Glutamate Induced Astrocytic Metabolism Biology Essay

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Cellular metabolism is intimately associated with intracellular oxidation-reduction balance. Here, we describe cytoplasmatic redox changes in hippocampal neurons in culture exposed to glutamate. Neurons were transfected with HyPer, a genetically encoded redox biosensor for hydrogen peroxide which allows real-time imaging of the redox state. HyPer increases its signal with hydrogen peroxide in a dose-dependent manner meanwhile the decay of the fluorescence informs about the reducing capability of the cytoplasm. The rate of decay was found to be augmented by low doses of glutamate (10 mM) as well as by pharmacological stimulation of NMDA glutamate receptors. Acute chelation of extracellular Ca2+ abolished the glutamate-induced effect observed on HyPer fluorescence. Further experiments indicated that mitochondrial function and hence energetic substrate availability commands the redox state of neurons and is required for the glutamate effect observed on the biosensor signal. Finally, our work pointed out that astrocytic metabolism is involved in the changes of neuronal redox state observed with glutamate.

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Keywords: hippocampal neurons, HyPer, redox, astrocytes, glutamate, fluorescency microscopy.

Introduction

The brain is one of the most metabolically expensive tissues of the body. It poses a high rate of glucose and oxygen consumption, although corresponding only to 2% of the whole body weight (Leslie C.Aiello and Peter Wheeler 1995;Passingham 1973. The metabolic demand of the brain shows regional and spatial patterns determined by neuronal transmission, a feature that has been exploited in clinical diagnosis and basic research.

At resting conditions, brain cells supply their energy demand from glucose as the unique metabolite available for oxidation (Sokoloff 1977. A different scenario is established when glutamate, the main excitatory neurotransmitter, is released to the synaptic cleft. First, a high energy demand occurs in brain cells due to the recovering of ionic gradients dissipated during the synaptic transmission. For instance, astrocytes respond to glutamate by stimulating their glucose transport and glycolytic flux to ensure the continuous ATP production, a phenomenon coupled to Na+ gradient-driven glutamate reuptake and plasma membrane Na+/K+ ATPase pump (Pellerin and Magistretti 1994;Loaiza et al. 2003;Porras et al. 2004;Barros et al. 2009;Chuquet et al. 2010. On the other hand, neurons seem to do not activate their glycolytic flux under neurotransmission. Recent evidence provided by Chuquet et col. indicates that neurons from the barrel cortex do not increase their glucose metabolism as astrocytes do upon whisker stimulation in rats (Chuquet et al. 2010, confirming differential patterns of glucose metabolism observed before in primary cultures and brain slices (Porras et al. 2004;Barros et al. 2009. These findings indicate that neurons, despite their higher energetic demand (Sokoloff 1977;Attwell and Laughlin 2001, do not take the energy from metabolizing glucose, they rather prefer an alternative carbon source to fuel the costs of synaptic transmission, likely lactate. Indeed, many reports in vitro have shown that lactate and glucose are equally effective to sustain synaptic function (Schurr et al. 1988;Izumi et al. 1997;Morgenthaler et al. 2006. All these findings fit well with the idea proposed by Magistretti and col. where glutamate increases the astrocytic glycolysis resulting in a net release of lactate to the extracellular space, which is taken up by neurons to be oxidized by mitochondria. This notion is better known as astrocyte to neuron shuttle hypothesis, ANLSH (Pellerin and Magistretti 1994.

In the brain, blood-borne glucose metabolism produces the primary energy source, adenosine triphosphate (ATP), and reduced nicotinamide adenine dinucleotide (NADH). Oxidative phosphorilation of glucose-derived metabolites, either lactate or pyruvate, also render NADH and ATP in the Krebs cycle. Alternatively, glucose-6-phosphate (glc-6-P) can also be metabolized by pentose phosphate pathway to produce biosynthetic molecules and reduced nicotinamide dinucleotide phosphate (NADPH), the cofactor necessary for the regeneration of reduced glutathione (GSH) by glutathione reductase (Meister 1988;Bolanos and Almeida 2010;Aoyama et al. 2008. Despite of NADH and NADPH share similar redox characteristics their functions are divergent. Whereas NADH is destined to energetic functions, NADPH plays a role in cellular redox functions. NADPH is not only essential for the regeneration of all antioxidant defense systems, such as GSH, thioredoxins and peroxiredoxins, but also participates in detoxification with cytocrome p450 and in the "oxidative burst" mediated by NADPH oxidase in immune cells (Pollak et al. 2007;Agledal et al. 2010;Petry et al. 2010. These compartmentalized roles find convergence with the NAD+ kinase (NADK), which phosphorylates NAD+ and has shown to exert control over the cellular content of NADPH in mammalian cells (Pollak et al. 2007;Outten and Culotta 2003.

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The intracellular redox state is basically determined by the ratio of redox-active pairs NADH/NAD+, NADPH/NADP+, GSH/glutathione disulfide (GSSG) and thioredoxin/oxidized thioredoxin. Among those pairs, GSH with an abundance of 2 to 3 mM and a relative ratio of GSH/GSSG around 100:1 is the major anti-oxidant in the brain cells (Dringen 2000. Healthy cells maintain the cytoplasmatic environment at reducing potential of -250 mV, this balance is held despite of the endogenous generation of reactive species of oxygen (ROS) generated by mitochondria as by products from reduction of O2 and H2O during oxidative phosphorilation. Moderate shifts toward more oxidative potentials are thought to promote cell differentiation. For instance, Sundaresan and colleagues demonstrated that increases in hydrogen peroxide were essential for signaling induced by platelet derived growth factor on vascular smooth cells (Sundaresan et al. 1995, many other reports have established a physiological role for ROS in cellular events as well (Noble et al. 2003, for a review, please refer to (D'Autreaux and Toledano 2007. More severe increases in ROS formation become harmful to cellular components. DNA damage, lipid peroxidation and unspecific thiol oxidation of proteins are known factors which ultimately affect structural and enzymatic functions leading to cell collapse and death. Together, the evidence indicates that redox state is a dynamic variable in the cell and can be affected by metabolic preferences and signaling triggered by external factors such as growth factors.

Remarkable improvements in measuring fluctuations in the redox state in real-time mode have been possible due to fluorescent protein-based redox probes. Basically, green and yellow fluorescent proteins (FPs) have been chosen as template to place cysteine pairs able to form disulfide bonds under oxidative environments, which in turn produce changes in the fluorescent spectrum of the protein, giving the property to follow changes in the redox state with changes in fluorescent intensity of FPs, revised by Meyer and Dick (Meyer and Dick 2010. Moreover, genetically encode biosensors have been targeted to organelles in order to obtain real time measurements of redox state of endoplasmic reticulum lumen (Merksamer et al. 2008 and mitochondria (Belousov et al. 2006;Gutscher et al. 2008. Herein, we took advantage of the fluorescent protein-based redox biosensor, HyPer, which was designed to monitor specifically intracellular hydrogen peroxide and allows ratiometric measurements (Belousov et al. 2006. However, once HyPer molecule is oxidized, its disulfide bond is susceptible to be reduced back to free thiol groups by the action of reducing mechanisms present in the cell rather than a mere hydrogen peroxide clearance. Thus, HyPer does not only informs about intracellular hydrogen peroxide formation, but gives valuable information about the reducing capacity of the environment as well. Based on those properties, we were able to follow the acute changes in the redox tone of cytosol in neurons exposed to glutamate, because this neurotransmitter changes the metabolic preference of neurons towards lactate consumption we hypothesize that such change should also induce an increment in the reducing capability in hippocampal neurons.

Materials and methods

Reagents and plasmids. DL-Lactate, sodium piruvate, potassium valinomycin, sodium monensin, reduced glutathione and standard chemicals were purchased from Sigma (St. Louis, MO). Fetal bovine serum, Minimal Essential Medium, trypsin, penicillin-streptomycin, Glutamax, Neurobasal and B27 supplement and fluorescent probes (Fluo-3 AM, Fura-Red, BCECF) were purchased from Invitrogen (Carlsbad, CA, USA). Hydrogen peroxide was obtained from Merck (Darmstadr, Germany). pHyPer-cito plasmid was purchased from Evrogen JSC (Moscow, Russia).

Brain cell culture. Sprague-Dawley rats were obtained from the Universidad de Chile; the entire procedure with animals was approved by the bioethical committee of the Universidad de Chile under protocol CBA #0264 FMUCH. Mixed cultures of neuronal and glia cells were prepared from brains 1-3 days old neonates following the same indication as in Loaiza et al. with minor modifications (Loaiza et al. 2003. In brief, hippocampi were dissected in Hank´s buffer, incubated in 1.25% trypsin for 10 min at 37°C, and mechanically dissociated in MEM-10% fetal bovine serum: The cells were plated on poly-L-lysine coated (0.1 mg/ml) coverslips and maintained at 37°C in a humidified atmosphere with 5% CO2/95% air. After one hour, the media was replaced by Neurobasal/B27 and the cell preparation was maintained by renewal of media every 3 days. Experiments were performed after 7 to 14 days in KRH buffer (in mM, 140 NaCl, 4.7 KCl, 20 HEPES, 1.25 MgSO4 and 1.25 CaCl2, pH 7.4).

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PC12 Cell culture. Rat pheochromocytoma PC12 cells (from American Type Culture Collection) were cultured in DMEM supplemented with heat-inactivated 10% horse serum, 5% FBS, 50 units/mL penicillin and 100 mg/mL streptomycin (Invitrogen, Carlsbad, CA). PC12 differentiated neurons were obtained by plating cells onto poly-L-Lysine coated coverslips in the presence of 50 ng/mL NGF (Sigma-Aldrich, St Louis, MO) for a period of 5 days in DMEM medium supplemented with 1% heat-inactivated horse serum and 1% FBS.

HyPer imaging. Hippocampal co-cultures or PC12 cells were transfected with pHyPer-cyto, a plasmid encoding a specific hydrogen peroxide sensor with cytosolic expression for mammalian cells. The transfection methods used were lipofectamine 2000 (Invitrogen, Carlsbad, CA) or calcium phosphate, both rendering similar percent of neuronal cells expressing HyPer. Cultures were imaged using an inverted Olympus IX81 microscope with a 40X objective [numerical aperture, N.A. 1.3]. The preparation was excited at 403/12 nm and 480/20nm (MT20E emission wheel filter) and emission was detected at 555/28 nm with a CCD camera (XM10, Olympus). The experiments were conducted 48 hours after transfection at room temperature (23-27°C) in KRH buffer.

Calcium and pH measurements. Cells were loaded with the fluorescent cell-permeant calcium indicators Fura-Red or Fluo-4 at 5 mM for 30 minutes at room temperature in KRH-glc containing 0.02% pluronic acid followed by 20 minutes of de-esterification time. Experiments were carried out at room temperature in KRH buffer. Fura-Red and Fluo-4 were imaged at 480/20 nm excitation and emission for Fluo-4 was collected at 555/28 nm, whereas for Fura-Red was at 605/52 nm. For the intracellular pH measurements cells were loaded with 2 ,7 -bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein (BCECF) at 0.5 mM in its esterificated form for 5 min in KRH-glc containing 0.02% pluronic acid. In order to gain stability in the fluorescent signal through the experiments, cells were allowed to de-esterify for at least 30 min. Intracellular BCECF calibration was achieved clamping the pH using a combination of 100 mM nigericin with 25 mM valinomycin in a high K+ buffer (in mM: 130 KCl, 10 NaCl, 10 HEPES, 1.25 MgSO4 and 1.25 CaCl2, pH was adjusted to values ranging from 6 to 8). This method allows a rapid equilibration of protons between extracellular media and cytosolic compartment, the interchange of H+ for K+ by nigericin would be limited by the highly controlled K+ permeability present at the plasma membrane, adding valinomycin the potassium permeability is not longer rate limiting for the proton interchange by nigericin. This technique allows fast extracellular [H+] equilibration with the cytoplasm.

Statistical analysis. Regression analysis was carried out with Sigmaplot 5.0 (Jandel Corporation, Erkrath, Germany). Data are presented as means ± SEM throughout the entire manuscript. Differences in mean values were evaluated with paired Student's t-test for comparisons of before and after conditions and repeated measurement ANOVA (RM-ANOVA) when more than two conditions were present during the experiment.

Results

Glutamate induces a rapid decrease in HyPer fluorescent signal in hippocampal neurons.

The cytoplasmatic redox state was monitored by means of HyPer imaging in transfected neurons. Increments in the biosensor ratio have proven to be specific for hydrogen peroxide (Belousov et al. 2006. As depicted in Supplemental Figure 1, HyPer ratio increased in a dose-dependent manner showing saturation at H2O2 doses over 50 mM. Doses as high as 1 mM were tested without further increments in the fluorescence ratio of HyPer compared with the maximum reached with 100 mM H2O2 (not shown). Additionally, sequential pulses of hydrogen peroxide (100 mM) induced comparable increments in the fluorescence ratio although with some loss in the signal amplitude during the time course. In addition, the biosensor was sensitive to exogenous glutathione (1 and 5 mM) decreasing its signal, either if glutathione was added at basal level or if it was assayed over a pre-oxidized biosensor (Supplemental Figure 1).

In order to investigate the impact of glutamate on the reducing capacity of neurons, a pre-pulse of H2O2 100 mM was applied to oxidize the biosensor, after this procedure the fluorescence decreased with a constant rate of 0.04 ± 0.01 min-1 (21 exps., slope a, Figure 1A). The addition of glutamate (10 mM) significantly accelerated the signal decay (slope b, Figure 1A). This effect was observed in ~80% of the neurons studied and, in average the neurotransmitter increased the slope by almost four fold. It is important to clarify that the reduction in the fluorescence from the oxidized HyPer molecule does not obey to a cytoplasmatic clearance of hydrogen peroxide, but rather to disulfide bonds reduction mechanisms present in the cells, as it was suggested by Meyer and Dick (Meyer and Dick 2010 and also confirmed by the pre-treatment with the catalase inhibitor, 3-amino-triazol, which did not interfere with the glutamate effect in the HyPer signal (data not shown).

Next, we tested whether the pharmacological stimulation of the NMDA-sensitive glutamate receptors could mimic the glutamate effect observed above. As is shown in Figure 1C, NMDA was equally effective as glutamate to decrease the HyPer signal in a neuron previously treated with H2O2. Complementarily, stimulation of NMDA receptors with D-aspartate also accelerated the rate of HyPer signal decay. At the contrary, the pretreatment with the ionotropic glutamate receptor blockers, amino-5-phosphonovaleric acid and 6,7-dinitroquinoxaline-2,3-dione, prevented the effect of glutamate on the HyPer signal (Figure 1D). These results suggest that activation of NMDA-sensitive glutamate receptors is involved in the mechanisms that connect glutamate sensing with the rapid increase in the reducing tone in neuronal cytoplasm.

Ionotropic glutamate receptor activation, particularly NMDA-sensitive receptors, leads to Na+ and Ca2+ influx. In order to bypass complex molecular signals triggered by glutamate receptor activation, we tested whether an independent Na+ entry could trigger a reduction in the biosensor fluorescence as well. Neurons were exposed to monensin, a Na+/H+ antiporter which promotes an increase in intracellular Na+ at expense of proton extrusion. Figure 2A shows that exposure to monensin triggered a rapid decrease of the fluorescent signal of HyPer. In excitable cells membrane potential can be perturbed by a discrete cation entry, which in turn will open voltage dependent channels, among those Ca2+ permeant channels. Considering this issue, cytosolic Ca2+ imaging in neurons was performed to confirm that a massive increase in cytoplasmatic Ca2+ occurred in the presence of monensin (see supplemental figure 2A). Also, cytosolic pH was recorded in neurons loaded with the cell permeant pH-sensitive dye BCECF. As it was expected, monensin induced a rapid and transient alkalinization, see supplementary Figure 2B.

Alternatively, we tried a different manner to induce a Na+ influx without the pH perturbations triggered by monensin. To that end, we treated neurons with veratridine, an alkaloid that interacts with voltage-dependent Na+ channels only in their open state, promoting a slow and gradual Na+ influx to finally reach the threshold to open voltage dependent Ca2+ channels and generate a cytosolic Ca2+ increase (Porras et al. 2004. Veratridine also changed the rate of decay in HyPer signal as it can be observed in the figure 2C. Due to the results with the alkaloid were similar to those obtained with monensin and that, both compounds have different mechanisms to generate a Na+ influx, it can be inferred that augments in the reducing tone are independent of changes of intracellular pH. Additional experiments replacing extracellular Na+ with the non-permeant monovalent cation N-methyl-D-glucamine (NMDG+) showed that glutamate was able to accelerate the HyPer ratio decay from a basal average of 0.024 ± 0.004 to 0.084 ± 0.018 (min-1, p<0.05), indicating that Na+ influx is not absolutely necessary to glutamate evokes changes in the reducing tone in neurons.

Considering that a rise in the cytosolic Ca2+ levels is a common factor in the experimental maneuvers described above, we decided to test directly the role of Ca2+ influx in the glutamate effect, performing experiments with a Ca2+-free solution plus 5 mM EGTA. Under this condition, glutamate was unable to evoke a decrease in the HyPer fluorescence. However, the glutamate effect was reestablished upon Ca2+ reintroduction to the extracellular media (Figure 3A). On the other hand, the application of a Ca2+ ionophore as ionomycin augments the rate of decay of HyPer signal, mimicking the glutamate effect (Figure 3C).Together, these results indicate that Ca2+ influx is a central component in the glutamate-evoked reduction of the biosensor in the cytoplasmatic neuronal environment.

The astrocytic metabolism is connected with the neuronal redox status

Intriguingly, the decrease in the fluorescence signal of the biosensor either induced by Ca2+ influx with ionophores or by exposure to glutamate could be observed in the absence of any metabolic fuel (e.g. glucose). Considering that neurons do not store glycogen, it is reasonable to assume that after ten minutes without glucose the glycolitic flux had diminished to negligible levels. However, mitochondria still can be produce reducing power by feeding the Krebs cycle with intermediaries. To unveil if the mitochondrial machinery was involved in the biosensor signal decrease evoked by glutamate, neurons were pre-incubated with the mitochondrial poison antimycin A. Figure 4 shows that under this condition, glutamate was unable to affect the fluorescence of the biosensor. This finding suggests that mitocondrial function is a necessary component for the observed glutamate effect.

The co-culture setting used here allowed us to investigate whether astrocytic metabolism is connected in some way with neuronal redox status. In order to impair only glial metabolism, cells were treated with fluoroacetate (5 mM) for at least 2 hours before the experimental procedure. This compound is only metabolized by astrocytes where it is converted in fluorocitrate, which finally exerts the toxic action specifically on aconitase thus stopping the Krebs cycle (Muir et al. 1986;Clarke 1991. In order to ensure glycogen depletion in astrocytes, thirty minutes before the experiment started, glucose was washed out and the experiment was carried out. Figure 5A shows a time course of HyPer signal recorded from cells which were energetically deprived with fluoroacetate. As depicted, this condition was effective in avoiding the glutamate effect on HyPer described before. No significant changes in the basal rate of HyPer recovery before and after glutamate exposure were detected. The addition of exogenous lactate at the end of the experiment augmented the rate of decay in HyPer fluorescency, indicating that the astrocytic metabolic machinery, together with the availability of lactate in the media, is pivotal in the decrease of the HyPer ratio at neurons.

To explore the transfer of energy substrates, we switched to PC12 cells, a cell line that can be differentiated to a neuronal phenotype with NGF (Casado et al. 1996. Initially, the dose of glutamate (10 mM) used on neurons during this study was ineffective to trigger an intracellular Ca2+ rise in differentiated PC12 cells and thus, higher doses, 0.1 and 1 mM, were tried and their effectivity to mobilize Ca2+ was confirmed by calcium imaging (not shown). As depicted in Figure 5C, both doses of glutamate showed to be ineffective at decreasing the HyPer fluorescence. However, adding lactate resulted in a drop in the HyPer signal with similar characteristics as those evoked by glutamate in neurons. These results strongly suggest that although differentiated PC12 cells exhibit the glutamate-sensing machinery, glutamate-induced redox changes require extra-neuronal (PC12 cells) factor(s) to evoke an increase in PC12 cytosolic reducing capacity. The effect of lactate 5 mM on the biosensor signal was further quantified in naïve neurons and PC12 cells. In both cellular types, the presence of this monocarboxylate increased the rate of HyPer fluorescence decay from a basal value of 0.18 ± 0.04 to 0.46 ± 0.07 min-1 in hippocampal neurons, whereas in PC12 cells was from a basal rate of 0.30 ± 0.08 to 0.65 ± 0.08 min-1 (supplemental figure 3). Another mitochondrial fuel such as pyruvate also accelerated the rate of decay of HyPer ratio from 0.07 ± 0.03 to 0.21 ± 0.05 min-1 (p < 0.05, 10 cells from 4 experiments). The effects of those energetic substrates on the reducing tone confirm that the availability of energetic substrates has an impact on the cellular redox state.

Discussion

Many cellular changes take place in neurons upon exposure to glutamate. Among those, changes in the metabolic preference of neurons have been a controversial point ever since the ANLS hypothesis was proposed in 1994. Since then, several reports have supported the notion that brain cells adapt their metabolism under glutamatergic neurotransmission (Pellerin and Magistretti 1994;Loaiza et al. 2003;Porras et al. 2004;Kasischke et al. 2004;Suzuki et al. 2011. Here, we have shown that metabolic coupling between neurons and astrocytes affects the rate of disulfide bonds reduction at neuronal cytoplasm is modulated by glutamate and that, this effect occurs at expenses of astrocytic metabolism.

This work presents real time monitoring of the reducing capabilities of the neuronal intracellular environment, specifically the cytoplasm. Our particular experimental strategy consisted in pre-oxidize the biosensor with a hydrogen peroxide pulse in order to monitor the rate of fluorescent ratio decay as a reflex of the reducing capability of the cytoplasm. The intracellular reduction of HyPer molecule on its disulfide bond is a process not related to hydrogen peroxide clearance, hence does not mediated by catalase activity, but rather related to enzymatic-mediated reduction mechanism (Meyer and Dick 2010. A recent study showed that roGFP1, another biosensor carrying a cysteine pair to sense oxidation/reduction reactions (Dooley et al. 2004;Funke et al. 2011 displayed no significant decrease in its basal fluorescence in HeLa cells exposed acutely to aminotriazole, an inhibitor of catalase. Only, the inhibition of thioredoxin reductase or dihydrolipoamide dehydrogenases was effective to reduce the roGFP1 signal reduction after a transient pre-oxidation with aldrithiol (Funke et al. 2011, confirming the idea that reduction of disulfide bonds depend of cellular enzymatic mechanism.

There is an overall consensus about that excitatory neurotransmission is highly expensive in terms of energy. Most of the ATP is consumed by neurons to power ionic pumps in order to restore the dissipated ionic gradients (Attwell and Laughlin 2001;Alle et al. 2009. Although glucose and lactate could provide sufficient energy to allow recycling of neurotransmitter in isolated nerves (Tarasenko et al. 2006 and also, allow synaptic plasticity in hippocampal brain slices (Izumi et al. 1997;Yang et al. 2003. Evidence from rat sensorial cortex indicates that neurons do not modify their glucose uptake during whisker stimulation as astrocytes do (Chuquet et al. 2010. In fact, active neurons seem to prefer lactate to function properly, a statement that showed to be truth in hippocampal formation, where long-term memory formation was dependent of lactate derived from glycogen breakdown, a process that only occurs at glia counterpart (Suzuki et al. 2011. In consequence, neurons should metabolize lactate to piruvate, a process catalyzed by the enzyme lactate dehydrogenase, which renders a NADH molecule. After this cytosolic step, piruvate would be further oxidized by mitochondria in the Krebs cycle, where electrons will finally reduce O2 to H2O producing more NADH and ATP, rendering neurons more aerobic than astrocytes during neurotransmission. Our results indicate that besides the metabolic changes evoked by glutamate, an accelerated reduction of disulfide bonds present in HyPer molecules depends of mitochondrial functioning as well. A process that fits well with the subcellular source described for the increments of NAD(P)H-derived intrinsic fluorescency measured in electrically stimulated brain slices (Brennan et al. 2006). Since HyPer is localized at cytoplasm and the reducing power is produced at mitochondria, it is clear that there are efficient mechanisms allowing redox transfer between these two compartments. In HeLa cells, mitochondrial depolarization and cytosolic GSH oxidation showed reciprocal dynamics, both cellular parameters measured by fluorescent microscopy of TMRM and glutaredoxin-roGFP biosensor, respectively (Gutscher et al. 2008. Additionally, exists many mechanisms implicated in the redox transfer between mitochondrial and cytosolic compartments (McKenna et al. 2006. Besides the pentose phosphate pathway, enzymatic mechanism that interchanges NADH for NADPH in eukaryotic cells (Outten and Culotta 2003 plus other sources of NADPH generation probably contribute to maintain the redox equivalents interchange and add complexity to the intracellular redox scenery(Pollak et al. 2007.

The co-culture of astrocytes and neurons used in this work was useful to study some aspects of metabolic interactions between those cells. Since the effect on the HyPer signal was obtained without extracellular glucose and was dependent of mitochondrial function, was natural to seek the source of the fuel that was powering neuronal mitochondria. At variance with astrocytes, neurons do not store glucose in the form of glycogen (Brown 2004;Magistretti and Allaman 2007 and have been reported that this cellular type responds releasing lactate to the media upon glutamate stimulation (Tsacopoulos and Magistretti 1996;Bittner et al. 2011). Therefore, astrocytes are the natural candidates to be responsible for energy transferring in our experimental conditions. With this in mind, we energetically deprived astrocytes using fluoroacetate, a specific astrocytic toxin used before to prove that astrocyte-derived lactate was able to maintain spontaneous firing in orexin neurons from hypothalamic brain slices (Parsons and Hirasawa 2010. In this study, glutamate lost its effect on HyPer reduction in both, co-cultures submitted to energetic depletion and in a pure cell line which lacks of supporting cells as astrocytes. Moreover, the fact that exogenous lactate was able to elicit a reduction in HyPer signal in both cellular systems aims to fueling mitochondria leads to an increase in the reducing tone at the cytoplasm.

Lactate metabolization by neurons necessarily implies lactate uptake via monocarboxylate transporters (MCTs), mainly the isoform MCT2. Due to translocation of monocarboxylates is accomplished in its protonated form, an intracellular acidification is expected. HyPer fluorescence is sensitive to pH changes, thus the natural acidification rate could be a component of the biosensor reduction rate. We consider that pH changes exert little interference in our case, because a controlled acidification of 0.5 pH units renders a drop in HyPer ratio of 0.8 units, whereas neurons exposed to glutamate 10 mM only presented a drop of 0.1 pH units within a time window where most of the effects where visualized (~150 seconds), suggesting that during the time window where the decrease of HyPer-derived fluorescence were monitored, the acidification of the cytoplasm is insufficient to explain the robust decrease of the HyPer fluorescence ratio. Another argument against the acidification is given by the effect obtained with the Na+/H+ antiporter, monensin. Even though this ionophore produces a rapid alkalinization, HyPer ratio diminished to similar levels as observed upon glutamate exposure. These observations allows us to rule out the acidification as a major contributor in the HyPer fluorescence ratio decrease evoked by glutamate, suggesting instead that a shift in neuronal metabolism induced by glutamate generates an increase in the cytoplasmatic reducing potential.

Our results indicate that the mitochondrial function is a key component to neurons increase the reducing potential at their cytoplasm. This observation is contradictory to the generalized notion of mitochondria as the major ROS source in the cell. Experimental evidence indicates that H2O2 production by liver mitochondria accounts only for 1-2% of the total oxygen consumption. These values were obtained with mitochondria powered by succinate at nonphysiological oxygen conditions, meaning that even under "forced" conditions mitochondria are extremely efficient to transfer electrons and avoid ROS generation (Boveris et al. 1972;Kudin et al. 2008. Additionally, there are reports that point out mitochondria as a sink for cytosolic ROS. Mitochondrial scavenger systems such as superoxide dismutase and catalase enzymes located at intermembrane space and matrix confer a high scavenging capacity per se. However, Zoccarato et al. showed that energized mitochondria are twenty-fold more efficient to remove extra-mitochondrial H2O2 than de-energized, suggesting that the production of reducing equivalents are linked to the metabolic function of this organelle (Zoccarato et al. 2008. Noteworthy, Ca2+ signaling might be working as a feed-forward mechanism to prepare neurons to efficiently metabolize lactate and piruvate by the mitochondria. Physiological Ca2+ increases at the cytoplasm have measurable impact in the levels of free Ca2+ in mitochondria; such connection has been recorded by mitochondria-targeted aequorin in beating cardiomyocytes (Bell et al. 2006. Moreover, calcium is a well known stimulating factor of oxidative metabolism in cardiomyocytes. Calcium Increases have been associated to stimulation of mitochondrial dehydrogenase enzymes, with the concomitant increase in NADH (Garcia-Perez et al. 2008 and ATP production (Bell et al. 2006.

In summary, we propose that neuronal work is associated with a metabolic adaptation with consequences in the reducing potential of cytosolic environment. If this elevated reducing capability is able to reduce a disulfide bond on the biosensor molecule probably affects in the same way other proteins with cysteine residues prompt to be modified by the redox status. The maintenance of this reducing tone at the cytoplasm under a metabolic demanding stage might be acting as a mechanism to avoid the oxidation of glutamate transporter (EAATs), which activity is impaired by oxidant agents and could lead to an excitotoxicity by failure to maintain extracellular glutamate at safe levels (Yun et al. 2007). Other redox-sensitive targets involved in excitotoxicity comprise the ryanodine isoform 2 channel, RyR2, which experiences S-glutathionylation in the endoplasmic reticulum from isquemic brains resulting in an increase in calcium release, a perturbation of the calcium homeostasis that may contribute to neuronal death (Bull et al. 2008. Calcium-activated potassium channels also show GSH sensitivity, their activity decrease at low levels of GSH in hippocampal neurons (Soh et al. 2001. Finally, activation of TRPM7-mediated inward currents by ROS have shown to be a key component of the vicious loop between ROS generation, increased cytosolic Ca2+ and TRPM7 stimulation that results in cortical neuron demise (Aarts et al. 2003.

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