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J. Biol. Chem., Vol. 279, Issue 31, 32989-33000, July 30, 2004

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In Situ Respiration and Bioenergetic Status of Mitochondria in Primary Cerebellar Granule Neuronal Cultures Exposed Continuously to Glutamate^*

Mika B. Jekabsons and David G. Nicholls

From the Buck Institute for Age Research, Novato, California 94945

Received for publication, February 11, 2004 , and in revised form, May 17, 2004.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Mitochondria play a central role in neuronal death during pathological exposure to glutamate (excitotoxicity). To investigate the detailed bioenergetics of the in situ mitochondria, a method is described to monitor continuously the respiration of primary cerebellar granule neuron cultures while simultaneously imaging cytoplasmic Ca²⁺ and mitochondrial membrane potential. Coverslip-attached cells were perfused in an imaging chamber with upstream and downstream flow-through oxygen electrodes. The bioenergetic consequences of chronic glutamate exposure were investigated, including ATP supply and demand, proton leak, and mitochondrial respiratory capacity during chronic glutamate exposure. In 25 mM K⁺ medium supplemented with 10% dialyzed serum, cells utilized 54% of their respiratory capacity in the absence of receptor activation (37% for ATP generation, 12% to drive the mitochondrial proton leak, and the residual 5% was nonmitochondrial). glutamate initially increased mitochondrial respiration from 51 to 68% of capacity, followed by a slow decline. It was estimated that 85% of this increased respiration was because of increased ATP demand, whereas 15% was attributable to a transient mitochondrial proton leak. N-Methyl-D-aspartate receptor activation was only responsible for 62% of the increased respiration. When adjusted for cell death over 3 h of glutamate exposure, respiration of the viable cells remained near basal and protonophore stimulated respiration to the same extent as control cells. Pyruvate-supplemented media protected cells from glutamate excitotoxicity, although this was associated with mitochondrial dysfunction. We conclude that excitotoxicity under these conditions is not because of an ATP deficit or uncoupling. Furthermore, mitochondria maintain the same respiratory capacity as in control cells.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Although glutamate is the dominant excitatory neurotransmitter in the central nervous system, excessive glutamate exposure can cause neuronal cell death by a process known as glutamate excitotoxicity (1, 2). This phenomenon amplifies brain damage following stroke, traumatic brain injury, and epilepsy where massive glutamate release from neurons in the affected region causes pathological activation of N-methyl-D-aspartate (NMDA)¹-selective glutamate receptors and contributes to neuronal cell death in the surrounding tissue (3). In primary neuronal culture models, necrosis is not instantaneous but rather occurs after a latent period followed by an uncontrolled rise in cytoplasmic free Ca²⁺, ([Ca²⁺]_c), known as delayed Ca²⁺ deregulation (DCD) (4, 5).

A central role for mitochondria in cerebellar granule neuron (CGN) excitotoxicity is firmly established, based on fluorescent monitoring of changes in mitochondrial membrane potential (_m) (6-8), mitochondrial Ca²⁺ accumulation (9, 10) and morphology (11), and on the protection afforded by mitochondrial depolarization prior to glutamate under conditions where glycolysis is adequate (12-14). However, there is very limited information on in situ mitochondrial respiration (15-18), even though the classic oxygen electrode chamber has proven to be the single most important apparatus for monitoring the function of isolated mitochondria. This is primarily because of the requirement for large numbers of cells in suspension when using conventional oxygen electrodes.

The approach taken in the present study was to develop a "cell respirometer" that retains the cells on their coverslips, reduces the dead volume as far as possible, and continuously superfuses media over the cells, detecting the difference in the upstream and downstream oxygen concentrations with sensitive flow-through micro oxygen electrodes. Because the assembly is mounted on a digital imaging microscope, a representative sample of the entire cell field can be continuously imaged to monitor cell morphology and survival together with fluorescent signals from one or more indicators, such as FURA-PE3 for [Ca²⁺]_c and TMRM⁺ for mitochondrial and plasma membrane potentials. In experiments to monitor mitochondrial function during potentially excitotoxic glutamate exposure, it is important to distinguish between a change in population respiration because of stochastic cell death and one that reflects a uniform decline in single cell respiration. By imaging a representative field, cell viability can be continuously assessed and used to correct the response.

The present study tested hypotheses that chronic glutamate exposure increases ATP demand, that oxidative phosphorylation becomes rate-limiting, that mitochondrial uncoupling is an early event (7, 19), and that mitochondrial respiratory capacity is impaired (15).

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Reagents—TMRM was from Molecular Probes (Eugene, OR). Fura-PE3-AM was from Tef Labs (Austin, TX). 10 kDa of dialyzed fetal bovine serum and all other reagents were from Sigma.

Cerebellar Granule Neuron Preparation and Indicator Loading—CGNs were prepared from 5- to 7-day-old rat Wistar pups essentially as described previously (20). 3 x 10⁶ cells were allowed to settle for 30-40 min onto 22 x 40-mm coverslips previously coated with 33 µg/ml polyethyleneimine and then maintained at 37 °C in a 95/5% air/CO₂ incubator. Cells were used after 7-10 days in culture. Prior to assembling the perfusion chamber, cells in culture media were incubated at 37 °C with 3 nM TMRM⁺ plus 1 µM tetraphenylboron for 2-2.5 h and with 2 µM FURA-PE3-AM for 50-80 min. Under these conditions, TMRM⁺ fluorescence was in nonquench mode (21), meaning that a decrease in either plasma membrane potential (_p) or _m is reflected in a decrease in whole-cell fluorescence. Fura-PE3-AM was used to monitor [Ca²⁺]_c because it was retained sufficiently by the granule neurons to provide reliable 340/380 nm signal over 2-5 h, whereas FURA2 AM was essentially lost after 3 h.

Respiration Measurements Using the Cell Respirometer—The respirometer is shown schematically in Fig. 1. A closed RC-30 imaging chamber (Warner Instruments, Hamden, CT) was assembled submerged in standard buffer without serum (to avoid cell drying or bubble trapping) with a lower 22 x 40-mm coverslip with attached cells, a 250-µm gasket (yielding a chamber volume of 130 µl), and a blank upper coverslip. The chamber was mounted on an Olympus IX 70 inverted fluorescence microscope equipped with a 40x oil-immersion objective. Miniature polarographic Clark-type oxygen electrodes constructed for perfusion systems with 1/16 inch fittings (Microelectrodes, Inc., Bedford, NH) monitored upstream and eluant oxygen tension. A peristaltic pump placed distal to the downstream oxygen electrode was used to set the system flow rate, which was typically 30-60 µl/min to provide adequate oxygen depletion for detection by the downstream electrode. The standard perfusion medium used for respiration measurements consisted of (in mM) 105 NaCl, 25 KCl, 20 TES, 15 glucose, 1.3 MgCl₂, 1.3 CaCl₂, 1.2 Na₂SO₄, 0.4 KH₂PO₄, 0.2 NaHCO₃, 10% dialyzed heat-inactivated fetal bovine serum, 3 nM TMRM⁺, and 1 µM tetraphenylboron, pH 7.3, at 37 °C. Variations of the standard buffer are specified under "Results." The upstream electrode controlled for changes in oxygen tension in the inflow and was used to check the downstream electrode base line at the beginning and end of the perfusion, via a tubing shunt that allowed buffer flow to by-pass the chamber (Fig. 1).

View larger version (24K): [in this window] [in a new window]   FIG. 1. Schematic illustrating essential features of the apparatus used to measure respiration of CGNs cultured on coverslips. 3 x 106 CGNs isolated from the cerebella of 7-day-old rats were plated onto 22 x 40-mm polyethyleneimine-coated coverslips. After 7-10 days of incubation, each coverslip with attached cells was preloaded with 2 µM FURA-PE3 and 3 nM TMRM+ in standard perfusion medium and then mounted into the base of a submerged Warner RC30 closed-bath chamber. The chamber was assembled submerged in standard buffer using a 250-µm gasket, resulting in an 130-µl chamber volume. The inlet and outlet ports of the chamber were connected to upstream and downstream flow-through polarographic oxygen electrodes (Microelectrodes Inc.) resting on the microscope stage. Each oxygen meter delivered a polarizing voltage to each electrode and had outputs that were connected to a dual-pen chart recorder for monitoring the signals over time. A tubing manifold was used to connect different buffers to the system. Cells were perfused with a peristaltic pump at 30-60 µl/min in the standard buffer (or supplemented according to each experiment) for a 30-min equilibration period and then a 30-min basal period to establish base-line respiration, FURA-PE3, and TMRM+. During the 30-min equilibration period, buffer flow to the chamber was redirected to the shunt tubing for 3-5 min to check that both oxygen electrodes gave the same air-saturated buffer readings. The shunt was also opened at the end of each experiment to assess downstream electrode drift over time. Cells were imaged with a 40x oil-immersion objective, and a single field-of-view was kept throughout each experiment.

View larger version (57K): [in this window] [in a new window]   FIG. 2. Representative experiment to assess cerebellar granule neuron mitochondrial function. 3 x 106 cells cultured 7-10 days in vitro on 22 x 40-mm coverslips were assembled as described in Fig. 1. A, upstream and downstream oxygen electrode outputs over a 2.5-h experiment to determine total cell respiration (a), reserve ATP-generating capacity (b), respiration supporting ATP turnover (c), respiration driving mitochondrial H+ leak (d), and nonmitochondrial respiration (e). Toward the end of the 30-min equilibration, the shunt was opened to check and adjust the downstream electrode base line. The shunt was closed, resulting in the downstream electrode signal going off-scale momentarily because of transient depletion of oxygen in the chamber while the shunt was open; the chamber did not, however, go anoxic, as the oxygen meter provided an independent means of monitoring the absolute oxygen tension. Numbers below the downstream oxygen electrode trace represent the mean ± S.E. (n = 4 independent preparations) of the respiration in nmol of O/min x 106 cells at each steady state. The perfusion rate was 44 µl/min. Basal respiration was monitored for 30 min, followed by perfusion with 2 µM FCCP for 10 min to assess maximal respiration. Cells were given 50 min of recovery in standard buffer and then sequentially perfused for 15 min with 0.2 µg/ml oligomycin and 20 min with 2 µM myxothiazol plus 1 µM rotenone (myx/rot). Flow was stopped to prime upstream tubing with each compound, resulting in a transient oxygen depletion. The shunt was opened to assess electrode drift. B, representative TMRM+, FURA-PE3 340 nm, FURA-PE 380 nm, and transmission images. C, time course of TMRM+ (dashed line) and FURA-PE3 340/380 (solid line) changes of a representative single cell soma during the experiment in A. FU, fluorescence units.

Once assembled into the chamber, cells were equilibrated in the standard medium for 30 min. Control data were collected for 30 min, with TMRM⁺, FURA-PE3, and transmission images taken every 10 min. The perfusing medium could be changed to allow responses to various additions to be followed over the next 1-5 h, as detailed in the figures.

The choice of tubing was important. TMRM⁺ adsorbed to Tygon tubing to such an extent that this tubing could not be used upstream of the chamber, otherwise the TMRM⁺ concentration exposed to the cells was difficult to assess and potentially variable; silastic tubing was used upstream of the chamber as it did not adsorb TMRM⁺. However, because silastic tubing is highly permeable to oxygen, it was not appropriate to connect the chamber to the downstream electrode, otherwise oxygen in the air rapidly equilibrated with the oxygen-depleted buffer exiting the chamber. Tygon tubing, which is less permeable to oxygen, was used for this connection to minimize oxygen diffusion into the oxygen-depleted buffer, because probe binding was immaterial downstream of the chamber. Additionally, the tubing length between chamber and downstream electrode was kept as short as possible.

Most important, the oxygen electrodes exhibited significant responses to temperature variations. To avoid this, a custom-made acrylic incubator was constructed to surround and maintain the entire microscope stage, objective and incubation reservoirs at 37 °C, eliminating buffer cooling and temperature artifacts.

Statistics—Analyses were performed using GraphPad Prism version 4.0 for Windows (GraphPad Software, San Diego, CA). Tests used were two-way ANOVA, one-way ANOVA with Tukey's post test, and un-paired Student's t test. The level of statistical significance was taken as p < 0.05.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Assessing Neuronal Bioenergetics Using Flow-through Oxygen Electrodes—Fig. 2 illustrates some of the considerable advantages of the respirometer over traditional closed-chamber, stirred-suspension oxygen electrodes to monitor coverslip-attached cell respiration. First, the cells remain attached to their coverslips; second, the perfusion mode permits long term experiments and medium exchange; and third, simultaneous monitoring of cell viability and function can be performed.

During steady-state perfusion, the respiratory rate is the product of the flow rate multiplied by the difference between the upstream and downstream oxygen tensions. Fig. 2 illustrates basic bioenergetic information obtained from a respirometer trace (Fig. 2A), and the associated images obtained by using the potentiometric probe TMRM⁺ and the Ca²⁺ probe FURA-PE3 (Fig. 2B). The following respiratory rates were determined: basal (V_basal), maximal uncontrolled respiration in the presence of 2 µM FCCP (V_FCCP), oligomycin (0.2 µg/ml)-insensitive (V_oligo), and residual nonmitochondrial (V_rot/myx) respiration in the presence of 2 µM myxothiazol plus 1 µM rotenone. The concentration of FCCP was just sufficient to produce uncontrolled respiration, although this rate often was not sustained, probably because of ATP depletion.

Assuming the myxothiazol plus rotenone-insensitive rate is nonmitochondrial, mitochondria accounted for 89 ± 2% ((V_basal - V_rot/myx)/V_basal) of basal respiration. Initial estimates (not taking account of the voltage dependence of the mitochondrial proton leak (22)) are that ATP synthesis is responsible for 69 ± 4% ((V_basal - V_oligo)/(V_basal - V_rot/myx)) of basal mitochondrial respiration, whereas the remaining 31 ± 4% oligomycin-insensitive rate ((V_oligo - V_rot/myx)/(V_basal - V_rot/myx)) drives H⁺ leaks across the mitochondrial inner membrane. The mitochondria exhibit a pseudo-respiratory control ratio of 6.3 ± 0.9 ((V_FCCP - V_rot/myx)/(V_oligo - V_rot/myx)) and in the basal state operate at 51 ± 3% ((V_basal - V_rot/myx)/(V_FCCP - V_rot/myx)) of maximal respiration.

Fig. 2C shows the effects of these additions on FURA-PE3 and TMRM⁺ fluorescence of a representative single CGN soma. Brief FCCP perfusion decreased TMRM⁺ fluorescence by 50% because of mitochondrial release of TMRM⁺ and consequent re-equilibration across the plasma membrane (6), and modestly increased the FURA-PE3 340/380 nm ratio. Cytoplasmic Ca²⁺ levels rapidly recovered during FCCP washout, whereas TMRM⁺ fluorescence took 50 min to return close to pre-FCCP values. The kinetics of TMRM⁺ recovery are primarily because of slow re-equilibration of probe across the plasma membrane, which has a low surface-to-volume ratio compared with the mitochondrial inner membrane (6, 21).

Under the present conditions (elevated KCl in the presence of serum) ATP synthase inhibition by oligomycin perfusion resulted in rapid Ca²⁺ deregulation. Thus glycolysis alone is not sufficiently active to maintain ATP levels necessary for cytoplasmic Ca²⁺ regulation. The oligomycin-insensitive respiration was stable for only 5-10 min, after which it slowly declined (Fig. 3A, solid line). The decline is a consequence of cell death, as correcting for dead cells produced a constant state 4 rate (Fig. 3A, dashed line; see next section for an explanation of this correction). Cell viability assessed morphologically was about 62% at the time of FCCP addition. However, even after this correction respiration only reached 84% of basal, contrasting with 184% of basal when protonophore was added in the absence of oligomycin (Fig. 2A and Fig. 4A). Because two steps in glycolysis require ATP and because ATP appears to be depleted in the presence of oligomycin, it is possible that the decreased uncontrolled respiration seen in Fig. 3A reflects compromised glycolytic pyruvate supply to the mitochondrion rather than a defect in mitochondrial substrate oxidation. To test this, the experiment was repeated in the presence of 10 mM exogenous pyruvate (Fig. 3B). Viable cell respiration following oligomycin addition was stable for a longer period and following FCCP was increased to 116% of basal. However, ATP depletion was still apparent judged by the failure to maintain cytoplasmic Ca²⁺ homeostasis and by the slow decline in TMRM⁺ signal (Fig. 3D). Under these nonquenching conditions the TMRM⁺ signal is equally affected by changes in _p and _m values (6). Because there is no increase in state 4 respiration following oligomycin (Fig. 3B), this indicates that the decline in TMRM⁺ fluorescence largely reflects plasma membrane depolarization because of ATP deficiency rather than mitochondrial depolarization. This is confirmed by the retained respiratory control revealed by the FCCP addition, although maximal respiratory capacity is reduced. Thus oligomycin addition in the absence of exogenous pyruvate causes an ATP depletion that results in impaired glycolysis and subsequent mitochondrial respiratory inhibition because of a failure of substrate supply. This can be prevented by exogenous pyruvate even though the substrate cannot contribute to cellular ATP generation in the presence of oligomycin.

View larger version (22K): [in this window] [in a new window]   FIG. 3. Effect of oligomycin and FCCP on basal respiration. Cells were equilibrated for 30 min in standard buffer (A and C) or buffer supplemented with 10 mM pyruvate (B and D). The basal state was recorded over a 15-min period. This was followed by perfusion with 0.2 µg/ml oligomycin (oligo) followed by 2 µM FCCP to assess uncontrolled respiration. A and B, respiration was normalized to the basal rate (A, 0.96; B, 1.11 nmol of O/min x 106 cells) and expressed per total (solid lines) or viable (dashed lines) cell. C and D, time course of TMRM+ fluorescence (dashed lines; FU, fluorescence units) and FURA-PE3 340/380 nm ratio (solid lines) changes in a representative cell soma from each experiment. Fractions represent the proportion of cells examined that failed to maintain [Ca2+]c homeostasis after oligomycin. Data are from a single CGN preparation cultured 7 days in vitro.

View larger version (20K): [in this window] [in a new window]   FIG. 4. Effects of 3 h of continuous glutamate/glycine perfusion on CGN respiration, TMRM+ fluorescence, and FURA-PE3 340/380 nm ratio. 3 x 106 cells were equilibrated for 30 min in standard buffer. Respiration for each preparation was normalized to the 30-60-min basal period. A, initial respiratory capacity assessed by perfusion with 2 µM FCCP (n = 6). Error bar represents ±1 S.D. of peak uncontrolled respiration. B, 180-min perfusion with standard buffer containing 250 µM glutamate plus 25 µM glycine (glu), followed by standard buffer with 2 µM FCCP to assess final respiratory capacity (n = 4). Basal respiration was 1.23 ± 0.29 nmol of O/min x 106 cells (mean ± S.E.). Solid line, normalized respiration per 106 total cells; dashed line, normalized respiration per 106 viable cells. Bars show S.Es. C-E, representative cell soma TMRM+ fluorescence (dashed lines; FU, fluorescence units) and FURA-PE3 340/380 nm ratio (solid lines) of CGNs divided into three groups as follows: those that survived 3 h of continuous glutamate (C), those that Ca2+-deregulated immediately (D), and those that recovered from initial Ca2+ load imposed by glutamate but deregulated after a variable lag period (E). Twenty cells from each experiment were randomly selected for quantitative fluorescence analysis. Fractions are the number of cells in each group.

Consequently, respiration was corrected for viable cell number with each image taken (Fig. 4B, dashed line). In the corrected trace, viable CGNs maintain respiration at or slightly above basal for the entire 3-h glutamate exposure. Because experimental variation in viability was large, it is uncertain if the apparent rise in viable cell respiration during the final 60 min of glutamate reflects a true increase in viable cell metabolism above basal. Additionally, the maximal respiratory capacity of the viable cells with FCCP was within the range of that determined for cells not exposed to glutamate (Fig. 4A). Thus surviving cells do not undergo a loss of respiratory capacity, and contradictory reports in the literature (15) are most probably because of cell death.

Viability was significantly lower after 3 h of continuous glutamate exposure compared with 4 h of control perfusions (Table I). After correcting for viability, each replicate showed a reserve ATP synthesis capacity after 3 h of continuous glutamate, although this was variable (100 ± 60%) and was not significantly different from that determined in control perfusions after 1 (84 ± 10%; Fig 4A) or 4 h (76 ± 15%; Table I).

View larger version (22K): [in this window] [in a new window]   FIG. 5. Oligomycin-insensitive respiration of CGNs and effects of acute and chronic glutamate. A, following 70 min of standard buffer, buffer with 0.2 µg/ml oligomycin was perfused followed at 85 min by 2 µM myxothiazol plus 1 µM rotenone (myx/rot). B, at 60 min cells were perfused with 250 µM glutamate plus 25 µM glycine (glu) for 10 min, followed by oligomycin plus glutamate/glycine at 70 min and myxothiazol plus rotenone at 85 min. C, at 60 min cells were perfused with glutamate/glycine, followed at 200 min by oligomycin plus glutamate/glycine and at 215 min by rotenone plus myxothiazol. Respiration data are normalized to the average basal rate for each preparation (n = 4 for each condition; values in nmol of O/min x 106 cells: A, 0.94 ± 0.15; B, 0.90 ± 0.12; C, 1.39 ± 0.21). Cells were used at 7-10 days in vitro. Rates next to each steady-state are expressed as percent of basal viable cell respiration; means sharing common superscripts are not significantly different, as assessed by 1-way ANOVA. D-F, representative cell soma TMRM+ fluorescence (dashed line; FU, fluorescence units) and FURA-PE3 340/380 nm ratio over the time course of each experiment. Arrows above traces correspond to additions specified in A-C.

To determine whether the increased H⁺ leak persists during longer glutamate exposure, CGNs were perfused for 140 min with glutamate followed by 15 min with oligomycin plus glutamate (Fig. 5C). After correction for nonviable cells, oligomycin-insensitive respiration returned to control levels and was significantly lower than after 25 min of total glutamate exposure (Fig. 5, C versus B). However, variation after additionally correcting for nonmitochondrial respiration meant that the H⁺ leak after 155 min of glutamate exposure was not significantly different from either the control or 25-min glutamate leak rates (one-way ANOVA, control, 33.7 ± 1.8%; 25-min glutamate exposure, 44.5 ± 0.6%; 155-min glutamate exposure, 35.7 ± 3.7%).

The NMDA Receptor Is the Primary Contributor to glutamate-stimulated Respiration—Although glutamate activates multiple receptors, it is the NMDA receptor that is responsible for excitotoxicity under the present conditions (23). To assess the NMDA receptor contribution to the metabolic demand imposed by glutamate, cells were sequentially exposed to glutamate and NMDA (Fig. 6A) or NMDA prior to glutamate (Fig. 6B). The NMDA and glutamate responses were independent of the order of addition (Fig. 6, A versus B). Respiration stimulated by NMDA was 62% of that stimulated by glutamate. Consistent with this finding, glutamate-stimulated respiration was reduced by 67% in the presence of the selective NMDA receptor antagonist MK801 (Fig. 6B). 5 µM MK801 completely blocked the NMDA response, confirming that this concentration was saturating under the conditions used (Fig. 6A). The responses elicited via the NMDA and non-NMDA components were virtually additive compared with the total glutamate response (Fig. 6B).

View larger version (17K): [in this window] [in a new window]   FIG. 6. Contribution of the NMDA receptor to the glutamate-induced respiratory response. CGNs were equilibrated for 45-50 min in standard buffer before recording. Cells were perfused with 250 µM glutamate plus 25 µM glycine (glu), 300 µM NMDA plus 25 µM glycine (NMDA), 5 µM MK801, NMDA plus MK801, or glutamate/glycine plus MK801 as shown. Numbers on each trace represent peak percent increase in respiration above the basal 100% period. Data are from a single granule neuron preparation cultured either 7 (A) or 8 (B) days in vitro. Basal respiration averaged 1.08 (A) and 1.19 (B) nmol of O/min x 106 cells. Cell viability, assessed as described in Fig. 4 and the text, was used to correct the raw traces for dead cells.

View larger version (20K): [in this window] [in a new window]   FIG. 7. Effects of pyruvate supplementation on CGN respiration during 3 h of continuous glutamate/glycine perfusion. Cells were equilibrated for 30 min in standard buffer supplemented with 10 mM pyruvate. A, at 60 min, initial uncontrolled respiration was assessed by perfusion with 2 µM FCCP plus pyruvate (n = 3). Error bar represents ±1 S.D. of peak uncontrolled rate. B, from 60 to 240 min CGNs were perfused with pyruvate-supplemented buffer containing 250 µM glutamate plus 25 µM glycine (glu). At 240 min, final uncontrolled respiration was assessed with standard buffer containing pyruvate and 2 µM FCCP (n = 4, cells 8-10 days in vitro). The mean basal respiration was 1.21 ± 0.29 nmol of O/min x 106 cells. Solid line, normalized respiration per 106 cells; dashed line, normalized respiration per 106 viable cells. Error bars represent S.E. C-E, representative cell soma TMRM+ fluorescence (dashed lines; FU, fluorescence units) and FURA-PE3 340/380 nm ratio (solid lines) of CGNs divided into groups that survived 3 h of continuous glutamate (C), those that deregulated immediately (D), and those that recovered from the initial Ca2+ load imposed by glutamate but deregulated after a variable lag period (E). Twenty cells from each experiment were randomly selected for quantitative fluorescence analysis. Fractions above fluorescence time courses are the number of cells in each group.

Although the initial respiratory response to glutamate was not enhanced, cells in 10 mM pyruvate-supplemented buffer exhibited greater initial [Ca²⁺]_c responses to glutamate, with mean peak FURA-PE3 340/380 nm increasing by 0.64 ± 0.03 (Fig. 7, C-E; n = 80 cells). Similarly, TMRM⁺ dropped to a greater extent after glutamate (71 ± 1%; n = 80 cells) than when using the standard buffer without pyruvate, indicating a greater plasma and/or mitochondrial depolarization.

Whereas pyruvate relieves substrate limitation, and hence increases uncontrolled respiration in the presence of oligomycin (Fig. 3B) or in the basal state (cf. Fig. 7A and Fig. 4A), the decrease in uncontrolled respiration following 3 h of glutamate exposure in the presence of pyruvate (Fig. 7B) is an apparent contradiction. More cells survive glutamate exposure in the presence of pyruvate; however, sustained [Ca²⁺]_c levels are higher (cf. Fig. 7C and Fig. 4C), suggesting that the respiratory capacity of viable cells may be compromised by Ca²⁺.

Submaximal FCCP Facilitates Cell Death during Initial Glutamate Exposure—In the presence of 10% serum, 1 µM FCCP is largely bound to albumin and is not sufficient to fully relieve CGN respiratory control (data not shown). Perfusion of cells with 1 µM FCCP starting 15 min prior to glutamate exposure increased respiration 41% above basal (Fig 8B). This was associated with a significant 22 ± 2% TMRM⁺ reduction but no significant FURA-PE3 340/380 change (basal, 1.59 ± 0.02, n = 40; 1 µM FCCP, 1.62 ± 0.02, n = 40 cells; Student's t test). Subsequent glutamate perfusion in the continued presence of 1 µM FCCP resulted in a significantly lower respiratory increase (8%, n = 2) compared with experiments in the absence of protonophore (32 ± 4%; n = 4). Additionally, in the presence of FCCP, the average peak FURA-PE3 340/380 increase following glutamate was significantly greater than without FCCP (0.50 ± 0.05, n = 40 cells versus 0.32 ± 0.02, n = 80 cells; p < 0.0001 by Student's t test). Viable cell respiration was sustained at or very close to the initial 1 µM FCCP-stimulated rate throughout the experiment. During the 1st h of glutamate/FCCP exposure, the presence of 1 µM FCCP was the only condition that resulted in a significantly lower viability when compared with controls (Table I).

View larger version (21K): [in this window] [in a new window]   FIG. 8. Effect of submaximal FCCP treatment on the response to 3 h of continuous glutamate/glycine. Cells were equilibrated for 30 min in standard buffer. A, initial uncontrolled respiration in the presence of 2 µM FCCP for control cells. Data from Fig. 4A. B, starting at 45 min, cells were perfused in standard buffer containing 1 µM FCCP. From 60 to 240 min, the perfusion medium additionally contained 250 µM glutamate plus 25 µM glycine (glu). At 240 min perfusion was switched to standard buffer with 2 µM FCCP to assess final uncontrolled respiration. Data are from two independent cell preparations (aged 9 and 10 days in vitro). Basal respiration averaged 0.96 ± 0.26 nmol of O/min x 106 cells (mean ± S.D.). Solid line, normalized respiration per 106 total cells; dashed line, normalized respiration per 106 viable cells. C-E, representative cell soma TMRM+ fluorescence (dashed lines; FU, fluorescence units) and FURA-PE3 340/380 nm ratio (solid lines)of CGNs divided into groups that survived 3 h of continuous glutamate (C), that Ca2+ deregulated immediately (D), and that recovered from initial Ca2+ load imposed by glutamate but deregulated after a variable lag period (E). Twenty cells from each experiment were randomly selected for quantitative fluorescence analysis. Fractions above the fluorescence time courses are the number of cells in each group.

View larger version (67K): [in this window] [in a new window]   FIG. 9. Effect of excess FCCP on the response to continuous glutamate/glycine. Cells were perfused for 45 min in standard buffer (30-min equilibration and 15-min control period) followed by 45 min with 5 µM FCCP and 90 min with 5 µM FCCP, 250 µM glutamate, and 25 µM glycine as indicated. A, respiration was normalized to the basal rate (0.35 nmol of O/min x 106 cells). Peak respiration with FCCP plus glutamate was 0.62 nmol of O/min x 106 cells. Following FCCP/glutamate perfusion, 2 µM myxothiazol, 1 µM rotenone, and 0.2 µg/ml oligomycin (myx/rot) was added to assess remaining mitochondrial respiration. This respiration, indicated by the arrow, was 19% of peak rate following initial glutamate addition. B, representative cell soma TMRM+ fluorescence (dashed lines; FU, fluorescence units) and FURA-PE3 340/380 nm ratio (solid lines) during the experiment. All 20 cells examined exhibited immediate loss of Ca2+ homeostasis following glutamate addition. C, transmission image taken during the control period prior to FCCP addition. D, transmission image taken after 90 min of FCCP/glutamate perfusion; about 98% of cells exhibited morphological changes at this time.

View larger version (35K): [in this window] [in a new window]   FIG. 10. Time-dependent changes in viable cell FURA-PE3 (A) and TMRM+ (B). Data were obtained from respirometer experiments during the initial basal period (30-60 min, dark bars) and just before FCCP addition (220-240 min, light bars). Pyruvate (10 mM) and MK801 (0.5 µM) where indicated were present throughout the experiment. Except in the control experiment, 250 µM glutamate plus 25 µM glycine (glu) was added at 60 min (i.e. after the initial period). The number of cells per group is in parentheses below designated treatments. Error bars show S.E. For viable cell FURA-PE3, 2-way ANOVA yielded significant main effects of time (p = 0.01) and treatment (p < 0.0001), and a significant time x treatment interaction (p < 0.0001). Similarly, 2-way ANOVA of TMRM+ yielded significant effects of time (p < 0.0001), treatment (p < 0.0001), and time x treatment (p < 0.0001). One-way ANOVA with Tukey's post-hoc test was used to determine which means within a treatment differed by time (* indicates 220-240-min mean significantly different from corresponding 30-60-min mean).

View larger version (19K): [in this window] [in a new window]   FIG. 11. The change in respiratory capacity of viable cells during the experiment correlates with the final FURA-PE3 ratio. Data were collected from respirometer experiments in the presence of glutamate (), glutamate plus 0.5 µM MK801 (), glutamate plus 10 mM pyruvate (), and control perfusions () (n = 4 independent preparations for each condition). Viable cells at the end of each individual experiment (prior to FCCP) were grouped by averaging their FURAPE3 signals (± S.E.). These values were plotted as a function of the change in respiratory capacity (in the presence of 2 µM FCCP) during the experiment, i.e. from 60 to 240 min ( FCCP rate). Negative values indicate that the FCCP response was lower than expected, and values 0 or greater indicate the response was as expected or better. The line was fitted to the data by least squares linear regression (r2 = 0.31, p = 0.02).

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Bioenergetic parameters measure the extents of displacement from equilibrium (e.g. Gibbs free energy, protonmotive force, membrane potential, redox state, and ATP/ADP ratios), the rates of energy flux (e.g. metabolic rates, ATP turnover, proton flux, and respiration), or combinations of these parameters (e.g. proton conductance that is determined as proton leak per mV of proton electrochemical gradient). Although there is considerable literature detailing the rates of glucose or pyruvate utilization and lactate or CO₂ production in CGN cultures (reviewed in Refs. 25-27), technical problems have limited the investigation of the primary mitochondrial flux parameter, namely the rate of electron transport or proton cycling as monitored by respiration. The classic oxygen electrode chamber has proven to be the single most important apparatus for monitoring the function of isolated mitochondria, because careful experimental design allows determination of ATP turnover, proton leaks, maximal electron transport activity, and other extensive parameters (for review see Ref. 28). However, a conventional cylindrical chamber is entirely unsuitable for monitoring the respiration of a neuronal monolayer attached to a substrate. This study shows that the respiration of a neuronal monolayer can reliably be measured over extended periods using flow-through polarographic oxygen electrodes.

Basal Respiration—In the present study, CGNs respire at 37 °C in 25 mM K⁺ medium with 10% dialyzed fetal bovine serum at a basal rate of 0.8-1.2 nmol of O/min x 10⁶ cells. This rate slightly underestimates true oxygen utilized under these conditions because not all the 3 x 10⁶ cells initially used attach to the coverslip, and some plated cells do not end up in the perfusion area once the chamber is assembled.

Most values in the existing literature are expressed per mg of cell protein. Hertz and Peng (26) report a value of 44 nmol of O/min x mg of cell protein for CGNs in 55 mM K⁺. Based on a value of 76 µg of protein/10⁶ cells (29), this translates to 3.3 nmol of O/min x 10⁶ cells. Slightly higher values (62 nmol of O/min x mg of cell protein, or 4.7 nmol of O/min x 10⁶ cells) were reported by Atlante et al. (15) for CGNs in 18 mM K⁺, whereas Almeida et al. (30) measured a basal rate of 6.5 nmol of O/min x mg of cell protein for cortical neuronal cells at 30 °C. In studies expressed per 10⁶ cells, Bal-Price et al. (17, 31) reported value of 3.9-4 nmol of O/min x 10⁶ CGNs in 25 mM K⁺. However, with the exception of the present study, it must be remembered that all cited investigations required scraping or trypsinization of the cells to remove them from their substrates, almost certainly resulting in some cell damage. Significantly, we are aware of no studies that report the extent of protonophore-relieved respiratory control seen in such preparations.

In the basal state, in situ CGN mitochondria operate at about 50% of maximal respiration (Fig. 2A). Oligomycin reduced respiration by 61%, suggesting that ATP turnover accounts for this percentage of basal cell respiration. However, this is an underestimate because oligomycin hyperpolarizes mitochondria, enhancing the activity of the highly potential dependent constitutive inner membrane proton leak (22). Rolfe et al. (32) showed that oligomycin reduced hepatocyte respiration by 64%; however, when corrected for membrane potential changes in H⁺ leak, ATP turnover was calculated to account for 69% of basal cell respiration. If CGN H⁺ leak is similarly dependent on _m, then the reported oligomycin-insensitive respiration is about 20% higher than it would be in the absence of inhibitor (Fig. 12). By using this correction, a better estimate is that 68% of CGN respiration supports ATP turnover, 21% supports H⁺ leak, and the remaining 11% (in the presence of rotenone plus myxothiazol) supports nonmitochondrial reactions.

View larger version (36K): [in this window] [in a new window]   FIG. 12. Schematic illustration of interrelationships between CGN respiratory parameters and m. Points were experimentally determined rates as follows: a, nonmitochondrial (Fig. 5); c, oligomycin (Fig. 5); d, oligomycin plus glutamate (Fig. 5); e, basal (Fig. 5); f, 25 min of glutamate exposure (Fig. 5); and g, uncontrolled respiration (Fig. 4A). Point b is the estimated respiration supporting the proton leak in the basal state, assuming that the oligomycin rate exceeds that in the basal state by 20% because of the increased m (32). Mitochondrial membrane potential with oligomycin was assumed to be similar to that determined for mitochondria within synaptosomes (43). A 20-mV potential drop was assumed sufficient for maximal respiration (43). Because substrate oxidation rate is a linear function of membrane potential (22, 44), potentials associated with points d-f were set according to this slope (i). The nonlinear control proton leak curve (ii) was drawn based on points a-c, whereas the curve after glutamate treatment (iii) was drawn based on points a and d, assuming that respiration driving the leak is similar to the control curve at potentials giving maximal respiration.

Acute Effects of Glutamate—A key finding from this study is that under the present conditions glutamate exposure acutely stimulates CGN respiration submaximally. From first principles it would be predicted that NMDA receptor activation would lead to an increase in CGN mitochondrial respiration on three counts as follows: utilization of the proton gradient to drive Ca²⁺ uptake into the matrix, increased plasma membrane Ca²⁺-ATPase activity to expel Ca²⁺ entering the cell, and increased Na⁺/K⁺-ATPase activity in response to increased Na⁺ entry. In fact, a 32% increase in basal respiration was observed immediately following glutamate infusion (Fig. 4B). This increase utilized about 40-50% of the reserve ATP-generating capacity of CGN mitochondria. Pyruvate supplementation did not further increase the response to glutamate (Fig. 7B). Thus, glycolysis was capable of supplying necessary substrate to sustain the higher respiratory chain activity, and oxidative phosphorylation was capable of supplying sufficient ATP to meet the energy demand imposed by glutamate.

Atlante et al. (15) exposed CGNs to glutamate/glycine in Mg²⁺-free low K⁺ medium for 30 min, washed, and suspended the neurons by scraping and determined respiration in the presence of 10 mM glucose. They reported a 20% decrease in respiration relative to control cells; however, because cell viability was not monitored, it is unclear whether this represented a change in respiration per viable cell. Surprisingly, they observed a total inhibition of respiration by ouabain both in control and glutamate-exposed neurons, suggesting that their assay was insufficiently sensitive to detect the respiration under conditions of low ATP demand. In their study, classic respiratory control was performed on homogenates from parallel cell preparations, and a 50% decrease in state 3 respiration was observed with succinate as substrate with little change in state 4 rate (15). This was incorrectly interpreted as indicating rapid "uncoupling" of oxidative phosphorylation, i.e. an increased H⁺ leak, which would have manifested as an increased state 4 respiration. The decreased state 3 rate could reflect either an inhibited respiratory chain or a decreased yield of mitochondria because of cell death.

In a related study, Almeida et al. (34) exposed cortical neurons cultured in 78-cm² dishes to glutamate for 15 min, restored the culture medium, and monitored respiration after 24 h by trypsinizing the cells, washing, storing on ice, and assaying in a conventional Clark-type oxygen electrode chamber. They reported a modest decrease in respiration, but the ability of the preparation to respond to succinate indicated extensive damage resulting from the manipulations.

The glutamate-induced initial increase in respiration can be analyzed both to determine the proportion of the response because of NMDA- and non-NMDA-mediated effects and also to distinguish between increased ATP demand and increased utilization of the H⁺ current via H⁺ leaks (classic uncoupling) or to drive matrix Ca²⁺ accumulation. Comparison of the effects of glutamate and the selective agonist NMDA (Fig. 6) shows that 62% of the increase in respiration is because of NMDA receptor activation. Clearly ion influx through this receptor stimulates ATP turnover but not to such a degree that demand exceeds supply. Thus, the protective effect of MK801 (Table I; Fig. 10) is likely a consequence of limiting one or more Ca²⁺-dependent processes rather than reducing ATP demand. The non-NMDA-driven respiration is additive to the NMDA effect and could be because of the metabolic effects of the metabotropic glutamate receptor, AMPA/kainate receptor activation, or glutamate uptake into neurons or contaminating astrocytes causing an increased ATP demand by the Na⁺/K⁺-ATPase. However, no respiratory inhibition was observed on addition of the glutamate transport inhibitor DL-threo--benzyloxyaspartic acid (100 µM, data not shown).

The analysis to distinguish ATP turnover from uncoupled respiration was performed. The second analysis was performed by acute addition of oligomycin to inhibit ATP synthase (Fig. 5). Again, it should be noted that under these conditions these CGN preparations do not possess sufficient glycolytic activity to maintain Ca²⁺ homeostasis in the absence of mitochondrial ATP synthesis, and rapid cell death ensues. The slight 10% increase in oligomycin-insensitive respiration observed 25 min following glutamate addition (Fig. 5) does not persist, confirming the absence of a significant population of rapidly respiring "uncoupled" mitochondria in the glutamate-exposed cells. The transient increase could (from first principles) be due to the utilization of the H⁺ current to drive matrix Ca²⁺ accumulation during the initial period of glutamate exposure. However, in this case a similar increase would be predicted following oligomycin alone, which causes cytoplasmic Ca²⁺ to rise to a similar extent as that in glutamate-treated cells (Fig. 5).

The CGN preparations used in Fig. 4B and Fig. 5, B and C, responded similarly to glutamate, with a peak respiration occurring 6 min after glutamate and reaching the same extent (30 ± 5% in Fig. 5B, 39 ± 7% in Fig. 5C, and 32 ± 4% in Fig. 4B). Oligomycin blocked oxidative phosphorylation after a short delay (Figs. 3 and 5). During this time, glutamate-stimulated respiration declined to 19-21% above basal or about 0.18 nmol of O/min x 10⁶ viable cells (Fig. 5C and Fig. 4B), suggesting that 50% of the glutamate-induced increase is attributable to a mitochondrial proton leak. However, because oligomycin hyperpolarizes mitochondria and glutamate slightly depolarizes mitochondria (6), it is necessary to correct the leak for changes in _m. A more realistic estimate is that H⁺ leak accounts for about 15% of the increase with glutamate, whereas the remaining 85% would be attributable to ATP turnover (Fig. 12). Because this uncoupling rapidly reverses upon glutamate washout, it does not seem to reflect a small minority of cells entering a fully uncoupled state but rather may be a consequence of activating a regulated H⁺ conductance (e.g. an uncoupling protein).

Chronic Glutamate Exposure—An important finding of this study is that mitochondria in surviving CGNs do not show diminished respiratory capacity after 3 h of continuous glutamate exposure. Previous attempts to monitor the effects of glutamate exposure on neuronal respiration (e.g. Ref. 35) have not allowed for stochastic cell death, and so it has been unclear whether reported chronic decreases in respiration were because of cell death or a uniform respiratory inhibition. If the assumption is made that Ca²⁺deregulated cells having altered morphology and no TMRM⁺ fluorescence do not respire, then the 45% remaining viable cells after 3 h glutamate show no decrease in respiration, or even a statistically insignificant increase (Fig. 4B). These cells retain protonophore-releasable respiratory control. Moreover, viable cell respiration with 1 µM FCCP is maintained at the pre-glutamate level throughout 3 h glutamate exposure (Fig. 8B), again suggesting no loss of mitochondrial function.

The analysis of cell respiration makes the assumption that Ca²⁺-deregulated, morphologically altered cells lacking TMRM⁺ fluorescence do not consume oxygen. This was tested by using oligomycin to cause a synchronous Ca²⁺ deregulation in the whole-cell population that was monitored in the respirometer (Fig. 3). No prominent increase in oligomycin-insensitive respiration was seen despite an initial FURA 340/380 nm ratio increase from about 1.4 to 3. Furthermore, extended incubation with oligomycin resulted in further respiratory inhibition, reduction in TMRM⁺ signal, and altered cell morphology. From this perspective, the assumption that glutamate-treated cells that undergo these changes do not respire seems valid. However, when 98% of cells were considered dead (i.e. nonrespiring) following treatment with excess FCCP plus glutamate, respiration declined by 81% rather than the expected 98% (Fig. 9). This indicates that (a) some cells considered "dead" do consume oxygen, and (b) the viable cell respiration traces overestimate the true rate at which these cells consume oxygen. Although this uncertainty does not affect our conclusions that cells exposed to glutamate retain normal mitochondrial function until cell death, it is clear that more experiments are required to determine the variable(s) that more accurately differentiate an oxygen-consuming cell from one that is truly dead.

Because rat brain mitochondria possess a high Ca²⁺ uniport activity sufficient to utilize the full capacity of the respiratory chain when extramitochondrial [Ca²⁺] is elevated (36), the lack of respiratory increase associated with the increased [Ca²⁺]_c following oligomycin was initially surprising (Fig. 3), suggesting that elevated [Ca²⁺]_c was not sufficient to activate the uniporter significantly, that the uniporter was inhibited, or that mitochondrial respiration was severely limited because of lack of substrate. Glycolytic failure did occur with oligomycin because uncontrolled respiration with pyruvate supplementation was much greater than with glucose alone (Fig. 3). Despite this, the failure to observe a prominent increase in respiration after oligomycin-induced Ca²⁺ deregulation cannot be attributed to substrate supply problems because bypassing glycolysis with exogenous pyruvate did not stimulate state 4 respiration.

The resting [Ca²⁺]_c of granule neurons in 25 mM KCl is about 100 nM (37), well below the threshold of about 500 nM for initiating mitochondrial Ca²⁺ accumulation (36). Even with 70 mM KCl to fully activate L-type Ca²⁺ channels, steady-state [Ca²⁺]_c only reaches about 200 nM (37). Therefore, the basal rate of Ca²⁺ entry in 25 mM KCl is quite low relative to influx necessary to achieve mitochondrial loading. In Fig. 3, C and D, it is seen that oligomycin caused FURA 340/380 nm to plateau at about 3.0; this is more readily seen in Fig. 3D with pyruvate present. The FURA plateau at 3 likely represents the critical concentration of about 500 nM at which the mitochondria began accumulating Ca²⁺ at a slow rate that matched the continued rate of Ca²⁺ entry across the plasma membrane (36). Subsequent FCCP addition further increased this ratio to >4 in Fig. 3D. Because FCCP depolarizes mitochondria, thus inhibiting Ca²⁺ accumulation, the further FURA increase is consistent with the plateau at 3 representing the point at which mitochondria began accumulating Ca²⁺. The plateau was not sustained in the cell depicted in Fig. 3C, suggesting failure of mitochondrial Ca²⁺ accumulation. This is consistent with the rapid loss of TMRM⁺ in this cell. Careful inspection of the respiration traces in Fig. 3, A and B, provides an indication as to the amount of oxygen utilized for Ca²⁺ uptake. Oligomycin addition initially reduced respiration to 33 and 34% of basal rate, respectively. This was followed by a very small increase (about 4% in both cases) to a final constant state 4 rate. In both cases the increase amounted to 0.04 nmol of O/min x 10⁶ cells. We suggest that this small increase represents the utilization of the protonmotive force to drive Ca²⁺ accumulation under these conditions. This is an exceedingly small rate of Ca²⁺ uptake, consistent with the idea that Ca²⁺ entry through L-type Ca²⁺ channels is relatively low under the current conditions. Thus, [Ca²⁺]_c did not increase to such an extent to prominently activate the Ca²⁺ uniporter.

Taken together, the data indicate that glutamate-induced DCD is associated with a loss of respiration rather than pronounced uncoupling. Does Ca²⁺ deregulation precede or follow loss of respiration? This remains an unresolved issue because these alternatives cannot adequately be resolved in these experiments, particularly given the synchronous changes in TMRM⁺ and FURA-PE3 signals as cells begin to lose Ca²⁺ homeostasis. Although there is no indication of a slow decline in mitochondrial function of viable cells after glutamate, it may be that mitochondrial respiration rapidly shuts down, resulting in Ca²⁺ deregulation. This possibility cannot be distinguished from the alternative that Ca²⁺ deregulation results in loss of respiration.

Chronic Glutamate Exposure with Pyruvate—Pyruvate supplementation significantly attenuates glutamate excitotoxicity (Table I). This protection cannot be attributed to enhanced mitochondrial substrate supply because glutamate did not maximally stimulate respiration and pyruvate did not further enhance glutamate-stimulated rate, in contrast to its effects on uncontrolled respiration. Surprisingly, cells that remained viable after 3 h of continuous glutamate exposure exhibited a pronounced reduction in maximal respiratory capacity (Fig. 7B), indicating an impairment downstream of pyruvate supply to the mitochondria. Four observations indicate that elevated cytoplasmic Ca²⁺ levels may be involved. First, there was a significant negative correlation between [Ca²⁺]_c and mitochondrial responsiveness to FCCP (Fig. 11). Second, Ca²⁺ transients immediately following glutamate addition were significantly greater with pyruvate present (Fig. 7). Third, cytoplasmic Ca²⁺ levels after 3 h of glutamate exposure were significantly higher than before glutamate addition (Fig. 10). Fourth, oligomycin-induced Ca²⁺ deregulation resulted in impaired uncontrolled respiration (cf. Fig. 3B and Fig. 7A). Clearly, pyruvate protection from glutamate excitotoxicity cannot be ascribed to improved mitochondrial function, as indicated by respiratory capacity. This is not surprising given that chronic glutamate exposure imposed very little energy demand above the basal state (Fig. 4). Reduction in the reserve ATP-generating capacity is therefore not an event associated with DCD. Previous studies (18, 38-42) have shown that pyruvate-supplemented media protect neurons from glutamate- or NMDA-mediated excitotoxicity. However, it is unclear how pyruvate paradoxically increases Ca²⁺ influx and protects against excitotoxicity.

Conclusions—The oxygen electrode has proven over the years to be the most versatile apparatus for investigating the function of isolated mitochondria and cells that exist in suspension. The lack of a similar device for cultured neurons and other cells attached to a substrate has restricted investigations of in situ mitochondrial function in such preparations. By combining continuous flow oxygen electrodes with a digital fluorescence imager, it is possible to obtain quantitative information on mitochondrial and cellular energy supply and demand while simultaneously viewing a representative field to monitor cell viability together with parameters such as membrane potentials and [Ca²⁺]_c. The hypotheses that glutamate stimulates CGN respiration maximally, that the increase is entirely the result of increased ATP demand, and that excitotoxicity is associated with a loss of reserve ATP-generating capacity are not valid for the present experimental conditions, although the data support the hypothesis that the NMDA receptor is a significant contributor to the overall energy demand imposed by glutamate. It has not escaped our attention that studies to investigate neuronal bioenergetic changes associated with apoptosis and other stresses are equally feasible with this technique.

	FOOTNOTES

 
^* This work was supported by a National Institutes of Health Grant R01 AG21440. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore ^<