Abbreviations
-
- 2-APB
-
- 2-aminoethoxydiphenyl borate
-
- ADPR
-
- ADP-ribose
-
- DAG
-
- diacylglycerols
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- DCD
-
- delayed calcium deregulation
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- KGDHC
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- α-ketoglutarate dehydrogenase complex
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- NMDA
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- N-methyl-d-aspartate
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- PTP
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- permeability transition pore
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- RNS
-
- reactive nitrogen species
-
- ROS
-
- reactive oxygen species
-
- siRNA
-
- short interfering RNA
-
- SOC channel
-
- store-operated Ca2+ channel
Background
A long-standing perception is that upon activation of glutamate receptors followed by a robust Ca2+ influx, in situ mitochondria generate reactive oxygen species (ROS) [1–6]. These studies inferred that mitochondrial Ca2+ sequestration is a prerequisite for production of ROS: abolition of mitochondrial membrane potential (ΔΨm) by mitochondrial poisons, and thus, electrophoretic calcium uptake or direct inhibition of the uniporter with ruthenium red prevented ROS generation. Parallel to these reports, the response of isolated mitochondria to calcium loading in terms of ROS production has also been scrutinized; it was found that mitochondrial Ca2+ uptake led to free radical production [7–12]. On the other hand, it was shown that ROS formation depends steeply on ΔΨm [13–15], and from a thermodynamic point of view, Ca2+ uptake occurring at the expense of membrane potential should result in a decrease in ROS production (in the absence of respiratory chain inhibitors), as it has also been demonstrated (reviewed in [16,17]). Nevertheless, brain mitochondria also generate ROS in a ΔΨm-independent manner [18–20]. The reason behind the opposing observations that mitochondrial ROS production increases or decreases upon Ca2+ uptake is not entirely clear; a plausible explanation lies in the condition in which mitochondria are probed for ROS, specifically whether or not the organelles undergo permeability transition pore (PTP) formation. Among the many features accompanying mitochondrial permeability transition (for a full list see [16] and references therein) loss of glutathione, cytochrome c, substrates and pyridine nucleotides are characteristic. This leads to an increase in ROS production from the impaired mitochondria by multiple means: (a) loss of glutathione from the matrix decreases the antioxidant capacity resulting in a net ‘steady-state’ increase in the amount of ROS [21]; (b) loss of cytochrome c impairs the flow of electrons in the respiratory chain inducing over-reduction of the complexes, favouring the generation of ROS [16,17,22]; (c) reduction in the matrix concentration of electron acceptors, i.e. NAD+, results in ROS emission from the α-ketoglutarate dehydrogenase complex (KGDHC) [23,24].
Mitochondrial formation of ROS-the role of KGDHC
The first observation of ROS production in mitochondrial fragments was reported in 1966 by Jensen [25]. Subsequent studies by Britton Chance's group, established that mitochondria generate ROS [26,27]. The sites of ROS formation within the organelle have been extensively reviewed elsewhere [17,20,28]. Among them, complex I [29–31] and III [32–35] of the respiratory chain have attracted most attention. However, in light of recent results on the substantial contribution of matrix enzymes (especially KGDHC) on ROS generation, we believe that in addition to the respiratory chain, the components of the Krebs cycle should also be considered as a possible important source of ROS in mitochondria.
Almost all studies have used respiratory chain inhibitors as tools to maximize and to identify potential sites of ROS production in isolated mitochondria. They revealed that inhibition of complexes I and III, respectively, with specific mitochondrial toxins such as rotenone and antimycin A, results in high rates of ROS production [29,36,37]. For complex I in particular, the ‘reverse electron transport’ mode of ROS production has gained momentum throughout the past four decades [38]; reverse electron transport requires high ΔΨm and is abolished by the complex I inhibitor, rotenone [18], but the pathophysiological relevance of this mode of ROS generation is questionable. Similar approaches have been used successfully to study ROS production in in situ brain mitochondria present in isolated nerve terminals (synaptosomes) [39], but no information is yet available regarding the specific sites or mechanisms of ROS generation in the absence of respiratory chain inhibitors.
Numerous reports in isolated or in situ mitochondria support complex I being regarded as a major site of ROS production, however, a lingering assumption remains that all ROS production caused by complex I inhibitors occurs at the complex I site. There are other sources of ROS within the mitochondrial matrix that are in equilibrium with the ratio NAD(P)H/NAD(P)+, such as the dihydrolipoyl dehydrogenase (Dld) component of KGDHC [40]. In intact mitochondria, complex I inhibition by any means, inevitably results in over-reduction of most if not all NAD+-linked matrix enzymes.
Among the NAD+-linked dehydrogenases that generate ROS, KGDHC deserves special attention. KGDHC is a mitochondrial enzyme tightly bound to the inner mitochondrial membrane on the matrix side [41]. It (as well as other but not all dehydrogenases) binds to complex I of the mitochondrial respiratory chain [42] and may form a part of the TCA cycle enzyme supercomplex [43]. Mammalian KGDHC is composed of multiple copies of three enzymes: α-ketoglutarate dehydrogenase (E1; EC 1.2.4.2), dihydrolipoamide succinyltransferase (E2; EC 2.3.1.61), and dihydrolipoamide dehydrogenase (E3 or Dld; EC 1.8.1.4). Dld is also a part of other multienzyme complexes such as the pyruvate dehydrogenase complex (PDHC), the branched chain ketoacid dehydrogenase complex, and the glycine cleavage system [44–47]. The catalytic mechanism of the α-ketoacid dehydrogenase complex was reviewed by Bunik [40].
Isolated KGDHC [23] as well as PDHC [24] in isolated and in in situ mitochondria respectively produce superoxide and H2O2. Quantitatively, it seems likely that KGDHC generates the majority of ROS among dehydrogenases: under conditions of maximum respiration induced with either ADP or an uncoupler, α-ketoglutarate supports the highest rate of H2O2 production [24]. The Dld component of KGDHC, and to a lesser degree of PDHC, generate ROS in isolated mouse brain mitochondria [24]. The reasons behind this quantitative discrepancy among the Dld-containing dehydrogenases regarding ROS production are at present, unknown. The isolated Dld subunit is able to form H2O2 and superoxide radical, accompanying NADH oxidation [40,48,49]. This observation is important as to the mechanisms and sites of ROS production in mitochondria because the flavin of the Dld subunit is abundant and possesses a sufficiently negative redox potential (Em 7.4 = −283 mV) to allow superoxide formation [50,51]. Moreover, H2O2 production by brain mitochondria isolated from heterozygous knockout mice deficient in Dld is significantly diminished, as compared to wild-type littermates [24].
Within KGDHC, it is the flavin or the neighbouring disulfide bridge in the catalytic centre of the Dld component that could act as an electron donor for superoxide formation [52]. KGDHC is activated by low concentrations of Ca2+ and matrix ADP [53–56]. Considering that KGDHC-mediated ROS production requires a fully active complex with all the cofactors and substrates (except NAD+), the fact that the enzyme activity is stimulated by Ca2+ and ADP may perhaps account for previous findings that mitochondrial ROS production was increased by Ca2+[7–11,14] and ADP [30]. Results obtained in our laboratory [23] demonstrate that Ca2+ activates ROS production by isolated KGDHC both in the presence and in the absence of pyridine nucleotides. Still, the reduced Dld subunit is the most likely source of ROS under conditions of an elevated NADPH/NADP+ ratio in the mitochondrial matrix [23,24]. The conditions promoting KGDHC-mediated ROS production may be any that increase the intramitochondrial NADH/NAD+ ratio (e.g. inhibition of oxidative phosphorylation or inhibition of any segment of the mitochondrial electron transport chain). This hypothesis is favoured by our results showing that ROS production by isolated KGDHC is strongly dependent on the NADH/NAD+ ratio [23].
The relationship of ROS to KGDHC is extended in an ‘ouroboros’ fashion to the self-inactivation of the enzyme by ROS. We demonstrated previously, that KGDHC is sensitive to inhibition by H2O2[57]. That inevitably leads to a decrease in complex I function, as repeatedly demonstrated [57–61], since KGDHC which is the rate-limiting step of the TCA cycle provides NADH as a substrate for the respiratory chain complex.
It is difficult to establish the extent of contribution of KGDHC and other enzymes to overall ROS production in mitochondria, as this is prone to be condition-dependent (e.g. choice of substrate), in addition to heavily reliant on non-Krebs cycle enzyme mediated ROS formation through the respiratory chain; i.e. both complex I and KGDHC are in equilibrium with the NAD(P)H/NAD(P)+ ratio, and therefore interdependent on each other concerning ROS formation. Thus, in organello it might not be possible to accurately estimate the degree of contribution of each ROS-forming site, because inhibition of ROS production in the one may aggravate ROS formation in the other, and vice versa.
The observation that KGDHC generates and is also self-inactivated by ROS, is of paramount importance in neuronal pathology. A compelling body of evidence indicates that mitochondria are the major source of ROS in several neurodegenerative conditions [37,62]. Also, KGDHC activity is severely reduced in a variety of neurodegenerative diseases associated with impaired mitochondrial functions, specifically, Alzheimer's disease [63–67], Parkinson's disease [68–71], progressive supranuclear palsy [72,73] and Wernicke–Korsakoff syndrome [74]. It is not known if the physical association of KGDHC with complex I (see above) plays a role in the dual deficiency of these protein complexes in Parkinson's disease. It appears that neuronal pathology is preferentially associated with KGDHC deficiency: in an animal model of diminished KGDHC activity caused by thiamine deprivation in the diet, neurons are dying, while endothelial cells, astrocytes and microglia are not affected. In fact, KGDHC activity is increased in these non-neuronal cell types [63], which might indicate that KGDHC deficiency has an etiologic role in the manifestation of some neurodegenerative diseases [75,76]. It must be emphasized that this multienzyme is the rate-limiting step of the Krebs cycle, and if altered that would inpact on the overall energy production in the affected tissue. Moreover, in vivo studies suggested that reduced activity of KGDHC predisposes to damage by toxins, such as 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP) or malonate, reducing the capacity of neurons to respond to stress [77,78]. In addition, it was shown recently that reduction in the E2 subunit of KGDHC is associated with diminished growth of cells and impaired antioxidant defence systems, without a reduction in the overall activity of the complex [79]. This finding should come at no surprise: several enzymes of the TCA cycle (and at least one glycolytic enzyme [80]) have roles beyond those of just being cycle participants for the provision of reducing equivalents: aconitase, isocitrate dehydrogenase and kgd2p (a subunit of KGDHC in yeast equivalent to E2 in mammals), have two or more different functions, in addition to having supporting functions for oxidative defences [79], involving the thioredoxin system [40]. Aconitase acts also as an iron-responsive element binding protein, isocitrate dehydrogenase is an RNA-binding protein, while kgd2p is a mitochondrial DNA binding protein [81–84].
Mitochondria from different brain regions contain different amounts of KGDHC [85,86], which may account for regional vulnerability. For instance, the cholinergic neurons of the nucleus basalis of Meynert have high levels of KGDHC, and these neurons are particularly vulnerable in Alzheimer disease [64].
Nevertheless, the relationship between KGDHC activity and mitochondrial damage per se is much less clear. One can speculate that KGDHC-mediated oxidative stress predisposes the cell to succumb to concomitant adverse conditions; in addition, a diminished KGDHC activity will lead to insufficient provision of reducing equivalents, lowering the energetic capacity of the mitochondria of the affected cell. However, studies with the KGDHC inhibitor KMV (alpha-keto-beta-methyl-n-valeric acid) suggest that inhibition of the enzyme might contribute to cell death by induction of permeability transition [87].
Permeability transition pore in situ
Permeability transition pore is considered to be a channel with a large conductance provided by proteins residing in both the inner and outer mitochondrial membrane, that is activated by mitochondrial Ca2+ overloading and other factors including oxidative stress [88,89]. In neurons the presence of PTP in situ has not gained wide acceptance among investigators and results published in the literature support views of both its presence and absence in several in vitro models of neurodegeneration [90–98]. One of the possible reasons for this discrepancy is that sensitivity to cyclosporin A is considered pathognomonic for mitochondrial PTP (see also [90]). Cyclosporin A is a potent inhibitor of PTP in isolated liver mitochondria [99] that has been demonstrated to be effective also in situ in this and other organs [100–103]. The sensitivity of isolated brain mitochondria to cyclosporin A depends highly on the conditions: in the absence of adenine nucleotides and magnesium, cyclosporin A mitigates Ca2+-induced mitochondrial pore formation [104,105] however, in the presence of 3 mm ATP plus 1 mm free Mg2+, cyclosporin A is only marginally effective, provided that mitochondria are challenged by boluses of CaCl2[104]. In the case that Ca2+ loading occurs slowly, cyclosporin A delays onset of PTP in brain mitochondria extensively, even in the presence of adenine nucleotides and magnesium [106]. The caveat here is that despite the decreased ATP levels to less than the millimolar range during ischemic deenergizing, ADP levels approximate 400 µm[107], and the Ki for inhibition of the PTP by ADP is in the low micromolar range [108]. Moreover, in situ neuronal mitochondria are exposed to bolus-like additions of Ca2+[109] during intense glutamate receptor stimulation for the duration of seizure activity or reversal of glutamate transporters throughout ischemia [110]. Ca2+ cycling across the mitochondrial inner membrane ensues subsequently [111]. On the other hand, intense stimulation of N-methyl-d-aspartate (NMDA) receptors on cultured cerebellar granule and hippocampal neurons causes major ultrastructural alterations of mitochondria, implying the activation of some form of PTP [112,113]. Mitochondrial alterations suggestive of pore opening is also demonstrated in vivo, during the postischemic period in the gerbil brain [114]. Yet, to identify these in situ mitochondrial alterations as the PTP on the basis of the functional/morphological/pharmacological criteria applied for isolated mitochondria is rather hasty.
Collectively, the sensitivity of glutamate-induced neuronal damage to cyclosporin A as diagnostic for PTP occurrence is unreliable. This ambiguity is also nurtured by the complex pharmacology of cyclosporin A and its affinity to non-PTP targets [90,115] that could be involved in the manifestation of neuronal injury [116], in addition to the fact that PTP may not have a causal role in excitotoxic cell death. It is to be noted that the magnitude of the literature involving cyclosporin A unrelated to mitochondria is 12 times larger than that implicating PTP! The nonimmunosuppressant analogue, N-methyl-valine-4-cyclosporin also gave contrasting results, conferring neuronal protection against excitotoxicity in some studies [92,117,118], but not in others [94].
What could be important though, is the role of the in situ mitochondrial pore formation in dictating the type of death that the ill-fated neuron will follow. A most simplistic view is that this pore will promote apoptosis due to release of cytochrome c followed by activation of caspases [119,120], provided that pertaining conditions divert the type of cell death from the necrotic to the apoptotic pathway [121,122]. The role of mitochondria in apoptosis and necrosis has been extensively reviewed elsewhere [121,123–131]. Recently however, a blow was delivered to the conception that PTP contributes to apoptotic cell death by three almost simultaneous and independent reports using cyclophilin D knockout mice [132–134]. Cyclophilin D is a component of the PTP complex [135,136] and it is the target for cyclosporin A. As expected, mitochondria isolated from the cyclophilin D knockout mice were much less susceptible to various PTP-inducing regimes, that are otherwise sensitive to cyclosporin A treatment (see also [137]). Unexpectedly though, tissues obtained from mutant mice were not more resistant to several apoptotic stimuli than those from their wild-type littermates; however, the resistance of the mutant mice to treatments known to result in necrotic cell death was much higher than in control mice.
Mitochondrial Ca2+-flux pathways and relation to signal transduction
In general, the contribution of mitochondria to intracellular Ca2+ homeostasis is ascribed to uptake and release through the uniporter, the mitochondrial Na+/Ca2+ exchanger, the PTP (both high- and low-conductance mode) and other less well characterized pathways, such as the ‘Na+-independent pathway for Ca2+ efflux’ and a H+/Ca2+ antiporter [89,138]. With the exception of the high-conductance mode of PTP and the uniporter, none of these molecular complexities have been described to be modulated by any signal transduction mediators. High-conductance PTP is known to be affected by matrix Ca2+ and ROS [89]. Also the uniporter is supposed to be activated only if extramitochondrial Ca2+ levels exceed a certain threshold concentration, termed the ‘set-point’[139]; however, this has been challenged recently, showing that in situ mitochondria accumulate Ca2+ well below the set-point, in permeabilized rat adrenal glomerulosal cells [140]. Nonetheless, despite that mitochondria are increasingly viewed as active mediators of [Ca2+]c regulation, the pathways that these organelles use to achieve this task are rather passive.
To this repertoire of Ca2+ influx and efflux mechanisms across the mitochondrial membranes, a novel Ca2+-efflux-only machinery has been recently added: a channel located in the inner membrane activated by diacylglycerols (DAGs) [141]. This is either a single channel with numerous substates (mean conductance ≈ 200 pS), or multiple channels with unequal conductance. DAGs cause a biphasic form of Ca2+ efflux in Ca2+-loaded mitochondria: the first wave of efflux is attributed to the activation of the DAG-sensitive nonselective cationic channels; the second wave is due to opening of the PTP. It is not yet known how activation of the former leads to induction of the latter. One is tempted to hypothesize that the initial Ca2+ efflux through DAG-sensitive channels causes intense Ca2+ cycling due to reuptake by the uniporter, leading to PTP. However, cyclosporin A fails to defend against the secondary Ca2+ efflux in liver mitochondria in the presence of DAGs, in which the immunosuppressant otherwise confers significant protection against PTP induction.
The role of DAG-sensitive mitochondrial channels in physiological [Ca2+]c regulation can easily be envisaged: upon phosphatidylinositol (4,5) bisphosphate (PIP2) hydrolysis, inositol-1,4,5-triphosphate (IP3) diffuses in the cytosol to activate IP3 receptors on the endoplasmic reticulum releasing Ca2+ to the cytoplasm, followed by triggering of Ca2+ influx from the extracellular space [142]. The role of mitochondria in shaping Ca2+ transients during such events is recognized in limiting Ca2+ diffusion, and secondarily relieving Ca2+-mediated negative feedback on the Ca2+ flux pathways themselves [143]. However, the other obligatory metabolite of PIP2 catabolism − DAG − may regulate the role of mitochondria in shaping those [Ca2+]c transients: mitochondrial DAG-sensitive channels would re-release sequestered matrix Ca2+ only in the vicinity where DAGs are formed most likely in microdomains, since this second messenger is extremely lipophilic and does not diffuse into the aqueous cytosol.
Mitochondrial permeabilization and the delayed calcium deregulation
The association of ROS to a possible PTP induction prior to neuronal cell death has received much attention in relation to the delayed, irreversible rise in [Ca2+]c following a prolonged glutamate stimulus, coined by Nicholls' group as ‘delayed calcium deregulation, DCD’[144] that commits a neuron to die [145–148]. DCD was originally described by Manev and colleagues [149], further characterized by the groups of Thayer [150] and Tymianski [146]. However, credit should also be given to an earlier work by Connor and colleagues, showing that a short exposure (1–3 s) of CA1 hippocampal neurons to NMDA causes an abrupt elevation in [Ca2+]c that returns to baseline; a subsequent exposure to NMDA of the same duration a few minutes later leads to an irreversible and sustained increase in intracellular [Ca2+]c in apical dendrites [151]. DCD is invariably demonstrated in every neuronal cell type studied, i.e. spinal [146], hippocampal [150], cerebellar granule [152], striatal [117] and cortical neurons [93,153]. The phenomenon is not observed if high extracellular K+ is alternatively employed to elevate [Ca2+]c; this led to the proposal of a ‘source specificity’ of Ca2+-induced neurotoxicity [146]. However, this was subsequently challenged by studies demonstrating that activation of NMDA receptors produces much larger Ca2+ entry than activation of voltage-dependent Ca2+ channels by high extracellular K+[154].
This secondary [Ca2+]c rise is not inhibitable by postglutamate addition of antagonists of NMDA or non-NMDA receptors [94,145,149,150], nor by blocking voltage-dependent Ca2+ or Na+ channels [145,149,150,155]. Results supporting views that DCD is comprised of an active Ca2+ influx pathway [93,146,149,150,155–159] as well as those indicating a failure in Ca2+ efflux mechanisms [160–162], are available in the literature. It is anticipated that these seemingly opposing observations represent two-facets of the same problem: even in the earliest report on DCD by Manev and colleagues [149] it was shown that during the postglutamate period neurons still accumulate 45Ca2+ within 30 s exposure to the isotope, without any statistically significant difference seen in the presence or absence of N-methyl-d-aspartate receptors/non-N-methyl-d-aspartate receptors/voltage dependent Ca2+ channels (NMDAR/non-NMDAR/VDCC blockers). That attests to the presence of a discrete pathway for Ca2+ influx. Yet, it was recently demonstrated that in an almost identical paradigm of excitotoxicity, the plasmalemmal Na+/Ca2+ exchanger (in particular the NCX3 isoform) is cleaved by calpain, severing the high capacity Ca2+ efflux pathway in neurons [161]. Provided that the Ca2+ influx pathway is most likely a channel, it must saturate [163] imposing a continuous load of calcium to the neuron. The turning point upon which the cell looses the ability to buffer the incoming calcium resulting in an abrupt, sustained and irreversible increase in [Ca2+]c, probably coincides with the cleavage of the exchanger (but see [164]). Therefore, inhibition of the, as yet unidentified, Ca2+ influx pathway or prevention of NCX proteolysis should thwart DCD. The question arises: what is the nature of the Ca2+ influx pathway?
Non-selective cationic channel(s) and the DCD
As mentioned above, inhibition of NMDAR/non-NMDAR/voltage-dependent Ca2+ or Na+ channels after the initial Ca2+ and Na2+ influx through the glutamate receptors, failed to prevent DCD. Yet, DCD demands the existence of a discrete pathway as it precedes, and eventually leads to, plasma membrane leakiness and cell death [145,146,148]. The notion that DCD is not attributed to the ‘traditionally’ recognized Ca2+ channels, such as glutamate receptor-operated or voltage-gated Ca2+ channels has been proposed previously [157,158]. Along this line, it was shown that a secondary activation of a nonselective cation conductance, termed postexposure current (Ipe), is induced subsequent to excitotoxic application of NMDA to hippocampal neurons that probably contributes to the delayed Ca2+ rise [156].
Relevant to the inability of the glutamate receptor blockers to prevent DCD, antiexcitotoxic therapy utilizing these compounds failed to produce a better outcome in clinical trials concerning stroke treatment [165–167]. To address this setback, Aarts and colleagues [159] examined the possibility that an overlooked neurotoxic process was occurring in a well-established in vitro model of excitotoxicity, by subjecting cultured neurons to oxygen–glucose deprivation. This treatment results in neuronal demise through NMDAR activation [168,169]. It was found that a member of the melastatin branch of the transient receptor potential channel (TRP) family, TRPM7 [170], mediates a lethal cation current loading the neurons with Ca2+ and Na+. This nonselective current was activated by ROS and reactive nitrogen species (RNS), and its abolition permitted the survival of neurons previously destined to die from prolonged anoxia, regardless of the presence or absence of NMDAR blockers.
In a subsequent study, we explored the hypothesis that a TRP channel contributes to the manifestation of DCD [93]. A pharmacological approach was used, applying 2-aminoethoxydiphenyl borate (2-APB) or La3+ to cultured cortical neurons challenged by prolonged glutamatergic stimulation. We observed that 2-APB and La3+ diminished the delayed Ca2+ rise with a 50% inhibitory concentration of 62 ± 9 µm and 7.2 ± 3 µm, respectively. Both substances are known to inhibit TRP channels in addition to acting on many other targets; 2-APB blocks store-operated Ca2+ (SOC) channels [171], the IP3 receptor [172], the sarco-endoplasmic reticulum Ca2+ ATPase (SERCA) pump [173], voltage-dependent K+ channels [174], gap junctions [175] and the cyclosporin A-insensitive PTP [104], while La3+ blocks SOC [176] and voltage-dependent Ca2+ channels [177]. Almost all non-TRP targets are irrelevant or have been previously excluded concerning the origin of DCD, except for the cyclosporin A-insensitive PTP that is abolished by 2-APB in isolated brain mitochondria [104]. However, in our hands, bongkrekic acid ameliorated the cyclosporin A-insensitive PTP but not the DCD [93,104]. From this study we concluded that a TRP channel could be responsible for the Ca2+ influx part of DCD. In general, the two inhibitors that we used do not distinguish among individual members of the TRP family, but for reasons explained below, it is tempting to speculate that it is the TRPM7. Unfortunately, we could not achieve silencing of TRPM7 expression in our cultures with short interfering RNA (siRNA); primary neurons are notoriously vulnerable to transfection techniques, as opposed to the ease and the high efficiency of the procedure in cell lines. Hopefully, the development of novel approaches such as the conjugation of siRNA to penetratins [178,179] will assist transfection protocols and allow research on primary neuronal cultures to benefit from the tremendous potential of siRNA.
The connection of TRPM7 to DCD may lie in the observation that this channel is activated by ROS and RNS [159]. For a long time, ROS were considered to be responsible for DCD [180]; however, in a recent study it was deduced that the increased ROS production is a consequence, rather than a cause of DCD [181]. In the latter study the authors also demonstrated that the increase in superoxide radical formation is predominantly associated with extramitochondrial phospholipase A(2) (PLA2) activation, and it does not emanate from mitochondria. That may be in contrast with previous reports claiming that ROS are the inducers of DCD. However over the years concerns have arisen as for the reliability of ROS-detecting dyes, given that some are affected by confounding parameters such as mitochondrial membrane potential (see discussion in [181]). The development of new dyes described recently will no doubt contribute to the clarification of these matters [182].
In light of the recent observations though, one could argue that TRPM7 is not the Ca2+ influx pathway of DCD, as the increase in superoxide radical appears after the secondary [Ca2+]c rise. However, the exact species activating TRPM7 is not known, and the extent of ROS production necessary to activate the channel maybe less than the detection level of the probes used. In addition, ROS/RNS could be just one of the many activators of the channel [183], while others that might play a significant role could be also mobilized upon prolonged glutamate exposure. We have found that by elevating intracellular [Mg2+]i DCD is abolished in cultured cortical neurons [93], and it is known that TRPM7 receives strong negative feedback by intracellular Mg2+[170]. In addition, TRPM7 currents induced by oxygen–glucose deprivation promote further ROS production [159], and this could partially explain the results of Vesce and colleagues, detecting an increase in superoxide formation after the delayed secondary [Ca2+]c rise [181]. In our opinion, TRPM7 is one of the best possible candidates for the Ca2+ influx part of DCD; other good candidates are TRPM2 (see below) and the calcium-permeable acid-sensing ion channel [184] (not reviewed here).
Nonselective cationic channels and the ’Ca2+ paradox’
In spite of the widely accepted role of [Ca2+]c deregulation in the manifestation of neurodegeneration, exactly how Ca2+ ions mediate neural cell death is less clear [185]. One of the most important unresolved issues is the mechanism by which [Ca2+]c increases to excessively high levels in neurons following periods of intense neuronal activation. Reaching further from the possibility of the involvement of TRP channels in the delayed calcium deregulation, these proteins could participate in an additional overlooked pathway of Ca2+ influx that may pertain during ischemia/reperfusion or other type of pathology. Large [Ca2+]c increases are known to be triggered by reintroduction of ‘normal’ Ca2+ concentrations to the extracellular milieu after the tissue has experienced a [Ca2+]e-free challenge, or at least a severe reduction in extracellular calcium concentration, termed ‘Ca2+ paradox’. The free extracellular calcium concentration falls dramatically in several brain disease states: (a) during or after ischemia (0.1–0.28 mm[186–189]); (b) traumatic brain injury (0.1 mm[190]); (c) severe hypoglycemia (0.12 mm[191]); and (d) spreading depression (0.06–0.08 mm[192]). Reduction of extracellular Ca2+ is mostly due to robust influx of the cation to the intracellular milieu, although the appearance of lactate in the interstitium during ischemia, with the ability to chelate divalent ions significantly, also plays a role [193,194].
The Ca2+ paradox
Paradoxical Ca2+ increases were originally described in isolated heart preparations [195] and subsequently shown to be associated with tissue damage in this and other organs, including the kidney and skeletal muscle [196,197], but not in others, i.e. liver [198]. Interestingly, the possibility that paradoxical Ca2+ influx contributes to neuronal degeneration was put forward almost 20 years ago [199], but the vast majority of subsequent work on [Ca2+]c elevation during excitotoxicity has since concentrated on other Ca2+ entry routes, including glutamate receptors and voltage-gated Ca2+ channels. Unfortunately, this emphasis has not resulted in any clinically useful intervention to limit the neuronal damage following ischemia/reperfusion or other brain injury. Inescapably, within a context of ischemia/reperfusion in which a Ca2+ paradox is encompassed [200], concomitant adverse conditions, e.g. oxygen–glucose deprivation, associated ROS production and many more − reviewed in [201] − contribute to irreversible tissue damage. Nevertheless, the paradoxical Ca2+ rise per se remains a poorly understood phenomenon. What is known though, is that abolition of in situ mitochondrial respiration and oxidative phosphorylation protects against the Ca2+ paradox [202]. The reasons behind this unexpected finding are not yet understood. A number of theories were put forward, including the deleterious effect of overloading mitochondria with Ca2+ that can only happen in respiring mitochondria.
Possible mechanisms underlying neuronal paradoxical Ca2+-increases
While multiple mechanisms could contribute to paradoxical Ca2+ increases, the most current interest is the activation of novel nonselective cation channels. It is known that reduction of [Ca2+]e activates nonselective cation currents in hippocampal neurons [203] and neocortical nerve terminals [204] termed csNSC and NSC, respectively, as well as in thalamic neurons [205], vagal afferent nerves [206] and ventricular myocytes [207]. Such currents may underlie paradoxical Ca2+ increases activated by transient [Ca2+]e removal. We have also observed the appearance of a nonselective, noninactivating cation conductance upon reducing extracellular Ca2+ and Mg2+ in cultures of cortical neurons, as well as in cortical and hippocampal neurons in brain slices from adult mice, raising the possibility that such currents are readily available in these cells (C. Chinopoulos, unpublished data). Furthermore, we have recently reported that cultured cortical neurons exhibit paradoxical Ca2+ entry [93] and it is conceivable that the [Ca2+]c rise is a result of the ‘tails’ of these currents. Alternative mechanisms for paradoxical Ca2+ rise lie in a diversity of molecular complexities: lowering [Ca2+]e reduces the shielding of negatively charged groups located at the membrane surface affecting the voltage-dependent activation of various ion channels [163,208]. In addition, it is the biophysical property of many types of channels to conduct monovalents in a less controlled manner in the absence of divalent cations, such as the Icrac-conducting channel [209,210], voltage-gated Ca2+ channels [211–215], Na+ channels [216,217], K+ channels [218], other unidentified channels [203–207] and many members of the TRP family of channels (see below). In extreme cases, channel selectivity is lost when [Ca2+]e is reduced to ultra-low (<1 µm) concentrations [219].
Apart from this biophysical property of channels, a number of receptor-based mechanisms are modulated by [Ca2+]e: (a) the Ca2+-sensing receptor is activated by millimolar changes in [Ca2+]e, and is widely distributed in mammalian tissues including brain [220]; (b) hemi-gap channels in horizontal cells of the catfish retina are activated by [Ca2+]e decreases [221] and it is likely that gap junctional regulation could be strongly modified by [Ca2+]e in the central nervous system [222]; (c) metabotropic glutamate receptors 1, 3 and 5 [223] are activated by physiological [Ca2+]e fluctuations in the synaptic cleft [224]; and (d) the Gamma-aminobutyric acid (B) GABABreceptor also possesses Ca2+ sensing properties, potentiating GABA responses upon increase of [Ca2+]e[225]. It is not yet known whether these additional Ca2+-sensing mechanisms may act alone or in concert with nonselective Ca2+ channels in producing significant excitotoxic Ca2+ increases following ischemic insults.
TRP channels as candidates for paradoxical Ca2+-increases
TRP channels are widely expressed in mammalian tissues, especially in neurons of the central nervous system [226]. With a few notable exceptions, the physiological roles of TRP channels in neurons remain largely unknown [226–231]. Diverse neuropathological conditions were also found to implicate TRP family members: (a) mucolipidosis type IV [232] involving a channel from the distant polycystin branch (TRPP); (b) TRPV4 in neuropathic pain [233], and – as discussed above − (c) TRPM7 in neuronal death caused by oxygen–glucose deprivation [159]; the latter study also proposed the possibility of TRPM2 involvement, a view supported by more recent observations on oxidative stress-induced cell death [234]. Furthermore, ROS were specifically shown to trigger the opening of TRPC3 [235], TRPM2 [236–238] and TRPM7 [159]. In preliminary experiments, we have observed that the presence of ROS abolishes [Ca2+]c decay during the paradoxical Ca2+ rise and converts it to a progressive [Ca2+]c rise (C. Chinopoulos, unpublished data).
Of particular interest however, are the observations that a number of TRP channels are activated by a decrease in [Ca2+]e, raising the possibility that they could contribute to paradoxical Ca2+ increases. Recent descriptions have included the Drosophila TRP channel [239], TRPC1 and TRPC3 [240], TRPC6 [241], TRPC7 [242,243], and TRPM7 [159].
Mitochondrial permeabilization and a possible link to TRP channel activation
Among the known activators of some members of the TRP family, NAD+ and its catabolite ADP-ribose (ADPR) were described to activate TRPM2 [244–247], in addition to the fact that the channel is stimulated by ROS/RNS [236,238,246]. Furthermore, it was demonstrated that the major source of free ADPR mediating the activation of TRPM2 in cultured cells were the mitochondria [248]. One could link these observations to the fact that opening of the PTP causes the release of mitochondrial NAD+ followed by its hydrolysis by an extramitochondrial NAD+ glycohydrolase to ADPR [103,249]. It is tempting to speculate that this ADPR in conjunction with ROS produced upon loss of mitochondrial integrity, activates the nonselective TRPM2 allowing a large Ca2+ and Na+ load to enter the cytosol. Since both high [Ca2+]c and ROS promote mitochondrial pore formation, it seems that the order of appearance of a pore or TRPM2 activation is trivial; what is probably more important is that activation of the one can lead to activation of the other, completing a vicious cycle. Intriguingly, silencing the expression of TRPM7 with siRNA, led to an accompanying decrease in TRPM2 expression. This suggests that the two transcripts might be coordinately regulated, raising the possibility that a fraction of the oxygen–glucose deprivation-induced current recorded earlier [159] is mediated by TRPM2 or TRPM7 heteromultimers, a structural arrangement commonly occurring among TRP channels [250,251]. Further implications of TRP channels in relation to the overall metabolic state of the cell in hypoxia have been reviewed elsewhere [252].
Trp channels and ionic homeostasis
In view of the fact that most TRP channels are nonselective, in addition to allowing Ca2+ ions to enter the cytosol they also permit Na+ influx and K+ efflux [226,253,254]. The ominous effects of an elevated [Na+]i are mostly associated with cell swelling and activation of the Na+/Ca2+ exchanger causing Ca2+ influx. However, it is possible that the effect of an increased [Na+]i may be directly on mitochondria as recently demonstrated, diminishing the half-life of mitochondrially encoded mRNA, without involving Ca2+[255,256]. In addition it was recently shown that in mature hippocampal slices, NAD(P)H transients during postsynaptic neuronal activation are not mediated by Ca2+, but rather reflect alterations in [Na+]i. That may explain our previous results in isolated nerve terminals showing that in the presence of an oxidative stress a concomitant elevation in [Na+]i acts deleteriously on in situ mitochondria [257]. The effect of K+ loss from the cytoplasm is commonly ignored; however, it was shown that it can promote neuronal apoptosis [258–260]. To what extent − if any − the activation of TRP channels is associated with alterations of Na+ and K+ homeostasis in neurodegeneration, is currently unknown. Nevertheless, the fact that these proteins are intensely expressed in the central nervous system [251,254,261] and their ever-increasing roles in physiology and pathology being discovered [253,262], identify them as excellent novel targets amenable to pharmacological manipulation [254,263,264].
References
- 1 Reynolds IJ & Hastings TG (1995) Glutamate induces the production of reactive oxygen species in cultured forebrain neurons following NMDA receptor activation. J Neurosci 15, 3318–3327.
- 2 Dugan LL, Sensi SL, Canzoniero LM, Handran SD, Rothman SM, Lin TS, Goldberg MP & Choi DW (1995) Mitochondrial production of reactive oxygen species in cortical neurons following exposure to N-methyl-D-aspartate. J Neurosci 15, 6377–6388.
- 3 Luetjens CM, Bui NT, Sengpiel B, Munstermann G, Poppe M, Krohn AJ, Bauerbach E, Krieglstein J & Prehn JH (2000) Delayed mitochondrial dysfunction in excitotoxic neuron death: cytochrome c release and a secondary increase in superoxide production. J Neurosci 20, 5715–5723.
- 4 Carriedo SG, Sensi SL, Yin HZ & Weiss JH (2000) AMPA exposures induce mitochondrial Ca2+ overload and ROS generation in spinal motor neurons in vitro. J Neurosci 20, 240–250.
- 5 Sengpiel B, Preis E, Krieglstein J & Prehn JH (1998) NMDA-induced superoxide production and neurotoxicity in cultured rat hippocampal neurons: role of mitochondria. Eur J Neurosci 10, 1903–1910.
- 6 Gunasekar PG, Kanthasamy AG, Borowitz JL & Isom GE (1995) NMDA receptor activation produces concurrent generation of nitric oxide and reactive oxygen species: implication for cell death. J Neurochem 65, 2016–2021.
- 7 Dykens JA (1994) Isolated cerebral and cerebellar mitochondria produce free radicals when exposed to elevated CA2+ and Na+: implications for neurodegeneration. J Neurochem 63, 584–591.
- 8 Kowaltowski AJ, Netto LE & Vercesi AE (1998) The thiol-specific antioxidant enzyme prevents mitochondrial permeability transition. Evidence for the participation of reactive oxygen species in this mechanism. J Biol Chem 273, 12766–12769.
- 9 Kowaltowski AJ, Castilho RF & Vercesi AE (1995) Ca2+-induced mitochondrial membrane permeabilization: role of coenzyme Q redox state. Am J Physiol 269, C141–C147.
- 10 Kowaltowski AJ, Castilho RF & Vercesi AE (1996) Opening of the mitochondrial permeability transition pore by uncoupling or inorganic phosphate in the presence of Ca2+ is dependent on mitochondrial-generated reactive oxygen species. FEBS Lett 378, 150–152.
- 11 Kowaltowski AJ, Naia, -d, a-Silva ES, Castilho RF & Vercesi AE (1998) Ca2+-stimulated mitochondrial reactive oxygen species generation and permeability transition are inhibited by dibucaine or Mg2+. Arch Biochem Biophys 359, 77–81.
- 12 Maciel EN, Vercesi AE & Castilho RF (2001) Oxidative stress in Ca2+ -induced membrane permeability transition in brain mitochondria. J Neurochem 79, 1237–1245.
- 13 Korshunov SS, Skulachev VP & Starkov AA (1997) High protonic potential actuates a mechanism of production of reactive oxygen species in mitochondria. FEBS Lett 416, 15–18.
- 14 Starkov AA, Polster BM & Fiskum G (2002) Regulation of hydrogen peroxide production by brain mitochondria by calcium and Bax. J Neurochem 83, 220–228.
- 15 Starkov AA & Fiskum G (2003) Regulation of brain mitochondrial H2O2 production by membrane potential and NAD(P) H redox state. J Neurochem 86, 1101–1107.
- 16 Starkov AA, Chinopoulos C & Fiskum G (2004) Mitochondrial calcium and oxidative stress as mediators of ischemic brain injury. Cell Calcium 36, 257–264.
- 17 Andreyev AY, Kushnareva YE & Starkov AA (2005) Mitochondrial metabolism of reactive oxygen species. Biochemistry (Moscow) 70, 200–214.
- 18 Votyakova TV & Reynolds IJ (2001) DeltaPsi (m)-dependent and -independent production of reactive oxygen species by rat brain mitochondria. J Neurochem 79, 266–277.
- 19 Sipos I, Tretter L & Adam-Vizi V (2003) The production of reactive oxygen species in intact isolated nerve terminals is independent of the mitochondrial membrane potential. Neurochem Res 28, 1575–1581.
- 20 Adam-Vizi V (2005) Production of reactive oxygen species in brain mitochondria: contribution by electron transport chain and non-electron transport chain sources. Antioxid Redox Signal 7, 1140–1149.
- 21 Anderson MF & Sims NR (2002) The effects of focal ischemia and reperfusion on the glutathione content of mitochondria from rat brain subregions. J Neurochem 81, 541–549.
- 22 Cai J & Jones DP (1998) Superoxide in apoptosis. Mitochondrial generation triggered by cytochrome c loss. J Biol Chem 273, 11401–11404.
- 23 Tretter L & Adam-Vizi V (2004) Generation of reactive oxygen species in the reaction catalyzed by alpha-ketoglutarate dehydrogenase. J Neurosci 24, 7771–7778.
- 24 Starkov AA, Fiskum G, Chinopoulos C, Lorenzo BJ, Browne SE, Patel MS & Beal MF (2004) Mitochondrial alpha-ketoglutarate dehydrogenase complex generates reactive oxygen species. J Neurosci 24, 7779–7788.
- 25 Jensen PK (1966) Antimycin-insensitive oxidation of succinate and reduced nicotinamide-adenine dinucleotide in electron-transport particles. I. pH dependency and hydrogen peroxide formation. Biochim Biophys Acta 122, 157–166.
- 26 Loschen G, Flohe L & Chance B (1971) Respiratory chain linked H2O2 production in pigeon heart mitochondria. FEBS Lett 18, 261–264.
- 27 Boveris A & Chance B (1973) The mitochondrial generation of hydrogen peroxide. General properties and effect of hyperbaric oxygen. Biochem J 134, 707–716.
- 28 Turrens JF (2003) Mitochondrial formation of reactive oxygen species. J Physiol 552, 335–344.
- 29 Lenaz G (2001) The mitochondrial production of reactive oxygen species: mechanisms and implications in human pathology. IUBMB Life 52, 159–164.
- 30 Barja G (1999) Mitochondrial oxygen radical generation and leak: sites of production in states 4 and 3, organ specificity, and relation to aging and longevity. J Bioenerg Biomembr 31, 347–366.
- 31 Herrero A & Barja G (2000) Localization of the site of oxygen radical generation inside the Complex I of heart and nonsynaptic brain mammalian mitochondria. J Bioenerg Biomembr 32, 609–615.
- 32 Turrens JF & Boveris A (1980) Generation of superoxide anion by the NADH dehydrogenase of bovine heart mitochondria. Biochem J 191, 421–427.
- 33 Loschen G, Azzi A, Richter C & Flohe L (1974) Superoxide radicals as precursors of mitochondrial hydrogen peroxide. FEBS Lett 42, 68–72.
- 34 Dionisi O, Galeotti T, Terranova T & Azzi A (1975) Superoxide radicals and hydrogen peroxide formation in mitochondria from normal and neoplastic tissues. Biochim Biophys Acta 403, 292–300.
- 35 Boveris A, Cadenas E & Stoppani AO (1976) Role of ubiquinone in the mitochondrial generation of hydrogen peroxide. Biochem J 156, 435–444.
- 36 Turrens JF (1997) Superoxide production by the mitochondrial respiratory chain. Biosci Report 17, 3–8.
- 37 Murphy AN, Fiskum G & Beal MF (1999) Mitochondria in neurodegeneration: bioenergetic function in cell life and death. J Cereb Blood Flow Metab 19, 231–245.
- 38 Hinkle PC, Butow RA, Racker E & Chance B (1967) Partial resolution of the enzymes catalyzing oxidative phosphorylation. XV. Reverse electron transfer in the flavin-cytochrome beta region of the respiratory chain of beef heart submitochondrial particles. J Biol Chem 242, 5169–5173.
- 39 Sipos I, Tretter L & Adam-Vizi V (2003) Quantitative relationship between inhibition of respiratory complexes and formation of reactive oxygen species in isolated nerve terminals. J Neurochem 84, 112–118.
- 40 Bunik VI (2003) 2-Oxo acid dehydrogenase complexes in redox regulation. Eur J Biochem 270, 1036–1042.
- 41 Maas E & Bisswanger H (1990) Localization of the alpha-oxoacid dehydrogenase multienzyme complexes within the mitochondrion. FEBS Lett 277, 189–190.
- 42 Sumegi B & Srere PA (1984) Complex I binds several mitochondrial NAD-coupled dehydrogenases. J Biol Chem 259, 15040–15045.
- 43 Lyubarev AE & Kurganov BI (1989) Supramolecular organization of tricarboxylic acid cycle enzymes. Biosystems 22, 91–102.
- 44 Koike K, Hamada M, Tanaka N, Otsuka KI, Ogasahara K & Koike M (1974) Properties and subunit composition of the pig heart 2-oxoglutarate dehydrogenase. J Biol Chem 249, 3836–3842.
- 45 Patel MS & Roche TE (1990) Molecular biology and biochemistry of pyruvate dehydrogenase complexes. FASEB J 4, 3224–3233.
- 46 Reed LJ & Hackert ML (1990) Structure-function relationships in dihydrolipoamide acyltransferases. J Biol Chem 265, 8971–8974.
- 47 Kochi H, Seino H & Ono K (1986) Inhibition of glycine oxidation by pyruvate, alpha-ketoglutarate, and branched-chain alpha-keto acids in rat liver mitochondria: presence of interaction between the glycine cleavage system and alpha-keto acid dehydrogenase complexes. Arch Biochem Biophys 249, 263–272.
- 48 Huennekens FM, Basford RE & Gabrio BW (1955) An oxidase for reduced diphosphopyridine nucleotide. J Biol Chem 213, 951–967.
- 49 Gazaryan IG, Krasnikov BF, Ashby GA, Thorneley RN, Kristal BS & Brown AM (2002) Zinc is a potent inhibitor of thiol oxidoreductase activity and stimulates reactive oxygen species production by lipoamide dehydrogenase. J Biol Chem 277, 10064–10072.
- 50 Kunz WS & Gellerich FN (1993) Quantification of the content of fluorescent flavoproteins in mitochondria from liver, kidney cortex, skeletal muscle, and brain. Biochem Med Metab Biol 50, 103–110.
- 51 Kunz WS & Kunz W (1985) Contribution of different enzymes to flavoprotein fluorescence of isolated rat liver mitochondria. Biochim Biophys Acta 841, 237–246.
- 52 Bunik VI & Sievers C (2002) Inactivation of the 2-oxo acid dehydrogenase complexes upon generation of intrinsic radical species. Eur J Biochem 269, 5004–5015.
- 53 Hamada M, Koike K, Nakaula Y, Hiraoka T & Koike M (1975) A kinetic study of the alpha-keto acid dehydrogenase complexes from pig heart mitochondria. J Biochem (Tokyo) 77, 1047–1056.
- 54 Mcminn CL & Ottaway JH (1977) Studies on the mechanism and kinetics of the 2-oxoglutarate dehydrogenase system from pig heart. Biochem J 161, 569–581.
- 55 Wan B, Lanoue KF, Cheung JY & Scaduto RC Jr (1989) Regulation of citric acid cycle by calcium. J Biol Chem 264, 13430–13439.
- 56 Kiselevsky YV, Ostrovtsova SA & Strumilo SA (1990) Kinetic characterization of the pyruvate and oxoglutarate dehydrogenase complexes from human heart. Acta Biochim Pol 37, 135–139.
- 57 Tretter L & Adam-Vizi V (2000) Inhibition of Krebs cycle enzymes by hydrogen peroxide: a key role of [alpha]-ketoglutarate dehydrogenase in limiting NADH production under oxidative stress. J Neurosci 20, 8972–8979.
- 58 Kumar MJ, Nicholls DG & Andersen JK (2003) Oxidative a-ketoglutarate dehydrogenase inhibition via subtle elevations in monoamine oxidase B levels results in loss of spare respiratory capacity: Implications for Parkinson's disease. J Biol Chem 278, 46432–46439.
- 59 Chinopoulos C, Tretter L & Adam-Vizi V (1999) Depolarization of in situ mitochondria due to hydrogen peroxide-induced oxidative stress in nerve terminals: inhibition of alpha-ketoglutarate dehydrogenase. J Neurochem 73, 220–228.
- 60 Chinopoulos C, Tretter L & Adam-Vizi V (2000) Reversible depolarization of in situ mitochondria by oxidative stress parallels a decrease in NAD(P)H level in nerve terminals. Neurochem Int 36, 483–488.
- 61 Chinopoulos C & Adam-Vizi V (1999) Depolarization of in situ mitochondria by hydrogen peroxide in nerve terminals. Ann NY Acad Sci 893, 269–272.
- 62 Beal MF (1996) Mitochondria, free radicals, and neurodegeneration. Curr Opin Neurobiol 6, 661–666.
- 63 Gibson GE, Blass JP, Beal MF & Bunik V (2005) The alpha-ketoglutarate-dehydrogenase complex: a mediator between mitochondria and oxidative stress in neurodegeneration. Mol Neurobiol 31, 43–64.
- 64 Gibson GE, Sheu KF, Blass JP, Baker A, Carlson KC, Harding B & Perrino P (1988) Reduced activities of thiamine-dependent enzymes in the brains and peripheral tissues of patients with Alzheimer's disease. Arch Neurol 45, 836–840.
- 65 Butterworth RF & Besnard AM (1990) Thiamine-dependent enzyme changes in temporal cortex of patients with Alzheimer's disease. Metab Brain Dis 5, 179–184.
- 66 Mastrogiacomo F, Bergeron C & Kish SJ (1993) Brain alpha-ketoglutarate dehydrogenase complex activity in Alzheimer's disease. J Neurochem 61, 2007–2014.
- 67 Terwel D, Bothmer J, Wolf E, Meng F & Jolles J (1998) Affected enzyme activities in Alzheimer's disease are sensitive to antemortem hypoxia. J Neurol Sci 161, 47–56.
- 68 Mizuno Y, Matuda S, Yoshino H, Mori H, Hattori N & Ikebe S (1994) An immunohistochemical study on alpha-ketoglutarate dehydrogenase complex in Parkinson's disease. Ann Neurol 35, 204–210.
- 69 Mizuno Y, Ikebe S, Hattori N, Nakagawa-Hattori Y, Mochizuki H, Tanaka M & Ozawa T (1995) Role of mitochondria in the etiology and pathogenesis of Parkinson's disease. Biochim Biophys Acta 1271, 265–274.
- 70 Jimenez-Jimenez FJ, Molina JA, Hernanz A, Fernandez-Vivancos EBF, Barcenilla B, Gomez-Escalonilla C, Zurdo M, Berbel A & Villanueva C (1999) Cerebrospinal fluid levels of thiamine in patients with Parkinson's disease. Neurosci Lett 271, 33–36.
- 71 Gibson GE, Kingsbury AE, Xu H, Lindsay JG, Daniel S, Foster OJ, Lees AJ & Blass JP (2003) Deficits in a tricarboxylic acid cycle enzyme in brains from patients with Parkinson's disease. Neurochem Int 43, 129–135.
- 72 Albers DS, Augood SJ, Park LC, Browne SE, Martin DM, Adamson J, Hutton M, Standaert DG, Vonsattel JP, Gibson GE & Beal MF (2000) Frontal lobe dysfunction in progressive supranuclear palsy: evidence for oxidative stress and mitochondrial impairment. J Neurochem 74, 878–881.
- 73 Park LC, Albers DS, Xu H, Lindsay JG, Beal MF & Gibson GE (2001) Mitochondrial impairment in the cerebellum of the patients with progressive supranuclear palsy. J Neurosci Res 66, 1028–1034.
- 74 Butterworth RF, Kril JJ & Harper CG (1993) Thiamine-dependent enzyme changes in the brains of alcoholics: relationship to the Wernicke–Korsakoff syndrome. Alcohol Clin Exp Res 17, 1084–1088.
- 75 Mastrogiacomo F, Lamarche J, Dozic S, Lindsay G, Bettendorff L, Robitaille Y, Schut L & Kish SJ (1996) Immunoreactive levels of alpha-ketoglutarate dehydrogenase subunits in Friedreich's ataxia and spinocerebellar ataxia type 1. Neurodegeneration 5, 27–33.
- 76 Gibson GE, Zhang H, Sheu KF, Bogdanovich N, Lindsay JG, Lannfelt L, Vestling M & Cowburn RF (1998) Alpha-ketoglutarate dehydrogenase in Alzheimer brains bearing the APP670/671 mutation. Ann Neurol 44, 676–681.
- 77 Klivenyi P, Starkov AA, Calingasan NY, Gardian G, Browne SE, Yang L, Bubber P, Gibson GE, Patel MS & Beal MF (2004) Mice deficient in dihydrolipoamide dehydrogenase show increased vulnerability to MPTP, malonate and 3-nitropropionic acid neurotoxicity. J Neurochem 88, 1352–1360.
- 78 Sheu KF, Calingasan NY, Lindsay JG & Gibson GE (1998) Immunochemical characterization of the deficiency of the alpha-ketoglutarate dehydrogenase complex in thiamine-deficient rat brain. J Neurochem 70, 1143–1150.
- 79 Shi Q, Chen HL, Xu H & Gibson GE (2005) Reduction in the E2k subunit of the alpha-ketoglutarate dehydrogenase complex has effects independent of complex activity. J Biol Chem 280, 10888–10896.
- 80 Sun YJ, Chou CC, Chen WS, Wu RT, Meng M & Hsiao CD (1999) The crystal structure of a multifunctional protein: phosphoglucose isomerase/autocrine motility factor/neuroleukin. Proc Natl Acad Sci USA 96, 5412–5417.
- 81 Kennedy MC, Mende-Mueller L, Blondin GA & Beinert H (1992) Purification and characterization of cytosolic aconitase from beef liver and its relationship to the iron-responsive element binding protein. Proc Natl Acad Sci USA 89, 11730–11734.
- 82 Elzinga SD, van Bednarz ALOK, Dekker PJ & Grivell LA (1993) Yeast mitochondrial NAD(+)-dependent isocitrate dehydrogenase is an RNA-binding protein. Nucl Acids Res 21, 5328–5331.
- 83 Jeffery CJ (1999) Moonlighting proteins. Trends Biochem Sci 24, 8–11.
- 84 Kaufman BA, Newman SM, Hallberg RL, Slaughter CA, Perlman PS & Butow RA (2000) In organello formaldehyde crosslinking of proteins to mtDNA: identification of bifunctional proteins. Proc Natl Acad Sci USA 97, 7772–7777.
- 85 Calingasan NY, Baker H, Sheu KF & Gibson GE (1994) Distribution of the alpha-ketoglutarate dehydrogenase complex in rat brain. J Comp Neurol 346, 461–479.
- 86 Park LC, Calingasan NY, Sheu KF & Gibson GE (2000) Quantitative alpha-ketoglutarate dehydrogenase activity staining in brain sections and in cultured cells. Anal Biochem 277, 86–93.
- 87 Huang HM, Ou HC, Xu H, Chen HL, Fowler C & Gibson GE (2003) Inhibition of alpha-ketoglutarate dehydrogenase complex promotes cytochrome c release from mitochondria, caspase-3 activation, and necrotic cell death. J Neurosci Res 74, 309–317.
- 88 Zoratti M & Szabo I (1995) The mitochondrial permeability transition. Biochim Biophys Acta 1241, 139–176.
- 89 Bernardi P (1999) Mitochondrial transport of cations: channels, exchangers, and permeability transition. Physiol Rev 79, 1127–1155.
- 90 Duchen MR (2000) Mitochondria and Ca2+ in cell physiology and pathophysiology. Cell Calcium 28, 339–348.
- 91 Nicholls DG & Budd SL (2000) Mitochondria and neuronal survival. Physiol Rev 80, 315–360.
- 92 Vergun O, Keelan J, Khodorov BI & Duchen MR (1999) Glutamate-induced mitochondrial depolarisation and perturbation of calcium homeostasis in cultured rat hippocampal neurones. J Physiol 519 Part 2, 451–466.
- 93 Chinopoulos C, Gerencser AA, Doczi J, Fiskum G & Adam-Vizi V (2004) Inhibition of glutamate-induced delayed calcium deregulation by 2-APB and La3+ in cultured cortical neurones. J Neurochem 91, 471–483.
- 94 Castilho RF, Hansson O, Ward MW, Budd SL & Nicholls DG (1998) Mitochondrial control of acute glutamate excitotoxicity in cultured cerebellar granule cells. J Neurosci 18, 10277–10286.
- 95 Dubinsky JM & Levi Y (1998) Calcium-induced activation of the mitochondrial permeability transition in hippocampal neurons. J Neurosci Res 53, 728–741.
- 96 Hoyt KR, Mclaughlin BA, Higgins DS Jr & Reynolds IJ (2000) Inhibition of glutamate-induced mitochondrial depolarization by tamoxifen in cultured neurons. J Pharmacol Exp Ther 293, 480–486.
- 97 Scanlon JM & Reynolds IJ (1998) Effects of oxidants and glutamate receptor activation on mitochondrial membrane potential in rat forebrain neurons. J Neurochem 71, 2392–2400.
- 98 Hoyt KR, Sharma TA & Reynolds IJ (1997) Trifluoperazine and dibucaine-induced inhibition of glutamate-induced mitochondrial depolarization in rat cultured forebrain neurones. Br J Pharmacol 122, 803–808.
- 99 Fournier N, Ducet G & Crevat A (1987) Action of cyclosporine on mitochondrial calcium fluxes. J Bioenerg Biomembr 19, 297–303.
- 100 Johans M, Milanesi E, Franck M, Johans C, Liobikas J, Panagiotaki M, Greci L, Principato G, Kinnunen PK, Bernardi P, Costantini P & Eriksson O (2005) Modification of permeability transition pore arginine(s) by phenylglyoxal derivatives in isolated mitochondria and mammalian cells. Structure-function relationship of arginine ligands. J Biol Chem 280, 12130–12136.
- 101 Soriano ME, Nicolosi L & Bernardi P (2004) Desensitization of the permeability transition pore by cyclosporin a prevents activation of the mitochondrial apoptotic pathway and liver damage by tumor necrosis factor-alpha. J Biol Chem 279, 36803–36808.
- 102 Petronilli V, Penzo D, Scorrano L, Bernardi P & Di Lisa F (2001) The mitochondrial permeability transition, release of cytochrome c and cell death. Correlation with the duration of pore openings in situ. J Biol Chem 276, 12030–12034.
- 103 Di Lisa F, Menabo R, Canton M, Barile M & Bernardi P (2001) Opening of the mitochondrial permeability transition pore causes depletion of mitochondrial and cytosolic NAD+ and is a causative event in the death of myocytes in postischemic reperfusion of the heart. J Biol Chem 276, 2571–2575.
- 104 Chinopoulos C, Starkov AA & Fiskum G (2003) Cyclosporin A-insensitive permeability transition in brain mitochondria: inhibition by 2-aminoethoxydiphenyl borate. J Biol Chem 278, 27382–27389.
- 105 Kristian T, Weatherby TM, Bates TE & Fiskum G (2002) Heterogeneity of the calcium-induced permeability transition in isolated non-synaptic brain mitochondria. J Neurochem 83, 1297–1308.
- 106 Chalmers S & Nicholls DG (2003) The relationship between free and total calcium concentrations in the matrix of liver and brain mitochondria. J Biol Chem 278, 19062–19070.
- 107 Ekholm A, Katsura K, Kristian T, Liu M, Folbergrova J & Siesjo BK (1993) Coupling of cellular energy state and ion homeostasis during recovery following brain ischemia. Brain Res 604, 185–191.
- 108 Haworth RA & Hunter DR (1980) Allosteric inhibition of the Ca2+-activated hydrophilic channel of the mitochondrial inner membrane by nucleotides. J Membr Biol 54, 231–236.
- 109 Peng TI, Jou MJ, Sheu SS & Greenamyre JT (1998) Visualization of NMDA receptor-induced mitochondrial calcium accumulation in striatal neurons. Exp Neurol 149, 1–12.
- 110 Rossi DJ, Oshima T & Attwell D (2000) Glutamate release in severe brain ischaemia is mainly by reversed uptake. Nature 403, 316–321.
- 111 Wang GJ & Thayer SA (2002) NMDA-induced calcium loads recycle across the mitochondrial inner membrane of hippocampal neurons in culture. J Neurophysiol 87, 740–749.
- 112 Isaev NK, Zorov DB, Stelmashook EV, Uzbekov RE, Kozhemyakin MB & Victorov IV (1996) Neurotoxic glutamate treatment of cultured cerebellar granule cells induces Ca2+-dependent collapse of mitochondrial membrane potential and ultrastructural alterations of mitochondria. FEBS Lett 392, 143–147.
- 113 Pivovarova NB, Nguyen HV, Winters CA, Brantner CA, Smith CL & Andrews SB (2004) Excitotoxic calcium overload in a subpopulation of mitochondria triggers delayed death in hippocampal neurons. J Neurosci 24, 5611–5622.
- 114 Dux E, Mies G, Hossmann KA & Siklos L (1987) Calcium in the mitochondria following brief ischemia of gerbil brain. Neurosci Lett 78, 295–300.
- 115 Hallak H, Ramadan B & Rubin R (2001) Tyrosine phosphorylation of insulin receptor substrate-1 (IRS-1) by oxidant stress in cerebellar granule neurons: modulation by N-methyl-D-aspartate through calcineurin activity. J Neurochem 77, 63–70.
- 116 Dawson TM, Steiner JP, Dawson VL, Dinerman JL, Uhl GR & Snyder SH (1993) Immunosuppressant FK506 enhances phosphorylation of nitric oxide synthase and protects against glutamate neurotoxicity. Proc Natl Acad Sci USA 90, 9808–9812.
- 117 Alano CC, Beutner G, Dirksen RT, Gross RA & Sheu SS (2002) Mitochondrial permeability transition and calcium dynamics in striatal neurons upon intense NMDA receptor activation. J Neurochem 80, 531–538.
- 118 Khaspekov L, Friberg H, Halestrap A, Viktorov I & Wieloch T (1999) Cyclosporin A and its nonimmunosuppressive analogue N-Me-Val-4-cyclosporin A mitigate glucose/oxygen deprivation-induced damage to rat cultured hippocampal neurons. Eur J Neurosci 11, 3194–3198.
- 119 Liu X, Kim CN, Yang J, Jemmerson R & Wang X (1996) Induction of apoptotic program in cell-free extracts: requirement for dATP and cytochrome c. Cell 86, 147–157.
- 120 Hengartner MO (2000) The biochemistry of apoptosis. Nature 407, 770–776.
- 121 Kim JS, He L & Lemasters JJ (2003) Mitochondrial permeability transition: a common pathway to necrosis and apoptosis. Biochem Biophys Res Commun 304, 463–470.
- 122 Lemasters JJ, Nieminen AL, Qian T, Trost LC, Elmore SP, Nishimura Y, Crowe RA, Cascio WE, Bradham CA, Brenner DA & Herman B (1998) The mitochondrial permeability transition in cell death: a common mechanism in necrosis, apoptosis and autophagy. Biochim Biophys Acta 1366, 177–196.
- 123 Polster BM & Fiskum G (2004) Mitochondrial mechanisms of neural cell apoptosis. J Neurochem 90, 1281–1289.
- 124 Kroemer G (2003) Mitochondrial control of apoptosis: an introduction. Biochem Biophys Res Commun 304, 433–435.
- 125 Van Loo G, Saelens X, Van Gurp M, Macfarlane M, Martin SJ & Vandenabeele P (2002) The role of mitochondrial factors in apoptosis: a Russian roulette with more than one bullet. Cell Death Differ 9, 1031–1042.
- 126 Zamzami N & Kroemer G (2001) The mitochondrion in apoptosis: how Pandora's box opens. Nat Rev Mol Cell Biol 2, 67–71.
- 127 Desagher S & Martinou JC (2000) Mitochondria as the central control point of apoptosis. Trends Cell Biol 10, 369–377.
- 128 Green DR & Reed JC (1998) Mitochondria and apoptosis. Science 281, 1309–1312.
- 129 Susin SA, Zamzami N & Kroemer G (1998) Mitochondria as regulators of apoptosis: doubt no more. Biochim Biophys Acta 1366, 151–165.
- 130 Green D & Kroemer G (1998) The central executioners of apoptosis: caspases or mitochondria? Trends Cell Biol 8, 267–271.
- 131 Kroemer G, Dallaporta B & Resche-Rigon M (1998) The mitochondrial death/life regulator in apoptosis and necrosis. Annu Rev Physiol 60, 619–642.
- 132 Schinzel AC, Takeuchi O, Huang Z, Fisher JK, Zhou Z, Rubens J, Hetz C, Danial NN, Moskowitz MA & Korsmeyer SJ (2005) Cyclophilin D is a component of mitochondrial permeability transition and mediates neuronal cell death after focal cerebral ischemia. Proc Natl Acad Sci USA 102, 12005–12010.
- 133 Baines CP, Kaiser RA, Purcell NH, Blair NS, Osinska H, Hambleton MA, Brunskill EW, Sayen MR, Gottlieb RA, Dorn GW, Robbins J & Molkentin JD (2005) Loss of cyclophilin D reveals a critical role for mitochondrial permeability transition in cell death. Nature 434, 658–662.
- 134 Nakagawa T, Shimizu S, Watanabe T, Yamaguchi O, Otsu K, Yamagata H, Inohara H, Kubo T & Tsujimoto Y (2005) Cyclophilin D-dependent mitochondrial permeability transition regulates some necrotic but not apoptotic cell death. Nature 434, 652–658.
- 135 Woodfield K, Ruck A, Brdiczka D & Halestrap AP (1998) Direct demonstration of a specific interaction between cyclophilin-D and the adenine nucleotide translocase confirms their role in the mitochondrial permeability transition. Biochem J 336, 287–290.
- 136 Crompton M, Virji S & Ward JM (1998) Cyclophilin-D binds strongly to complexes of the voltage-dependent anion channel and the adenine nucleotide translocase to form the permeability transition pore. Eur J Biochem 258, 729–735.
- 137 Basso E, Fante L, Fowlkes J, Petronilli V, Forte MA & Bernardi P (2005) Properties of the permeability transition pore in mitochondria devoid of Cyclophilin D. J Biol Chem 280, 18558–18561.
- 138 Nicholls D & Akerman K (1982) Mitochondrial calcium transport. Biochim Biophys Acta 683, 57–88.
- 139 Nicholls DG (1978) The regulation of extramitochondrial free calcium ion concentration by rat liver mitochondria. Biochem J 176, 463–474.
- 140 Pitter JG, Maechler P, Wollheim CB & Spat A (2002) Mitochondria respond to Ca2+ already in the submicromolar range: correlation with redox state. Cell Calcium 31, 97–104.
- 141 Chinopoulos C, Starkov AA, Grigoriev S, Dejean LM, Kinnally KW, Liu X, Ambudkar IS & Fiskum G (2005) Diacylglycerols activate mitochondrial cationic channel(s) and release sequestered Ca2+. J Bioenerg Biomembr 37, 237–247.
- 142 Mikoshiba K & Hattori M (2000) IP3 receptor-operated calcium entry. Sci STKE 2000, E1.
- 143 Bianchi K, Rimessi A, Prandini A, Szabadkai G & Rizzuto R (2004) Calcium and mitochondria: mechanisms and functions of a troubled relationship. Biochim Biophys Acta 1742, 119–131.
- 144 Nicholls DG & Budd SL (1998) Mitochondria and neuronal glutamate excitotoxicity. Biochim Biophys Acta 1366, 97–112.
- 145 Tymianski M, Charlton MP, Carlen PL & Tator CH (1993) Secondary Ca2+ overload indicates early neuronal injury which precedes staining with viability indicators. Brain Res 607, 319–323.
- 146 Tymianski M, Charlton MP, Carlen PL & Tator CH (1993) Source specificity of early calcium neurotoxicity in cultured embryonic spinal neurons. J Neurosci 13, 2085–2104.
- 147 Witt MR, Dekermendjian K, Frandsen A, Schousboe A & Nielsen M (1994) Complex correlation between excitatory amino acid-induced increase in the intracellular Ca2+ concentration and subsequent loss of neuronal function in individual neocortical neurons in culture. Proc Natl Acad Sci USA 91, 12303–12307.
- 148 Limbrick DD Jr, Churn SB, Sombati S & Delorenzo RJ (1995) Inability to restore resting intracellular calcium levels as an early indicator of delayed neuronal cell death. Brain Res 690, 145–156.
- 149 Manev H, Favaron M, Guidotti A & Costa E (1989) Delayed increase of Ca2+ influx elicited by glutamate: role in neuronal death. Mol Pharmacol 36, 106–112.
- 150 Randall RD & Thayer SA (1992) Glutamate-induced calcium transient triggers delayed calcium overload and neurotoxicity in rat hippocampal neurons. J Neurosci 12, 1882–1895.
- 151 Connor JA, Wadman WJ, Hockberger PE & Wong RK (1988) Sustained dendritic gradients of Ca2+ induced by excitatory amino acids in CA1 hippocampal neurons. Science 240, 649–653.
- 152 Budd SL & Nicholls DG (1996) Mitochondria, calcium regulation, and acute glutamate excitotoxicity in cultured cerebellar granule cells. J Neurochem 67, 2282–2291.
- 153 Rajdev S & Reynolds IJ (1994) Glutamate-induced intracellular calcium changes and neurotoxicity in cortical neurons in vitro: effect of chemical ischemia. Neuroscience 62, 667–679.
- 154 Hyrc K, Handran SD, Rothman SM & Goldberg MP (1997) Ionized intracellular calcium concentration predicts excitotoxic neuronal death: observations with low-affinity fluorescent calcium indicators. J Neurosci 17, 6669–6677.
- 155 Hartley DM & Choi DW (1989) Delayed rescue of N-methyl-D-aspartate receptor-mediated neuronal injury in cortical culture. J Pharmacol Exp Ther 250, 752–758.
- 156 Chen QX, Perkins KL, Choi DW & Wong RK (1997) Secondary activation of a cation conductance is responsible for NMDA toxicity in acutely isolated hippocampal neurons. J Neurosci 17, 4032–4036.
- 157 Limbrick DD Jr, Pal S & Delorenzo RJ (2001) Hippocampal neurons exhibit both persistent Ca2+ influx and impairment of Ca2+ sequestration/extrusion mechanisms following excitotoxic glutamate exposure. Brain Res 894, 56–67.
- 158 Limbrick DD Jr, Sombati S & Delorenzo RJ (2003) Calcium influx constitutes the ionic basis for the maintenance of glutamate-induced extended neuronal depolarization associated with hippocampal neuronal death. Cell Calcium 33, 69–81.
- 159 Aarts M, Iihara K, Wei WL, Xiong ZG, Arundine M, Cerwinski W, Macdonald JF & Tymianski M (2003) A key role for TRPM7 channels in anoxic neuronal death. Cell 115, 863–877.
- 160 Khodorov B, Pinelis V, Golovina V, Fajuk D, Andreeva N, Uvarova T, Khaspekov L & Victorov I (1993) On the origin of a sustained increase in cytosolic Ca2+ concentration after a toxic glutamate treatment of the nerve cell culture. FEBS Lett 324, 271–273.
- 161 Bano D, Young KW, Guerin CJ, Lefeuvre R, Rothwell NJ, Naldini L, Rizzuto R, Carafoli E & Nicotera P (2005) Cleavage of the plasma membrane Na+/Ca2+ exchanger in excitotoxicity. Cell 120, 275–285.
- 162 Ward MW, Rego AC, Frenguelli BG & Nicholls DG (2000) Mitochondrial membrane potential and glutamate excitotoxicity in cultured cerebellar granule cells. J Neurosci 20, 7208–7219.
- 163 Hille B (2001) Ionic Channels of Excitable Membranes. Sinauer Associates, Inc, Sunderland, MA.
- 164 Kiedrowski L, Czyz A, Baranauskas G, Li XF & Lytton J (2004) Differential contribution of plasmalemmal Na/Ca exchange isoforms to sodium-dependent calcium influx and NMDA excitotoxicity in depolarized neurons. J Neurochem 90, 117–128.
- 165 Ikonomidou C & Turski L (2002) Why did NMDA receptor antagonists fail clinical trials for stroke and traumatic brain injury? Lancet Neurol 1, 383–386.
- 166 Muir KW & Lees KR (2003) Excitatory amino acid antagonists for acute stroke. Cochrane Database Syst Rev CD001244.
- 167 Birmingham K (2002) Future of neuroprotective drugs in doubt. Nat Med 8, 5.
- 168 Goldberg MP, Weiss JH, Pham PC & Choi DW (1987) N-methyl-D-aspartate receptors mediate hypoxic neuronal injury in cortical culture. J Pharmacol Exp Ther 243, 784–791.
- 169 Goldberg MP & Choi DW (1993) Combined oxygen and glucose deprivation in cortical cell culture: calcium-dependent and calcium-independent mechanisms of neuronal injury. J Neurosci 13, 3510–3524.
- 170 Nadler MJ, Hermosura MC, Inabe K, Perraud AL, Zhu Q, Stokes AJ, Kurosaki T, Kinet JP, Penner R, Scharenberg AM & Fleig A (2001) LTRPC7 is a Mg. ATP-regulated divalent cation channel required for cell viability. Nature 411, 590–595.
- 171 Ma HT, Venkatachalam K, Li HS, Montell C, Kurosaki T, Patterson RL & Gill DL (2001) Assessment of the role of the inositol 1,4,5-trisphosphate receptor in the activation of transient receptor potential channels and store-operated Ca2+ entry channels. J Biol Chem 276, 18888–18896.
- 172 Maruyama T, Kanaji T, Nakade S, Kanno T & Mikoshiba K (1997) 2APB, 2-aminoethoxydiphenyl borate, a membrane-penetrable modulator of Ins(1,4,5)P3-induced Ca2+ release. J Biochem (Tokyo) 122, 498–505.
- 173 Bilmen JG, Wootton LL, Godfrey RE, Smart OS & Michelangeli F (2002) Inhibition of SERCA Ca2+ pumps by 2-aminoethoxydiphenyl borate (2-APB). 2-APB reduces both Ca2+ binding and phosphoryl transfer from ATP, by interfering with the pathway leading to the Ca2+-binding sites. Eur J Biochem 269, 3678–3687.
- 174 Wang Y, Deshpande M & Payne R (2002) 2-Aminoethoxydiphenyl borate inhibits phototransduction and blocks voltage-gated potassium channels in Limulus ventral photoreceptors. Cell Calcium 32, 209–216.
- 175 Harks EG, Camina JP, Peters PH, Ypey DL, Scheenen WJ, Van Zoelen EJ & Theuvenet AP (2003) Besides affecting intracellular calcium signaling, 2-APB reversibly blocks gap junctional coupling in confluent monolayers, thereby allowing measurement of single-cell membrane currents in undissociated cells. FASEB J 17, 941–943.
- 176 Kwan CY & Putney JW Jr (1990) Uptake and intracellular sequestration of divalent cations in resting and methacholine-stimulated mouse lacrimal acinar cells. Dissociation by Sr2+ and Ba2+ of agonist-stimulated divalent cation entry from the refilling of the agonist-sensitive intracellular pool. J Biol Chem 265, 678–684.
- 177 Nelson MT, French RJ & Krueger BK (1984) Voltage-dependent calcium channels from brain incorporated into planar lipid bilayers. Nature 308, 77–80.
- 178 Davidson TJ, Harel S, Arboleda VA, Prunell GF, Shelanski ML, Greene LA & Troy CM (2004) Highly efficient small interfering RNA delivery to primary mammalian neurons induces MicroRNA-like effects before mRNA degradation. J Neurosci 24, 10040–10046.
- 179 Derossi D, Chassaing G & Prochiantz A (1998) Trojan peptides: the penetratin system for intracellular delivery. Trends Cell Biol 8, 84–87.
- 180 Nicholls DG & Ward MW (2000) Mitochondrial membrane potential and neuronal glutamate excitotoxicity: mortality and millivolts. Trends Neurosci 23, 166–174.
- 181 Vesce S, Kirk L & Nicholls DG (2004) Relationships between superoxide levels and delayed calcium deregulation in cultured cerebellar granule cells exposed continuously to glutamate. J Neurochem 90, 683–693.
- 182 Setsukinai K, Urano Y, Kakinuma K, Majima HJ & Nagano T (2003) Development of novel fluorescence probes that can reliably detect reactive oxygen species and distinguish specific species. J Biol Chem 278, 3170–3175.
- 183 Takezawa R, Schmitz C, Demeuse P, Scharenberg AM, Penner R & Fleig A (2004) Receptor-mediated regulation of the TRPM7 channel through its endogenous protein kinase domain. Proc Natl Acad Sci USA 101, 6009–6014.
- 184 Xiong ZG, Zhu XM, Chu XP, Minami M, Hey J, Wei WL, Macdonald JF, Wemmie JA, Price MP, Welsh MJ & Simon RP (2004) Neuroprotection in ischemia; blocking calcium-permeable Acid-sensing ion channels. Cell 118, 687–698.
- 185 Arundine M & Tymianski M (2003) Molecular mechanisms of calcium-dependent neurodegeneration in excitotoxicity. Cell Calcium 34, 325–337.
- 186 Kristian T, Katsura K, Gido G & Siesjo BK (1994) The influence of pH on cellular calcium influx during ischemia. Brain Res 641, 295–302.
- 187 Ekholm A, Kristian T & Siesjo BK (1995) Influence of hyperglycemia and of hypercapnia on cellular calcium transients during reversible brain ischemia. Exp Brain Res 104, 462–466.
- 188 Silver IA & Erecinska M (1992) Ion homeostasis in rat brain in vivo: intra- and extracellular [Ca2+] and [H+] in the hippocampus during recovery from short-term, transient ischemia. J Cereb Blood Flow Metab 12, 759–772.
- 189 Harris RJ, Symon L, Branston NM & Bayhan M (1981) Changes in extracellular calcium activity in cerebral ischaemia. J Cereb Blood Flow Metab 1, 203–209.
- 190 Nilsson P, Laursen H, Hillered L & Hansen AJ (1996) Calcium movements in traumatic brain injury: the role of glutamate receptor-operated ion channels. J Cereb Blood Flow Metab 16, 262–270.
- 191 Zhang ET, Hansen AJ, Wieloch T & Lauritzen M (1990) Influence of MK-801 on brain extracellular calcium and potassium activities in severe hypoglycemia. J Cereb Blood Flow Metab 10, 136–139.
- 192 Hansen AJ & Zeuthen T (1981) Extracellular ion concentrations during spreading depression and ischemia in the rat brain cortex. Acta Physiol Scand 113, 437–445.
- 193 Martell A & Smith R (1977) Critical Stability Constants. Plenum, New York.
- 194 Immke DC & Mccleskey EW (2001) Lactate enhances the acid-sensing Na+ channel on ischemia-sensing neurons. Nat Neurosci 4, 869–870.
- 195 Zimmerman AN & Hulsmann WC (1966) Paradoxical influence of calcium ions on the permeability of the cell membranes of the isolated rat heart. Nature 211, 646–647.
- 196 Duncan CJ & Morton JW (1996) Membrane damage and the Ca2+-paradox in the perfused rat kidney. Kidney Int 49, 639–646.
- 197 Soza M, Karpati G & Carpenter S (1986) Calcium paradox in skeletal muscles: physiologic and microscopic observations. Muscle Nerve 9, 222–232.
- 198 Okuda M, Lee HC, Chance B & Kumar C (1992) Depletion and repletion of Ca2+ in the perfused rat liver. J Laboratory Clin Med 120, 57–66.
- 199 Young W (1986) Ca paradox in neural injury: a hypothesis. Cent Nerv Syst Trauma 3, 235–251.
- 200 Siemkowicz E & Hansen AJ (1981) Brain extracellular ion composition and EEG activity following 10 minutes ischemia in normo- and hyperglycemic rats. Stroke 12, 236–240.
- 201 Schaller B & Graf R (2004) Cerebral ischemia and reperfusion: the pathophysiologic concept as a basis for clinical therapy. J Cereb Blood Flow Metab 24, 351–371.
- 202 Ruigrok TJ, Boink AB & Zimmerman AN (1976) Influence of ATP or oxygen plus substrate on occurrence of the calcium paradox. Recent Adv Stud Cardiac Struct Metab 11, 565–569.
- 203 Xiong Z, Lu W & Macdonald JF (1997) Extracellular calcium sensed by a novel cation channel in hippocampal neurons. Proc Natl Acad Sci USA 94, 7012–7017.
- 204 Smith SM, Bergsman JB, Harata NC, Scheller RH & Tsien RW (2004) Recordings from single neocortical nerve terminals reveal a nonselective cation channel activated by decreases in extracellular calcium. Neuron 41, 243–256.
- 205 Formenti ASA, Arrigoni E & Martina M (2001) Changes in extracellular Ca2+ can affect the pattern of discharge in rat thalamic neurons. J Physiol 535, 33–45.
- 206 Undem BJ, Oh EJ, Lancaster E & Weinreich D (2003) Effect of extracellular calcium on excitability of guinea pig airway vagal afferent nerves. J Neurophysiol 89, 1196–1204.
- 207 Mubagwa K, Stengl M & Flameng W (1997) Extracellular divalent cations block a cation non-selective conductance unrelated to calcium channels in rat cardiac muscle. J Physiol 502 (2), 235–247.
- 208 Frankenhaeuser B & Hodgkin AL (1957) The action of calcium on the electrical properties of squid axons. J Physiol 137, 218–244.
- 209 Hoth M & Penner R (1993) Calcium release-activated calcium current in rat mast cells. J Physiol 465, 359–386.
- 210 Su Z, Shoemaker RL, Marchase RB & Blalock JE (2004) Ca2+ modulation of Ca2+ release-activated Ca2+ channels is responsible for the inactivation of its monovalent cation current. Biophys J 86, 805–814.
- 211 Kostiuk PG, Mironov SL & Shuba I (1983) [2 selective filters in the calcium channel of the somatic membrane of mollusk neurons]. Neirofiziologiia 15, 420–427.
- 212 Almers W, Mccleskey EW & Palade PT (1984) A non-selective cation conductance in frog muscle membrane blocked by micromolar external calcium ions. J Physiol 353, 565–583.
- 213 Fukushima Y & Hagiwara S (1985) Currents carried by monovalent cations through calcium channels in mouse neoplastic B lymphocytes. J Physiol 358, 255–284.
- 214 Hess P, Lansman JB & Tsien RW (1986) Calcium channel selectivity for divalent and monovalent cations. Voltage and concentration dependence of single channel current in ventricular heart cells. J Gen Physiol 88, 293–319.
- 215 Polo-Parada L & Korn SJ (1997) Block of N-type calcium channels in chick sensory neurons by external sodium. J Gen Physiol 109, 693–702.
- 216 Armstrong CM & Cota G (1999) Calcium block of Na+ channels and its effect on closing rate. Proc Natl Acad Sci USA 96, 4154–4157.
- 217 Armstrong CM (1999) Distinguishing surface effects of calcium ion from pore-occupancy effects in Na+ channels. Proc Natl Acad Sci USA 96, 4158–4163.
- 218 Johnson JP Jr, Balser JR & Bennett PB (2001) A novel extracellular calcium sensing mechanism in voltage-gated potassium ion channels. J Neurosci 21, 4143–4153.
- 219 Xiong ZG & Macdonald JF (1999) Sensing of extracellular calcium by neurones. Can J Physiol Pharmacol 77, 715–721.
- 220 Brown EM & Macleod RJ (2001) Extracellular calcium sensing and extracellular calcium signaling. Physiol Rev 81, 239–297.
- 221 Devries SH & Schwartz EA (1992) Hemi-gap-junction channels in solitary horizontal cells of the catfish retina. J Physiol 445, 201–230.
- 222 Ebihara L, Liu X & Pal JD (2003) Effect of external magnesium and calcium on human connexin 46 hemichannels. Biophys J 84, 277–286.
- 223 Kubo Y, Miyashita T & Murata Y (1998) Structural basis for a Ca2+-sensing function of the metabotropic glutamate receptors. Science 279, 1722–1725.
- 224 Vassilev PM, Mitchel J, Vassilev M, Kanazirska M & Brown EM (1997) Assessment of frequency-dependent alterations in the level of extracellular Ca2+ in the synaptic cleft. Biophys J 72, 2103–2116.
- 225 Wise A, Green A, Main MJ, Wilson R, Fraser N & Marshall FH (1999) Calcium sensing properties of the GABA(B) receptor. Neuropharmacology 38, 1647–1656.
- 226 Moran MM, Xu H & Clapham DE (2004) TRP ion channels in the nervous system. Curr Opin Neurobiol 14, 362–369.
- 227 Gunthorpe MJ, Benham CD, Randall A & Davis JB (2002) The diversity in the vanilloid (TRPV) receptor family of ion channels. Trends Pharmacol Sci 23, 183–191.
- 228 Liman ER, Corey DP & Dulac C (1999) TRP2: a candidate transduction channel for mammalian pheromone sensory signaling. Proc Natl Acad Sci USA 96, 5791–5796.
- 229 Stowers L, Holy TE, Meister M, Dulac C & Koentges G (2002) Loss of sex discrimination and male-male aggression in mice deficient for TRP2. Science 295, 1493–1500.
- 230 Greka A, Navarro B, Oancea E, Duggan A & Clapham DE (2003) TRPC5 is a regulator of hippocampal neurite length and growth cone morphology. Nat Neurosci 6, 837–845.
- 231 Li HS, Xu XZ & Montell C (1999) Activation of a TRPC3-dependent cation current through the neurotrophin BDNF. Neuron 24, 261–273.
- 232 Sun M, Goldin E, Stahl S, Falardeau JL, Kennedy JC, Acierno JS Jr, Bove C, Kaneski CR, Nagle J, Bromley MC, Colman M, Schiffmann R & Slaugenhaupt SA (2000) Mucolipidosis type IV is caused by mutations in a gene encoding a novel transient receptor potential channel. Hum Mol Genet 9, 2471–2478.
- 233 Alessandri-Haber N, Dina OA, Yeh JJ, Parada CA, Reichling DB & Levine JD (2004) Transient receptor potential vanilloid 4 is essential in chemotherapy-induced neuropathic pain in the rat. J Neurosci 24, 4444–4452.
- 234 Fonfria E, Marshall IC, Benham CD, Boyfield I, Brown JD, Hill K, Hughes JP, Skaper SD & Mcnulty S (2004) TRPM2 channel opening in response to oxidative stress is dependent on activation of poly (ADP-ribose) polymerase. Br J Pharmacol 143, 186–192.
- 235 Balzer M, Lintschinger B & Groschner K (1999) Evidence for a role of Trp proteins in the oxidative stress-induced membrane conductances of porcine aortic endothelial cells. Cardiovasc Res 42, 543–549.
- 236 Wehage E, Eisfeld J, Heiner I, Jungling E, Zitt C & Luckhoff A (2002) Activation of the Cation Channel Long Transient Receptor Potential Channel 2 (LTRPC2) by Hydrogen Peroxide. A splice variant reveals a mode of activation independent of ADP-ribose. J Biol Chem 277, 23150–23156.
- 237 Kraft R, Grimm C, Grosse K, Hoffmann A, Sauerbruch S, Kettenmann H, Schultz G & Harteneck C (2004) Hydrogen peroxide and ADP-ribose induce TRPM2-mediated calcium influx and cation currents in microglia. Am J Physiol Cell Physiol 286, C129–C137.
- 238 Hara Y, Wakamori M, Ishii M, Maeno E, Nishida M, Yoshida T, Yamada H, Shimizu S, Mori E, Kudoh J, Shimizu N, Kurose H, Okada Y, Imoto K & Mori Y (2002) LTRPC2 Ca2+-permeable channel activated by changes in redox status confers susceptibility to cell death. Mol Cell 9, 163–173.
- 239 Hardie RC, Raghu P, Moore S, Juusola M, Baines RA & Sweeney ST (2001) Calcium influx via TRP channels is required to maintain PIP2 levels in Drosophila photoreceptors. Neuron 30, 149–159.
- 240 Lintschinger B, Balzer-Geldsetzer M, Baskaran T, Graier WF, Romanin C, Zhu MX & Groschner K (2000) Coassembly of Trp1 and Trp3 proteins generates diacylglycerol- and Ca2+-sensitive cation channels. J Biol Chem 275, 27799–27805.
- 241 Jung S, Strotmann R, Schultz G & Plant TD (2002) TRPC6 is a candidate channel involved in receptor-stimulated cation currents in A7r5 smooth muscle cells. Am J Physiol Cell Physiol 282, C347–C359.
- 242 Okada T, Inoue R, Yamazaki K, Maeda A, Kurosaki T, Yamakuni T, Tanaka I, Shimizu S, Ikenaka K, Imoto K & Mori Y (1999) Molecular and functional characterization of a novel mouse transient receptor potential protein homologue TRP7. Ca2+-permeable cation channel that is constitutively activated and enhanced by stimulation of G protein-coupled receptor. J Biol Chem 274, 27359–27370.
- 243 Shi J, Mori E, Mori Y, Mori M, Li J, Ito Y & Inoue R (2004) Multiple regulation by calcium of murine homologues of transient receptor potential proteins TRPC6 and TRPC7 expressed in HEK293 cell. J Physiol 561, 415–432.
- 244 Perraud AL, Fleig A, Dunn CA, Bagley LA, Launay P, Schmitz C, Stokes AJ, Zhu Q, Bessman MJ, Penner R, Kinet JP & Scharenberg AM (2001) ADP-ribose gating of the calcium-permeable LTRPC2 channel revealed by Nudix motif homology. Nature 411, 595–599.
- 245 Heiner I, Eisfeld J, Halaszovich CR, Wehage E, Jungling E, Zitt C & Luckhoff A (2003) Expression profile of the transient receptor potential (TRP) family in neutrophil granulocytes: evidence for currents through LTRPC2 induced by ADP-ribose and NAD. Biochem J 371, 1045–1053.
- 246 Kolisek M, Beck A, Fleig A & Penner R (2005) Cyclic ADP-ribose and hydrogen peroxide synergize with ADP-ribose in the activation of TRPM2 channels. Mol Cell 18, 61–69.
- 247 Harteneck C (2005) Function and pharmacology of TRPM cation channels. Naunyn Schmiedebergs Arch Pharmacol 371, 307–314.
- 248 Perraud AL, Takanishi CL, Shen B, Kang S, Smith MK, Schmitz C, Knowles HM, Ferraris D, Li W, Zhang J, Stoddard BL & Scharenberg AM (2004) Accumulation of free ADP-ribose from mitochondria mediates oxidative stress-induced gating of TRPM2 cation channels. J Biol Chem 280, 6138–6148.
- 249 Ziegler M (2000) New functions of a long-known molecule. Emerging roles of NAD in cellular signaling. Eur J Biochem 267, 1550–1564.
- 250 Strubing C, Krapivinsky G, Krapivinsky L & Clapham DE (2001) TRPC1 and TRPC5 form a novel cation channel in mammalian brain. Neuron 29, 645–655.
- 251 Strubing C, Krapivinsky G, Krapivinsky L & Clapham DE (2003) Formation of novel TRPC channels by complex subunit interactions in embryonic brain. J Biol Chem 278, 39014–39019.
- 252 Toescu EC (2004) Hypoxia sensing and pathways of cytosolic Ca2+ increases. Cell Calcium 36, 187–199.
- 253 Montell C (2005) The TRP superfamily of cation channels. Sci STKE 2005 re3.
- 254 Montell C (2001) Physiology, phylogeny, and functions of the TRP superfamily of cation channels. Sci STKE 2001, RE1.
- 255 Chandrasekaran K, Mehrabian Z, Li XL & Hassel B (2004) RNase-L regulates the stability of mitochondrial DNA-encoded mRNAs in mouse embryo fibroblasts. Biochem Biophys Res Commun 325, 18–23.
- 256 Mehrabian Z, Liu LI, Fiskum G, Rapoport SI & Chandrasekaran K (2005) Regulation of mitochondrial gene expression by energy demand in neural cells. J Neurochem 93, 850–860.
- 257 Chinopoulos C, Tretter L, Rozsa A & Adam-Vizi V (2000) Exacerbated responses to oxidative stress by an Na+ load in isolated nerve terminals: the role of ATP depletion and rise of [Ca2+] (i). J Neurosci 20, 2094–2103.
- 258 Yu SP, Yeh CH, Sensi SL, Gwag BJ, Canzoniero LM, Farhangrazi ZS, Ying HS, Tian M, Dugan LL & Choi DW (1997) Mediation of neuronal apoptosis by enhancement of outward potassium current. Science 278, 114–117.
- 259 Yu SP, Yeh C, Strasser U, Tian M & Choi DW (1999) NMDA receptor-mediated K+ efflux and neuronal apoptosis. Science 284, 336–339.
- 260 Yu SP (2003) Regulation and critical role of potassium homeostasis in apoptosis. Prog Neurobiol 70, 363–386.
- 261 Montell C, Birnbaumer L & Flockerzi V (2002) The TRP channels, a remarkably functional family. Cell 108, 595–598.
- 262 Nilius B, Voets T & Peters J (2005) TRP channels in disease. Sci STKE 2005, re8.
- 263 Inoue R, Hanano T, Shi J, Mori Y & Ito Y (2003) Transient receptor potential protein as a novel non-voltage-gated Ca2+ entry channel involved in diverse pathophysiological functions. J Pharmacol Sci 91, 271–276.
- 264 Wissenbach U, Niemeyer BA & Flockerzi V (2004) TRP channels as potential drug targets. Biol Cell 96, 47–54.
- 265 Shuttleworth CW, Brennan AM & Connor JA (2003) NAD(P)H fluorescence imaging of postsynaptic neuronal activation in murine hippocampal slices. J Neurosci 23, 3196–3208.