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Zentrum für Physiologie und Pathophysiologie, Abteilung Neuro- und Sinnesphysiologie, Georg-August-Universität Göttingen, Göttingen, Germany
Submitted 18 March 2005; accepted in final form 27 April 2005
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ABSTRACT |
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INTRODUCTION |
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). Accordingly, changes in mitochondrial activity affect the cytosolic redox status, which is determined by the cellular H2O2 content as well as the two redox couples NAD+/NADH and reduced/oxidized glutathione (Dröge 2001). It is now clear that ROS are not only involved in cellular damage resulting from metabolic disturbances (Chan 2001), status epilepticus (Kova
s et al. 2002), and various neurodegenerative diseases such as Alzheimer disease, parkinsonism, and amyotrophic lateral sclerosis (Agar and Durham 2003; Halliwell 1992), but also serve as subtle intracellular messengers under physiological conditions (Dröge 2001). The primary site of modulation of cellular proteins by redox signaling are the sulfhydryl (SH) groups of the amino-acid cysteine. Their reduction or oxidation critically influences the activity of the protein affected because disulfide bonds and consequently protein structure might be modulated (Dröge 2001; Lipton et al. 2002).
In hippocampal neurons, various ion channels and receptors have been shown to be sensitive to redox modulation. Among these are N-methyl-D-aspartate NMDA)-type glutamate receptors (Sucher and Lipton 1991), persistent Na+ channels (Hammarström and Gage 2000), K+ channels (Seutin et al. 1995), and also Schaffer collateral/CA1 neuron synapses are modulated by H2O2 (Pellmar 1987). Accordingly, there are various possibilities for redox modulation to control neuronal excitability. However, so far it is unknown to what degree redox signaling is involved in the response of neurons and glia to metabolic insults such as anoxia and ischemia. Could cellular redox signaling possibly be part of the oxygen-sensing process in hippocampal neurons and trigger their immediate response to oxygen withdrawal? Could redox signaling even modulate the susceptibility of neural tissue against metabolic insults, could it possibly bear neuroprotective potential?
To address these questions, we focused on the very end of the redox signaling cascade, the SH groups: we analyzed to what degree the susceptibility of the hippocampal CA1 field toward anoxic insults, e.g., the generation of hypoxia-induced spreading depression (HSD), is affected by the modulation of SH groups. The specific targeting of SH groups was achieved by the SH oxidizing agent 5,5'-dithiobis 2-nitrobenzoic acid (DTNB), the SH-reducing drug 1,4-dithio-DL-threitol (DTT), and the SH-alkylating compound N-ethylmaleimide (NEM) (for review, see Lipton et al. 2002). The effects of these compounds were analyzed on the network level, the cellular level, and on the subcellular level.
Some of the results have been published as abstracts (Hepp et al. 2004, 2005).
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METHODS |
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Hippocampal tissue slices were prepared from ether-anesthetized Sprague-Dawley rats of 150–300 g body wt (4–8 wk old, mostly males). After decapitation, the brain was rapidly removed from the skull and placed in chilled artificial cerebrospinal fluid (ACSF) for 1–2 min. The two hemispheres were separated, and 400-µm slices were cut using a tissue slicer (Campden Instruments, 752M Vibroslice). On a few occasions, the hippocampus was first isolated and then chopped (400 µm slices) using a custom-made tissue chopper. Slices were transferred to an interface recording chamber of the Oslo style and were left undisturbed for 
90 min. The recording chamber was kept at a temperature of 35–36°C. It was continuously aerated with 95% O2-5% CO2 (400 ml/min), and perfused with oxygenated ACSF (3–4 ml/min). Hypoxia was induced by switching the chamber’s gas supply to 95% N2-5% CO2. To protect the slices from drying out and to prevent oxygenation from the air during hypoxic episodes, the slice chamber was covered by a lid with a small (2 cm2) opening for the positioning of the electrodes.
Cell cultures of hippocampal neurons were prepared from 2- to 4-day-old Sprague-Dawley rats. After decapitation, the brain was removed and placed in ice-cold HBSS (Hanks’ balanced-salt solution) containing 20% FCS (Biochrom). The dentate gyrus region was removed from the isolated hippocampi, and the remaining tissue was cut into smaller pieces and trypsinated (5 mg/ml) for 10 min at 37°C. Cells were then dissociated by gentle trituration, and the suspension was centrifuged for 10 min (1,500 rpm, 4°C). The pellet was re-suspended and plated on Matrigel (BD Biosciences)-coated glass cover slips, which were transferred to four-well culture plates (Nunc). Cultures were incubated at 37°C in a humidified, 5% CO2-containing atmosphere. After 24 h, half of the 600 µl medium in each well was exchanged with growth-medium containing 4 µM cytosine arabinoide (Sigma-Aldrich). Cultures were kept for 
2 wk, refreshing medium and growth factors after 4 days. Within 2–3 days in culture, cells fully recovered well-pronounced dendritic processes and were suitable for experiments; most experiments were performed between 3 and 7 days in culture.
Solutions
Chemicals, unless otherwise mentioned, were obtained from Sigma-Aldrich. The ACSF had the following composition (in mM): 130 NaCl, 3.5 KCl, 1.25 NaH2PO4, 24 NaHCO3, 1.2 CaCl2, 1.2 MgSO4, and 10 dextrose; aerated with 95% O2-5% CO2 to adjust pH to 7.4.
Minimum essential cell culture medium (MEM medium, Invitrogen) was supplemented with 5 mg/ml glucose, 0.2 mg/ml NaHCO3, and 0.1 mg/ml transferrin (Calbiochem). For initial plating, it also contained 10% FCS (fetal calf serum), 2 mM L-glutamine, and 25 µg/ml insulin. The medium used after day 4 in culture contained 5% FCS, 0.5 mM L-glutamine, 20 µl/ml B27 50x supplement (Invitrogen), and 100 µg/ml penicillin-streptomycin (Biochrom).
DTNB, DTT, and tetraethylammonium chloride (TEA) were directly dissolved in ACSF. DL-2-amino-5-phosphonovaleric acid (DL-AP5, Tocris) was prepared as 100 mM stock solution in 1 M NaOH and was kept frozen. Rhodamine 123 (Molecular Probes) was dissolved in absolute EtOH (20 mg/ml). Charybdotoxin (Sigma-Aldrich and Alomone Labs) and iberiotoxin (Alomone Labs) were dissolved in 100 mM NaCl, 10 mM HEPES plus 0.1% BSA. Paxilline, 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX), and tolbutamide were dissolved in DMSO as 10, 50, and 100 mM stocks, respectively, and stored at 4°C. NEM was prepared freshly as 250 mM stock in DMSO. To allow for sufficient diffusion of drugs into the interfaced slices, all drugs were applied for 20 min. Final DMSO and EtOH concentrations were 0.2%.
Microelectrodes
Single-barreled glass microelectrodes for extracellular recordings were pulled from thin-walled borosilicate glass (GC150TF-10, Harvard Apparatus) using a horizontal puller (P-97 Flaming/Brown Micropipette Puller, Sutter Instruments). They were filled with ACSF and their tips were broken to a final resistance of 5–10 M
. Sharp microelectrodes for current-clamp recordings were made from thick-walled borosilicate glass (GC 150F-10, Harvard Apparatus) and filled with 2 M K-acetate +5 mM KCl +10 mM HEPES; pH 7.4. Their resistances were 
80 M.
Hypoxia-protocol and electrical recordings
Severe hypoxia was induced by switching the recording chamber’s gas supply from carbogen (95% O2-5% CO2) to 95% N2-5% CO2, while the carbogen aeration of the experimental solutions was continued. Such treatment resulted within a few minutes in the occurrence of HSD. The time point of reoxygenation critically determines the reversibility of the hypoxia-induced changes. Therefore we resubmitted oxygen soon (15–20 s) after the onset of HSD, which was indicated by a sudden drop in extracellular DC potential. Within that time the extracellular DC potential shift had reached its nadir.
All signal amplitudes were measured between the prehypoxia baseline and the maximal change. Only rapid negative extracellular DC potential changes (
Vo) of 10-mV amplitude were considered as HSD, and only those slices were accepted for the experiments, which produced an HSD no earlier than 90 s after oxygen withdrawal (no earlier than 60 s in Ca2+-free solutions). HSD onset was defined as the occurrence of the sudden Vo.
Evoked responses were elicited by 0.1-ms unipolar stimuli delivered via microwire electrodes made from bare stainless steel wire (50-µm diam, AM-Systems) and recorded as described in detail earlier (Müller and Somjen 1998). Orthodromic responses were elicited by stimulation of Schaffer collaterals and recorded in stratum radiatum of the CA1 region; antidromic responses were elicited by stimulating pyramidal cell axons in the alveus and recorded in s. pyramidale of the CA1 region. All extracellular recordings were performed with a locally constructed extracellular DC potential amplifier.
Current-clamp recordings from CA1 neurons were performed with an intracellular recording amplifier (SEC-05L, NPI Instruments, Tamm, Germany). Bridge balance and electrode-capacitance compensation were adjusted before insertion of the electrode and continuously controlled during the entire recording. CA1 pyramidal neurons were identified by their location in s. pyramidale, membrane potential, spontaneous activity, action-potential shape and input resistance (Morin et al. 1996). Glial cells (astrocytes) lining the pyramidal cell layer were identified by their negative membrane potential of –80 to –90 mV, low input resistance, the absence of spikes in response to depolarizing stimuli (D'Ambrosio et al. 1998). Successful cell impalement was achieved by slowly advancing the electrode into the slice and applying a brief high-frequency current pulse ("buzz") to the electrode. Only CA1 neurons and glial cells with stable membrane potentials of at least –55 and –80 mV, respectively, were accepted. Neuronal input resistance was probed every 10 s by a hyperpolarizing current of 400- to 600-pA amplitude and 300-ms duration. Data were sampled at 2.5 kHz using a TL-1/Labmaster acquisition system and the Axotape V2 software (Axon Instruments). Input resistance was measured at the steady-state level of the voltage deflections and averaged over 10 successive current injections. Changes in input resistance were expressed as percent of pretreatment value.
Optical recordings
Optical recordings of intracellular Ca2+ concentrations, mitochondrial membrane potential, and NADH/FAD autofluorescence were performed with a standard computer-controlled fluorescence imaging system composed of a monochromatic light source (Polychrome II; Till Photonics, Gräfelfing, Germany) and a highly sensitive CCD camera (Imago QE, PCO Imaging, Kelheim, Germany) attached to an upright microscope (Axiotech vario, Zeiss, Oberkochen, Germany). The computer controlled monochromatic light source selects the predefined wavelength(s) by a galvanometer-mounted grating that projects the desired range of the spectrally dispersed light onto a narrow slit. Therefore the final excitation light shows a Gaussian wavelength distribution around the selected center-wavelength; bandwidth at half-maximum is 15 nm. In the following, just the center wavelengths of excitation light are mentioned. Rh123 was excited at 485 nm and fluorescence was recorded using a 505-nm beamsplitter and a 535/35-nm band-pass filter. Cellular NADH and FAD were excited alternately at 360 nm (NADH) and 460 nm (FAD), which are close to their respective absorption maxima, and the resulting autofluorescence was recorded simultaneously using a 505-nm beamsplitter and a 510/40-nm band-pass filter. This beamsplitter/emitter constellation used absorbed a large part of NADH autofluorescence (which is highest around 400–450), yet it made possible the simultaneous detection of NADH and FAD autofluorescence. For imaging experiments, tissue slices and cell cultures were placed in a submersion-style chamber at 33–35°C.
Statistics
The data were obtained from 70 rats, and because most experiments did not last longer than 1–2 h, up to five slices could be used from each brain. However, to ensure independence of observations, each experimental treatment was performed on at least three different rats. All numerical values are represented as means ± SD. Significance of the observed changes was tested using a two-tailed, unpaired Student’s t-test and a significance level of 5% (unpaired observations). Drug-induced changes were compared with untreated control slices that also underwent the corresponding number of HSD episodes. In the case of paired observations, a one-sample t-test was used to compare normalized drug effects against pretreatment control conditions, defined as unity or as 100%. In the diagrams significant changes are marked by asterisks (*P < 0.05; **P < 0.01). Statistical calculations were done with the Excel software (Office 2000).
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RESULTS |
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Severe hypoxia induces within a few minutes a nearly complete depolarization of hippocampal neurons and glial cells, which is associated by the occurrence of a negative shift in the extracellular DC potential, termed hypoxia-induced spreading depression (for review, see: Somjen 2001, 2004). In untreated slices HSD occurred within 156.6 ± 54.5 s, the associated negative shift of the extracellular DC potential (Vo) averaged –16.1 ± 4.2 mV and measured at its half-amplitude level it lasted 45.6 ± 10.4 s (n = 115). As we have shown previously, HSD of short duration can be induced repeatedly in a given slice, and with a sufficient recovery time of 30 min in between the hypoxic episodes, the characteristic parameters are not significantly affected for the first three HSD episodes (Müller and Somjen 1998).
Modulation of sulfhydryl groups did, however, result in characteristic changes of HSD. As compared with the previously induced control HSD, application of the sulfhydryl oxidizing agent DTNB (2 mM, 20 min) postponed the onset of HSD by 29.2 ± 18.8% and decreased its amplitude by 15.7 ± 8.8% (n = 9); HSD duration was not affected (Fig. 1, A and C). By contrast, treatment with the SH-reducing agent DTT (2 mM, 20 min) shortened the time to HSD onset by 25.0 ± 17.3% and decreased its amplitude by 26.1 ± 14.9% (n = 8), the duration of HSD was not affected (Fig. 1, B and C). The sulfhydryl alkylating agent NEM (500 µM, 20 min) shortened the time to HSD onset by 48.4 ± 10.4% and reduced its duration by 29.9 ± 5.5%; HSD amplitude was not affected (n = 7, Fig. 1C). Application of the rather unspecific oxidizing agent H2O2 (1 mM, 20 min) caused a moderate, less consistent postponement of HSD by 12 ± 19% (n = 7).
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Withdrawal of extracellular Ca2+ (n = 7, Fig. 2, A and C) or application of 2 mM Ni2+ (n = 5, Fig. 2, B and C) prevented the DTNB-induced postponement of HSD, indicating the requirement of Ca2+ influx from extracellular space for the delayed HSD onset. In the presence of the NMDA antagonist DL-AP5 (150 µM, n = 6) or the AMPA/kainate antagonist CNQX (25 µM, n = 8), the DTNB-induced postponement of HSD remained unchanged (Fig. 2C).
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), or treatment of slices with the KATP channel blocker tolbutamide (200 µM, n = 6) did not affect the DTNB-induced postponement of HSD (Fig. 3B). However, pretreating slices with 20–25 nM charybdotoxin (CTX, n = 5 Fig. 3A) or 50 nM iberiotoxin (IBX, tested in 3 trials only) consistently abolished the DTNB-induced postponement of HSD (Fig. 3, A and B). It therefore seems that sulfhydryl modulation activates Ca2+-activated, high-conductance K+ channels of the BK type.
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). In our trials, paxilline (10 µM) did not prevent the hastening of HSD onset by DTT (n = 5, Fig. 4B), and it also failed to prevent the postponement of HSD by DTNB (n = 4, Fig. 4B). It therefore seems questionable whether paxilline is effective under our experimental conditions at all.
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To explore whether DTNB and DTT affect other neuronal functions that are likely to be influenced by BK channels, we performed sharp electrode recordings from single CA1 neurons and glial cells in interfaced slices. In addition, we probed for changes in synaptic function by recording extracellular field potentials.
Because pyramidal cells and interneurons respond to hypoxia identically (Müller and Somjen 2000a), the data of both types of cells were pooled (11 pyramidal cells, 9 interneurons). Average membrane potential and input resistance of the recorded neurons were –65.9 ± 5.0 mV and 25.5 ± 7.5 M (n = 20). In untreated control slices, oxygen withdrawal triggered within 1–2 min an initial hyperpolarization of CA1 neurons that then turned into a slow depolarization and finally triggered (after 177 ± 69 s, n = 7) an explosive depolarization close to 0 mV (for details, see Müller and Somjen 2000a,b) (Fig. 5A).
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Incubation with DTT (2 mM, 20–25 min) caused a moderate depolarization in four cells and a moderate hyperpolarization in the two others; the input resistance did not change significantly (Fig. 5C). However, in the presence of DTT spontaneous spike discharges became more frequent, and within 15 min well-pronounced burst discharges developed (Fig. 5D). During these burst discharges single spikes became progressively broadened (Fig. 5D). In the presence of DTT, oxygen withdrawal triggered the massive hypoxic depolarization within 97 ± 45 s. The initial hypoxic hyperpolarization was usually blocked, and the neurons directly started to depolarize slowly when oxygen was withdrawn (Fig. 5C).
By incidence in hippocampal cell cultures, we observed the retraction of glial processes in response to DTT (Fig. 6A), whereas neuronal processes remained intact. This retraction started within 7–10 min of DTT application and was completed within another 30–40 min. DTNB application did not cause such changes in cell shape. To elucidate, whether DTT disturbs glial function, which would severely interfere with neuronal and synaptic function, we performed sharp electrode recordings from CA1 glial cells in hippocampal slices. The average glial membrane potential was –89.7 ± 5.6 mV (n = 6); their input resistance was too low to be probed reliably. In response to DTT glial cells showed a slow but continuous depolarization which averaged 19.2 ± 7.5 mV after 15–20 min of DTT treatment (Fig. 6B, n = 6). The synchronized burst discharges elicited by DTT in pyramidal neurons, caused—apparently due to an increased extracellular K+ concentration—slow transient depolarizations of glial cells by 20 mV (Fig. 6B). Continuing DTT treatment beyond 30 min usually caused spontaneous SD episodes during which glial cells underwent the characteristic massive depolarization (see also Müller and Somjen 2000a).
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Because direct effects of DTNB and DTT on mitochondria would be expected to modulate neuronal susceptibility to hypoxia, we elucidated whether these compounds affect mitochondrial membrane potential and/or metabolism. In detail, we probed for changes in rhodamine 123 (Rh123) fluorescence, a marker for mitochondrial membrane potential (
), and for changes in autofluorescence, representing the cellular redox couples NADH/NAD+ and FADH2/FAD. These recordings were performed in a submersion style chamber (33–35°C).
In Rh123 labeled slices (2–5 µg/ml Rh123 loading for 25 min) as well as in cultured CA1 neurons, the protonophore FCCP (1–2 µM) induced a clear increase in Rh123 fluorescence, 65 ± 21% in s. radiatum of acute slices (n = 7) and 63 ± 33% in the cytosol (soma) of cultured CA1 neurons (n = 8), thereby indicating pronounced mitochondrial depolarization (Fig. 9, A and B). In contrast, DTNB and DTT (2 mM each, applied for 7–9 min) did not markedly affect mitochondrial membrane potential—neither in cultured CA1 neurons nor in acute hippocampal slices (Fig. 9, A and B). Also, recording autofluorescence from acute slices did not reveal any pronounced effects of DTNB or DTT (2 mM each) on the cellular levels of NADH and FAD. Autofluorescence excited at 360 nm represents reduced NADH, whereas autofluorescence excited at 460 nm corresponds to oxidized flavins (FAD). As expected, in response to mitochondrial inhibition by anoxia, 1 mM CN– caused opposite changes in these simultaneously recorded measures (Duchen and Biscoe 1992), shifting NADH to its reduced and FAD to its oxidized form (Fig. 9, C and D). Accordingly, NADH autofluorescence increased by 9.2 ± 1.9%, while FAD autofluorescence decreased by 7.4 ± 1.8% (n = 6 each). DTT induced only barely noticeable changes in autofluorescence levels. DTNB, which absorbs light in the range of 300–450 nm, caused an artificial drop in NADH fluorescence due to competitive absorption of excitation light (Fig. 9D). The time course of these changes as well as the absence of a corresponding, opposite signal in FAD autofluorescence do confirm the artifactual nature of the DTNB-induced changes in NADH autofluorescence.
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DISCUSSION |
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How is SD postponement achieved?
The generation of HSD reflects the concerted activation of various types of ion channels and glutamate receptors enabling massive, self-regenerative Ca2+ and Na+ influx into neurons that is paralleled by K+ release into extracellular space (Aitken et al. 1991; Müller and Somjen 1998, 2000b; Somjen 2001, 2004). Computer simulations, based on experimental data (Kager et al. 2002), as well as current source density analyses (Wadman et al. 1992) provided evidence that SD is ignited in the dendrites. The ignition point is reached, when the sum of all inward currents exceeds the total outward current, i.e., when the net dendritic current turns inward (Kager et al. 2002). Inhibition of voltage-gated Na+ channels, Ca2+ channels, or glutamate receptors does not prevent SD but just postpones its onset. No matter what maneuvers are taken, in single neurons the anoxic depolarization, once triggered, evolves until completion in an all-or-none fashion (Müller and Somjen 2000b; Kager et al. 2002). If the hypoxic depolarization is less synchronized within the neuronal population, this might result in a reduced extracellular DC potential shift but again without dampening the massive depolarization in individual pyramidal cells and interneurons. But how does oxidation of SH groups and the resulting postponement of HSD onset fit into this scenario?
The immediate, early response of hippocampal CA1 neurons to hypoxia is an initial hyperpolarization, which is generated by activation of K+ channels and subsequent K+ release into interstitial space (Hansen et al. 1982; Donnelly et al. 1992; Müller and Somjen 2000a). Whether solely BK-type channels or KATP channels are activated or whether both types of channels are involved, with their contribution being weighed depending on the severity of an insult and the experimental conditions, is still under discussion (Erdemli et al. 1998; Fujimura et al. 1997; Zawar and Neumcke 2000). Under our experimental conditions (slices of adult rats placed in an interface chamber at 35–36°C), the initial hyperpolarization seems to be mediated mostly by BK-type channels: DTNB was found to hyperpolarize CA1 neurons by activation of BK channels and subsequent oxygen withdrawal did not cause a further hyperpolarization of these cells. That the postponement of HSD by DTNB was abolished by charybdotoxin, iberiotoxin, Ni2+, and Ca2+ withdrawal further supports the activation of BK channels. By contrast, tolbutamide was without effect, ruling out an involvement of KATP channels—at least during the early phase of hypoxia. Also the time to onset of HSD in untreated control slices (167 ± 63 s, n = 19) and in slices treated with 200 µM tolbutamide (143 ± 42 s, n = 12) did not differ.
We therefore conclude that the DTNB-mediated oxidation of SH groups activates BK channels, thereby postponing the onset of HSD. Whether DTNB directly acts on BK channels—modulating their gating properties or rendering them more Ca2+ sensitive—cannot be decided yet on the basis of our data. Indications for an increased Ca2+ sensitivity of hippocampal BK channels were found during the postischemic period (Gong et al. 2002). Alternatively, DTNB could as well modulate voltage-gated Ca2+ channels and enhance Ca2+ influx during the initial phase of hypoxia, thereby activating BK channels indirectly. Such an activation of Ca2+ channels, leading to subsequent activation of a charybdotoxin-sensitive KCa current, was reported in dissociated hippocampal neurons by Nowicky and Duchen (1998) in response to cyanide poisoning of mitochondria or their uncoupling by FCCP. The fact is that in our experiments Ca2+ influx from extracellular space was required for the activation of BK channels and that it was carried via voltage-gated Ca2+ channels, not via glutamate receptors, because DL-AP5 and CNQX failed to antagonize the DTNB-induced postponement of HSD.
The different time course of BK channel activation in the presence of DTNB or the enhanced BK-mediated outward current resulted in a postponement of SD. This could be due to more efficient glial K+ buffering, which was demonstrated in computer simulations to postpone or even prevent SD (Somjen 2004; p. 309). BK channels were already open before the onset of hypoxia, thereby causing an earlier hyperpolarization and allowing the neurons to withstand the generation of SD for a longer time period. Interestingly, BK channels were also reported to control the generation of Ca2+ spikes in pyramidal cell dendrites (Golding et al. 1999). Accordingly, an increased BK channel activity could successfully shift the ratio of inward versus outward current more to favor of the outward current, thereby preventing dendritic depolarization and thus HSD ignition within the dendritic tree just a little bit longer. As BK channels are assumed to be expressed in astrocytes as well (Barres et al. 1990), they could facilitate K+ uptake and K+ buffering within the glial syncytium. However, because ignition of SD occurs in neurons while glial cells just follow passively the extracellular increase in K+ (Müller and Somjen 2000a; Somjen 2001), we rather believe that the postponement of HSD due to modulation of BK channels is more likely to take place in neurons rather than glia.
Redox modulation of hippocampal circuitry and excitability
DTNB tended to decrease synaptic efficacy only slightly, while axonal conduction was not affected (Figs. 7A and 8A). The slight dampening of synaptic function is probably the result of the moderate hyperpolarization of single CA1 neurons in response to DTNB, i.e., a reduced postsynaptic excitability due to the increased resting K+ conductance or a dampening effect of DTNB on the NMDA component of the EPSP (Tauck 1992). Paired-pulse facilitation, which arises from residual Ca2+ remaining in the presynaptic terminal after the first stimulus (Zucker 1989), was not reduced by DTNB, suggesting that presynaptic Ca2+ cycling was not affected.
By contrast, DTT markedly reduced synaptic efficacy but did not depress antidromic responses. In addition, it increased postsynaptic excitability (Figs. 7B and 8B). The moderate depolarization observed in single hippocampal neurons and the induction of burst discharges by DTT (Fig. 5D) correspond to earlier reports obtained in guinea pigs (Tolliver and Pellmar 1988), and they indicate an enhanced excitability of CA1 neurons. Obviously these changes in CA1 neurons are also responsible for the firing of multiple population spikes in response to single antidromic stimuli (Fig. 8B) as well as for the observed depolarization of glial cells (Fig. 6B), which suggests an increase in extracellular K+. Thus the observed depression of synaptic function rather seems to be due to presynaptic effects of DTT in the Schaffer collateral terminal. If also induced presynaptically, the DTT-mediated depolarization or the increased extracellular K+ could inactivate presynaptic Na+ and Ca2+ channels, thereby reducing Ca2+ influx and transmitter release from the synaptic terminals (Eccles et al. 1963; MacDermott et al. 1999). As glutamine failed to prevent synaptic depression by DTT, glial poisoning by DTT can be excluded as a possible explanation for synaptic failure.
Besides the proven modulation of BK channels, additional channels and receptors are known to be sensitive to sulfhydryl modulation. DTT increases, while DTNB decreases NMDA currents (Sanchez et al. 2000; Tauck 1992), thereby enhancing NMDA receptor-mediated toxicity in hippocampal slice cultures (Pringle et al. 2000). In contrast, DTNB suppresses spontaneous ictal activity due to inhibition of NMDA receptors (Sanchez et al. 2000). Yet for our data, redox modulation of glutamate receptors can be ruled out because the effects of DTNB and DTT on HSD persisted in the presence of NMDA and non-NMDA antagonists (Figs. 2C and 4B).
Persistent Na+ channels are potentiated by hypoxia (Hammarström and Gage 1998) and an increase in extracellular K+ (Somjen and Müller 2000), and they play a key role in the triggering of normoxic SD and HSD (Kager et al. 2002; Somjen 2001). Interestingly INap is also sensitive to SH modulation, being increased by NO donors in a DTT-sensitive manner (Hammarström and Gage 1999). Nevertheless, persistent Na+ channel modulation cannot explain our findings either because it was BK-channel inhibition that abolished the DTNB-mediated postponement of SD. Also, if Na+ channels were modulated by DTNB or DTT, one would expect effects on axonal conduction as well. Antidromic responses were, however, found not to be modulated by either DTT or DTNB (Fig. 8).
Changes in cellular redox state during hypoxia
The oxidation of SH groups was found to postpone HSD. Severe hypoxia itself causes a decrease of mitochondrial metabolism by inhibiting complex IV of the respiratory chain, which should decrease the formation of ROS (Boveris and Chance 1973; Votyakova and Reynolds 2001). In addition, the redox couple NAD+/NADH is shifted more to the reducing form, thus shifting the cellular redox status toward reducing conditions (Mills and Jöbsis 1972; Riepe et al. 1996) (Fig. 9). In contrast, reoxygenation, rotenone, antimycin, Ca2+-induced mitochondrial depolarization, increased ATP-synthesis and mitochondrial uncoupling by, e.g., FCCP increase ROS production (Bindokas et al. 1996; Boveris and Chance 1973).
Could reducing conditions possibly favor the generation of HSD? NMDA receptor-mediated currents are increased under reducing conditions (Sanchez et al. 2000; Tauck 1992; Tolliver and Pellmar 1988), i.e., neuronal excitability increases. Also, BK channels are blocked by reducing conditions but activated by oxidizers such as DTNB and H2O2. In addition to our findings, Nowicky and Duchen (1998) reported activation of Ca2+-sensitive K+ channels in dissociated CA1 neurons in response to mitochondrial depolarization by either FCCP or chemical anoxia induced by cyanide. So one might ask whether sulfhydryl/redox modulation could be the missing link of how mitochondrial failure affects K+ channel activity, how mitochondrial dysfunction finally affects membrane conductances. The fact is that neurons respond immediately to metabolic compromise, and they respond—with protective mechanisms—way before cellular ATP is depleted (Hansen et al. 1982; Lipton and Whittingham 1982; Müller et al. 2002). The detailed molecular events involved in the sensing of hypoxia by central neurons are only partly understood, but it seems that SH modulation may play a pivotal role in these processes.
Neuroprotective potential of SH oxidation
The postponement of HSD was found to depend on the activation of BK channels. In general, these channels decrease neuronal excitability, and others reported them to reduce cell death from ischemic insults in CA1 pyramidal cell cultures and to regulate presynaptic glutamate release (Runden-Pran et al. 2002). We found that—even though BK channels are activated anyway during the early phase of hypoxia—their increased activity already before oxygen withdrawal succeeded in stabilizing the membrane potential and preventing the ignition of HSD for some more time—on average 30% longer than in untreated slices. As a result, the dramatic disturbance of ionic homeostasis and especially the massive Ca2+ load being associated with the hypoxic depolarization (Hansen 1985; Martin et al. 1994; Müller and Ballanyi 2003) were delayed. The decreased amplitude of HSD might indicate desynchronization of the anoxic depolarization in CA1 neurons. Both postponement and desynchronization may prevent HSD during short-term insults and possibly also reduce its spreading speed and range. Accordingly, parts of the neuronal population could be affected later or might even be spared from the loss of membrane potential and the threatening Ca2+ overload. Stroke is of course rarely anticipated, but peri-infarct SD waves reaching out in the penumbra and aggravating neuronal damage of surrounding tissue that was initially not affected (Busch et al. 1996; Kempski et al. 2000) could be dampened or depressed by well-directed SH modulation. Therefore oxidizing SH groups might mediate neuroprotection during metabolic insults.
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Address for reprint requests and other correspondence: M. Müller, Zentrum Physiologie und Pathophysiologie, Universität Göttingen, Humboldtallee 23, D-37073 Göttingen, Germany (E-mail: mike{at}neuro-physiol.med.uni-goettingen.de)
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