Introduction

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Excitotoxicity has been causally linked with glutamate-elicited Ca²⁺ influx (Choi, 1987; Hartley et al., 1993; Eimerl and Schramm, 1994) and with Ca²⁺ accumulation in mitochondria (Kiedrowski and Costa, 1995; White and Reynolds, 1996; Schinder et al., 1996; Stout et al., 1998). Although exposure to glutamate also greatly elevates cytoplasmic Na⁺ concentrations ([Na⁺]_C) in cultured neurons (Kiedrowski et al., 1994; Pinelis et al., 1994), it is yet unclear what role the increase in [Na⁺]_C plays in excitotoxicity. Although excessive swelling caused by the influx of Na⁺, Cl, and water leads to neuronal death (Rothman, 1985; Choi, 1987), such a mechanism characterizes kainate- rather than glutamate- or NMDA-induced excitotoxicity (Kiedrowski, 1998). Earlier studies have shown that high [Na⁺]_C contributes to the NMDA-induced Ca²⁺ influx by activating the reverse operation of the plasma membrane Na⁺/Ca²⁺ exchanger (NaCaX) (Kiedrowski et al., 1994; Hoyt et al., 1998). The activation of the reverse NaCaX also requires that cytoplasmic Ca²⁺ be raised to micromolar levels (Hilgemann et al., 1992). This makes the channels that are permeable for both Na⁺ and Ca²⁺, such as NMDA receptor channels (Mayer and Westbrook, 1987), perfect triggers of the reverse NaCaX activation.

The role of the NaCaX in excitotoxicity was previously tested in vitro using Na-free media (Mattson et al., 1989; Storozhevykh et al., 1998) in which Na⁺ was substituted with a large cation, N-methyl-D-glucamine (NMG⁺), that very poorly, if at all, permeates NMDA receptor channels. These studies showed that in the presence of NMG⁺, the NMDA-induced Ca²⁺ influx and excitotoxicity are greatly enhanced, and the data were interpreted as indicating that such an outcome results from an inhibition of Ca²⁺ extrusion by the NaCaX (Mattson et al., 1989), or from an alleged excessive release of endogenous glutamate (Storozhevykh et al., 1998). Because these interpretations failed to consider that the substitution of Na⁺ with NMG⁺ might prevent the Na-dependent plasma membrane (PM) depolarization that normally accompanies activation of NMDA receptors, the aim of the present study was to test what impact such depolarization has on the NMDA-induced Ca²⁺ influx and excitotoxicity.

To this end, using digital fluorescence microscopy and fluorophores sensitive to the plasma membrane potential (E_m) and to cytoplasmic Ca²⁺ concentrations ([Ca²⁺]_C), E_m, and [Ca²⁺]_C were simultaneously monitored in primary cultures of cerebellar granule cells (CGCs) exposed to NMDA. NMDA receptors were activated by an agonist, whereas extracellular Na⁺ was present or substituted either with NMG⁺ or with Li⁺. Because neither NMG⁺ nor Li⁺ support Ca²⁺ transport via the NaCaX (Blaustein, 1977; Hilgemann 1989), but Li⁺, in contrast to NMG⁺, permeates NMDA receptor channels (Tsuzuki et al., 1994) and should depolarize the PM (Hösli et al., 1973), this experimental design was expected to isolate the effects of PM depolarization from the effects of Na-dependent activation of the NaCaX. Because substitution of extracellular Na⁺ with Li⁺ or NMG⁺ is also expected to affect the operation of the plasma membrane Na⁺/H⁺ exchanger (NaHX), which controls cytoplasmic pH (pH_C) (Aronson, 1985), and because the role of pH_C in Ca²⁺ homeostasis is obscure, pH_C was also monitored.

	Materials and Methods

Top Summary Introduction Materials and Methods Results and Discussion References

Neuronal Cultures. CGCs were prepared from 8-day-old Sprague-Dawley rats and suspended in culture medium consisting of basal Eagle's medium supplemented with 25 mM KCl, 10% bovine fetal serum, 2 mM glutamine, and 50 µg/ml gentamycin, as described previously (Kiedrowski et al., 1994). The cells were plated on poly-D-lysine (10 µg/ml)-coated 35-mm dishes at a density of 2 × 10⁶ cells/dish; for the imaging experiments, CGCs were plated on poly-D-lysine-coated 2.5-cm glass coverslips. Glial proliferation was curtailed by the addition of 10 µM cytosine arabinofuranoside 24 h after plating. Cultures at 8 to 11 days in vitro were used for the experiments.

Media. Experimental solutions were based on Locke's buffer (Na-Locke's) containing 154 mM NaCl, 5.6 mM KCl, 3.6 mM NaHCO₃, 1.3 mM CaCl₂, 1 mM MgCl₂, 5 mM glucose, and 10 mM HEPES, pH 7.4, adjusted with Tris. The Na-free Locke's buffer in which Na⁺ was replaced with Li⁺ (Li-Locke's) contained 157.6 mM LiCl, 2 mM KCl, and 3.6 mM KHCO₃, with the remaining ingredients the same as in Na-Locke's. When Na⁺ was replaced with NMG⁺ or Cs⁺, the buffer was identical with Li-Locke's except that Li⁺ was substituted with NMG⁺ or Cs⁺, respectively. Ca-free solutions contained 3 mM EGTA and no CaCl₂, with other components of Locke's buffer unchanged. Glutamate (100 µM) or NMDA (300 µM) were applied in Mg-free media containing 10 µM glycine. Glucose-free solutions contained 5 mM 2-deoxy-D-glucose instead of D-glucose. The mitochondria-depolarizing cocktail (MDC) was composed of a Ca- and glucose-free Na-Locke's supplemented with 10 µM cyanide m-chlorophenylhydrazone (CCCP), 3 µg/ml oligomycin, and 10 µM MK-801 [(+)5-methyl-10-11-dihydro-5H-dibenzocyclohepten-5,10-imine].

Viability Test. [Ca²⁺]_C levels were monitored in CGCs exposed to glutamate as described in the text. Following glutamate withdrawal, by washing with Na-Locke's, [Ca²⁺]_C levels were monitored for an additional 20 to 30 min. Then the cells were returned to a conditioned medium (CM), i.e., the medium (containing 25 mM KCl) in which the cells were cultured. Viability of the same cells was assessed after 20 to 24 h by comparing oblique illumination images of the neurons occupying the same microscopic field before and 20 to 24 h after the excitotoxic challenge; neurons with a damaged PM were identified using propidium iodide (Jones and Senft, 1985). To this end, the cells were incubated for about 10 min at room temperature with 10 µM propidium iodide dissolved in Na-Locke's, and the propidium iodide fluorescence emitted at over 520 nm after 488 nm excitation was imaged. Whether the positions of propidium iodide-positive spots corresponded to the positions of dead cells was examined by digitally subtracting, using the Attofluor software (Atto Instruments, Rockville, MD), the images of the propidium fluorescence from the oblique illumination images of the same microscopic field. This procedure allows one to distinguish false signals, i.e., the propidium iodide-positive spots representing cellular debris floating in the medium. In some experiments propidium iodide was added to the CM immediately after the challenge with glutamate, and the progression of the plasma membrane deterioration was followed for up to 70 h by monitoring the propidium iodide fluorescence.

Simultaneous Assay of [Ca²⁺]_C and E_m. The coverslips plated with CGCs were transferred to custom-made recording chambers, in which the cells were loaded for 60 min at 37°C with 4 µM of fura-2 acetoxymethyl ester dissolved in CM. The stock concentration of fura-2 acetoxymethyl ester was 1 mM, in dimethyl sulfoxide (DMSO). Following the loading, the cells were washed using CM without fura-2. Fluorescence data were acquired from the cells that, based on their neuronal appearance and the small size of the cell bodies (about 10 µm in diameter), could be identified as CGCs. Occasionally, larger cells (about 20 µm in diameter), most likely astrocytes, were also observed and monitored. All imaging experiments were carried out at 37°C using the TC-102 temperature controller and the LU-CB1 Leiden culture system from Medical Systems Corp. (Greenvale, NY). Monitoring of the fura-2 fluorescence was begun while the cells were still incubated in CM; the pH of the CM was maintained at physiological levels by delivering 6% CO₂ to the recording chamber. Then, the CM in the recording chamber was replaced with Na-Locke's supplemented with 100 nM bis(1,3-dibutylbarbituric acid)trimethine oxonol [DiBAC₄(3)] to monitor E_m (Bräuner et al., 1984; Laskey et al., 1992).

Simultaneous Assay of [Ca²⁺]_C and pH_C. The coverslips with CGCs were handled the same way as described above for the assay of [Ca²⁺]_C and E_m except that the cells were loaded for 60 min at 37°C with 4 µM fura-2 acetoxymethyl ester plus 0.2 µM BCECF (2',7'-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein) acetoxymethyl ester. The stock concentration of BCECF acetoxymethyl ester was 0.05 mM, in DMSO. Following the loading, the cells were washed using CM without fura-2 and BCECF. Images of the fluorescence emitted at over 520 nm at 334, 380, 440, and 488 nm excitations were saved every 10 to 20 s. [Ca²⁺]_C was calculated from the F₃₃₄/F₃₈₀ ratio as described earlier. The F₄₈₈/F₄₄₀ ratio was used to determine pH_C from the in situ calibrations performed after each experiment. The background F₄₄₀ and F₄₈₈ values were measured in cell-free areas at all time points and were subtracted from the fluorescence data before calculating the F₄₈₈/F₄₄₀ ratio. The pH calibrating solutions contained 10 µM nigericin, 10 µM CCCP, 134.2 mM K-gluconate, 25.4 mM KCl, 1.3 mM CaCl₂, 1 mM MgCl₂, 3.6 mM KHCO₃, and 10 mM 2-[N-morpholino]ethanesulfonic acid or 10 mM HEPES. 2-[N-morpholino]ethanesulfonic acid was used to adjust the pH of the calibrating solution to a range of 5.5 to 6.5; HEPES was used in the pH range of 7.0 to 8.0. The data were fitted to the four-parameter equation of a sigmoidal curve:
Representative experiments (n = 7) yielded the following parameters: y₀ = 0.39 ± 0.01, a = 1.18 ± 0.06, x₀ = 6.99 ± 0.03, and b = 0.41 ± 0.02.

Assay of Cytoplasmic K⁺ Concentration ([K⁺]_C). CGCs were loaded for 60 min at 37°C with 5 µM K⁺ binding benzofuran isophthalate (PBFI) acetoxymethyl ester dissolved in CM. The stock concentration of PBFI acetoxymethyl ester was 1.25 mM, in DMSO. PBFI fluorescence was monitored at 37°C using the same excitation and emission settings as described for fura-2. The F₃₃₄/F₃₈₀ ratio was calibrated for [K⁺]_C in situ at the end of the experiments. The [K⁺]_C calibrating buffers were prepared by appropriate mixing of two solutions containing high and low concentrations of K⁺. The high-concentration K⁺ solution contained 5 µM gramicidin D, 134.2 mM K-gluconate, 25.4 mM KCl, 3.6 mM KHCO₃, 1 mM MgCl₂, and 10 mM HEPES, pH 7.2, adjusted with Tris. In the low-concentration K⁺ solution, 134.2 mM Li-gluconate and 25.4 mM LiCl were used instead of the respective K⁺ salts, and the remaining ingredients were unchanged. [K⁺]_C values were calculated using a nonlinear least-squares fit of the data to the Michaelis-Menten equation as described by Kasner and Ganz (1992).

⁴⁵Ca²⁺ Uptake. CGCs were incubated for 15 min at 37°C with 1 ml of experimental media containing 1 µCi of ⁴⁵Ca²⁺. The extracellular ⁴⁵Ca²⁺ was then removed by triple washing with 2 ml of an ice-cold buffer that contained 154 mM NaCl, 5.6 mM KCl, 3.6 mM NaHCO₃, 1 mM MgCl₂, 2 mM EGTA, and 10 mM HEPES, pH 7.4, adjusted with Tris. The cells were then dissolved in 1 ml of 0.5 M NaOH; neutralized aliquots of this solution were used for scintillation spectroscopy and for protein determination.

Statistical Analysis. All averaged data are expressed as the means ± S.E.M. The statistical tests used are indicated in the text. All statistical analyses of the [Ca²⁺]_C data were performed using the F₃₃₄/F₃₈₀ ratios. The [Ca²⁺]_C data could not be used for statistical analysis because when [Ca²⁺]_C approached the fura-2 saturating levels, the calculated [Ca²⁺]_C values became very imprecise. In some experiments, the fura-2 fluorescence was monitored using different sets of neutral density filters and/or gains, which affected the absolute values of the F₃₃₄/F₃₈₀ ratios (for example, note the differences in F₃₃₄/F₃₈₀ ratios between Figs. 2 and 6). To normalize the data, the F₃₃₄/F₃₈₀ ratios were converted to [Ca²⁺]_C values using the calibrating parameters R_max, R_min, and  obtained in sister cultures in the same experimental sessions; then, the [Ca²⁺]_C values from various experiments were converted to the normalized F₃₃₄/F₃₈₀ ratio using a single representative set of the calibrating parameters R_max', R_min', and ', according to the formula:

Materials. Fura-2 acetoxymethyl ester, PBFI acetoxymethyl ester, BCECF acetoxymethyl ester, and DiBAC₄(3) were obtained from Molecular Probes (Eugene, OR). MK-801 was purchased from Research Biochemicals Inc. (Natick, MA). The culture media and all other chemicals were from Sigma Chemical Co. (St. Louis, MO).

	Results and Discussion

Top Summary Introduction Materials and Methods Results and Discussion References

[Ca²⁺]_C Levels in CGCs Incubated in CM. Although the very low (about 20 nM) [Ca²⁺]_C levels in CGCs incubated in Na-Locke's are very homogeneous, a small number of CGCs show a marked heterogeneity in glutamate-induced [Ca²⁺]_C transients and fail to efficiently buffer cytoplasmic Ca²⁺ (see Fig. 4C in Kiedrowski and Costa, 1995). In the present study, it was routinely observed that it is possible to predict which neurons will fail to buffer the glutamate-induced [Ca²⁺]_C transients. From among 1713 CGCs (10 different platings) incubated in CM (containing 25 mM K⁺), a subpopulation of 72 neurons could be distinguished that maintained [Ca²⁺]_C at much higher levels (>600 nM) than the majority of CGCs (300-400 nM), but that nevertheless promptly decreased [Ca²⁺]_C to about 20 to 30 nM when CM was replaced with Locke's buffer and the extracellular K⁺ ([K⁺]_E) concentration was reduced from 25 to 5.6 mM (Fig. 1). Such neurons failed to buffer the [Ca²⁺]_C transients elicited by NMDA (see representative data in Fig. 1) or glutamate (data not shown). Because this subpopulation of neurons appears to have distinctly different properties, in the present study these neurons were singled out, and their data were excluded when average responses, typical for the majority of CGCs, were calculated. This abnormal subpopulation of CGCs requires an additional, separate, characterization.

View larger version (19K): [in this window] [in a new window]   Fig. 1.   Basal [Ca2+]C levels and NMDA-induced [Ca2+]C transients in CGCs. CGCs were loaded with fura-2 while incubated in a CM. As indicated by the bars, CM was replaced with Na-Locke's (Na-L) and then the cells were exposed to NMDA (300 µM NMDA + 10 µM glycine in the absence of Mg2+). Shown are the [Ca2+]C traces recorded from 39 individual neurons in a representative experiment. The two neurons that maintained very high [Ca2+]C while incubated in CM are the same that failed to buffer the NMDA-induced [Ca2+]C transient. Similar results were routinely observed in over 20 different preparations of CGCs.

Substitution of Na⁺ with NMG⁺ Exacerbates Glutamate-Induced Destabilization of Ca²⁺ Homeostasis and Excitotoxicity. When the Na⁺ in the extracellular medium of CGCs was replaced with NMG⁺(NMG-Locke's), a 15-min exposure to glutamate (100 µM glutamate plus 10 µM glycine in the absence of Mg²⁺) at 37°C resulted in a permanent destabilization of Ca²⁺ homeostasis (Fig. 2A). When the test of neuronal viability based on propidium iodide staining was performed 24 h after the neuronal challenge with glutamate, it was observed that many neurons were propidium iodide-negative in spite of evident morphological changes, such as a prominent shrinkage of the cell body (see white arrowheads in Fig. 2, A2 and E2). The lack of propidium iodide staining in such neurons complicated the quantitation of excitotoxicity. Obviously, 24 h after the challenge with glutamate the process of neurodegeneration was not yet accomplished. Therefore, additional experiments were performed in which neuronal viability was monitored for up to 70 h after the glutamate exposure (Fig. 2E). It was observed that of 182 neurons exposed to glutamate and NMG-Locke's, 115 died within the first 24 h, and an additional 45 during the next 46 h (Fig. 2F).

View larger version (99K): [in this window] [in a new window]   Fig. 2.   Replacement of Na+ with NMG+ exacerbates the glutamate-induced destabilization of Ca2+ homeostasis and excitotoxicity. A, B, and C, relationships between Ca2+ homeostasis and neuronal survival. [Ca2+]C was monitored in the CGCs indicated by the arrowheads in the oblique illumination images (A1, B1, and C1). Cells were exposed for 15 min at 37°C to 100 µM glutamate + 10 µM glycine (GLU) applied in a Mg-free Locke's buffer in which Na+ was replaced with NMG+ (NMG-L) when 1.3 mM extracellular Ca2+ was present (A), or absent (B), or in a buffer in which both Na+ and Ca2+ were present (C). Cell viability was tested 24 h after the exposure by inspecting the same microscopic field (A2, B2, and C2). The cells with a damaged PM were identified using propidium iodide fluorescence (A3, B3, and C3). Using a procedure of digital subtraction described in Materials and Methods, the images of propidium iodide fluorescence were superimposed over the oblique illumination images (A4, B4, and C4). Note that negatives of the propidium iodide fluorescence are created, and therefore the previously bright spots become black. This procedure allows one to distinguish the "false" propidium iodide signals that represent cellular debris floating in the medium. Such debris is visible in images B3 and C3. A5, B5, and C5, show the GLU-induced [Ca2+]C transients (F334/F380 ratio) recorded from the cells indicated by the arrowheads in A1, B1, and C1, respectively. All experiments were repeated at least three times with similar results on different preparations of CGCs. Note that many neurons showing a profound morphological neurodegeneration (see arrowheads in A2) were still impenetrable to propidium iodide (A3), which may indicate that the process of PM disintegration in these neurons was not yet accomplished. D, E, F, progress of excitotoxicity in time. CGCs have been exposed for 15 min to GLU while incubated in Na-L (D) or NMG-L (E) ([Ca2+]C was not monitored in this experiment). The superimposed images of oblique illumination and propidium iodide fluorescence D2, D3, E2, and E3 were constructed as described in Materials and Methods and illustrated in A-C. The scale bars shown in A1, D1, and E1 represent 50 µm. Average data from experiments analogous to those shown in D and E are presented in F. Data in F are means ± S.E.M. from three different dishes; each microscopic field was occupied by 53 to 85 neurons and a total of 190 to 239 neurons per each treatment was counted. Note that in many neurons that die following the exposure to GLU/NMG-L, the PM is impermeable for propidium iodide for at least 24 h (white arrowheads in E2 and E3). Note also that during the time course of the observation some neurons lost the propidium iodide stain due to disintegration (black arrowheads in D2 and E2). **p < .01 compared with respective control, p < .01 compared with neuronal viability 24 h after the GLU/NMG-L challenge (one-way ANOVA followed by Newman-Keuls test).

View larger version (54K): [in this window] [in a new window]   Fig. 3.   The glutamate-induced Ca2+ influx and excitotoxicity in CGCs incubated in NMG-Locke's is blocked by MK-801. A, 1-4 show the F334/F380 ratio images of fura-2 fluorescence from CGCs exposed to glutamate and NMG-Locke's the same way as described in Fig. 2 A. 5 and 6, show the oblique illumination images of the same cells at various times during the experiment. B, cells treated as in A but glutamate is applied with 10 µM MK-801. F334/F380 ratio images were taken at the following times: A1 and B1, CGCs incubated in CM; the arrowhead points to the cell that represents the abnormal population of CGCs with high basal [Ca2+]C described in the text; A2 and B2, 8 min after application of Na-Locke's; A3, 15 min after addition of Mg-free NMG-Locke's containing 100 µM glutamate and 10 µM glycine; B3, same as A3 but in the presence of MK-801; A4 and B4, 20 min after glutamate/glycine withdrawal, the cells incubated in Na-Locke's. Times at which the oblique illumination images were taken: A5 and B5, cells incubated in CM before the experiments; A6 and B6, 19 h after the challenge with glutamate, cells incubated in CM containing 10 µM propidium iodide; A7 and A8, 29 and 44 h after the challenge, respectively; B7, 68 h after the challenge, B8, same cells as shown in B7 but 2 min after addition of 0.01% Triton X-100. Cells indicated by arrowheads in A4 and B3 are described in the text. Scale bars in A5 and B5 represent 20 µm. Images 6, 7, and 8 of A and B are overlays of the propidium iodide fluorescence images over the oblique illumination images and were created as described in Fig. 2. Note that the CGCs that died first (A6) were those that failed to promptly restore low [Ca2+]C following glutamate withdrawal (A4) and that MK-801 completely suppressed the glutamate-induced Ca2+ influx (B3) and excitotoxicity (B7) in the majority of CGCs.

Simultaneous Monitoring of E_m and [Ca²⁺]_C. We have previously observed that activation of NMDA or kainate/-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid receptors in CGCs results in an elevation of [Na⁺]_C up to 60 mM or higher (Kiedrowski et al., 1994), which depolarizes the PM. While evaluating various means of monitoring E_m changes, I was concerned that a direct electrophysiological approach might not faithfully reflect the E_m changes characteristic for an intact neuron exposed to glutamate because the ionic content of the intracellular electrode, in particular high [K⁺], might obscure the glutamate-induced changes in [Na⁺]_C and [K⁺]_C characteristic for an intact neuron. Therefore, to monitor E_m for the purpose of this study, a noninvasive method was chosen, which makes use of the fluorescence of an E_m-sensitive dye, DiBAC₄(3), an anionic fluorescent probe that accumulates in the cytoplasm of depolarized cells via a Nernst equilibrium-dependent uptake (Bräuner et al., 1984). [Ca²⁺]_C and E_m were monitored simultaneously according to the approach described by Laskey et al. (1992). Cells were first loaded with fura-2 and then exposed to 100 nM DiBAC₄(3) (see Materials and Methods for details). After illumination at 334 and 380 nm to excite fura-2, there was no fluorescence emission by DiBAC₄(3), and inversely, after 488 nm illumination to excite DiBAC₄(3), there was no fluorescence emission by fura-2 (Fig. 4, B and C); therefore [Ca²⁺]_C and E_m could be monitored selectively in a single cell. PM depolarization was associated with a robust increase in DiBAC₄(3) fluorescence, which was very intense in peripheral parts of cell bodies but virtually absent in the nuclear area (Fig. 4B). The apparent lack of DiBAC₄(3) fluorescence in the nucleus of depolarized cells is consistent with earlier observations (Bräuner et al., 1984) and was confirmed using confocal microscopy (data not shown). Therefore, the peripheral parts of cell bodies were chosen to monitor DiBAC₄(3) fluorescence as an index of E_m. To test the reliability of these E_m and [Ca²⁺]_C measurements, the E_m changes were imposed on CGCs by applying an unselective monovalent cation ionophore, gramicidin D (5 µM), and a Na- and Ca-free solution in which the K⁺ concentration was changed from 163.2 to 3.6 mM by substituting K⁺ with NMG⁺. Under these conditions, 163.2 mM K⁺ is expected to completely depolarize the PM, whereas 3.6 mM K⁺ is expected to create a large, negative inside, K⁺ diffusion potential. Repetitive applications of depolarizing pulses of 163.2 mM K⁺ or hyperpolarizing pulses of 3.6 mM K⁺ were associated with respective increases or decreases in DiBAC₄(3) fluorescence intensity, indicative of the expected E_m changes (Fig. 4E). These changes in E_m failed to affect [Ca²⁺]_C in the majority of CGCs (Fig. 4E). In the above-described abnormal 4% subpopulation of CGCs with high [Ca²⁺]_C while incubated in the CM, however, the hyperpolarizing pulses of 3.6 mM K⁺ were associated with [Ca²⁺]_C transients (data not shown), which, considering that the extracellular medium was Ca-free, represented Ca²⁺ release from intracellular stores. Similar Ca²⁺ transients were observed in cells that were about twice as large as CGCs and probably represented astrocytes (Fig. 4D). The lack of such intracellular Ca²⁺ release in the majority of CGCs confirms our earlier observations that intracellular Ca²⁺ stores in unstimulated CGCs are virtually depleted (Kiedrowski and Costa, 1995). The mechanism of the PM hyperpolarization-induced Ca²⁺ release from intracellular stores requires further investigation.

View larger version (39K): [in this window] [in a new window]   Fig. 4.   The method of simultaneously monitoring [Ca2+]C and Em. A, a magnified portion of an oblique illumination image of granule neurons in which [Ca2+]C and Em were simultaneously monitored. The scale bar represents 10 µm. B, image of DiBAC4(3) fluorescence (left) and the overlay of DiBAC4(3) over fura-2 fluorescences (right) of CGCs shown in A depolarized with a solution containing 163 mM K+ and 5 µM gramicidin. Note that the DiBAC4(3) fluorescence is intense in the peripheral parts of cell bodies. C, image of the same CGCs hyperpolarized with a solution containing 3.6 mM K+, 159.6 mM NMG+, and 5 µM gramicidin. Note that the DiBAC4(3) fluorescence in C is much less intense than in B. D, an oblique illumination image of a group of cerebellar granule cells and of a larger cell, indicated by the arrowhead, probably an astrocyte. [Ca2+]C and Em data from this presumed astrocyte are shown in E. The scale bar represents 10 µm. E, shown are average changes ± S.E.M. in [Ca2+]C and in the DiBAC4(3) fluorescence intensity (F488) monitored in 17 CGCs (including the three cells shown in A, B, and C) and in a single larger cell morphologically resembling an astrocyte (see D). The data were obtained in a single experiment representative of three such experiments performed on different preparations of CGCs. Arrows marked B and C indicate the times at which the images shown in B and C, respectively, were taken. The cells were incubated in CM, which was replaced with Na-Locke's containing 100 nM DiBAC4(3), and then the cells were exposed to Ca-free media containing 5 µM gramicidin D (Gramic/Ca-free) that either depolarized or hyperpolarized the PM. The depolarizing medium contained 5 µM gramicidin D, 3 mM EGTA, 134.2 mM K-gluconate,25.4 mM KCl, 1 mM MgCl2, 3.6 mM KHCO3, 10 mM HEPES, pH 7.2, adjusted with Tris (K 163.2). In the hyperpolarizing medium [K+]E was reduced to 3.6 mM by substituting K+ with NMG+ (K 3.6/NMG). Note that increases or decreases in the DiBAC4(3) fluorescence intensity faithfully reflect the expected changes in Em. Note also that hyperpolarization of the PM in the presumed astrocyte, but not in CGCs, is associated with a [Ca2+]C transient that represents Ca2+ release from an internal store. Similar [Ca2+]C transients induced by PM hyperpolarization were observed in four other astrocytes.

Substitution of Na⁺ with NMG⁺ Increases NMDA-Mediated Ca²⁺ Influx by Affecting E_m. The above-described approach of a simultaneous monitoring of [Ca²⁺]_C and E_m was used to test whether the NMG-induced destabilization of Ca²⁺ homeostasis (Figs. 2A and 3A) was caused by a dysfunction of the NaCaX or by a difference in E_m. Because in the overwhelming majority of CGCs incubated in NMG-Locke's glutamate elicited Ca²⁺ influx and excitotoxicity by activating NMDA receptors (Fig. 3 B), in subsequent experiments NMDA rather than glutamate was used as an excitatory stimulus. CGCs were exposed to NMDA (300 µM NMDA plus 10 µM glycine, in the absence of Mg²⁺); for the first 2 min, NMDA was applied in a standard Locke's buffer containing Na⁺ (Na-Locke's), and the expected [Ca²⁺]_C transient and depolarization of the PM promptly occurred (Fig. 5A). Then Na⁺ was replaced with Li⁺ or NMG⁺, neither of which supports the NaCaX, but the remaining components of the previous solution were not changed. If the destabilization of Ca²⁺ homeostasis observed in CGCs exposed to glutamate when Na⁺ is replaced with NMG⁺ is caused by an inhibition of Ca²⁺ extrusion via the NaCaX, a replacement of Na⁺ with Li⁺ should yield an effect similar to that obtained by replacement of Na⁺ with NMG⁺. The switch from Na⁺ to NMG⁺ affected [Ca²⁺]_C and E_m differently, however, than the switch from Na⁺ to Li⁺ (Fig. 5A). In the presence of Ca²⁺, NMG⁺ elicited a robust increase in [Ca²⁺]_C, and transiently hyperpolarized the PM; in the absence of Ca²⁺, the NMG-dependent hyperpolarization persisted for as long as it was studied (Fig. 5A, lower). By contrast, when Na⁺ was replaced with Li⁺, NMDA depolarized the PM similarly as if Na⁺ were present (Fig. 5A, lower), but [Ca²⁺]_C decreased, within 5 min, to significantly lower levels, i.e., 360 ± 8 nM in the presence of Li⁺ (72 cells in four experiments) versus 665 ± 22 nM in the presence of Na⁺ (74 cells, four experiments), respectively (P < .001, Mann-Whitney rank sum test; see Fig. 5A, upper for representative data).

View larger version (24K): [in this window] [in a new window]   Fig. 5.   Replacement of Na+ with NMG+ during neuronal exposure to NMDA hyperpolarizes the PM and stimulates Ca2+ influx. A, effects of NMG+ and Li+ on [Ca2+]C (upper) and on Em (lower) in CGCs exposed to NMDA; NMDA (300 µM NMDA + 10 µM glycine, Mg-free medium), Li-L (Na-free Locke's buffer, Na+ replaced with Li+). Data are the means ± S.E.M. from about 20 neurons in each experiment and were reproduced four times using two different preparations of CGCs. B, substitution of Na+ with NMG+ enhances NMDA-elicited 45Ca2+ accumulation, but substitution of Na+ with Li+ has the opposite effect. 45Ca2+ accumulation was measured as described in Materials and Methods in CGCs exposed for 15 min to 300 µM NMDA + 10 µM glycine applied in Mg-free Locke's buffers supplemented with the indicated concentrations of Li+ (NMDA/Li) or NMG+ (NMDA/NMG); Na+ concentration was, respectively reduced to maintain osmoticity. To measure the basal 45Ca2+ accumulation in the presence of Li+ (basal/Li) or NMG+ (basal/NMG), NMDA and glycine were omitted and 1 mM Mg2+ + 10 µM MK-801 were added to the medium. Data are the means ± S.E.M. from three experiments on three different preparations of CGCs.

Relationships among E_m, [Ca²⁺]_C, and pH_C. Because NMG⁺, in contrast to Li⁺, does not support the operation of the NaHX (Aronson, 1985), one might speculate that a cytoplasmic acidification, caused by a tonic inhibition of NaHX by NMG⁺ (Raley-Susman et al., 1991), might affect Ca²⁺ homeostasis and contribute to the observed effects of substituting of Na⁺ with NMG⁺ versus Li⁺. To test whether this is the case, the effects of Li⁺ or NMG⁺ on NMDA-induced changes in [Ca²⁺]_C and pH_C were measured simultaneously using the Ca²⁺- and the pH-sensitive fluorescent dyes fura-2 and BCECF, respectively (see Materials and Methods for details). Basal pH_C in CGCs incubated in a standard Locke's buffer was 7.04 ± 0.01 (n = 340), which is consistent with previous observations (Raley-Susman et al., 1991; Hartley and Dubinsky, 1993; Irwin et al., 1994). While performing a simultaneous calibration of fura-2 and BCECF data in situ, it was observed that when pH_C dropped below 7.0, the maximal F₃₃₄/F₃₈₀ ratio of fura-2 (R_max) progressively decreased (Fig. 6 A). It appears that a drop of pH_C below 7.0 affects the fura-2 fluorescence properties and, therefore, the in situ calibration of fura-2 performed at a pH_C of about 7.2 to 7.4 is not valid when pH_C drops below 7.0. Although in this report it was not attempted to correct for this artifact, the pH_C and the [Ca²⁺]_C data have been displayed simultaneously, and one can determine when the fura-2 F₃₃₄/F₃₈₀ ratio might have been affected by the excessive drop in pH_C.

View larger version (22K): [in this window] [in a new window]   Fig. 6.   Relations between pHC and [Ca2+]C in CGCs. A, simultaneous in situ calibration of fura-2 and BCECF fluorescences. Cells were exposed to CM, Na-L, and then to the buffers adjusting the fura-2 F334/F380 ratio to the minimal (Rmin), or maximal (Rmax) value (see Materials and Methods for details). Then the plasma membrane was permeabilized for H+ by application of 10 µM nigericin and 10 µM CCCP, and the pH of the Rmax buffer was changed as indicated. The black ([Ca2+]C) and the white (pHC) circles represent means from 56 neurons. S.E.M.s in this and several other panels of Figs. 6 and 7 are not visible because they are smaller than the areas covered by the symbols representing data points. Note that a drop in pH below 7.0 affects Rmax. B, destabilization of [H+] equilibria between cytoplasmic organelles and cytosol and between cytosol and extracellular medium by application of 10 µM CCCP + oligomycin (3 µg/ml) affects pHC but not[Ca2+]C. Data are the means from 47 neurons. The [Ca2+]C values next to the abscissa in this as well as other panels were calculated based on an in situ calibration that was performed at pH 7.2. It has to be stressed, that in all cases when pHC dropped below 7.0, such [Ca2+]C values are underestimated, and are given only for the purpose of comparison with other experiments in which pHC was not monitored. C, An influx of Na+ or Cs+ via the NMDA receptor channels rapidly removes the CCCP/oligomycin-induced acidification of the cytoplasm in CGCs incubated in NMG-L. Shown are overlapping data from two representative experiments testing the effects of Na+ (60 neurons) or Cs+ (56 neurons) on pHC and [Ca2+]C. Because before the application of NMDA with Na-L or a Na-free Locke's buffer in which Na+ was replaced with Cs+ (Cs-L) similar patterns of pHC and [Ca2+]C changes were observed in both experiments, therefore, for clarity, the initial parts or the pHC and [Ca2+]C traces from the experiment in which Cs-L was applied (white and black circles) have been omitted. Data are the means ± S.E.M. Note that the application of Cs-L affects pHC the same way as the application of Na-L does; in the presence of Cs-L, however, [Ca2+]C decreases more rapidly. An interpretation of these results is given in the text. D, an NMDA-induced Ca2+ influx causes a more profound and more rapid drop in pHC as compared with the drop in pHC caused by an inhibition of the plasma membrane Na+/H+ exchanger with NMG+. At the end of this experiment, CGCs were exposed to the MDC, which was Ca-free and contained 10 µM CCCP + 3 µg/ml oligomycin (for details see Materials and Methods). Data are the means from 38 neurons. Note that MDC causes a rapid alkalization of the cytoplasm.

Relationships among pH_C, Mitochondrial Ca²⁺ Overload, Plasma Membrane Na⁺/Ca²⁺ Exchange Operation, and Energy Metabolism in CGCs Exposed to NMDA. Because glutamate excitotoxicity is associated with Ca²⁺ sequestration in mitochondria (Kiedrowski and Costa, 1995; Schinder et al., 1996; White and Reynolds, 1996), which has been causally linked to neuronal death (Stout et al., 1998), it was necessary to determine how substitution of extracellular Na⁺ with NMG⁺ versus Li⁺ would affect the amounts of Ca²⁺ diverted to mitochondria. For this purpose, at the end of the 5-min period of monitoring [Ca²⁺]_C and pH_C in CGCs exposed to NMDA under various ionic conditions, the cells were treated with MDC, a Na-Locke's buffer that was Ca-free and glucose-free and contained 10 µM CCCP, 3 µg/ml oligomycin, and 10 µM MK-801. Any increase in [Ca²⁺]_C in response to the MDC application was expected to result from Ca²⁺ released from the depolarized cytoplasmic Ca²⁺ stores, including mitochondria.

View larger version (25K): [in this window] [in a new window]   Fig. 7.   Relationships between mitochondrial Ca2+ and pHC in CGCs exposed to NMDA. A, in CGCs exposed to NMDA for 5 min only, MDC fails to elicit an increase in [Ca2+]C, which indicates that the exposure to NMDA was not associated with a mitochondrial Ca2+ overload. Data are the means from 43 neurons from a representative experiment. Compare with Fig. 6D and note that the NMDA-induced cytoplasmic acidification and the MDC-induced alkalization are much smaller in CGCs incubated in Na-L than in CGCs incubated in NMG-Locke. B, when MDC was applied to CGCs exposed to NMDA for 15 min, three populations of neurons could be distinguished based on the MDC effects on [Ca2+]C and pHC: 1) an instant increase in [Ca2+]C and the most rapid alkalization of the cytoplasm (12 cells, ); 2) a delayed increase in [Ca2+]C and a slower rate of cytoplasmic alkalization (16 cells, ); and 3, a rapid drop in [Ca2+]C not associated with any major change in pHC (16 cells, ). Data are the means ± S.E.M. from a representative experiment. C, when CGCs were deprived of glucose and exposed to NMDA for 5 min, MDC caused an instant increase in [Ca2+]C and a rapid alkalization of the cytoplasm in all of CGCs. Data are the means from 46 neurons from a representative experiment. D, when the glucose-deprived CGCs were exposed to NMDA in the presence of Li-L, MDC caused a drop in [Ca2+]C and failed to alkalize the cytoplasm. Data are the means ± S.E.M. from 57 neurons from a representative experiment.

Mitochondrial Ca²⁺ Overload in Energetically Challenged Neurons Results from Ca²⁺ Influx via Reverse Operation of Plasmalemmal NaCaX. Because the principal goal of in vitro studies of excitotoxicity is an understanding of the mechanisms of neurodegeneration characteristic of an ischemic brain, in which glutamate receptors are activated while the system is deprived of energy, it was of interest to examine how mitochondria would contribute to Ca²⁺ homeostasis and how pH_C would change under conditions of energy deprivation. To this end, [Ca²⁺]_C and pH_C were monitored in CGCs exposed to NMDA under glucose-free conditions. In such CGCs, following NMDA addition the [Ca²⁺]_C transient did not stabilize at a plateau level as observed when glucose was present (Fig. 7A) but after 2 to 3 min started to increase and during the 5th min of exposure [Ca²⁺]_C approached 1040 ± 57 nM, n = 46 (Fig. 7C); the steady increase in [Ca²⁺]_C was more clearly visible when the exposure to NMDA was longer than 5 min (data not shown). The pH_C changes in the majority of CGCs exposed to NMDA for 5 min were not affected by glucose deprivation (compare Fig. 7, A and C); however, glucose deprivation dramatically changed the effect of MDC addition on [Ca²⁺]_C and pH_C. When MDC was applied, [Ca²⁺]_C abruptly increased to fura-2 saturating levels, and this was accompanied by a rapid cytoplasmic alkalization that overshot the basal pH_C levels in 46 of 49 neurons (Fig. 7C). Note that the opposite result was observed in CGCs exposed to NMDA in the presence of glucose (Fig. 7A). In only three of 49 glucose-deprived neurons MDC caused a decrease in [Ca²⁺]_C accompanied by only a slight increase in pH_C (data not shown).

NMDA Elicits a Ca-Dependent K⁺ Efflux. From Fig. 5A (bottom) it is apparent that when all extracellular Na⁺ was replaced with NMG⁺, NMDA was still able to depolarize the PM, provided that Ca²⁺ was present in the medium. A possible mechanism might involve a Ca-dependent collapse of the K⁺ concentration gradient across the PM, because the NMDA receptor channels are permeable to K⁺ (Mayer and Westbrook, 1987; Tsuzuki et al., 1994). For this to be considered a valid explanation, one has to demonstrate first that the K⁺ concentration gradient indeed collapses. To test this possibility, [K⁺]_C were measured using a K⁺-sensitive fluorescent dye, PBFI (Minta and Tsien, 1989). The suitability of PBFI as a probe to measure [K⁺]_C was tested by monitoring PBFI fluorescence in CGCs exposed to glutamate. When glutamate was applied to CGCs incubated in Na- Locke's, no decrease in the F₃₃₄/F₃₈₀ ratio of PBFI fluorescence was observed (Fig. 8A). Such an outcome might be caused by the poor selectivity of PBFI for K⁺ over Na⁺. The in vitro determined dissociation constant (K_d) of PBFI for K⁺ is 8 mM and for Na⁺ is 21 mM (Minta and Tsien, 1989), therefore, one may expect only a small change in PBFI fluorescent properties when intracellular K⁺ is replaced by Na⁺. Indeed, as soon as Na⁺ was substituted with Li⁺, an ion of low affinity for PBFI, K_d = 380 mM (Minta and Tsien, 1989), a prompt decrease was observed in the F₃₃₄/F₃₈₀ ratio (Fig. 8A), which indicates an efflux of K⁺ plus Na⁺ from the cells and a simultaneous influx of Li⁺. Using the in situ calibration as shown in Fig. 8A, the K_d of PBFI for K⁺ was calculated as being 17.5 ± 0.8 mM (29 experiments on seven different preparations of CGCs). Due to this high affinity of PBFI for K⁺, when [K⁺]_C exceeded 100 mM, PBFI was practically saturated with K⁺, and changes in [K⁺]_C could not be detected unless [K⁺]_C dropped to 80 mM or lower.

View larger version (19K): [in this window] [in a new window]   Fig. 8.   [K+]C in CGCs exposed to glutamate. A, method of monitoring [K+]C using PBFI. CGCs loaded with PBFI (see Materials and Methods for details) were exposed to 100 µM glutamate + 10 µM glycine under Mg-free conditions (GLU) as described in Fig. 2. At the end of the experiment, [K+]C was calibrated in situ using 5 µM gramicidin D and buffers containing the indicated K+ concentrations (see Materials and Methods for details). Data are the means ± S.E.M. from 40 neurons. Note that glutamate failed to decrease the F334/F380 ratio when applied in the presence of Na+, but a prompt decrease in the F334/F380 ratio was observed when Na+ was replaced with Li+, which reflects the fact that PBFI has a high affinity for K+ and Na+, but not for Li+. B, NMDA-elicited K+ efflux is blocked by MK-801 (MK). CGCs were exposed to NMDA, as described in Fig. 5, in the presence or absence of 10 µM MK-801. Data are the means ± S.E.M. from about 40 neurons in each experiment. C, when Na+ is replaced with Li+, [K+]C in CGCs exposed to NMDA adjusts to ambient K+ concentrations. In CGCs exposed to NMDA as in B, the [K+]E was increased from 5.6 mM to 60 mM (K 60), followed by the addition of 1 mM ouabain (Ouab). Data are the means ± S.E.M. from 26 neurons. Note that the addition of ouabain failed to affect [K+]C, which indicates that the K+ pumping by Na+/K+ ATPase was already inhibited by Li+ alone. D, when Na+ is replaced with NMG+, glutamate elicits a slow and Ca-dependent K+ efflux from CGCs. CGCs were exposed to 100 &#