It has been suggested that augmented nerve cell death in neurodegenerative diseases might result from an impairment of mitochondrial function. To test this hypothesis, we investigated age-dependent changes in neuronal survival and glutamate effects on Ca2+ homeostasis and mitochondrial energy metabolism in cultured hippocampal neurons from diploid and trisomy 16 (Ts16) mice, a model of Down’s syndrome. Microfluorometric techniques were used to measure survival rate, [Ca2+]ilevel, mitochondrial membrane potential, and NAD(P)H autofluorescence. We found that Ts16 neurons die more than twice as fast as diploid neurons under otherwise identical culture conditions. Basal [Ca2+]i levels were elevated in Ts16 neurons. Moreover, in comparison to diploid neurons, Ts16 neurons showed a prolonged recovery of [Ca2+]iand mitochondrial membrane potential after brief glutamate application. Glutamate evoked an initial NAD(P)H decrease that was found to be extended in Ts16 neurons in comparison to diploid neurons. Furthermore, for all age groups tested, glutamate failed to cause a subsequent NAD(P)H overshoot in Ts16 cultures in contrast to diploid cultures. In the presence of cyclosporin A, an inhibitor of the mitochondrial membrane permeability transition, NAD(P)H increase was observed in both diploid and Ts16 neurons. The results support the hypothesis that Ca2+ impairs mitochondrial energy metabolism and may play a role in the pathogenesis of neurodegenerative changes in neurons from Ts16 mice.
- trisomy 16
- Down’s syndrome
- Alzheimer’s disease
- hippocampal culture
- mitochondrial membrane potential
The presence of an extra copy of human chromosome 21 [trisomy 21 (Ts21)] leads to the genesis of Down’s syndrome, a disorder associated with mental retardation, facial dysmorphology, and congenital heart disease. Individuals with Down’s syndrome are known to have a tendency to develop neuropathological features of Alzheimer’s disease in the third decade of life. This observation has led to the suggestion that the overexpression of gene products from chromosome 21 is responsible for the early onset of Alzheimer’s disease (Richards et al., 1991). Indeed, plaques containing β-amyloid protein (βAP) have been found in the brain of Ts21 individuals (Rumble et al., 1989), presumably as a result of overexpression of the β-amyloid precursor protein (βAPP), which is encoded on chromosome 21.
The trisomy 16 mouse (Ts16) is a model of Down’s syndrome (Ts21) and to some extent also of Alzheimer’s disease (Coyle et al., 1988; Colton et al., 1990). At least nine genes mapped on human chromosome 21 are also located on mouse chromosome 16, including the genes for superoxide dismutase (SOD) and βAPP (Richards et al., 1991; Holtzman et al., 1992). As with individuals with Down’s syndrome, Ts16 fetal mice exhibit edema of the neck, inner ear anomalies, congenital heart disease, and retardation in CNS development (for review, see Epstein, 1986). The Ts16 mouse model is limited, however, by the fact that Ts16 fetuses usually die at gestation day 18–20 (Richards, 1991) because of Ts16-induced disturbances in the cardiovascular system. To overcome the limitation presented by death in utero and to increase neuronal survival times, we used dissociated hippocampal cell cultures for our studies.
Previous studies suggested that cultured neurons from Ts16 mice show altered electrogenesis possibly associated with augmented Ca2+ loading of cells (Orozco et al., 1988; Ault et al., 1989; Galdzicki et al., 1993). In such cultures, neuronal cell loss has been shown to be accelerated (Stabel-Burow et al., 1997). Furthermore, cultured hippocampal Ts16 mice neurons display an inherited defect in survival response mediated by glutamate in low concentration (Bambrick et al., 1995). Among the overexpressed proteins in this model is βAPP, which has been shown to have a role in the regulation of intracellular Ca2+ (Mattson et al., 1993b). In contrast, the βAPP product βAP, which had been demonstrated to occur in Ts16 hippocampal neurons (Richards et al., 1991), is suspected to destabilize intracellular calcium homeostasis and render neurons more vulnerable to excitotoxicity (Mattson et al., 1992).
Disturbed Ca2+ homeostasis may lead to neuronal cell loss by various mechanisms (Choi, 1995; Mattson et al., 1995). One of the possibilities is that increased [Ca2+]i causes depolarization of mitochondrial membrane and thereby disturbs the respiratory chain and the subsequent production of ATP (Aw et al., 1987; Richter and Kass, 1991; Duchen and Biscoe, 1992). Reduced ATP supply will ultimately interfere with electrolyte homeostasis and the transport processes of cellular substrates. Furthermore, a dysfunction of the mitochondrial electron transport chain will lead to a disturbance in the NAD(P)+/NAD(P)H ratio (Hansford, 1980). The associated shift in the mitochondrial redox balance will perturb normal citrate cycle metabolism, which because it is both a route of disposal and a source of the synthesis of glutamate and other neurotransmitters may have consequences for neurotransmitter synthesis and degradation (Hansford, 1985).
In this study we have investigated changes in intracellular [Ca2+] levels, mitochondrial membrane potential, and NAD(P)H after brief exposures to glutamate in cultured diploid and Ts16 hippocampal neurons. Our results demonstrate that Ts16 neurons display alterations in Ca2+ signaling and mitochondrial functions, such as the membrane potential and NAD(P)+/NAD(P)H ratio.
MATERIALS AND METHODS
Preparation of cells. A breeding scheme was established between male mice with balanced bilateral Robertsonian translocations of chromosome 16 [Rb(16.17)32LUB and Rb(11.16)2H, kindly supplied by Professor H. Winking, Medizinische Hochschule Lübeck, Germany] that were mated with NMRI females (Bundesinstitut für gesundheitlichen Verbraucherschutz und Veterinärmedizin, BgVV, Berlin, Germany). Such matings result, on average, in one of three embryos with three copies of chromosome 16 (Gropp et al., 1975). Primary hippocampal cultures were prepared at gestation day 16 from Ts16 embryos and their diploid littermates (Banker and Cowan, 1977; Peacock et al., 1979). For this procedure, embryos were removed into ice-cold GBSS solution [Gey’s balanced salt solution, containing (in mm): NaCl 136, KCl 5, MgSO4 0.3, NaH2PO4 1, CaCl2 1.5, NaHCO3 2.7, KH2PO4 0.22, MgCl2 1, glucose 5, pH 7.4] after the maternal animal was decapitated under deep ether anesthesia. The Ts16 embryos were identified by edema of the neck, smaller size, and abnormal blood supply as a result of a hyperchrome liver (Lane et al., 1996) and confirmed in initial experiments by karyotyping (Fig. 1). After preparation of diploid and Ts16 hippocampi, cells were separately dispersed by repetitive trituration with Pasteur pipettes and plated on poly-d-lysine-coated 12 mm coverslips (5–6 × 104 cells/coverslip) and incubated at 36.5°C in a humidified atmosphere of 95% air and 5% CO2 with minimum essential medium (MEM; Life Technologies, Eggenstein, Germany) supplemented with 10% (and after 3 d in culture, with 2%) heat-inactivated horse serum (Life Technologies), 12 mmglucose, 2 mm glutamine, serum extender MITO (Schubert, Schwandorf, Germany), 1 μm arabinofuranosid, and 2.5 × 104 U penicillin/streptomycin per milliliter culture medium. Cells were maintained in culture for up to 4 weeks. Most experimental data were obtained within 2–21 d in culture.
Fluorescence measurements. Microfluorimetric experiments were performed using an imaging system based on a Zeiss Axioskop microscope with 10× and 40× water-immersion objectives (numerical aperture, 0.3 and 0.75, respectively; Zeiss, Jena, Germany), a xenon light source with a combination of two monochromators [Photon Technology Instruments (PTI), Wedel, Germany], a charge-coupled device camera (Hamamatsu, Herrsching, Germany), and a photomultiplier (Seefelder Messtechnik, Seefeld, Germany). Image hardware was controlled by an IBM-compatible computer running commercial software developed by PTI.
The cells were incubated in media containing the different fluorescence dyes (Molecular Probes Europe, Leiden, Netherlands) for 10–15 min at 36.5°C. After the cells were washed for 15 min at 36.5°C using fresh media, the dyes were retained for 3–5 hr. Rhodamine 123 (Rh123) was dissolved in aqueous solution (0.1% ethanol), and cells were loaded by incubation with a final concentration of 10 μg of dye per 1 ml of culture medium (26.3 μm). Rh123 fluorescence was excited at 490 nm and measured above 530 nm using a 515 nm dichroic mirror and a 530 long-pass filter. To differentiate between living and dead cells, double-staining with the intercalating dyes acridine orange (AO) (5 μm) and ethidium bromide (EB) (10 μm) dissolved in aqueous solution were used. The two dyes were applied simultaneously, excited at 490 nm, and measured using an optical combination of a 505 nm dichroic mirror and a 515 nm long-pass filter. The membrane-permeable AO interacts in living cells with DNA (emission 515 nm, green) and RNA (emission 650 nm, red), whereas the membrane-impermeable EB binds only in dead cells to DNA (emission 605 nm, red) (Oyama et al., 1994) (Fig.2 B). For measurements of [Ca2+]i, the acetoxymethyl (AM) ester of fura-2 (final concentration 2–3 μm) was used. Fura-2 was excited at 340 nm and 380 nm, and fluorescence was measured at 510 nm. The calibration of the fura-2 fluorescence signal was performed by using an in vitro calibration procedure. Fura-2 (1 μm) as the free acid was added to saline containing Ca2+-EGTA buffers, giving minimum and saturating levels of Ca2+ and hence the minimum (R min) and maximum (R max) fluorescence ratios and also the ratio of the Ca2+-free and Ca2+-saturated fluorescence excited at 380 nm (β), required for the equation (Grynkiewicz et al., 1985): [Ca2+]i =K Dβ(R −R min/R max −R).
The value of the in vitrodissociation constant K D in the described system was close to reported data [224 nm (Duchen, 1992a)]. Autofluorescence of NAD(P)H was monitored by exciting at 360 nm and measuring light emitted above 400 nm using a 390 nm dichroic mirror and a 400 nm long-pass filter (Aubin, 1979; Duchen, 1992a).
For all fluorescence measurements, neurons were differentiated from glial cells in phase-contrast illumination [relatively small phase-dark cell body (10–15 μm) with fine processes; see Fig.2 A].
Drugs and solutions. During the experiments, cells were continuously superfused with oxygenated (95% O2, 5% CO2) artificial CSF (ACSF), containing (in mm): NaCl 124, KCl 3, NaH2PO41.25, MgSO4 2, CaCl2 2, NaHCO3 26, glucose 10, pH 7.35. Sodium glutamate (Sigma, Deisenhofen, Germany) was applied with concentrations of 10 μm to 1 mm. For a number of experiments, the following drugs were added to culture media: 30 μm 2-amino-5-phosphonovalerate (APV), 10 μm 2,3-dihydroxy-6-nitro-7-sulfamoylbenzo(F) quinoxalin (NBQX), 50 μg/ml tocopherol (vitamin E) (all from Sigma), and 0.5–1.5 μm cyclosporin A (Biomol, Hamburg, Germany). In some experiments the solution was changed to a nominally Ca2+-free saline solution. In these cases, 2 mm MgSO4 substituted 2 mmCaCl2. All experiments were performed at 30–32°C.
Data acquisition. Living neurons were identified by phase-contrast illumination and AO/EB double staining. Moreover neurons were characterized in phase-contrast illumination measurements by a relatively small, phase-dark cell body (10–15 μm) with fine processes and therefore could be easily differentiated from glial cells. If it was not clear whether the fluorescent nucleus was from a neuron or an underlying glial cell, the subfield was excluded from analysis. The AO-positive and EB-negative neurons were counted in 20–30 subfields (diameter 400 μm) of each culture dish. To approximate the total number of living neurons on the dish, the subfield mean value of AO-positive and EB-negative neurons was multiplied by the ratio of the coverslip surface (113.10 mm2) and subfield surface (0.13 mm2).
CCD camera image frames were usually obtained from groups of 3–10 optically identified neurons, digitized with 8 bits (spatial resolution up to 512 × 480 pixels), and stored on the computer hard disk. The recording frequency was adapted for the different experiments at between 0.5 and 300 images/sec. For the off-line analysis, fluorescence signals from single neurons were measured by using median values from individually adjusted regions of interest. These data were finally presented as the change in fluorescence signal from the baseline level ΔF/F 0 (Rh123) or as an absolute change of Ca2+ concentration (fura-2).
Photomultiplier NAD(P)H measurements were acquired from clusters consisting of 10–15 optically identified neurons. The recording frequency was between 2 and 10 Hz. The data were normalized to change in autofluorescence signal from the baseline level ΔF/F 0.
Statistics. All values are given as means ± SEM. Statistical differences of individual data points were assessed by using a one-way ANOVA followed by Bonferroni/Dunn comparison. To analyze statistical differences in spontaneous neuronal cell death, the slopes of the linear regression of the semilogarithmical plot, e.g., the death rate constants, were assessed by using a one-way ANOVA followed by Bonferroni/Dunn comparison.
Tocopherol inhibits spontaneous neuronal cell death in diploid and Ts16 cultures
For a quantitative analysis of spontaneous neuronal cell death in diploid and Ts16 cultures, the total number of surviving neurons on coverslips was counted every second day starting at 2 d in vitro (DIV). For every second DIV, at least two coverslips from three different preparations were evaluated by counting the number of AO-positive/EB-negative (living) and AO-negative/EB-positive (dead) neurons in 20–30 subfields of the culture dish. Out of these data the total numbers of living and dead neurons on the coverslips were approximated. For all experimental sets, the percentage decline of surviving neurons over time in culture could be fitted using a single-exponential function f(t) ∼ exp(λt). Therefore, the death of each single neuron can be taken as an independent stochastic event (Dubinsky et al., 1995). In a semilogarithmical plot, the death rate constant λ results from the slope of the linear regression line. Figure 2 C–F shows the mean values ± SEM in percentage of living neurons in control conditions and with the addition of APV/NBQX, cyclosporin A, and tocopherol to the culture medium.
Under control conditions (Fig. 2 C), the linear fits indicate a death rate constant for diploid neurons of 10.1 ± 0.5% per day and 22.7 ± 0.9% per day for Ts16 neurons (slopes of the fits are significantly different at p < 0.001). Thus, Ts16 neurons die more than twice as fast as diploid neurons under otherwise identical culture conditions. The addition of glutamate receptor antagonists APV (30 μm) and NBQX (10 μm) to the culture medium (Fig. 2 D) had no significant effect on the survival of both diploid and Ts16 neurons. Compared with the situation under control conditions, the death rate constant did not change in diploid (10.2 ± 1.2%) or Ts16 cultures (18.8 ± 2.3%). With the addition of 0.5 μm cyclosporin A (Fig.2 E), an inhibitor of the mitochondrial permeability transition, analogous results were found: the neuronal death rate constant showed no significant change in diploid (11.9 ± 1.1%) and Ts16 cultures (19.4 ± 1.4%) in comparison to control conditions. Only the application of 50 μg/ml tocopherol (Fig.2 F), an antioxidant binding preferentially to the plasma membrane, led to a significant increase in neuronal survival. Whereas the neuronal death rate constant in diploid cultures showed a reduction to 6.6 ± 0.8%, the death rate constant in Ts16 cultures was halved (9.4 ± 1.1%) in comparison to control conditions. The observed reduction in neuronal death rate constant was significant in both diploid (p < 0.01) and Ts16 cultures (p < 0.001) compared with the situation under control conditions. Furthermore, the addition of tocopherol to the culture medium abolished the significant difference between the neuronal death rate constant in diploid and Ts16 cultures observed under control conditions and in the presence of APV/NBQX or cyclosporin A.
Inhibition of the mitochondrial permeability transition prevents glutamate-induced neuronal death in diploid and Ts16 neurons
To study differences in the vulnerability of cultured hippocampal diploid and Ts16 neurons to excitotoxic damage, we applied of 50 μm glutamate for 60 min to the culture medium. Before and every 2 hr after glutamate stimulation, the number of AO-positive/EB-negative and AO-negative/EB-positive neurons was evaluated from at least two coverslips from three different preparations. The experiments were performed on diploid and Ts16 cultures between 8 and 20 DIV. The total number of living as well as dead neurons on the coverslips was approximated as described above.
Figure 3 A shows the neuronal survival in diploid and Ts16 cultures for 24 hr under control conditions. The increased death of Ts16 neurons compared with diploid neurons was already recognizable but not significantly different at this time (24 hr: diploid 95.2 ± 4.2%, Ts16 85.7 ± 6.7%). The application of 50 μm glutamate for 60 min was followed by a strong intensification of neuronal death in diploid and Ts16 cultures. Twenty-four hours after the glutamate stimulation, the proportion of surviving neurons decreased to 66.5 ± 6.6% in diploid and 30.7 ± 7.6% in Ts16 cultures (diploid vs Ts16,p < 0.001) (Fig. 3 B). In comparison to diploid cultures and the results under control conditions, the neuronal death in Ts16 cultures was found to be enhanced even 24 hr after glutamate application.
The glutamate-induced intensification of neuronal death was dependent on the extracellular Ca2+ concentration. After nominal removal of Ca2+ from the extracellular medium, the increase in neuronal cell death that followed the application of 50 μm glutamate was widely suppressed in both diploid and Ts16 cultures (24 hr: diploid 81.9 ± 6.3%, Ts16 66.5 ± 7.3%) (Fig. 3 C). Diploid and Ts16 neurons were also successfully protected against glutamate-induced excitotoxicity in the presence of the glutamate receptor antagonists APV (30 μm) and NBQX (10 μm) (24 hr: diploid 87.8 ± 5.7%, Ts16 78.8 ± 7.4%) (Fig. 3 D). The removal of extracellular Ca2+ and the presence of APV/NBQX protected neurons from glutamate-induced increases of [Ca2+]i (Choi, 1988b; Bleakman et al., 1993). Elevation of [Ca2+]i leads to a reduction and subsequent collapse of mitochondrial membrane potential (Duchen, 1992b). To study the significance of [Ca2+]i and mitochondrial function for neuronal survival, we investigated the effects of cyclosporin A, an inhibitor of mitochondrial permeability transition (Starkov et al., 1994; Pastorino et al., 1995; Nicolli et al., 1996). Cyclosporin A delays mitochondrial depolarization (Nieminen et al., 1996) and presumably prevents glutamate-induced collapse of mitochondrial membrane potential (Schinder et al., 1996). The presence of 1.5 μm cyclosporin A protected diploid and Ts16 neurons to a similar extent as did nominally Ca2+-free or APV/NBQX-containing culture medium against glutamate-induced increase in neuronal cell death (24 hr: diploid 83.7 ± 6.5%, Ts16 72.8 ± 7.9%) (Fig. 3 E). Tocopherol, in contrast to the protecting effects in unstimulated cultures, had only minor effects on the glutamate-induced increase in the death of diploid and Ts16 neurons. In the presence of 50 μg/ml tocopherol, 24 hr after the application of 50 μm glutamate, 69.6 ± 5.4% and 45.3 ± 7.9% live neurons were counted in diploid and Ts16 cultures, respectively (diploid vs Ts16, p < 0.01) (Fig. 3 F). In Ts16 cultures, tocopherol reduced the neuronal cell death in comparison to the glutamate-only control condition (amount of surviving Ts16 neurons 24 hr after glutamate application without vs with tocopherol = p < 0.01).
Age-dependent increase of basal [Ca2+]i in Ts16 neurons
The age dependence of basal [Ca2+]i and the other investigated parameters were analyzed by subdividing our culture into four age groups: I, ≤6 DIV; II, 7–12 DIV; III, 13–18 DIV; IV, ≥19 DIV. Figure 4 illustrates the intracellular Ca2+ levels monitored with fura-2 AM for diploid and Ts16 neurons in these four different age groups. For each age group of diploid and Ts16 cultures, at least 600 neurons out of three separate cultures (four different coverslips with 50–60 neurons for each culture) were studied. Neurons with incomplete dye loading (<20 nm [Ca2+]i) and dying neurons (>500 nm[Ca2+]i) were excluded from the analysis (diploid 4.1%, Ts16 5.8% of all measured neurons). In the youngest age group (up to 6 DIV), the basal [Ca2+]i did not significantly differ between diploid (group I: 93.67 ± 3.57 nm) and Ts16 neurons (group I: 77.51 ± 3.71 nm). The basal [Ca2+]i in age group II was unchanged in diploid neurons (96.99 ± 1.81 nm) and increased in Ts16 neurons (113.08 ± 3.58 nm). Ts16 neurons of age group II showed a larger basal [Ca2+]iin comparison to diploid neurons, but the difference was not significant. With further aging, the basal [Ca2+]i was stable in diploid neurons but increased steadily in Ts16 neurons. The [Ca2+]i of Ts16 neurons of groups III and IV was significantly increased in comparison to diploid control neurons (group III: diploid 94.37 ± 8.3 nm, Ts16 140.44 ± 6.5 nm, p < 0.001; group IV: diploid 93.24 ± 5.36 nm, Ts16 149.76 ± 3.37 nm, p < 0.001).
Age-dependent changes in potassium- and glutamate-induced rises of [Ca2+]i in diploid and Ts16 neurons
Fura-2 AM-loaded diploid and Ts16 neurons were exposed for 10 sec to 50 mm K+ or 100 μmglutamate. Changes in [Ca2+]i were investigated by obtaining 340/380 nm ratio images with one image per 2 sec (0.5 Hz). Figure 5 shows plots of the average changes in [Ca2+]i after stimulation with K+ (at least 30 neurons for each age group out of three different preparations) (Fig. 5 A) or glutamate (45 neurons for each age group out of six different preparations) (Fig. 5 B).
The application with 50 mm K+ for 10 sec induced a rapid increase in [Ca2+]i in all age groups of both diploid and Ts16 neurons, presumably by depolarization of the plasma membrane and activation of voltage-gated Ca2+ channels. The maximum rise in [Ca2+]i showed only minor changes in the different age groups (diploid 496–558 nm; Ts16 521–598 nm). The [Ca2+]iincrease in Ts16 neurons showed a slight delay in comparison to diploid neurons in all age groups. Furthermore, the recovery of [Ca2+]i in Ts16 neurons was prolonged in comparison to diploid neurons of the same age groups. Figure5 C illustrates that the time integral of the [Ca2+]i signal after the stimulation with K+ increases as a function of age in both diploid and Ts16 neurons. The total [Ca2+]i integral in Ts16 neurons was significantly augmented in all age groups in comparison to diploid control neurons.
In contrast to the stimulation with K+, the glutamate-induced increase in [Ca2+]ishowed a strong growth with age in culture that corresponds to the expression of glutamate receptors by cultured neurons. Thus, cultured hippocampal neurons become sensitive to NMDA after 7–10 DIV (Mattson and Kater, 1988, 1989). Therefore the rise in [Ca2+]i during and after exposure to glutamate increased with the age of the neurons. Interestingly, the amplitude of [Ca2+]i increase in each age group was somewhat larger in the diploid neurons than in Ts16 neurons (Table 1). In contrast to diploid control neurons, Ts16 neurons show both a slower rise time of [Ca2+]i and a slower recovery to baseline. The rise time of [Ca2+]i was nearly constant with age in diploid and Ts16 neurons. Decay times of [Ca2+]i became longer with daysin vitro both in diploid and Ts16 neurons but were significantly longer in Ts16 neurons than in diploid neurons in all age groups. The [Ca2+]i declines with at least two different time constants. In Table 1 both time constants, τfast and τslow, are quantified for the different age groups. There was an elevation in the integral of [Ca2+]i signal over time in Ts16 neurons as compared with diploid neurons that was independent of the initial rise in [Ca2+]i (Fig.5 D). This is caused by a slowed Ca2+recovery kinetic that is also reflected in the significant elevated time constants of both phases of Ca2+ recovery in most age groups of Ts16 cultures. Only τfast of age group II showed no significant difference between diploid and Ts16 neurons (Table 1).
Delayed recovery of mitochondrial membrane potential after glutamate-induced depolarization in Ts16 neurons
Mitochondria are the only organelles known to have a significant negative membrane potential (Chen, 1989). This potential is driven by the respiratory electron transport chain and is required for the synthesis of ATP. The lipophilic cation rhodamine 123 (Rh123) is accumulated by mitochondria in response to the negative membrane potential (Johnson et al., 1980; Chen, 1989). Binding of the accumulated dye molecules to the mitochondrial matrix is associated with a fluorescence quench (Emaus et al., 1986). Depolarization of the mitochondrial membrane allows redistribution of the dye from the mitochondria into the cytosol. This event is correlated with an increase in the Rh123 fluorescence signal. In contrast, hyperpolarization of the mitochondrial membrane will increase the uptake of the dye from the cytosol into the mitochondria and thereby increase the fraction of quenched dye. Thus hyperpolarization of mitochondrial membrane will decrease the Rh123 fluorescence signal (Duchen et al., 1993). In this study, the distribution and quenching of the Rh123 fluorescent signal was used to monitor changes in mitochondrial membrane potential.
It has been reported previously that intracellular Ca2+ accumulation will depolarize mitochondrial membrane potential and thereby increase the Rh123 signals (Duchen, 1992b). Figure 6 illustrates that the application of glutamate leads to a depolarization of the mitochondrial membrane in the presence of extracellular Ca2+ in both diploid and Ts16 neurons. This effect was lost when glutamate was applied in the presence of nominally Ca2+-free medium. The effect was reversible after reapplication of Ca2+-containing medium and did not depend on age. The Ts16 neurons showed a larger increase in the Rh123 signal and depolarization of mitochondrial membrane in comparison to diploid neurons in all age groups. As a result, in Ts16 neurons we observed an elevated depolarization of mitochondrial membranes despite the reduced initial [Ca2+]i peak.
Figure 7 illustrates the depolarization of mitochondrial membranes, as monitored by Rh123, during application of 10 μm, 100 μm, and 1 mmglutamate to diploid and Ts16 neurons in all age groups. The illustrated plots are based on mean values from at least 24 Ts16 and 36 diploid neurons out of three different cultures for each age group. The glutamate stimulus of 30 sec was immediately followed by a depolarization of the mitochondrial membrane and a relatively slow recovery of polarization to baseline. In the first age group (≤6 DIV), Ts16 neurons showed a smaller but not significant difference in the maximum Rh123 signal in contrast to diploid neurons. In age group II (7–12 DIV), maximum Rh123 signal was doubled for glutamate concentrations >50 μm in Ts16 neurons and diploid neurons. In comparison to diploid control neurons, Ts16 neurons of age groups III and IV (≥13 DIV) were characterized by a larger maximum Rh123 signal that showed an elevation with age in culture. Figure7 B summarizes concentration–response curves for maximum Rh123 signal increase after glutamate stimulation for all age groups and glutamate concentrations. The concentration–response curves leveled off above 0.5 mm glutamate in all age groups and in both cell types. After 2 weeks in culture (group III, 13–18 DIV), Ts16 neurons show a significantly larger maximum Rh123 increase compared with diploid neurons after stimulation with 50, 75, and 100 μm glutamate (Table 2). In older Ts16 neurons (group IV, ≥19 DIV), the level of depolarization of mitochondrial membrane was also found to be increased above diploid neurons for all glutamate concentrations >25 μm. There were also significant differences in the kinetics of the Rh123 signal between diploid and Ts16 neurons. Generally the rise time and recovery of Rh123 signal increased with the applied glutamate concentration. The rise time and recovery of depolarization were larger in Ts16 neurons as compared with diploid neurons. The difference in the time to the maximum of the Rh123 signal became significant at concentrations ≥0.5 mm glutamate in all age groups. The recovery of mitochondrial membrane potential was significantly prolonged only in concentrations ≤100 μm glutamate, and this difference was particularly obvious in age groups II and III (Table3).
Glutamate induces changes in NAD(P)H/NAD(P)+ratio in diploid and Ts16 neurons
Increases in [Ca2+]i may lead to intramitochondrial Ca2+ accumulation, which in turn may increase respiration via activation of different intramitochondrial enzymes of the citrate cycle, namely pyruvate dehydrogenase, NAD+-isocitrate dehydrogenase, and α-ketoglutarate dehydrogenase (Moreno-Sánchez and Hansford, 1988; Richter and Kass, 1991). Increased activity of the citrate cycle results in an increase in the NADH/NAD+ ratio (Duchen et al., 1993). We therefore measured the autofluorescence that is mediated by the NAD(P)H fraction. The autofluorescence signal measured under these conditions is derived from both mitochondrial and cytosolic NADH and NADPH. Because the autofluorescence spectra overlap, it is not possible to differentiate between the signals originating from these two; for this reason we refer to NAD(P)H, indicating that the signals are derived from either NADH or NADPH or both. Under these conditions, an increase in autofluorescence signal indicates an increase in the reduced state of the pyridine nucleotide, i.e., NAD(P)H, and a decrease in autofluorescence signal indicates an increased oxidation to NAD(P)+.
Figure 8 represents changes in NAD(P)H autofluorescence induced by application of 100 μmglutamate in all age groups for diploid and Ts16 neurons (n ≥ 9 out of four different cultures for both diploid and Ts16 neurons). Single characteristic recordings are shown. The glutamate stimulus of 30 sec induced an immediate decrease of NAD(P)H autofluorescence signal in both diploid and Ts16 neurons. The amount of NAD(P)H decline showed no significant difference between diploid and Ts16 neurons (Fig. 8 C). However, in contrast to diploid neurons, Ts16 neurons displayed a prolonged recovery. The duration of NAD(P)H signal recovery increased with age in culture. Ts16 neurons of age groups II, III, and IV (≥7 DIV) required a significantly longer time for recovery to baseline than diploid neurons (II: diploid 190 ± 58 sec, Ts16 1090 ± 239 sec, p < 0.001; III: diploid 158 ± 48 sec, Ts16 853 ± 193 sec,p < 0.001; IV: diploid 340 ± 64 sec, Ts16 1768 ± 238 sec, p < 0.001) (Fig.8 D). In diploid neurons, an overshooting response with an increase in the NAD(P)H signal was noted, pointing to a secondary Ca2+ or glutamate-dependent stimulation of the citrate cycle. This overshooting response was dependent on age in culture in both amplitude of NAD(P)H autofluorescence increase and rise time to peak. Except for the youngest age group, such overshooting responses were not observed in Ts16 cultures, suggesting a reduction or loss of compensatory citrate cycle activation (Fig.8 E,F).
Effects of cyclosporin A on glutamate-induced changes in NAD(P)H/NAD(P)+ ratio
Depolarization of mitochondrial membranes after glutamate-induced rises in [Ca2+]i and Ca2+ accumulation in mitochondria may result in the opening of permeability transition pores. Mitochondrial membrane permeability transition is thought to mediate oxidative damage to mitochondria and therefore induce neuronal cell death (Takeyama et al., 1993; Ankarcrona et al., 1996; Schinder et al., 1996). The immunosuppressant drug cyclosporin A has been demonstrated to inhibit the nonspecific mitochondrial permeability transition (Broekemeier et al., 1992; Kass et al., 1992). Therefore we used cyclosporin A to investigate the role of mitochondrial membrane permeability transition for the glutamate-induced reduction in NAD(P)H autofluorescence signal in Ts16 neurons in comparison to diploid neurons.
Figure 9 A shows that in the presence of 1.5 μm cyclosporin A, no NAD(P)H decrease was observed in either diploid or Ts16 neurons in response to 100 μm glutamate. Instead, an initially slow and finally rapid increase in NAD(P)H autofluorescence signal was measured. In the presence of cyclosporin A, the maximum level and duration of the glutamate-induced rise in NAD(P)H signal were elevated. Furthermore, we observed no significant age dependence in NAD(P)H signal response to glutamate except for age group I (≤6 DIV). This supports the idea that a [Ca2+]i increase is required to change the NAD(P)H signal in the presence or absence of cyclosporin A. In Ts16 neurons (age group II–IV), however, the NAD(P)H maxima was reduced by ∼15% in comparison with diploid neurons (Fig.9 B). Moreover, Ts16 neurons showed a significant delay in NAD(P)H signal increase (mean time to maximum for age group II–IV: diploid 153 ± 21 sec, Ts16 262 ± 26 sec; p< 0.001 for each age group, diploid vs Ts16 neurons) (Fig.9 C). The findings imply that mitochondrial permeability transition may be involved in the absence of the NAD(P)H signal overshoot after application of glutamate in Ts16 neurons.
The present study demonstrates the important role of changes in calcium homeostasis and mitochondrial function in neuronal cell loss occurring in hippocampal cell cultures from Ts16 mice. In particular, our study describes alterations in the NAD(P)H autofluorescence signal, which served as a marker of mitochondrial energy metabolism. A connection between disturbances of intracellular Ca2+ regulation, mitochondrial dysfunction, and neuronal cell death in Ts16 cultures is suggested.
Survival of diploid and Ts16 hippocampal neurons in culture
Ts16 neurons display a significantly increased death rate in comparison to diploid control neurons under culture conditions. Glutamate receptor antagonists APV and NBQX as well as cyclosporin A, an inhibitor of the mitochondrial permeability transition, had only minor effects on the observed death rate in diploid and Ts16 cultures. In contrast, tocopherol (vitamin E) protected Ts16 cultures against the augmented neuronal loss. The neuroprotective effect from the membrane-localized antioxidant tocopherol indicates an elevated concentration of reactive oxygen species (ROS) in Ts16 cultures. An increased generation of ROS in cortical neurons from fetal Down’s individuals has been suggested to cause neuronal apoptosis in vitro (Busciglio and Yankner, 1995). Increased ROS levels in culture may result from increased production or a reduced disposal of ROS molecules or both. Several investigators have proposed that the triplication of Cu/Zn-SOD in Ts16 mice and Down’s individuals results in an elevated level of ROS (Sinet, 1982; Groner et al., 1994;Bar-Peled et al., 1996; Peled-Kamar et al., 1997). Furthermore, in transgenic mice with an elevated level of Cu/Zn-SOD, a disruption in cellular ROS metabolism has been demonstrated (Avraham et al., 1988,1991; Peled-Kamar et al., 1995; Lotem et al., 1996). This finding may result from the capability of Cu/Zn-SOD to catalyze the formation of ROS using anionic scavengers and H2O2 as substrates (Yim et al., 1993). In Ts16 cultures, an increased production of superoxide radicals by microglial cells has been shown (Colton et al., 1990). Furthermore, the observation that cultured Ts16 neurons possess a significantly reduced level of the intracellular ROS-scavenger glutathione in comparison to diploid neurons (Stabel-Burow et al., 1997) also points to elevated levels of ROS.
Age-dependent changes in basal [Ca2+]i in Ts16 neurons
Neuronal Ca2+ homeostasis is regulated by Ca2+ influx through voltage-activated and receptor-gated Ca2+ channels and Ca2+ efflux via the Na+/Ca2+ exchanger and ATP-dependent Ca2+ pumps. Furthermore, [Ca2+]i is buffered by ATP-dependent Ca2+ transport into intracellular stores and binding to intracellular proteins (for review, see Carafoli, 1987). We have not yet studied the mechanisms underlying the age-dependent basal [Ca2+]i increase in Ts16 neurons that we have described in this study. Changes in electrical properties in cultured hippocampal Ts16 neurons have been reported (Galdzicki et al., 1993). Ts16 neurons show an abnormal action potential and an increased plasma membrane Ca2+ conductance (Rapoport and Galdzicki, 1994). Ca2+ shift into mitochondria has been shown as an important part of intracellular Ca2+ regulation (Gunter et al., 1994; White and Reynolds, 1995). A reduced buffering capacity or elevated mitochondrial Ca2+ release may result in the observed [Ca2+]i increase in Ts16 neurons. Such mitochondrial Ca2+ dysregulation has been reported to be caused by an elevated ROS concentration in Ts16 neurons (Weis et al., 1994).
Previous studies have shown an abnormal calcium homeostasis in astrocytes from Ts16 cultures (Bambrick et al., 1997; Müller et al., 1997). Thus the average basal [Ca2+]i in Ts16 astrocytes was more than twice as high as in diploid astrocytes. Furthermore, elevated amounts of calcium were observed in endoplasmatic reticulum Ca2+ stores in Ts16 astrocytes that may result from increased intracellular Ca2+ load or augmented mitochondrial Ca2+ efflux (Bambrick et al., 1997).
An interesting possibility is a destabilized calcium homeostasis attributable to the overexpression of βAPP as reported by Mattson and colleagues (1993a), which is of particular interest in view of the fact that βAPP is overexpressed in Ts16 mice. It has been shown that βAPP expression is regulated by development (Holtzman et al., 1992). Therefore, changes in the expression of βAPP during neuronal development may result in the significantly increased basal [Ca2+]i in Ts16. To our knowledge there are no data available that describe an effect of βAPP on basal [Ca2+]i.
Considering that Ca2+-dependent elevation in neuronal death is reported when [Ca2+]i rises above 300 nmfor longer times (Johnson et al., 1992), we assume that increases in basal [Ca2+]i do not contribute directly to the increased death rate in Ts16 neurons.
Glutamate-induced changes of the neuronal death rate
Neuronal survival decreased drastically after application of glutamate in both diploid and Ts16 cultures. Glutamate-induced neurotoxicity is expected to occur gradually and to be triggered by a rapid increase in [Ca2+]i via NMDA receptors (Choi, 1988a, 1992, 1995). Thus, selective inhibition of NMDA receptors or absence of extracellular Ca2+ is known to protect neurons against glutamate-effected elevation in cell death (Choi, 1988b; Tymianski et al., 1993). This is in line with the current findings, where a reduction in neuronal death was observed after glutamate application in the absence of extracellular Ca2+ or in the presence of the NMDA antagonist APV in diploid and Ts16 cultures. In previous studies it has been shown that glutamate application induced large increases in [Ca2+]i, leading to a depolarization of mitochondrial membrane (Duchen et al., 1993; Hartley et al., 1993; Schinder et al., 1996). Both elevation in [Ca2+]i and depolarization of mitochondrial membrane are thought to induce opening of mitochondrial permeability transition pores (Hoek et al., 1995; Bernardi, 1996). The mitochondrial permeability transition leads to mitochondrial swelling, collapse of mitochondrial membrane potential, and uncoupling of oxidative phosphorylation (Broekemeier et al., 1992; Zazueta et al., 1994; Bernardi, 1996). It is of particular interest that a blocking of the mitochondrial permeability transition using cyclosporin A prevented glutamate-induced Ca2+-mediated neurotoxicity in both diploid and Ts16 cultures. The reduced neuronal cell loss in the presence of cyclosporin A indicates a key role of mitochondria in glutamate-induced elevated exitotoxicity, as suggested previously by several investigators (Bernardi, 1992; Reed and Savage, 1995; Schinder et al., 1996; Waring and Beaver, 1996; Zamzami et al., 1996).
Glutamate causes augmented neuronal cell death in Ts16 cultures
The results presented in this study show that glutamate caused an elevated neuronal death rate in Ts16 cultures in comparison with diploid cultures. We propose that disturbances in the interplay between Ca2+ homeostasis and mitochondrial function triggers the augmented neuronal death in Ts16 cultures. Furthermore, enhanced ROS generation in Ts16 cultures may contribute to the damaging neuronal cascade.
In Ts16 neurons, [Ca2+]i increases induced by either K+ or glutamate application displayed a prolonged recovery and an elevated Ca2+integral. As a consequence, we postulated a delayed repolarization of mitochondrial membranes after the prolonged increases in [Ca2+]i in Ts16 neurons. Indeed we found that depolarizations of mitochondrial membranes recovered more slowly in Ts16 than in diploid neurons. Elevated [Ca2+]i may be responsible for mitochondrial membrane potential disturbance (Gunter et al., 1994) and mitochondrial damage (Mattson et al., 1993c), and we found that Ca2+-induced depolarizations of mitochondrial membranes were not only prolonged but also increased in Ts16 neurons in comparison to diploid neurons. In addition to alterations in Ca2+ homeostasis, other factors may contribute to this increased depolarization and prolonged recovery of the mitochondrial membrane. ROS molecules are important candidates. Several studies have shown that glutamate causes an increase in ROS formation (Dugan et al., 1995; Reynolds and Hastings, 1995). Elevated ROS concentration is expected to increase direct NADH oxidation (Bandy and Davison, 1990; Duchen et al., 1993). Therefore, the prolonged initial reduction of NAD(P)H signal and the absence of the NAD(P)H signal overshoot that followed the application of glutamate in Ts16 neurons may indicate an elevated ROS generation. This suggestion is supported by the observed NAD(P)H signal overshoot that corresponds to the stimulation of mitochondrial Ca2+-dependent dehydrogenases in Ts16 neurons during blockade of mitochondrial permeability transition by cyclosporin A. This mitochondrial permeability transition is induced by Ca2+ in conjunction with other agents, particularly ROS (Gunter and Pfeiffer, 1990; Gunter et al., 1994; Hoek et al., 1995). Furthermore, Kowaltowski and colleagues (1996) showed that the mitochondrial permeability transition in the presence of Ca2+ is dependent on mitochondrial-generated reactive oxygen species. Thus an increased concentration of ROS may be involved in this mechanism and may amplify the Ca2+-induced mitochondrial permeability transition (Takeyama et al., 1993; Weis et al., 1994; Richter et al., 1995). Moreover, cyclosporin A has been reported to prevent further mitochondrial ROS production (Kass et al., 1992; Bernardi, 1996;Costantini et al., 1996).
Several findings may indicate a generally increased ROS generation in Ts16 neurons (Fig. 10): (1) reduced neuronal death rate in the presence of tocopherol, (2) prolonged reduction in NAD(P)H autofluorescence signal after depolarization of the mitochondrial membrane, (3) a Ca2+-induced NAD(P)H signal overshoot in the presence of cyclosporin A, and (4) reduced glutamate-induced neurotoxicity in the presence of cyclosporin A and, to some extent, in the presence of tocopherol. Furthermore, overexpression of Cu/Zn-SOD (Groner et al., 1994) and βAPP via the product βAP can cause an increased production of ROS (Behl et al., 1994). Elevated ROS concentration is known to cause mitochondrial damage (Beal et al., 1993; Dykens, 1994; Bolaños et al., 1995,1997) and disturb Ca2+ homeostasis (Richter and Kass, 1991; Gunter et al., 1994; Mattson, 1994; Bondy, 1995; Mattson et al., 1995; White and Reynolds, 1996). Further studies on the Ts16 mouse model are needed to determine whether the primary defect is linked to Ca2+ homeostasis or mitochondrial ROS production.
This study was supported by the Graduiertenkolleg “Schadensmechanismen im zentralen Nervensystem (ZNS): Einsatz bildgebender Verfahren” and the Sonderforschungsbereich (SFB) 507 TP C4. We thank Dr. H. Winking for kindly supplying the trisomy 16 mice. We gratefully acknowledge technical support from Dr. H. Siegmund, Dr. H. J. Gabriel, S. Latta, and A. Düerkop. Many thanks to Dr. D. Bilkey, R. Kemp, and Dr. D. Schmitz for critical reading of this manuscript.
Correspondence should be addressed to S. Schuchmann, Department of Neurophysiology, Institute of Physiology, Charité, Humboldt University Berlin, Tucholskystrasse 2, D-10117 Berlin, Germany.