neurons of AHS tissue, indicating a negative m and activation-dependent mitochondrial depolarization. Our data demonstrate severe metabolic dysfunction during neuronal activation in the hippocampus from chronic epileptic rats and humans, although mitochondria maintain negative m. Thus, our findings provide a cellular correlate for ‘hypometabolism’ as described for epilepsy patients and suggest mitochondrial enzyme defects in TLE.
Key Words: hypometabolism; mitochondria; NADPH; potassium [K]; temporal lobe epilepsy
Abbreviations:
m = mitochondrial membrane potential; ACSF = artificial cerebrospinal fluid; AHS = Ammon's horn sclerosis; CCCP = cyanide m-chlorophenyl hydrazone; [K+]o = extracellular potassium concentration; TLE = temporal lobe epilepsy
Received April 11, 2005. Revised May 15, 2005. Accepted May 19, 2005.
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Introduction |
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Temporal lobe epilepsy (TLE) is a prevalent form of focal epilepsy that is frequently resistant to anticonvulsants and, in some patients, progressive in nature (Engel, 2001
; Pitkänen and Sutula, 2002). The mechanisms underlying the pathogenesis of TLE remain unclear.
Hippocampal tissue from TLE patients is often characterized by neuronal loss and astrogliosis (Ammon's horn sclerosis, AHS) (Margerison and Corsellis, 1966; Babb et al., 1984), network reorganization (Sutula et al., 1989; Parent and Lowenstein, 1997), as well as alterations of receptor and ion channel functions (Steinhäuser and Seifert, 2002; Avanzini and Franceschetti, 2003). In addition, functional neuroimaging studies demonstrated a decrease in glucose utilization (‘hypometabolism’) in seizure foci and adjacent brain structures of epilepsy patients (Kuhl et al., 1980; O'Brien et al., 1997; Casse et al., 2002), which might be explained by dysfunction of mitochondrial oxidative and/or glycolytic energy metabolism. However, no record exists of detailed investigation of metabolic and mitochondrial functions during neuronal activation in the ex vivo hippocampus from TLE patients.
In the brain, the functional integrity of mitochondria is essential because these organelles generate most of the ATP that is used for ion transport, maintenance of excitability and neurotransmission (Ames, 2000). In mitochondria, reduced nucleotides, NADH and FADH2, mainly serve in energy transfer and connect dehydrogenase activity within the Krebs–Szentgyörgyi cycle to oxidative phosphorylation (McCormack et al., 1990; Hansford and Zorov, 1998). Monitoring NAD(P)H fluorescence provides insight into the status of both oxidative and glycolytic energy metabolism, e.g. in brain slice preparations (Lipton, 1973; Schuchmann et al., 2001; Kann et al., 2003a, b; Shuttleworth et al., 2003).
Neuronal activation results in the release of potassium to extracellular space (Heinemann et al., 1990). Electrophysiological recordings of the extracellular potassium concentration ([K+]o) indicate the degree of local activity from neurons neighbouring the recording site (Heinemann and Lux, 1975). Thus, simultaneous recordings of [K+]o and NAD(P)H fluorescence have been used to examine the coupling of neuronal activity and energy metabolism (Lewis and Schuette, 1975; Kann et al., 2003a, b).
The present study was designed to explore cellular metabolic and mitochondrial functions in the ex vivo hippocampus from pharmaco-resistant TLE patients during neuronal activation that was elicited by electrical stimulation or glutamate application. To be able to determine alterations of stimulus-induced NAD(P)H fluorescence transients, we controlled the induction of neuronal activation in tissue with varying degrees of neuronal cell loss by simultaneous monitoring of [K+]o. Using the cationic dye, Rhodamine-123 we examined the inner membrane potential of mitochondria (m) in slices and individual neurons. Because of the lack of human control tissue, we related our findings from TLE patients to tissue from chronic epileptic rats utilizing the ‘pilocarpine model’ of experimental TLE (Cavalheiro et al., 1991; Mello et al., 1993).
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Methods |
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Pilocarpine-treated rats and TLE patients
Status epilepticus was induced in male Wistar rats (115–130 g) by injection of pilocarpine hydrochloride (320 mg/kg, i.p.; Sigma-Aldrich, Taufkirchen, Germany) 30 min after pre-treatment with scopolamine hydrobromide (1 mg/kg, s.c.; Sigma-Aldrich) to reduce peripheral cholinergic effects. After 1.5 h, status epilepticus was terminated by diazepam injection (5–10 mg/kg, i.p.; Ratiopharm, Ulm, Germany). After 90 and 210 min, Ringer solution supplemented with lactate (27.2 mM; Sigma-Aldrich) and sodium hydrogen carbonate (71 mM; Sigma-Aldrich) was injected (i.p.) to improve survival. Control rats were treated with the same protocol, but with saline instead of pilocarpine. Rats were kept in separate cages under standard housing conditions. All rats were monitored by video recordings to assure development of spontaneous epileptic activity: a seizure-free interval of two to three weeks was followed by spontaneous recurrent seizures that lasted as long as the animals were alive (Mello et al., 1993
; Lehmann et al., 2001). Recurrent seizures were characterized by oral automatism, head and bilateral forelimb myoclonus, rearing, falling and loss of consciousness. Seizure frequency of pilocarpine-treated rats was 0.7 ± 0.4 (n = 8) per day. Rats (n = 15) were killed 7.7 ± 0.9 months after pilocarpine-induced status epilepticus (and sham treatment). Animal experiments were conducted in accordance with the guidelines of the European Communities Council and approved by the Regional Berlin Animal Ethics Committee (G0328/98, G0024/04).
All the 16 patients with pharmaco-resistant TLE involved in this study had undergone partial unilateral resection of the hippocampus (performed by T.-N. Lehmann), i.e. of the anterior half of the hippocampal body 
20 mm posterior from the tip of the inferior horn of the lateral ventricle. The diagnosis was made in the Epilepsy Center of Berlin-Brandenburg (Herzberge) according to European Guidelines for pre-surgical evaluation (European Federation of Neurological Societies Task Force, 2000). The study was approved by the Ethics Committee at Charité (EA125/2001, EA1/042/04) and in accordance with the ‘Declaration of Helsinki’. Each patient gave an informed consent in written form for studies on resected tissue.
After histopathological diagnosis (Institute for Neuropathology, Charité), tissue from TLE patients was assigned to a group characterized by AHS (Wyler grades III and IV) and a group without AHS (non-AHS group; Wyler grades I and II) (Wyler et al., 1992).
Tissue preparation
Resected human hippocampal tissue was immediately immersed in oxygenated solution (95% O2, 5% CO2, 4°C) and cut in tissue blocks of 2–4 mm to facilitate oxygen penetration during transport (30 min). The solution contained in mM: KCl 3, NaH2PO4 1.25, glucose 10, MgSO4 2, MgCl2 2, CaCl2 1.6, NaHCO3 21, sucrose 200 and 
-tocopherol 0.1 (pH 7.4). Sodium was reduced to prevent hypoxia-induced Na+-influx to neurons while -tocopherol was added for scavenging free radicals. After transport, tissue blocks were immediately cut into slices. Using this approach, we were able to record neuronal responses and evoked epileptiform activity in resected tissue from >60 TLE patients over hours after surgery (Kivi et al., 2000; Gabriel et al., 2004). Rats were decapitated under deep diethylether anaesthesia and their brains were quickly removed. Horizontal and coronal hippocampal slices (450 µm) were prepared from rat and human material, respectively, in ice-cold, oxygenated (95% O2, 5% CO2) artificial cerebrospinal fluid (ACSF), using a vibratome (Campden, Leicester, UK) and directly transferred to an incubation chamber. Slices were incubated in ACSF for 2–4 h to recover from the preparation procedure (room temperature) before being used for experiments (superfusion with ACSF, 4 ml/min, 33–35°C). ACSF contained (in mM): NaCl 129, KCl 3, NaH2PO4 1.25, MgSO4 1.8, CaCl2 1.6, NaHCO3 21 and glucose 10 (pH 7.4). All salts and -tocopherol were from Sigma-Aldrich.
Measurements of [K+]o and microfluorimetry of NAD(P)H and rhodamine-123
DC-coupled recordings of field potentials and changes in [K+]o were performed with double-barrelled K+-sensitive and reference microelectrodes (Heinemann and Arens, 1992). In brief, electrodes were pulled from double-barrelled theta glass (Science Products, Hofheim, Germany). The reference barrel was filled with 154 mM NaCl solution, the ion-sensitive barrel with potassium ionophore I cocktail A (60031; Fluka Chemie, Buchs, Switzerland) and 100 mM KCl. Electrodes with a sensitivity of 59 ± 2 mV to a 10-fold increase in [K+] were used for experiments. The amplifier was equipped with negative capacitance feedback control, which permitted recordings of changes in [K+]o with time constants of 50–200 ms.
The recording electrode was positioned in the neuronal cell somata layer and in the centre of the fluorescence recording area (x20 objective, 0.5 numerical aperture; Zeiss, Jena, Germany). The neuronal compartment was activated by electrical stimulation (20 Hz, 10 s, 1–10 V) of stratum radiatum for recordings in CA3, CA1 and the subiculum as well as of the perforant path for recordings in the dentate gyrus (DG). The bipolar stimulation electrode was positioned 600–700 µm apart from the recording electrode to avoid direct depolarization of the astrocytic compartment within the recording area. NAD(P)H was excited at 360 nm (monochromator; Photon Technology Instruments, Wedel, Germany) and fluorescence emission was recorded at 460 nm using a photomultiplier (Seefelder Messtechnik, Seefeld, Germany) mounted on an epifluorescence microscope (Axioskop, Zeiss). In contrast to oxidized forms, reduced nicotinamide dinucleotides, NADH and NADPH, fluoresce when excited by ultraviolet light (Aubin, 1979). In mitochondria, NAD+ is reduced to NADH by several dehydrogenases, and NADP+ is reduced to NADPH by isocitrate dehydrogenase as well as by transhydrogenase activity at the expense of NADH (Jackson, 2003). In the cytosol, NAD+ is reduced within the second part of the glycolysis. Because the emission spectra of NADH and NADPH overlap, NAD(P)H indicates that recorded fluorescence might have originated from either one or both (Schuchmann et al., 2001). For microfluorimetric rhodamine-123 recordings in AHS tissue, slices from patients with numbers 14, 15 and 16 were bulk-loaded with rhodamine-123 for 10 min (Sigma-Aldrich; stock solution in ethanol, 5 µM final concentration) and incubated for another 60 min in ACSF at room temperature. Rhodamine-123 was excited at 490 nm and fluorescence emission was recorded at 515–530 nm.
Fluorescence data and changes in voltage of the recording electrode were digitized at 10 Hz using a standard PC and FeliX software (Photon Technology Instruments). Changes in NAD(P)H and rhodamine-123 fluorescence are presented as F/F0 in %. Rhodamine-123 fluorescence baseline shift was corrected offline. Changes in [K+]o are presented in mM. To translate the recorded potential values (mV) in [K+]o, a modified Nernst equation was used:
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Intracellular recordings and rhodamine-123 imaging
Slices from patients with numbers 3, 4, 5, 6 and 7 were fixed with a platinum wire in the recording chamber mounted on an epifluorescence microscope (BX50WI, Olympus, Hamburg, Germany) and superfused with ACSF at 4 ml/min. Patch pipettes (4–5 M
resistance) were pulled with a horizontal puller (Sutter Instruments Corp., Novato, CA, USA). The pipette solution contained (in mM): KGlu 135, NaCl 10, CaCl2 0.05, EGTA 1, HEPES 10, MgATP 3 (Sigma-Aldrich) at pH 7.3 (275–290 mOsmol/kg). Current-clamp recordings of neurons were performed in the whole-cell configuration using a patch-clamp amplifier, EPC 7 (Heka Elektronik GmbH, Lambrecht/Pfalz, Germany), IBM-compatible PC and WinTida 4.0 (Heka). Series resistance and input resistance were 12 ± 1.2 M and 48 ± 3.8 M (mean ± SE; n = 8), respectively.
Recordings of changes in m were performed as described (Schuchmann et al., 2000). In brief, membrane-permeable, cationic fluorescent dye, rhodamine-123 (10 µg/ml, stock solution in ethanol) was added to the pipette solution. The dye was allowed to diffuse to the cytoplasm for 20–30 min, after establishing the whole-cell patch. Due to its cationic nature, rhodamine-123 accumulates in respiring mitochondria that maintain a negative m. In mitochondria, fluorescence intensity of rhodamine-123 is quenched due to matrix protein binding. Upon mitochondrial depolarization, the dye is released to the cytoplasm and de-quenched resulting in a cytoplasmic increase in rhodamine-123 fluorescence intensity (Bindokas et al., 1998; Ward et al., 2000). Fluorescence recordings were performed using a NORAN Oz confocal laser scanning microscope (Prairie Technologies Inc., Middleton, WI, USA) with acquisition software (Intervision, Irix 6.3, Silicon Graphics, Mountain View, CA, USA). Rhodamine-123 was excited using the 488 nm line of a krypton–argon laser (Omnichrom 643, Melles Griot, Bensheim, Germany) with an intensity of <10 µW as determined under the objective. Fluorescence was measured with a water immersion objective (x60, 0.9 numerical aperture, Olympus, Hamburg, Germany). The pinhole was set to 50–100 µm and averaged (8–16 frames) 8-bit images were captured at 1–2 Hz. The focal plane contained the soma and primary dendrites of a neuron. Changes in rhodamine-123 fluorescence were evaluated offline using the ImageJ (RSB, NIMH, Bethesda, MD, USA). Changes in fluorescence were determined in the cytoplasm by setting regions of interest and are expressed as F/F0 in % after baseline correction (Kovács et al., 2005).
Cresyl violet staining
Slices from rats and humans were fixed in paraformaldehyde (4%, 0.1 M phosphate buffer; Sigma-Aldrich), after the preparation procedure. For further processing, fixed slices were incubated with sucrose solution (30%, 0.15 M phosphate buffer; Sigma-Aldrich) for 12–16 h, and 30 µm sections were prepared using a freezing microtom (Leica Microsystems, Wetzlar, Germany). Sections were exposed to an ethanol series (6 steps, 3 min each) and rinsed in distilled water. Sections were then immersed in cresyl violet solution (0.5% cresyl violet in distilled water; Sigma-Aldrich) for 90 s, rinsed in distilled water, and transferred to differentiation solution (200 ml, 50% ethanol, 3 drops of acetic acid). Subsequently, sections were exposed to a series of ethanol (4 steps, 3 min each), 2-propanol (two times; Sigma-Aldrich), and xylol (two times; Sigma-Aldrich) and finally embedded with DePeX (Serva Electrophoresis GmbH, Heidelberg, Germany). Sections were analysed using an upright Axioskop (Zeiss) equipped with a CCD camera (DX30; Kappa optoelectronics, Gleichen, Germany). Images were processed with ImageBase Control® (Kappa).
Statistics
Experimental data (n) from different areas of the hippocampus were obtained from slices of 8 pilocarpine-treated rats, 7 control rats, 11 AHS and 5 non-AHS patients. Results of a particular experiment were pooled and are given as mean ± SE. For a given parameter (e.g. overshoot in NAD(P)H transient), two groups, either of the rat preparation (control versus pilocarpine-treated rats) or of the human preparation (non-AHS versus AHS), were compared in a given region (e.g. area CA1) using one way ANOVA.
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Results |
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Histopathology of chronic epileptic tissue
Eight pilocarpine-treated rats that had developed spontaneous recurrent seizures (grade 5 on the Racine scale) (Racine, 1972
) were included in the present study. In these animals, neuronal cell loss occurs predominantly in the medial entorhinal cortex and in the hippocampal formation (Du et al., 1995). Within the hippocampal formation, the hilus, area CA3 and area CA1 are significantly more affected than the subiculum (Mello et al., 1993; Fig. 1A–D). These findings correspond well to histopathological alterations of AHS tissue from TLE patients (Babb et al., 1984; Blümcke et al., 1999) making pilocarpine-treated rats a useful and reliable model for TLE (Cavalheiro et al., 1991; Mello et al., 1993).
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Histopathology of tissue from pharmaco-resistant TLE patients revealed AHS (Wyler grades III and IV) in 11 and less neuronal cell loss (non-AHS; Wyler grades I and II) in 5 of the resected hippocampi (Fig. 1E–H). The age of the patients ranged from 19 to 56 years. Mean age and mean duration of TLE within the AHS group (non-AHS group) were 33.6 ± 4.0 (29.6 ± 3.9; P = 0.5) and 26.1 ± 3.6 (13.9 ± 5.2; P = 0.08) years, respectively. Table 1 contains the details on patient data.
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Alterations of NAD(P)H transients in chronic epileptic tissue
We recorded NAD(P)H fluorescence to investigate cellular energy metabolism during neuronal activation in different areas of acute hippocampal slices from rats and humans. Neuronal activation was elicited by electrical stimulation (10 s, 20 Hz) and resulted in transient increases in [K+]o of up to 2.3 mM (Fig. 2), corresponding to the upper range of in vivo responses to sensory and electrical stimuli (Heinemann et al., 1990
; Somjen, 1995).
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As expected, stimulus-induced neuronal activation was closely associated with characteristic biphasic NAD(P)H fluorescence transients in control rats. The transients consisted of a brief initial ‘drop’ that was followed by a prolonged ‘overshoot’ of up to several minutes (Fig. 2A). Such biphasic NAD(P)H transients have been described in a variety of neuronal preparations (Lipton, 1973
; Duchen, 1992; Schuchmann et al., 1998; Kann et al., 2003a, b). In contrast, stimulus-induced NAD(P)H transients were altered in chronic epileptic tissue. The overshoot was smaller in area CA1 from chronic epileptic rats (Fig. 2B) and absent in many experiments in tissue from TLE patients, although increases in [K+]o reflected substantial neuronal activation (Fig. 2C, D and F). In Fig. 3, we summarize the quantification of drops and overshoots as well as of increases in [K+]o in area CA1 and the subiculum of the rat preparation (control versus pilocarpine-treated group) and the human preparation (non-AHS versus AHS group). To be able to compare the groups, we included all recordings with increases in [K+]o of <1.6 mM. Notably, the overshoots were significantly smaller in area CA1 of the pilocarpine-treated group, while the drops were similar as compared to the control (Fig. 3A). Similar increases in [K+]o in both groups broadly excluded the possibility that the difference in overshoots was because of lower neuronal activation in pilocarpine-treated rats, e.g. because of sclerosis. In the subiculum, we found no significant differences between both groups, indicating a region-specific alteration of NAD(P)H transients in chronic epileptic rats. In tissue from TLE patients, the findings were more complex. In a few experiments, neuronal activation was associated with overshoots of up to 2.3% (Fig. 2C and D, plots). Largest overshoots were observed in patients with numbers 1, 6 and 10. Interestingly, they only differed from the other patients (n = 13) in age of TLE onset (26.3 ± 3.5 versus 6.4 ± 1.7 years; P = 0) and seizure frequency (13.3 ± 4.9 versus 6.2 ± 1.1 per month; P = 0.04). We rate this to be a trend because of the small number of n. In the majority of experiments, we observed very small overshoots in both area CA1 and the subiculum that went along with large drops (Fig. 3B). Moreover, these alterations of NAD(P)H transients were independent of the degree of sclerosis, because there were no significant differences when comparing AHS and non-AHS groups in both areas. Alterations were also found in area CA3 (n = 6, 3 patients) (not shown) and the DG of human tissue (Fig. 2F).
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Stimulus-induced NAD(P)H transients indicate adaptation of mitochondrial oxidative and glycolytic metabolism (Lipton, 1973
; Schuchmann et al., 2001), and the alterations suggest severe metabolic dysfunction in chronic epileptic tissue. Because mitochondria are the major source of energy supply in the CNS (Ames, 2000), alterations of NAD(P)H transients may reflect chronic defects of mitochondrial enzymes and/or membrane properties.
Negative mitochondrial membrane potential in chronic epileptic tissue
Respiring mitochondria establish a negative m that is essential for substrate cycling, Ca2+-uptake via the uniporter, and ATP synthesis (McCormack et al. 1990; Hansford and Zorov, 1998; Nicholls and Budd, 2000), and the significant role of mitochondrial Ca2+-uptake in regulation of NAD(P)+ reduction has been reported in a variety of cell types (Robb-Gaspers et al., 1998; Kann et al., 2003a, b; Bruce et al., 2004). Therefore, monitoring the changes of m during neuronal activation with mitochondria-specific, voltage-sensitive dye (rhodamine-123) provided an opportunity to prove whether (chronically) collapsed m of mitochondria was one of the mechanisms underlying alterations of NAD(P)H transients. To sufficiently activate all neurons, we applied glutamate via the bath solution.
Glutamate elicited similar m responses in area CA1 of both chronic-epileptic and control rats (n = 8, two animals, each) that were bulk-loaded with rhodamine-123 (Fig. 4A). Subsequent application of mitochondrial uncoupler, carbonyl cyanide m-chlorophenyl hydrazone (CCCP) (Nicholls and Budd, 2000; Kann et al., 2003a), revealed the overall capacity of mitochondria to accumulate rhodamine-123 owing to negative m. The same experiment was performed in area CA1 of three AHS patients revealing m responses in the similar range (Fig. 4A). To control the degree of neuronal activation as elicited by glutamate, we tested its effects on NAD(P)H fluorescence and [K+]o in area CA1 of AHS tissue (Fig. 4B). As with electrical stimulation, glutamate application resulted in a transient increase in [K+]o (1.1 ± 0.5 mM, n = 3, three AHS patients) while the overshoot of NAD(P)H transients remained small (1.4 ± 0.3%). The different time course of changes in NAD(P)H and [K+]o was probably due to the prolonged duration of glutamate application (60 s). Nevertheless, regarding the increase in maxima of the transients, this experiment confirmed that both stimulation protocols elicited neuronal activation in a similar range.
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To verify mitochondrial responses in individual neurons of human tissue, we combined patch-clamp recordings and confocal laser scanning microscopy. Visually identified neurons (n = 8, five AHS patients) were investigated in area CA1 and the proximal subiculum in AHS tissue, where alterations of NAD(P)H transients were most prominent. Twenty to thirty minutes after establishing a whole-cell patch with pipette solution that contained rhodamine-123, a bright dotted staining pattern of mitochondrial structures developed while cytoplasmic rhodamine-123 fluorescence remained weak. Confocal imaging allowed for depiction of individual mitochondria in the somata of the neurons (Fig. 5A and B), which was a first indication of mitochondria that maintained a negative
m. All investigated neurons (n = 8) responded to glutamate with a depolarization of 48.1 ± 6 mV from a resting cell membrane potential of –57.4 ± 2 mV (Fig. 5C). Application of glutamate was repeated in four of these neurons after the cell membrane potential had recovered to baseline (308 ± 62 s), which resulted in similar depolarization (not shown). In all neurons (n = 8), cytoplasmic rhodamine-123 fluorescence intensity increased after a glutamate-induced train of action potentials (Fig. 5C). Fluorescence intensity reached maximum of 103.8 ± 29% almost simultaneously with the peak of cell membrane depolarization (after 131 ± 35.8 s) and recovered slowly. This range of fluorescence intensity corresponded to that obtained in activated neurons in a non-epileptic rat slice preparation applying the same recording and evaluation methods (Kovács et al., 2005), which strongly supports our data from bulk-loaded slices (Fig. 4A). Note that confocal imaging from regions of interest in individual neurons provides fluorescence signals in a higher order of magnitude as compared to the totalized microfluorimetric signal from neurons and glial cells of bulk-loaded slices (Kovács et al., 2002).
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These data showed that mitochondria in chronic epileptic tissue, even in individual neurons, maintained a negative
m and depolarized during neuronal activation; although NAD(P)+ reduction was significantly impaired in the respective hippocampal regions. This might indicate defects on the mitochondrial enzyme level rather than on mitochondrial membrane properties. |
Discussion |
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The main findings of the present study are significant alterations of NAD(P)H fluorescence transients during neuronal activation in acute hippocampal slices from chronic epileptic rats and humans, although mitochondria maintain negative
m. These findings provide a cellular correlate for ‘hypometabolism’ as described for epilepsy patients and, thereby, suggest mitochondrial enzyme defects in TLE.
Induction of similar neuronal activation
K+-sensitive microelectrodes measure accumulation of potassium in a restricted extracellular space (Heinemann and Lux, 1975). The electrodes detect changes in [K+]o in a distance of at least 150 µm, and were positioned in the centre of a region of interest. We assume that, thereby, monitoring of [K+]o widely reflected the degree of neuronal activation in the region where NAD(P)H fluorescence was recorded. For quantitative comparison of NAD(P)H transients, this enabled us to elicit similar neuronal activation in hippocampal slices with different degrees of sclerosis. However, since there is some evidence for an impairment of glial K+-uptake mechanisms in area CA1 of AHS tissue (Kivi et al., 2000; Hinterkeuser et al., 2000), neuronal activation might have been overestimated in this region. This might partially explain smaller overshoots of NAD(P)H transients as compared to non-AHS tissue (Fig. 3B). At present, we cannot estimate the extent of impaired glial K+-uptake that manifested itself during the range of neuronal activation as elicited in AHS tissue. Nevertheless, alterations of NAD(P)H transients were also observed in subiculum where impairment of glial K+-uptake has not been found.
Negative mitochondrial membrane potential
The cell-permeable, cationic fluorescent dye, rhodamine-123 has been successfully used to monitor mitochondrial responses in a variety of preparations (Bindokas et al., 1998; Ward et al., 2000; Kovács et al., 2002). Applying pharmacology and mitochondria-specific probes (MitoTracker®), we have further confirmed that rhodamine-123 is a specific and reliable tool for investigations on mitochondria (Schuchmann et al., 2000; Kovács et al., 2005).
Despite significant alterations in NAD(P)H transients in area CA1 from chronic epileptic rats and humans, rhodamine-123 accumulated in mitochondria of bulk-loaded slices and of patched neurons (Figs 4 and 5). Moreover, application of glutamate and CCCP elicited dye release to the cytoplasm in a similar range as compared to controls (Fig. 4) and to individual neurons in non-epileptic tissue (Kovács et al., 2005). Because activity-dependent release of rhodamine-123 from mitochondria is commonly considered as reflecting mitochondrial depolarization during increased mitochondrial Ca2+-uptake (Bindokas et al., 1998; Ward et al., 2000), our data indicate that mitochondria in chronic epileptic tissue maintained a negative m. In addition to mitochondrial Ca2+-uptake, this would allow substrate transport across the inner mitochondrial membrane, and these mechanisms, in concert with mitochondrial enzyme activities, are key determinants in the regulation of NAD(P)+ reduction and ATP generation (McCormack et al. 1990; Hansford and Zorov, 1998).
Metabolic dysfunction during neuronal activation
In pilocarpine-treated rats, we found unaltered drops but significantly smaller overshoots of NAD(P)H transients reflecting less effective NAD(P)+ reduction (Fig. 3A). This difference was unlikely to be on account of less neuronal activation in chronic epileptic tissue, because increases in [K+]o were virtually the same. Moreover, the difference was restricted to area CA1, which probably excluded an unspecific effect of pilocarpine treatment on metabolic functions in the animal model. Our findings are in line with observations that area CA1 is much more vulnerable to seizure-induced neuronal damage than the subiculum (Babb et al., 1984; Mello et al., 1993), and with the few reports that have described alterations of NAD(P)H transients in models of experimental epilepsy (Kunz et al., 1999; Kovács et al., 2002). In AHS and non-AHS tissue from TLE patients, we found very small overshoots of NAD(P)H transients (Fig. 3B). Together with the animal model data, we believe that this alteration is more a characteristic of the chronic epileptic human brain than of the human brain per se. At present, we cannot precisely determine whether the pronounced drops of NAD(P)H transients in human tissue exclusively reflect more severe alterations of NAD(P)+ reduction and/or pathological variances in NAD(P)H oxidation and/or neuronal volume regulation. Though employing strategies of containment (Methods section), we cannot rule out the possibility that factors like antiepileptic drugs, anaesthesia and, potentially, tissue hypoxia during neurosurgery might have partially contributed to this phenomenon in human tissue (Gao et al., 1995; Schuchmann et al., 2001). In contrast to pilocarpine-treated rats, we observed no significant differences in alterations of NAD(P)H transients in both area CA1 and the subiculum when comparing AHS and non-AHS tissues. This deviant finding might be explained by prolonged disease duration in TLE patients (Table 1) and might indicate a gradual as well as spatial progression of metabolic dysfunction that becomes independent of the degree of sclerosis at later stages of TLE. This view is supported by neuroimaging studies also revealing ‘hypometabolism’ in brain structures adjacent to epileptic foci in patients (Casse et al., 2002). Conversely, our data support the hypothesis that ‘hypometabolism’ is more a reflection of dysfunction in cellular energy metabolism than neuronal cell loss (see also Henry et al., 1994; O'Brien et al., 1997).
NAD(P)H fluorescence in living cells is a summation of cellular NAD(P)H oxidation and NAD(P)+ reduction (Aubin, 1979; Schuchmann et al., 2001). NAD(P)H oxidation reflects, e.g. enhanced cytochrome c oxidase activity upon a decrease in ATP/ADP ratio (Kadenbach, 2003) and/or enhanced mitochondrial generation of superoxide (Dringen, 2000; Nicholls, 2002) during neuronal activation. NAD(P)+ reduction reflects enhanced activity of enzymes of the Krebs–Szentgyörgyi cycle in mitochondria as well as cytosolic glycolysis (Lipton, 1973; Duchen, 1992; Schuchmann, 2001). Thus, characteristic biphasic NAD(P)H fluorescence transients occur during neuronal activation in brain slice preparations (Fig. 2A), irrespective of the mode of stimulation (Kann et al., 2003a, b; Shuttleworth et al., 2003). Only recently, it has been proposed that the initial drop of NAD(P)H transients originates from the neuronal compartment whereas the prolonged overshoot is primarily astrocytic (Kasischke et al., 2004). Nevertheless, biphasic NAD(P)H transients were described in individual sensory and hippocampal neurons (Duchen, 1992; Schuchmann et al., 1998) similar to other cell types (Pralong et al., 1994; Hajnóczky et al., 1995; Robb-Gaspers et al., 1998). These findings basically offer two intriguing options for interpreting our data from chronic epileptic tissue: firstly, there may be defects on the mitochondrial enzyme level in neurons (see also, Kunz et al., 2000; Kudin et al., 2002; Cock et al., 2002) because small overshoots primarily reflect less effective mitochondrial NAD(P)+ reduction, which are presumably not governed by collapsed m and mitochondrial Ca2+-cycling (Figs 3
–5) or decreased numbers of mitochondria (Blümcke et al., 1999). Secondly, there may be a severe dysfunction on the level of astrocytic glycolysis and/or neuronal-astrocytic metabolic coupling (Magistretti and Pellerin, 1999; Kasischke et al., 2004). At present, we cannot estimate the extent to which each of these mechanisms contributes to metabolic dysfunction during neuronal activation in chronic epileptic tissue.
Implications for temporal lobe epilepsy
Besides the central role in energy metabolism, NAD(P)H is a key element in a variety of intracellular signalling cascades (Berger et al., 2004) and, e.g. essential for biosynthesis and reduction of cellular antioxidant, glutathione (Dringen, 2000; Nicholls, 2002). Thus, metabolic dysfunction in neurones and glial cells might significantly affect ATP homeostasis and excitability as well as intrinsic anti-oxidative mechanisms. Under certain conditions, these disturbances might favour neuronal vulnerability and manifestation of seizures and status epilepticus. Future epilepsy research might focus on alterations of intracellular signalling cascades and free radical generation during neuronal activation in chronic epileptic tissue, including their role in secondary tissue damage, as well as on effects of antiepileptic drugs on energy metabolism. Development of pharmacological tools for protection of mitochondria and free radical scavenging might serve as adjuvant therapy.
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Acknowledgements |
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This study was supported by grants, ICA CT 2002 70007 (European Union), OTKA-F043589 (Hungary), Graduiertenkolleg 238, Sonderforschungsbereiche TR3 and 507 (Germany). The authors thank Drs K. Schulze, H. Siegmund and H.-J. Gabriel for excellent technical support, Drs K. Buchheim, S. Schuchmann, A. Friedman and M. J. Gutnick for very helpful discussions.
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