Abstract
The neurodegeneration of cerebellar granule cells, after low potassium induced apoptosis, is known to be temporally divided into an early and a late phase. Voltage-dependent anion channel-1 (VDAC1) protein, changing from the closed inactive state to the active open state, is central to the switch between the early and late phase. It is also known that: (i) VDAC1 can undergo phosphorylation events and (ii) AMP-activated protein kinase (AMPK), the sensor of cellular stress, may have a role in neuronal homeostasis. In the view of this, the involvement of AMPK activation and its correlation with VDAC1 status and activity has been investigated in the course of cerebellar granule cells apoptosis. The results reported in this study show that an increased level of the phosphorylated, active, isoform of AMPK occurs in the early phase, peaks at 3 h and guarantees an increase in the phosphorylation status of VDCA1, resulting in a reduced activity of this latter. However this situation is transient in nature, since, in the late phase, AMPK activation decreases as well as the level of phosphorylated VDAC1. In a less phosphorylated status, VDAC1 fully recovers its gating activity and drives cells along the death route.
Similar content being viewed by others
Avoid common mistakes on your manuscript.
Introduction
Cerebellar granule cells (CGCs), which undergo apoptosis when deprived of potassium, are an in vitro model for neurodegeneration since several events, resembling those happening in the course of Alzheimer’s Disease (AD), take place in this experimental system (see [1] and refs. therein). We have demonstrated that the apoptotic pathway can be formally divided into an early (0–3 h) and a late (3–8 h) phase, each one characterized by specific events acting sequentially [2]. In early-apoptosis, different neuroprotective defense mechanisms prevail [3,4,5], mainly sustained by a Warburg-like response. This is characterized by increased expression and activity of some glycolytic enzymes (i.e. glucose transporters 1 and 3 (GLUT1 and GLUT3), hexokinase (HK), phosphofructokinase and lactate dehydrogenase), HK interaction with the voltage-dependent anion channel-1 (VDAC1), closure of VDAC1 and consequent numbness of mitochondria [5]. In the late phase, the increase in glucose-6P induces HK dissociation from VDAC1. The activity of VDAC1 increases and, due to the opening of the channel protein, mitochondria restart working at full regime, increasing ROS production and leading cell to death [6].
AMP-activated protein kinase (AMPK), a hetero-trimeric complex composed of a catalytic alpha subunit and two regulatory beta and gamma subunits, is a serine/threonine kinase critical for maintaining cellular energy homeostasis. AMPK is a sensor of cellular stress, maximally activated when phosphorylated at Thr172 on its catalytic alpha subunit (P-AMPK) by one of the upstream kinases, liver kinase B1 (LK1) or calcium/calmodulin-dependent protein kinase kinase β (CamKKβ) [7]. Once activated, by phosphorylating key enzymes involved in different metabolic pathways, AMPK triggers catabolic reactions and simultaneously represses anabolic pathways in order to maintain energetic homeostasis [8]. Notwithstanding AMPK is highly expressed in neurons, its role in neuronal homeostasis and during neurodegeneration is still under debate with AMPK apparently endowed with the dual effect to promote either death or survival. The activation of AMPK has been shown to induce neuronal apoptosis by increasing the expression of the pro-apoptotic molecule Bim [9] or to be neuroprotective by increasing GLUT3 expression [8]. The kinetics of AMPK activation, as well as the cellular context in which it occurs, seems to be important factors for determining its cellular responses. While a transient activation of AMPK was described to promote neuroprotection, sustained AMPK activation induced apoptosis either in vivo and in vitro [10]. Furthermore, AMPK deregulation is central for the metabolic changes occurring in major neurodegenerative diseases, including Huntington, Parkinson as well as AD, albeit with a controversial role [11]. The active isoform of AMPK is highly elevated in postmortem AD brain [12] and downregulation of AMPK activity reduced the Aβ-induced loss of synapses [13]. On the other hand, it was described that AMPK activation repressed amyloidogenic route and tau phosphorylation in neurons as well as Aβ levels and amyloid deposition in transgenic mice [14, 15].
VDAC1, the most abundant VDAC isoform, is a protein considered to be the gate-keeper of mitochondria, responsible for the flux of small metabolites and ions across mitochondrial membranes. We have recently identified, in this protein, the cellular switch regulating the progression from the early to the late phase of CGC apoptosis [6]. It is known that VDAC1 can undergo post-translational modifications that may alter its activity and interaction with other proteins as well as influence its proteolytic degradation [16]. In particular, phosphorylation at threonine, serine and tyrosine residues has been described in various physiopathological conditions.
The aim of this study was to determine whether AMPK activation is involved in the apoptotic process of CGCs. It is known that VDAC1 can be regulated by post-translation modification such as phosphorylation, thus we investigated whether a link exists between AMPK activation and VDAC1 phosphorylation status in the course of neuronal apoptosis. Results obtained in this study suggest that AMPK is activated early in CGCs undergoing apoptosis peaking at 3 h, and its activation strictly correlates with the phosphorylation status of VDAC1 and hence with VDAC1 activity.
Materials and methods
Reagents
Compound C was obtained from Enzo Life Sciences (Farmingdale, NY, USA), 5′-aminoimidazole-4-carboxamide riboside (AICAR) was from SIGMA (St Louise, MO, USA) as well as all other reagents unless indicated otherwise.
Cell cultures and induction of apoptosis
Primary cultures of CGCs were obtained from dissociated cerebellar of 7-day-old Wistar rats as in [17]. Cells were plated in basal medium Eagle (BME; Life technologies, Gaithersburg, MD, USA) supplemented with 10% fetal calf serum, 25 mM KCl, 2 mM glutamine and 100 µg/ml gentamicin on dishes (Nunc, Roskilde, Denmark) coated with poly-l-lysine. 1b-Arabinofuranosylcytosine (10 µM) was added to the culture medium 18–22 h after plating to prevent proliferation of non-neuronal cells. All experiments were performed with fully differentiated neurons (7–8 DIV). To induce apoptosis, cells were washed twice and switched to serum-free BME, containing 5mM KCl (K5) supplemented with glutamine and gentamicin. Apoptotic cells are then referred to as S-K5 cells. Control cells were treated identically but maintained in serum-free BME medium supplemented with 25 mM KCl (K25) and then referred to as S-K25 cells. Cultures were treated with different substances at the concentrations and for the times specified in the figure legends.
Assessment of neuronal viability
Viable CGCs were quantified by counting the number of intact nuclei after lysing the cells in detergent-containing solution [18]. This method has been shown to be reproducible and accurate and to correlate well with other methods of assessing cell survival-death [19]. Cell counts were performed in triplicate and are reported as means ± SD.
Caspase-3 activity
Caspase activity was measured by using the Clontech ApoAlert Caspase-3 Assay Kit following manufacturer’s instructions (Clontech Laboratories Inc., Mountain View, CA, USA). DEVDpNA was used as a colorimetric substrate. The increase in protease activity was determined by the spectrophotometric detection at 405 nm of the chromophore p-nitroanilide (pNA) after its cleavage by caspase-3 from the labelled caspase-3-specific substrate (DEVDpNA).
Western blotting analysis
Cell lysis and Western blot analysis were performed mainly as in [3, 4]. Briefly, equal amounts of protein were subjected to SDS-PAGE on 11% Tricine-SDS-polyacrylamide gels, blotted onto PVDF membranes and then probed with polyclonal anti-AMPK and anti-Phospho-AMPK (Thr172) (Millipore, Temecula, CA, USA), and monoclonal anti-GADPH and anti-PARP antibodies (Sigma Chemical Co. St Louis, MO, USA). HRP-conjugated secondary antibodies were used for detection followed by enhanced chemiluminescence development.
Co-immunoprecipitation
Cells were lysed in 1% Triton X-100, 150 mM NaCl, 20 mM TrisHCl, 1 mM EGTA, 1 mM EDTA, 1 mM DTT pH 7.2, for 10 min at 4 °C in the presence of protease and phosphatase inhibitor cocktails. Lysates were then centrifuged at 4 °C for 10 min at 960×g. The protein extracts (500 μg) were immunoprecipitated by Protein A/G PLUS-Agarose according to the manufacturer’s instructions using 7 μg of polyclonal anti-VDAC antibody (Santa Cruz Biotechnology). After overnight incubation at 4 °C, the immunocomplexes were eluted with Laemmli buffer 2× and were next analyzed by immunoblotting with the monoclonal anti-Phospho-Threonine (P-Thr) and anti-VDCA1 antibodies (Millipore, Temecula, CA, USA).
VDAC1 activity assay
VDAC1 activity assay was performed, as described in [6], using an experimental strategy which allows to follow the ADP/ATP exchange measuring VDAC1 and adenine nucleotide translocator (ANT-1) activities in the same experiment. Briefly, the measurement was carried out in homogenates from cells in the absence or presence of compounds designed to block VDAC1, i.e. DIDS (20 μM), or ANT-1, i.e. ATR (2 μM). Appearance of ATP, in the exchange with ADP added to homogenate, was monitored by using the ATP detecting system—consisting of glucose (2.5 mM), HK (0.5 e.u.), G6PDH (0.5 e.u.) and NADP+ (0.2 mM)—and the rate of NADP+ reduction in the extramitochondrial phase was followed as the absorbance increase at 334 nm. NADPH formation decrease is recorded in the presence of DIDS or ATR, showing that (i) the exchange of ADPext with ATPint exclusively occurs through the outer and inner mitochondrial membranes, respectively; (ii) both the membranes are intact and (iii) the exchange is mediated by protein/s. Then, the determination of VDAC1 and ANT-1 activities is made possible by applying the control strength criterion and using the inhibitors at different concentrations. The channel activity was expressed as percent of S-K25 control cells.
Statistical analysis
All statistical analyses in this study were performed by using SPSS software (SPSS, Chicago, IL, USA). The data were representative of at least three independent neuronal preparations (with comparable results) each one in independent measurements and are reported as the mean with the standard deviation (SD). Statistical significance of the data was evaluated using the one-way analysis of variance (ANOVA) followed by the post-hoc Bonferroni test. p < 0.05 was considered as significant for all analyses.
Results
AMPK is activated in CGC apoptosis
It is well known that CGCs, referred to as S-K25 cells, can be maintained in culture in the absence of serum but in the presence of 25 mM KCl [20, 21]. On the other hand, CGCs undergo massive apoptosis if the potassium concentration is lowered to 5 mM [20,21,22,23]. Apoptotic CGCs, referred to as S-K5, show typical hallmarks of apoptosis as well as degenerative changes and neurite retraction [20, 22, 24]. We used this experimental model for all the experiments, and the pictures of both control (S-K25) and apoptotic (S-K5) cells were shown in Fig. 1a. In Fig. 1b the activity of caspase-3, an apoptotic hallmark, was measured at 8 h after apoptosis induction showing a sharp increase in S-K5 cells as compared to S-K25 cells. The cleavage of the nuclear protein PARP (116 kDa), often used to assess the apoptotic nature of the death [25,26,27], was shown in Fig. 1c and identified by Western blots as the appearance of the 85 kDa band in S-K5 cells (see also [3]). Used as a control, the broad range caspase inhibitor, z-VAD, proved to be effective in preventing both caspase-3 activation [see also 2, 28] and PARP cleavage (see also [3]).
In Fig. 2 cell viability of control and apoptotic cells was measured either in the presence or absence of Compound C (CC), a widely used AMPK inhibitor [29] and AICAR, a drug known to activate AMPK [30,31,32]. As shown in Fig. 2a, the survival of S-K5 cells decreases up to 42 ± 5% after 24 h in low potassium as compared to the 94 ± 2% observed in S-K25 cells. When increasing concentrations of CC were added to S-K5 cells, at the time of apoptosis induction, a progressive improvement of cell survival was obtained with a full recovery reached at 2 μM CC and about 95 ± 3% living cells. The same inhibitor concentration (2 μM) proved not to be toxic on S-K25 cells. On the other hand, further increasing CC concentration up to 15 μM, dramatically worsened cell viability both in control and apoptotic cells and survival dropped to 28 ± 2 and 9 ± 2% respectively (Fig. 2a). Then, CC 2 μM was chosen for further experiments, being the only concentration able to fully rescue apoptotic cells and non-toxic for control cells.
In a second set of experiments the AMPK inhibitor was added at increasing time after apoptosis induction—i.e. after 1, 3, 6 and 8 h from the potassium withdrawn—in order to delineate the time-window of the apoptotic process in which it exerts the prosurvival effect. As shown in Fig. 2b, CC completely rescues cells from death only when added at time 0. On the other hand, the pro-survival effect of CC progressively decreases when added after 1, 3 or 6 h after apoptosis induction and at 8 h it is no longer apparent.
The effect of the AMPK activator, AICAR, is then checked on S-K25 and S-K5 cells. At the concentration of 100 μM, AICAR was proved to induce a 15% cell death in control cells, as compared to S-K25 cells alone, and even more in apoptotic cells where cell survival decreased from 50 ± 5%, observed in untreated S-K5 cells, to 28 ± 3% in the presence of the activator (Fig. 2c).
P-AMPK level early increases in apoptotic CGCs
Increased phosphorylation of AMPK at Thr172 is a marker of AMPK activation. Then, to evaluate whether AMPK activity was modulated in the course of CGC apoptosis, total AMPK and P-AMPK were measured by Western blot analysis of CGC extracts taken at different time after apoptosis induction. As shown in Fig. 3a, a significant increase in P-AMPK was detected with apoptosis progression, starting as soon as 1 h after potassium withdrawn and reaching a maximum after 3 h. Later, the level of P-AMPK decreased, being roughly similar to that of control cells after 24 h. Densitometric analysis and calculation of the P-AMPK/AMPK ratio confirmed these results (Fig. 3b). Having previously verified that AMPK appears to be involved in the early phase of CGC apoptosis, the effect of both inhibitor and activator of AMPK was then checked at 1 and 3 h. At both times, AICAR was proved to be able to further increase the level of P-AMPK while CC reduced it (Fig. 3a, b).
VDAC is phosphorylated within the first 3 h of CGC apoptosis and kept partially inactive
The level of phosphorylated VDCA1 (P-VDAC1) was assessed with coimmunoprecipitation experiments and results were reported in Fig. 4a, b. In S-K5 cells, VDAC1 appeared to be in a phosphorylated status after 3 h of apoptosis with about 30% increase over S-K25 cells. After 8 h the level of P-VADC1 decreased up to that of control cells. When the effect of CC on S-K5 cells was verified (Fig. 4a), the level of P-VDAC1 decreased after 3 h of apoptosis from 137 ± 8 to 106 ± 9%. No significant variations were revealed after 8 h of apoptosis. As a control, the same experiment was performed in the presence of LiCl, a well-known inhibitor of the GSK3β kinase, which is able to phosphorylate VDAC and which totally rescues S-K5 from death (data not shown and [33]). Similar to the results obtained in the presence of CC, also 20 mM LiCl was able to reduce the level of P-VADC1 at 3 h but had no effect at 8 h (Fig. 4b). On the other hand, AICAR was able to further increase the level of P-VDAC1 after only 1 h of apoptosis rising up its value from 135 ± 7 to 175 ± 12% (Fig. 4c).
With the aim to verify whether the phosphorylated status of VDAC1 could be responsible for the modulation of its activity in the course of CGC apoptosis, the activity of VDAC1 was assessed as a function of time both in presence and absence of CC, LiCl and AICAR. Results are reported in Fig. 5 and expressed as percent of S-K25 cells. In S-K5 cells, at 1 and 3 h, VDAC1 activity was lower than in control cells, being 65 ± 4 and 53 ± 6% respectively, as already found in our previous paper [6]. In presence of CC, i.e. when the level of P-VDAC1 decreased, VDAC1 activity partially recovered rising up to 76 ± 5 and 73 ± 3%, after 1 and 3 h of apoptosis. LiCl produced similar results. Conversely, AICAR reduced the activity of VDAC1 of about 12% at both times. When the assay was performed after 8 h of apoptosis, i.e. when the P-VADC1 level in S-K5 cells decreased to a value similar to that of control cells, the activity of VDAC1 increased up to 84 ± 4% and both CC and LiCl proved to be ineffective to further increase it. Instead AICAR was still effective to keep constantly low VDAC1 activity.
Discussion
The AMPK role in influencing neuronal homeostasis is often contradictory: AMPK activation could induce cell death [9, 13, 34,35,36] as well as promote neuroprotection. [37,38,39]. Since the kinetics of AMPK activation might be crucial to determine its role inside neurons, whether in favor or in detriment to neuronal survival [8, 9, 40], it is intriguing to verify the involvement of AMPK in the time course of the apoptotic process of CGCs.
In the present study we have demonstrated that AMPK activation is an upstream event in CGC apoptosis. In S-K5 cells treated with the AMPK inhibitor, CC, a full recovery in survival is obtained only when CC is added simultaneously with apoptosis induction. On the other hand, CC—when added to S-K5 cells in subsequent times with respect to apoptosis induction—proves to progressively lose its pro-survival effect, suggesting that CC targets a crucial step in the very early phase of the apoptotic time-course. Consistently, analysis of the expression level of AMPK isoforms has shown a clear but transient increase in P-AMPK, which is peaked at 3 h, soon after apoptosis induction. Later, in the route leading to death, P-AMPK level decreases even below the level observed in control cells after 24 h. This trend clearly suggests the involvement of other control mechanisms such as dephosphorylation mediated by phosphatases, whose involvement has been documented in cell death regulation as well as in AD [41, 42]. In line with our findings, it was also recently demonstrated that the activation of AMPK signaling occurs in CGCs undergoing glutamate excitotoxicity [43]. Herein, the AMPK-dependent compensatory mechanisms (i.e. increased expression of GLUT3 at the plasma membrane and increased intracellular ATP levels) sustain the neuroprotective role of the FCCP-induced mild mitochondrial uncoupling against glutamate excitotoxicity and support the notion that AMPK could actually be activated prior to or at very early phase of cell death in order to prevent energy loss via upregulation of cellular energetic flux.
As far as CGCs undergoing low potassium induced-apoptosis are concerned, at a first glance AMPK activation seems to be detrimental to neuronal survival. In fact CC is able to fully rescue neurons from death, as measured 24 h after apoptosis induction, as well as to keep low the level of P-AMPK protein in the initial phase of the apoptotic process. In parallel experiments, AICAR, the AMPK activator, which further increases P-AMPK/AMPK ratio, induces only a slight worsen in cell survival. However, when analyzing these results, it should be kept in mind that neurodegeneration is a complex process, encompassing several molecular events that take place in the time interval between the early activation of AMPK (observed between 1 and 3 h) and the late detection of cell death (measured after 24 h). It is likely that—in this time frame—AMPK could phosphorylate different substrates that could have conflicting effects. In this contest, AMPK has been reported to be an activator of glucose uptake and glycolysis, two events that have been correlated with its neuroprotective nature [8]. Recently, we have also demonstrated that apoptotic CGCs undergo an early metabolic reprogramming characterized by increased activities of some glycolytic enzymes. This Warburg effect-like mechanism is one of the earliest attempt that CGCs put in place to resist the death process, at least until the accumulating glucose-6P doesn’t induce HK detachment from VDAC. At this point, the consequent increased activity of VDAC, the gate-keeper that controls mitochondrial function, behaves as a cellular switch that accelerates the progression from the early-to the late-phase of apoptosis [5, 6]. Therefore, it is tempting to speculate that the early activation of AMPK could be involved in, if not responsible for, the onset of the Warburg effect described in apoptotic CGCs. In this context, it is noteworthy that the pharmacological inhibition of VDAC proved to activate AMPK signaling pathway in endothelial cells [44].
Due to the central role played by VDAC1 and since it is known that, at least in vitro, VDAC1 can be phosphorylated by different kinases, we investigated whether VDAC1 phosphorylation occurs in the course of CGC apoptosis and whether it is somehow dependent on AMPK activation. Herein, we provide evidence that, parallel to the increased level of P-AMPK, the level of P-VDAC1 increases too and becomes higher in early apoptosis, at 3 h, than at 8 h. The inhibitor and the activator of AMPK—i.e. CC and AICAR—alter the phosphorylation status of VDAC1, by reducing or increasing it respectively, and thus influencing the activity of the channel protein. LiCl, the inhibitor of the GSK3β kinase which is known to phosphorylate VDAC1 [45, 46], has been used as a positive control. During the first 3 h of apoptosis, when VDAC1 is in a phosphorylated status, its activity decreases as compared to control S-K25 cells suggesting that phosphorylation partially closes the channel protein and limits its activity. Consistently, in the presence of AICAR, while P-VDAC1 level increases, VDAC1 activity further decreases remaining constantly low at all the times analyzed. On the other hand, CC, as well as LiCl, is able to reduce P-VDAC1 level and recover VADC1 activity at level even higher than that observed in apoptotic cells, at 1 and 3 h. At 8 h, VDAC1 activity increases in apoptotic cells, but both CC and LiCl were not able to induce further increment, presumably because VDAC1 phosphorylation has reached a plateau level and/or come into play other regulatory mechanisms that contribute to control VDAC1 opening, i.e. activity.
At present, the role and the effect of VDAC1 phosphorylation on signal transduction pathways and on cell fate is still not clear nothwidstanding a number of different kinases have been described to participate in the regulation of VDAC1 [47, 48], In AD brain, the level of VDAC1 dephosphorylation, presumably due to phosphatase activation, was found to correlate with the severity of the disease [49] and amyloid beta (Aβ) was reported either to induce dephosphorylation of VDAC1 in neuronal lipid rafts [49] or VDAC1 phosphorylation by GSK3β kinase [50]. Similarly, by using in vitro approaches, the phosphorylated state of VDAC1 was correlated with either the closed state of the protein [51, 52] or the open state [52] and the effect of phosphorylation has been interpreted as to be either proapoptotic [51] or antiapoptotic [52].
Herein, by using an in vivo approach, we hypothesize that AMPK could be involved in VDAC phosphorylation, thus contributing to maintain VDAC1 in a partially closed (i.e. inactive) state during early-apoptosis in CGCs. In support of this hypothesis, Strogolova et al. [54] reported, for the first time, a direct interaction between Snf1 protein kinase—the AMPK-homolog in Saccharomyces cerevisiae—with the yeast mitochondrial porin (Por 1) and suggested that Por1/VDAC can be as a sort of “functional niche” that blocks Snf1/AMPK close to mitochondria, i.e. in the better position to monitor glucose/energy failure. However, as far as the results herein reported are concerned, due the inherent limitations in the use of inhibitors, whose side effects must be taken into account, we cannot exclude a priori that AMPK, besides directly acting on VDAC, could also indirectly regulate its phosphorylation.
In conclusion, we suggest that AMPK is activated early in CGCs undergoing apoptosis and its activation strictly correlates with the phosphorylation status of VDAC1 and hence with VDAC1 activity. Furthermore, results obtained make sense that the controversial role on the activation/inhibition of AMPK in apoptosis could be a matter of time. In early-apoptosis, AMPK activation is endowed with a neuroprotective intent since AMPK could mediated, directly or indirectly, the phosphorylation of VADC1 thus contributing to keep its activity low and to sustain the prosurvival Warburg effect-like. However, in late-apoptosis, the ability of AMPK to support the cellular resistance to death decreases and while AMPK is no more able to sustain adequate level of VDAC1 phosphorylation, other AMPK-substrates can progressively disclose their toxicity. Future research aim will be to seek out and identify other substrates of AMPK that may play a key role in the route of CGC neurodegeneration.
References
Calissano P, Matrone C, Amadoro G (2009) Apoptosis and in vitro Alzheimer disease neuronal models. Commun Integr Biol (Camb) 2:163–169
Atlante A, Bobba A, Calissano P, Passarella S, Marra E (2003) The apoptosis/necrosis transition in cerebellar granule cells depends on the mutual relationship of the antioxidant and the proteolytic systems which regulate ROS production and cytochrome c release en route to death. J Neurochem 84:960–971
Bobba A, Atlante A, Moro L, Calissano P, Marra E (2007) Nitric oxide has dual opposite roles during early and late phases of apoptosis in cerebellar granule neurons. Apoptosis 12:1597–1610
Bobba A, Casalino E, Petragallo VA, Atlante A (2014) Thioredoxin/thioredoxin reductase system involvement in cerebellar granule cell apoptosis. Apoptosis 19:1497–1508
Bobba A, Amadoro G, La Piana G, Calissano P, Atlante A (2015) Glycolytic enzyme upregulation and numbness of mitochondrial activity characterize the early phase of apoptosis in cerebellar granule cells. Apoptosis 20:10–28
Bobba A, Amadoro G, La Piana G, Petragallo VA, Calissano P, Atlante A (2015) Glucose-6-phosphate acts as tip the balance in modulating apoptosis in cerebellar granule cells. FEBS Lett. 589:651–658
Steinberg GR, Kemp BE (2009) AMPK in health and disease. Physiol Rev 89:1025–10278
Weisová P, Concannon CG, Devocelle M, Prehn JH, Ward MW (2009) Regulation of glucose transporter 3 surface expression by the AMP-activated protein kinase mediates tolerance to glutamate excitation in neurons. J Neurosci 29:2997–3008
Concannon CG, Tuffy LP, Weisová P, Bonner HP, Dávila D, Bonner C, Devocelle MC, Strasser A, Ward MW, Prehn JH (2010) AMP kinase-mediated activation of the BH3-only protein Bim couples energy depletion to stress-induced apoptosis. J Cell Biol 189:83–94
Cardaci S, Filomeni G, Ciriolo MR (2012) Redox implications of AMPK-mediated signal transduction beyond energetic clues. J Cell Sci 125:2115–2125
Salminen A, Kaarniranta K (2012) AMP-activated protein kinase (AMPK) controls the aging process via an integrated signaling network. Ageing Res Rev 11:230–241
Vingtdeux V, Davies P, Dickson DW, Marambaud P (2011) AMPK is abnormally activated in tangle- and pre-tangle-bearing neurons in Alzheimer’s disease and other tauopathies. Acta Neuropathol 121:337–349
Mairet-Coello G, Courchet J, Pieraut S, Courchet V, Maximov A, Polleux F (2013) The CAMKK2-AMPK kinase pathway mediates the synaptotoxic effects of Aβ oligomers through Tau phosphorylation. Neuron 78:94–108
Greco SJ, Sarkar S, Johnston JM, Tezapsidis N (2009) Leptin regulates Tau phosphorylation and Amyloid through AMPK in Neuronal Cells. Biochem Biophys Res 380:98–104
Vingtdeux V, Giliberto L, Zhao H, Chandakkar P, Wu Q, Simon JE, Janle EM, Lobo J, Ferruzzi MG, Davies P, Marambaud P (2010) AMP-activated protein kinase signaling activation by resveratrol modulates amyloid-beta peptide metabolism. J Biol Chem 285:9100–9113
Martel C, Wang Z, Brenner C (2014) VDAC phosphorylation, a lipid sensor influencing the cell fate. Mitochondrion 19:69–77
Levi G, Aloisi F, Ciotti MT, Gallo V (1984) Autoradiographic localization and depolarization-induced release of acidic amino acids in differentiating cerebellar granule cell cultures. Brain Res 290:77–86
Volontè C, Ciotti T, Battistini L (1994) Development of a method for measuring cell number: application to CNS primary neuronal cultures. Cytometry 17:274–276
Stefanis L, Troy CM, Qi H, Greene LA (1997) Inhibitors of trypsin-like serine proteases inhibit processing of the caspase Nedd-2 and protect PC12 cells and sympathetic neurons from death evoked by withdrawal of trophic support. J Neurochem 69:1425–1437
D’Mello SR, Galli C, Ciotti T, Calissano P (1993) Induction of apoptosis in cerebellar granule neurons by low potassium: inhibition of death by insulin-like growth factor I and cAMP. Proc Natl Acad Sci USA 90:10989–10993
Galli C, Meucci O, Scorziello A, Werge TM, Calissano P, Schettini G (1995) Apoptosis in cerebellar granule cells is blocked by high KCl, forskolin, and IGF-1 through distinct mechanisms of action: the involvement of intracellular calcium and RNA synthesis. J Neurosci 15:1172–1179
Schulz JB, Weller M, Klockgether T (1996) Potassium deprivation-induced apoptosis of cerebellar granule neurons: a sequential requirement for new mRNA and protein synthesis, ICE-like protease activity, and reactive oxygen species. J Neurosci 16:4696–4706
Nardi N, Avidan G, Daily D, Zilkha-Falb R, Barzilai A (1997) Biochemical and temporal analysis of events associated with apoptosis induced by lowering the extracellular potassium concentration in mouse cerebellar granule neurons. J Neurochem 68:750–759
Armstrong RC, Aja TJ, Hoang KD, Gaur S, Bai X, Alnemri ES, Litwack G, Karanewsky DS, Fritz LC, Tomaselli KJ (1997) Activation of the CED3/ICE-related protease CPP32 in cerebellar granule neurons undergoing apoptosis but not necrosis. J Neurosci 17:553–562
Ciani E, Virgili M, Contestabile M (2002) Akt pathway mediates a cGMP-dependent survival role of nitric oxide in cerebellar granule cells. J Neurochem 81:218–228
Tewari M, Quan LT, O’Rourke K, Desnoyers S, Zeng Z, Beidler DR, Poirier GG, Salvesen GS, Dixit VM (1995) Yama/CPP32 beta, a mammalian homolog of CED-3, is a Crma-inhibitable protease that cleave the death substrate poly (ADP-ribose) polymerase. Cell 81:801–809
Stefanis L, Park DS, Friedman WJ, Greene LA (1999) Caspase-dependent and -independent death of camptothecin-treated embryonic cortical neurons. J Neurosci 19:6235–6247
Atlante A, Bobba A, Paventi G, Pizzuto R, Passarella S (2010) Genistein and daidzein prevent low potassium-dependent apoptosis of cerebellar granule cells. Biochem Pharmacol 79:758–767
Zhou G, Myers R, Li Y, Chen Y, Shen X, Fenyk-Melody J, Wu M, Ventre J, Doebber T, Fujii N, Musi N, Hirshman MF, Goodyear LJ, Moller DE (2001) Role of AMP-activated protein kinase in mechanism of metformin action. J Clin Invest 108:1167–1174
Corton JM, Gillespie JG, Hawley SA, Hardie DG (1995) 5-aminoimidazole-4-carboxamide ribonucleoside. A specific method for activating AMP-activated protein kinase in intact cells? Eur J Biochem 229:558–565
Sullivan JE, Brocklehurst KJ, Marley AE, Carey F, Carling D, Beri RK (1994) Inhibition of lipolysis and lipogenesis in isolated rat adipocytes with AICAR, a cell-permeable activator of AMP-activated protein kinase. FEBS Lett 353:33–36
Hardie DG (2015) AMPK: positive and negative regulation, and its role in whole-body energy homeostasis. Curr Opin Cell Biol 33:1–7
Tajes M, Yeste-Velasco M, Zhu X, Chou SP, Smith MA, Pallas M, Camins A, Casadesus G (2009) Activation of Akt by lithium: Pro-survival pathways in aging. Mech Ageing Dev 130:253–261
Li J, Zeng Z, Viollet B, Ronnett GV, McCullough LD (2007) Neuroprotective effects of adenosine monophosphate-activated protein kinase inhibition and gene deletion in stroke. Stroke 38:2992–2999
McCullough LD, Zeng Z, Li H, Landree LE, McFadden J, Ronnett GV (2005) Pharmacological inhibition of AMP-activated protein kinase provides neuroprotection in stroke. J Biol Chem 280:20493–20502
Nakatsu Y, Kotake Y, Hino A, Ohta S (2008) Activation of AMP-activated protein kinase by tributyltin induces neuronal cell death. Toxicol Appl Pharmacol 230:358–363
Culmsee C, Monnig J, Kemp BE, Mattson MP (2001) AMP-activated protein kinase is highly expressed in neurons in the developing rat brain and promotes neuronal survival following glucose deprivation. J Mol Neurosci 17:45–58
Ramamurthy S, Ronnett G (2012) AMP-activated protein kinase (AMPK) and energy-sensing in the brain. Exp Neurobiol 21:52–60
Poels J, Spasić MR, Callaerts P, Norga KK (2009) Expanding roles for AMP-activated protein kinase in neuronal survival and autophagy. Bioessays 31(9):944–952
Choi SY, Yang JH (2016) AMP-activated protein kinase is involved in perfluorohexanesulfonate-induced apoptosis of neuronal cells. Chemosphere 149:1–7
Sun H, Wang Y (2012) Novel Ser/Thr protein phosphatases in cell death regulation. Physiology (Bethesda) 27:1–18
Sontag JM, Sontag E (2014) Protein phosphatase 2 A dysfunction in Alzheimer’s disease. Front Mol Neurosci 11:7–16
Weisová P, Anilkumar U, Ryan C, Concannon CG, Prehn JH, Ward MW (2012) ‘Mild mitochondrial uncoupling’ induced protection against neuronal excitotoxicity requires AMPK activity. Biochim Biophys Acta 1817:744–753
Head SA, Shi W, Zhao L, Gorshkov K, Pasunooti K, Chen Y, Deng Z, Li RJ, Shim JS, Tan W, Hartung T, Zhang J, Zhao Y, Colombini M, Liu JO (2015) Antifungal drug itraconazole VDAC1 to modulate the AMPK/mTOR signaling axis in endothelial cells. Proc Natl Acad Sci USA 112:E7276–E7285
Das S, Wong R, Rajapakse N, Murphy E, Steenbergen C (2008) Glycogen synthase kinase 3 inhibition slows mitochondrial adenine nucleotide transport and regulates voltage-dependent anion channel phosphorylation. Circ Res 103:983–991
Pastorino JG, Hoek JB, Shulga N (2005) Activation of glycogen synthase kinase 3beta disrupts the binding of hexokinase II to mitochondria by phosphorylating voltage-dependent anion channel and potentiates chemotherapy-induced cytotoxicity. Cancer Res 65:10545–10554
Kerner J, Lee K, Tandler B, Hoppel CL (2012) VDAC proteomics: post-translation modifications. Biochim Biophys Acta 1818:1520–1525
Sheldon KL, Maldonado EN, Lemasters JJ, Rostovtseva TK, Bezrukov SM (2011) Phosphorylation of voltage-dependent anion channel by serine/threonine kinases governs its interaction with tubulin. PLoS ONE 6:e25539
Fernandez-Echevarria C, Díaz M, Ferrer I, Canerina-Amaro A, Marin R (2014) Aβ promotes VDAC1 channel dephosphorylation in neuronal lipid rafts. Relevance to the mechanisms of neurotoxicity in Alzheimer’s disease. Neuroscience 278:354–366
Reddy PH (2013) Amyloid beta-induced glycogen synthase kinase 3β phosphorylated VDAC1 in Alzheimer’s disease: implications for synaptic dysfunction and neuronal damage. Biochim Biophys Acta 1832:1913–1921
Gupta R, Ghosh S (2015) Phosphorylation voltage-dependent anion channel by c-Jun N-terminal Kinase-3 leads to closure of the channel. Biochem Biophys Res Commun 459:100–106
Chen Y, Gaczynska M, Osmulski P, Polci R, Riley DJ (2010) Phosphorylation by Nek1 regulates opening and closing of voltage dependent anion channel 1. Biochem Biophys Res Commun 394:798–803
Tewari SG, Zhou Y, Otto BJ, Dash RK, Kwok WM, Beard DA (2015) Markov chain Monte Carlo based analysis of post-translationally modified VDAC gating kinetics. Front Physiol 5:513. doi:10.3389/fphys.2014.00513
Strogolova V, Orlova M, Shevade A, Kuchin S (2012) Mitochondrial porin Por1 and its homolog Por2 contribute to the positive control of Snf1 protein kinase in Saccharomyces cerevisiae. Eukaryot Cell 11:1568–1572
Acknowledgements
The authors thank Mr Gaetano Devito for his skilful technical assistance and Dr A. Storelli for linguistic consultation. This research was supported by: Project FIRB-MERIT—RBNE08HWLZ_012 to M.N.G.
Author information
Authors and Affiliations
Corresponding author
Rights and permissions
About this article
Cite this article
Bobba, A., Casalino, E., Amadoro, G. et al. AMPK is activated early in cerebellar granule cells undergoing apoptosis and influences VADC1 phosphorylation status and activity. Apoptosis 22, 1069–1078 (2017). https://doi.org/10.1007/s10495-017-1389-8
Published:
Issue Date:
DOI: https://doi.org/10.1007/s10495-017-1389-8