Bioenergetic Impairment in Animal and Cellular Models of Alzheimer’s Disease: PARP-1 Inhibition Rescues Metabolic Dysfunctions

Abstract

Amyloid-beta peptide accumulation in the brain is one of the main hallmarks of Alzheimer’s disease. The amyloid aggregation process is associated with the generation of free radical species responsible for mitochondrial impairment and DNA damage that in turn activates poly(ADP-ribose)polymerase 1 (PARP-1). PARP-1 catalyzes the poly(ADP-ribosylation), a post-translational modification of proteins, cleaving the substrate NAD⁺ and transferring the ADP-ribose moieties to the enzyme itself or to an acceptor protein to form branched polymers of ADP-ribose. In this paper, we demonstrate that a mitochondrial dysfunction occurs in Alzheimer’s transgenic mice TgCRND8, in SH-SY5Y treated with amyloid-beta and in 7PA2 cells. Moreover, PARP-1 activation contributes to the functional energetic decline affecting cytochrome oxidase IV protein levels, oxygen consumption rates, and membrane potential, resulting in cellular bioenergetic deficit. We also observed, for the first time, an increase of pyruvate kinase 2 expression, suggesting a modulation of the glycolytic pathway by PARP-1. PARP-1 inhibitors are able to restore both mitochondrial impairment and pyruvate kinase 2 expression. The overall data here presented indicate a pivotal role for this enzyme in the bioenergetic network of neuronal cells and open new perspectives for investigating molecular mechanisms underlying energy charge decline in Alzheimer’s disease. In this scenario, PARP-1 inhibitors might represent a novel therapeutic intervention to rescue cellular energetic metabolism.

Keywords

Alzheimer’s disease bioenergetic metabolism mitochondria PARP-1 PKM2

INTRODUCTION

Alzheimer’s disease (AD) is a progressive neurodegenerative disorder representing the most common cause of dementia in the elderly population [1, 2]. AD pathology is characterized by progressive memory impairment and cognitive deficits, and by the presence of extracellular senile plaques mainly composed of amyloid-beta peptide (Aβ) and intraneuronal neurofibrillary tangles containing hyperphosphorylated tau protein [3].

Aβ derives from the sequential proteolytic processing of the amyloid-β protein precursor (AβPP) by beta- and gamma-secretases. When an unbalance between Aβ production and removal due to genetic and/or environmental factors occurs, Aβ oligomerization takes place producing different species of soluble supramolecular assemblies, and some of them finally converge toward fibrils formation [4]. Recent evidence suggests that oligomers, rather than insoluble fibrils, are implicated in Aβ primary neurotoxicity [5 –7] mediated by different mechanisms such as dysregulation of calcium homeostasis, inflammatory responses and generation of reactive oxygen species (ROS) [8 –10]. The relationship between ROS produced by Aβ accumulation and mitochondrial dysfunction in AD is well documented [11 –15]. Several pieces of evidence have indicated that many enzymes involved in glycolysis, in oxidative phosphorylation (OxPhos) and other metabolic pathways are affected at transcriptional level and their activities are robustly reduced in AD human brains [16, 17]. In particular, the decreased expression of proteins belonging to the electron transport chain complexes generates a progressive reduction in aerobic glucose metabolism within cerebral tissues. Previous studies demonstrated a downregulation of proteins mainly related to complexes I and IV of OxPhos in triple transgenic mice (pR5/APP/PS2) [18]. Moreover, mitochondrial bioenergetic deficit precedes the main pathological signs of AD in female triple transgenic mouse model of AD (3xTgAD) as indicated by the downregulation of expression and by the decrease in COX IV and PDH E1α activity in a time dependent manner [19].

When the mitochondrial deficit occurs, cells may rely mainly on glycolysis and other pathways for energy production. One of the major regulators of the glycolytic flux is pyruvate kinase (PK) that catalyzes the conversion of phosphoenolpyruvate to pyruvate in the rate-limiting ATP-generating step of glycolysis. PKM2 is an isoform expressed predominantly in embryonic tissue and in tumors [20] and differs from PKM1, mainly present in adult, for 23 amino acids within a 56-residue alternatively spliced exon (exon 10 for PKM2 and 9 for PKM1) [21]. PKM2 is able to slow down mitochondrial respiration and to divert glycolytic intermediates toward biosynthetic pathways, even in the presence of oxygen (Warburg effect) [22].

Although mechanisms responsible of bioenergetic unbalance due to Aβ are not yet clearly defined, more and more papers are focusing on NAD⁺, due to its emerging role not only in ATP production but also as substrate of different enzymes acting beyond energetic metabolic pathways. Among them poly(ADP-ribose)polymerase (PARP-1) catalyzes the poly(ADP-ribosylation), a post-translational modification of proteins, cleaving the substrate NAD⁺ and transferring the ADP-ribose (ADPr) moieties to the enzyme itself or to a protein acceptor to form branched polymers of ADPr.

The poly(ADP-ribose) chains (pADPr) may be released by the poly(ADP-ribose)glycohydrolase enzyme, whose endoglycosylase and exoglycosylase activities are extremely important for the pADPr release. PARPs and pADPr contribute not only to the genomic integrity and cell death pathways, but as was recently pointed out, to a broader spectrum of cellular functions such as inflammation and gene transcription regulation [23 –26].

The involvement of PARP-1 in controlling bioenergetics was suggested by Berger (1985) who first hypothesized that PARP-1 hyperactivation is responsible for NAD⁺/ATP depletion [27]. In particular, in the cytosol, the depletion leads to the glycolytic arrest and to neuron and glia death [28]. Recent papers demonstrated a key role of PARP-1 in controlling bioenergetics metabolism, underpinning a common mechanism in several pathologies [29, 30]. PARP-1 silenced cells exhibited a markedly enhanced cellular metabolic phenotype at baseline and under oxidative stress conditions [31]. Furthermore, several papers suggested the presence of PARP-1 in mitochondria, probably involved in the maintenance of DNA integrity and in mitochondrial energy failure [32 –34].

The involvement of PARP-1 in neurodegeneration has been investigated by several authors [35 –37]. Over-activation of PARP-1 was observed in primary mice microglia cultures and hippocampal rat slices by different research groups [38, 39]. A significant increase of PARP-1 activity in neuroblastoma-derived cell SH-SY5Y treated with Aβ, in 7PA2 cells overproducing amyloid peptide and in vivo in brain specimens of TgCRND8 transgenic mice has been observed [40].

In this scenario, the aim of this work is to elucidate the contribution of PARP-1 to energetic metabolism networks and to explore the possible capability of PARP-1 inhibitors to enhance mitochondrial functions in AD. Our results show that PARP-1 inhibition is able to relieve bioenergetic metabolic impairment. Therefore, the inhibition of PARP-1 might result in a successful therapeutic strategy for AD patients.

MATERIALS AND METHODS

Cell cultures

The human dopaminergic neuroblastoma cell line SH-SY5Y was obtained by ICLC (Genova, Italy) and maintained in DMEM/F12 supplemented with 10% heat-inactivated fetal bovine serum (FBS) and 2 mM glutamine, at 37°C and under 5% CO₂ humidified incubator. SH-SY5Y, an extremely versatile neuronal cell line, has been extensively utilized as an experimental model for the study of neuronal death processes induced by several agents [41]. SH-SY5Y cells were used in the range of 20–25 passage numbers.

7PA2, a Chinese hamster ovary (CHO) cell line that was stably transfected with a complementary DNA coding for APP751 containing the Val717Phe familial AD mutation that leads to Aβ_1–42 overproduction, was a kind gift of Prof. D. Walsh, USA [42] and Prof. A. Cattaneo. The produced Aβ peptide is the same that occurs in AD brain and its biological effect is exerted at low nanomolar to high picomolar concentrations similar to those found in cerebrospinal fluid and in brain [43]. CHO cells were used as control. Both cell lines were maintained in DMEM/F12 supplemented with 10% heat-inactivated fetal bovine serum, 10 mM glutamine, Pen/Strep 1X at 37°C and under 5% CO₂ humidified incubator (for 7PA2 200 μM G418 was added to select transfected cells). CHO and 7PA2 cells were used at the same passage number (within 8th passage).

Treatments of cell culture

SH-SY5Y cells were treated with amyloid-beta peptide 25–35 fragment (Aβ_25–35) 25 μM final concentration. Aβ_25–35 was dissolved in saline buffer (PBS) at a concentration of 1 mM o/n at 37°C and then added in culture medium. SH-SY5Y cells and 7PA2 cells were incubated with PARP inhibitors such as MC2050 or Olaparib. MC2050, a new PARP-1 inhibitor, was dissolved in water at an initial concentration of 50 mM and then diluted to a final concentration of 50 μM in culture medium [44]. Olaparib (AstraZeneca) was dissolved at a concentration of 100 mM in DMSO and then diluted to a final concentration of 3.3 μM in culture medium as tested and selected in previous experiments (data not shown). In SH-SY5Y PARP-1 inhibitors were added o/n before Aβ_25–35 treatment. 7PA2 cells were treated with the inhibitors after 24 h from seeding.

PARP activity was evaluated for each set of experiments by a colorimetric PARP assay kit from Trevigen. Aβ_25–35 cytotoxicity was detected in SH-SY5Y cells by MTT assay [40].

Animals

TgCRND8 mice (TgCRND8–129Sv), carrying the double mutant form APP695 (KM670/671NL+V717F), were obtained from Dr. David Westaway, Center of Research on Neurodegenerative Disease, University of Toronto, and maintained in the heterozygous status (TgCRND8+/–) by mating male Tg with female WT (TgCRND8–/–) 129Sv mice (Charles River Laboratories, Wilmington, MA, USA). These mice show robust amyloid peptide deposition at 3 months of age [45]. All animals were housed in an air-conditioned room (temperature 21±1°C, relative humidity 60±10%) with 12 : 12 h light:dark cycle (light on from 8 AM to 8 PM) and with food and water ad libitum.

Biochemical analyses were carried out on a total of 16 animals (8 WT and 8 Tg). At the first and third month of life, hippocampus and entorhinal cortex, sites of the major amyloid accumulation, were collected, stored at –80°C and successively used for biochemical and molecular analyses as previously described [45]. All the experiments were performed in such a way as to sacrifice the minimum number of animals required and were approved by the authors’ institution (Sapienza University of Roma) in accordance with the European Communities Council Directive (86/609/EEC) and formally approved by the Italian Ministry of Health (D.L. 92/116).

Mitochondria isolation

Mitochondria were isolated following the protocols of [46] with slight modifications. Briefly, 10⁷ SH-SY5Y treated with Aβ_25–35 and 7PA2 cells were incubated in presence or absence of PARP inhibitors. Cells were homogenized in Mitochondrial buffer A (MBA) (0.6 M Mannitol, 10 mM Tris HCl pH 7.4, 1 mM EGTA, 0.1%, BSA, proteases inhibitors, 1 mM PMSF) with a glass Potter-Elvehjem and spun at 400 g for 10 min. The pellet was resuspended in MBA and spun 400 g for 10 min. The two supernatants from both centrifugations were combined and spun again at 400 g for 10 min to get rid of cell debris. The supernatant was centrifuged at 11000 g for 10 min and the pellet containing purified mitochondria was resuspended in MBA. The sample was then washed twice and collected after centrifugation at 11000 g for 5 min and resuspended in 100 μl of Mitochondrial buffer B (0.6 M Mannitol, 10 mM Tris HCl pH 7.4, 1 mM EGTA, 1 mM protease inhibitors), stored on ice, and immediately proceeded. The quality of eachisolation was checked by western blot analyses with the antibodies anti-γ-tubulin (data not shown).

Citrate synthase activity

Cell lysates obtained from SH-SY5Y treated with Aβ_25–35 and 7PA2 cells incubated in presence or absence of PARP inhibitors were centrifuged at 13,000×g and assayed for citrate synthase activity as described in [47]. Untreated SH-SY5Y and CHO cells are regarded as control. Citrate synthase is determined by the appearance of the free SH group of the released CoASH. The SH is measured using 5, 5’-dithiobis-(2-nitrobenzoate) (DTNB) setting the spectrophotometer at 412 nm. Citrate syntase activity is expressed as Units/mg protein.

Western blotting

PARylated protein detection

Treated and untreated SH-SY5Y and 7PA2 cells were lysed in 100 μl RIPA buffer (50 mM Tris-HCl pH 7.4, 150 mM NaCl, 1% NP-40, 0.1% SDS, 1 mM EDTA, 0.5% NaDOC, 1 mM PMSF, proteases inhibitors). Lysed samples were incubated on ice for 30 min and centrifuged at 20,000 g for 15 min at 4°C. Supernatants were collected and proteins quantification were performed using a Bradford Assay (Bio-Rad). Equal amounts of protein from lysates cells (20 μg) were separated on 12% SDS-PAGE and transferred to nitrocellulose membrane (GE Healthcare) using Turbo Blot system (Bio-Rad). Membranes were probed with anti-ADPr polymer antibodies (1 : 1000, Trevigen). The protein bands were visualized with Crescendo (Millipore) using ChemiDoc system (Bio-Rad) and densitometric analyses were performed with ImageJ software and normalized to a reference protein.

COXIV, PKM1, and PKM2 detection

Treated and untreated SH-SY5Y and 7PA2 cells or mice brain tissues were lysed as reported above. Equal amounts of protein from lysates cells (20 μg) and mitochondrial proteins (5 μg) purified as above described, were separated on 12% SDS-PAGE and transferred to nitrocellulose membrane (GE Healthcare) using Turbo Blot system (Bio-Rad). The membranes were probed with the following primary antibodies: anti-PKM1 (1 : 1500, Sigma SAB4200094), anti-COX IV (1 : 2000 Abcam ab14744)), anti-PKM2 (1 : 1000, Cell Signaling #3198), anti-actin (1 : 5000, Millipore MAB1501).

The protein bands were visualized as reported above. Classical mitochondrial makers such as VDAC have not been utilized due to influence of Aβ on these proteins levels.

NAD⁺ measurement

Total cellular NAD⁺ was quantified using NAD/NADH Quantification kit (Sigma-Aldrich). Briefly, 2×10⁵ cells of SH-SY5Y treated with Aβ_25–35 and 7PA2 cells incubated in presence or absence of PARP inhibitors were lysed in NADH/NAD extraction buffer by thawing and freezing. Untreated SH-SY5Y and CHO cells are regarded as control. The assay was carried out according to the manufacturer’s and as described in [48].

Mitochondrial membrane potential assay

Mitochondrial membrane potential (Δψm) wasevaluated following the mitochondrial electro-phoretic accumulation of the membrane permeable, fluorescent cationic probe JC-1 (Cayman). JC-1 dye exhibits potential-dependent accumulation in mitochondria, indicated by a fluorescence emission shift from green (∼529 nm) to red (∼590 nm). Consequently, mitochondrial depolarization is indicated by a decrease in the red/green fluorescence intensity ratio [49].

SH-SY5Y, 7PA2, and CHO cells were seeded in 96 wells black plates with medium without phenol red, at a density of 10000 cells/well and 4000 cells/well, respectively. SH-SY5Y treated with 25 μM Aβ_25–35 for 24 h and 7PA2 were incubated with or without PARP-1 inhibitors. 1.5 μM JC-1 dye was added to each wells and incubated at 37°C for 20 min. After washing in PBS, cells were analyzed by flow cytometry and 30,000 events were collected for each sample. Hence, Δψm was determined on the same number of intact cells as the ratio between red lecture (λ_Ex = 570 nm e λ_Em = 590 nm) and green lecture (λ_Ex = 485 nm e λ_Em = 535 nm), using VICTOR^TM Multilabel Counter, Perkin Elmer.

Oxygen consumption

Cellular respiration was evaluated using the High Resolution Respirometry (2k-Oxygraph, OROBOROS Instrument) and according to [50].

SH-SY5Y treated with Aβ_25–35 for 24 h and 7PA2 cells were pre-incubated with or without PARP-1 inhibitors for 48 h. Untreated SH-SY5Y and CHO cells were used as control. Cells were scraped and washed once with PBS and once with respiration medium buffer (0.5 mM EGTA, 3 mM MgCl₂, 20 mM taurine, 10 mM KH₂PO₄, 20 mM HEPES, 0.1% BSA, 110 mM mannitol, 0.3 mM DTT, pH 7.1). After buffer equilibration, both 4×10⁶ intact cells were put in the chamber and cell respiration was evaluated under basal (not stimulated) metabolic condition for 20 min. Data are expressed as pmol O₂/(s*10⁶ cells).

Lactate and ATP measurement

Lactate production was evaluated according to [51]. Briefly, media from SH-SY5Y treated with 25 μM Aβ_25–35 for 24 h and from 7PA2 cells both incubated with or without PARP-1 inhibitors, were recovered and spun at 1500 rpm for 10 min at 4°C. 100 μl of the supernatant solutions was subjected to lactic acid dehydrogenase enzymatic assay. NAD⁺ reduction to NADH was followed by measurement of absorbance at 340 nm in a Beckman DU 650 spectrophotometer (Beckman Instruments, Fullerton, CA) for 1 min. Lactate contents in the samples were calculated from the changes in absorbance of NADH at 340 nm in 1 min and normalized to μg of total proteins.

ATP was evaluated according to [52]. Oligomycin (omy) (1 μg/ml), a respiratory chain (OxPhos) inhibitor, was added with the Aβ treatment in SH-SY5Y and after 24 h from PARP-1 inhibitors in the 7PA2. The difference between basal glycolytic lactate and lactate produced with oligomycin represents the Δlact. The ratio ATP glycolytic /ATP OxPhos production is measured as the ratio between basal lactate upon Δlact.

Real time PCR

Total RNA was extracted from cells and from homogenized brain regions using Trizol reagent (Invitrogen, Carlsbad, CA) in accordance with the manufacturer’s instructions. The extracted RNA pellet was resuspended in diethyl pyrocarbonate (DEPC)-treated water and cDNA was synthesized from 1 μg of total RNA with Tetro cDNA Synthesis Kit (Bioline) at 42°C for 1 h. 1 μg of total cDNA was used for each real-time reaction; analyses were performed in triplicate for each sample as previously described [53]. PKM1 and PKM2 expression were determined with qRT-PCR with the following set of primers [54]:

PKM1 forward: 5’-CGAGCCTCAAGTCACTCCAC-3’;

PKM1 reverse: 5’-GTGAGCAGACCTGCCAGACT-3’;

PKM2 forward: 5’-ATTATTTGAGGAACTCCGCCGCCT-3’;

PKM2 reverse: 5’-ATTCCGGGTCACAGCAATGATGG-3’;

RSP27A RefSeq Accession: NM_002954 (Qiagen)

The results were analyzed using the Gene Expression Macro 1.1 (Bio-Rad, Segrate), which recorded the threshold cycle (Ct). An untreated cell sample (CTR) was used as a control, and target gene Ct values were normalized against RPS27A (Qiagen). Data were analyzed by using the 2–DDCt method and expressed as fold change with respect to CTR (relative quantity).

Statistical analysis

Experiments were repeated at least three times and all the results are expressed as the mean value±standard deviation (SD). Data analyses were performed using GraphPAD prism 4 software. Statistical comparison between groups was made using either Student’s t test or the ANOVA and Bonferroni post hoc test. p values <0.05 were regarded as significant.

RESULTS

Amyloid-β peptide impairs bioenergetic metabolism in AD models

Expression and activity of cytochrome c oxidase (COX IV) significantly decrease in postmortem brain tissue from AD patients, indicating an impairment of OxPhos in neurons [19]. To determine whether a mitochondrial dysfunction occurs in TgCRND8 mice and whether this impairment shows a time course overlapping with AD progression, we analyzed COX IV protein levels in entorhinal cortex and hippocampal regions of TgCRND8 and WT mice at 1 and 3 months of age by western blot analyses. COX IV levels decreased in TgCRND8 mice in both regions with a reduction of 30% with respect to the WT mice at 1 month and of 40% and 50% in entorhinal cortex and hippocampus, respectively, at 3 months of age, confirming mitochondrial impairment in our mouse model (Fig. 1A, B) already at 1 month of age where soluble Aβ is present even in absence of plaques [55]. It is difficult to assess whether this decrease of COX IV level was due to neural loss or to a decreased expression but results reported below on cellular models prompt us to consider more sustainable the latter explanation.

We also analyzed COX IV protein levels in our cellular models. SH-SY5Y were treated with Aβ_25–35, a shorter and more manageable form of the full-length Aβ_1–42 peptide, able to induce inflammation and oxidative stress comparable to that induced by the full-length peptide and ensuring reliable and reproducible results [56, 57]. COX IV levels were monitored in untreated and treated SH-SY5Y at 24, 48, and 72 h and in CHO and 7PA2 cells at 48 h after seeding, by western blot analyses (Fig. 2A, B). A significant decrease of COX IV levels was observed both in Aβ-treated SH-SY5Y compared to control cells. The same data were obtained in 7PA2 that naturally secrete Aβ_1–42.

PARP-1 activity influences mitochondrial function and bioenergetic metabolism in cellular models of AD

Since several papers have shown that PARP triggers mitochondrial dysfunctions [58, 59] we investigated whether PARP-1 also contributes to the impairment of OxPhos in our cellular models. To achieve this aim we used two different PARP inhibitors namely MC2050 and Olaparib. This latter is now in global clinical trial III as therapeutic agent for different solid tumors [60]. It is worthy out of mention that both these molecules are able to inhibit not only PARP-1, but also PARP-2 although to a less extent. Considering PARP-1 as the main PARylating enzyme involved in neurodegeneration, in this paper we refer to the inhibitory effects as mostly acting on PARP-1 enzyme.

We assessed PARP-1 activation in presence and in absence of MC2050 by western blot. As shown in Fig. 3 there was an increased PARP-1 PARylation in SH-SY5Y after Aβ treatment and in 7PA2 with respect to their respective control cells. MC2050 inhibitor counteracted this effect directly influencing PARP-1 activity. As shown in our previous paper, PARP-1 upregulation is associated to an increase of general protein PARylation [40].

It is well known that PARP-1 hyperactivation leads to NAD⁺ depletion, and our data, in our cellular models, confirm this phenomenon. As shown in Fig. 4, Aβ exposure in SH-SY5Y and 7PA2 caused a significant decrease in NAD⁺ content compared to control cells. As expected, MC2050 and Olaparib significantly prevented Aβ-induced NAD⁺ reduction.

To assess the influence of PARP-1 on energetic metabolism upon Aβ challenge, we followed different metabolic parameters such as mitochondrial membrane potential (Δψm), oxygen consumption rate, COX IV levels, lactate and ATP production in the presence or absence of PARP-1 inhibitors.

We measured Δψm using the fluorescent dye JC-1. Figure 5A shows that Δψm was rapidly reduced in SH-SY5Y injured by Aβ at 24 h and that pretreatment with MC2050 and Olaparib, partially attenuated this loss. Similarly, in 7PA2 cells there was a decrease of Δψm with respect to CHO and both MC2050 and Olaparib were similarly helpful in relieving the damage (Fig. 5B). Control experiments performed on untreated SH-SY5Y and CHO cells revealed that PARP-1 inhibitors neither affect Δψm nor any other metabolic parameter analyzed in subsequent experiments (Fig. 5A, B).

OxPhos activity was further analyzed studying O₂ consumption rate (OCR) measured by high resolution respirometry. Basal respiration rate of untreated SH-SY5Y was 23 pmol O₂/sx10⁶ cells while this value dropped at 15 pmol O₂/sx10⁶ cells in Aβ-treated SH-SY5Y. When Aβ-treated SH-SY5Y cells were incubated in the presence of either MC2050 or Olaparib, OCR was restored to 23.6 and 25 pmol O₂/sx10⁶ cells, respectively (Fig. 6A). 7PA2 cells showed a similar behavior with a basal respiration rate of CHO, 7PA2 and 7PA2 incubated with MC2050 or Olaparib of 28, 17, 24 and 23 pmol O₂/sx10⁶ cells, respectively (Fig. 6B). The data showed an OCR reduction in both amyloid injured cells, and the ability of PARP-1 inhibitors in restoring controlvalues.

We extended our analysis on mitochondrial impairment investigating PARP-1 influence on COX IV protein levels. Western blot experiments were undertaken on isolated mitochondria obtained from both SH-SY5Y treated with Aβ_25–35 and from 7PA2 cells in the presence or absence of MC2050. A significant decrease of COX IV protein levels was observed in SH-SY5Y cells treated with Aβ_25–35 and in 7PA2 compared to control cells, which was prevented by PARP-1 inhibition (Fig. 6C, D).

The above reported data demonstrated an impairment of mitochondrial function in our AD cellular models, and therefore we investigated whether an increased glycolytic flux was ascertainable to compensate the decreased ATP production by OxPhos. Since the amount of lactate produced under basal conditions is an indication of glycolytic ATP [52], we compared lactate released in the culture medium of SH-SY5Y cells treated and untreated with Aβ_25–35 and in the medium of 7PA2 and CHO cells by enzymatic assay.

The amyloid peptide challenge raised lactate production in a slight but significant manner in treated SH-SY5Y cells, while a 2-fold increase was observed in 7PA2 compared to CHO cells (Fig. 7A, B) demonstrating an increment of anaerobic glycolysis in cells injured by Aβ. This increment however, was less than that measured in the presence of oligomycin (omy) which specifically inhibits the OxPhos. The difference between lactate detected in the presence and absence of oligomycin (Δlact) was lower in both Aβ-injured cells with respect to untreated SH-SY5Y and CHO, indicating a slight increase of anaerobic metabolism in the presence of amyloid peptides (Fig. 7A, B). The use of the PARP-1 inhibitor MC2050 on Aβ_25–35 treated cells and 7PA2 attenuated the increase in lactate production (Fig. 7A, B). The same results were obtained with Olaparib (data not shown).

Δlact is determined as the difference between basal glycolytic lactate and lactate produced in the presence of oligomycin. The former is indicative of ATP produced by glycolysis while the latter represents a measure of the effort made by cells in presence of oligomycin to partly compensate the depletion of ATP due to the oxidative phosphorylation inhibition and hence it is a generally accepted parameter used to evaluate ATPoxphos.

Therefore the basal lactate concentration value divided by Δlact was taken as indicative of the ATPgly/ATPOxPhos ratio. In SH-SY5Y treated with Aβ_25–35 and in 7PA2 cells this ratio is increased by 1.5 and 2.3-fold compared to the respective ratios observed in the control cells indicating a higher glycolytic efficiency to compensate the impairment of mitochondrial respiration in these cells. As expected, this increase was partially reversed by the use of PARP-1 inhibitors, and this effect was probably due to the partial reactivation of the oxidative phosphorylation (see above) (Fig. 7C, D).

It is worth noting that the above reported results on bioenergetic parameters are not affected by mitochondrial biogenesis/turnover deregulation since the analysis of citrate synthase activity yielded similar values for our cellular models and for control cells (data not shown).

PKM2 levels are affected by amyloid beta deposition in mice and in cellular models of AD

Amyloid treated SH-SY5Y cells and 7PA2 appeared to be able to compensate the impairment of OxPhos by an increase in glycolytic rate (Warburg effect). Since isoform 2 of pyruvate kinase (PKM2) contributes to the metabolic shift from oxidative phosphorylation to aerobic glycolysis [61], we investigated the expression of PKM2 along with the isoform 1 of pyruvate kinase (PKM1) both in cell cultures and also in transgenic mice TgCRND8. Western blot and RT-PCR analyses showed that in TgCRND8 mice PKM2 protein levels were increased 2-fold at 1 month of age both in entorhinal cortex (Fig. 8A, C) and hippocampus (Fig. 8B, D) compared to age-matched WT mice. This increase was completely reversed at 3 months of age in both brain regions where a significant decrease in PKM2 levels was observed. In contrast, PKM1 showed an increase of about 50% only at 3 months of age when compared with age matched WT mice in both tissues (Fig. 8A, B). We also analyzed PKM2 and PKM1 protein levels in amyloid treated SH-SY5Y and in 7PA2 cells at 24 and 48 h. Figure 9A shows that in SH-SY5Y cells treated with Aβ_25–35, PKM2 protein level increased by up to 60% in a time dependent manner while PKM1 remained constant. A similar effect was observed for PKM2 and PKM1 levels in 7PA2 cells (Fig. 9B).

Finally, to verify whether PARP-1 also modulates the levels of PKM2, western blot experiments were undertaken on both SH-SY5Y treated with Aβ_25–35 and on 7PA2 cells in presence or absence of MC2050. Figure 10 shows that the increase in PKM2 protein level observed both in SH-SY5Y cells treated with Aβ_25–35 and in 7PA2 compared to control cells was prevented by PARP-1 inhibition.

DISCUSSION

Several studies suggested that mitochondrial dysfunction significantly contributes to AD onset and progression, and it may be considered as the primary event in the pathogenesis of AD and then a target for therapeutic intervention [62, 63].

A major goal, in this paper was to study the impairment of oxidative metabolism in AD cellular and mice models and the role of PARP-1 in affecting this energy failure. Firstly, we analyzed whether our in vivo and cellular models recapitulate metabolic impairments monitoring three crucial parameters of OxPhos activity. We investigated the protein levels of COX IV that drives mitochondrial oxidative phosphorylation along with Δψm and oxygen consumption in SH-SY5Y treated with Aβ_25–35, in 7PA2 cells and in TgCRND8 mice. Results demonstrated a significant decrease of COX IV protein levels associated with a lower Δψm as well as in OCR, indicating that our models displayed a mitochondrial dysfunction similar to that found in human AD patients [64, 65], and in transgenic mice expressing AβPP with the Arctic mutation [66]. These data strongly confirm that the amyloid insult leads to the metabolic impairment and since this event is common even to other neurodegenerative diseases based on the accumulation of misfolded proteins, we can speculate that metabolic dysregulation is closely linked to the progression of these neuropathologies.

A causal relationship between PARP-1 and neurodegeneration has been already demonstrated in different studies [39, 67]. Since significant PARP-1 activation occurred in 7PA2 cells and SH-SY5Y cells treated with Aβ_25–35 and in TgCRND8 mice, here we investigated the existence of an association between the bioenergetic failure and PARP-1 [40]. Indeed, upon PARP-1 inhibition, SH-SY5Y cells treated with Aβ_25–35 and 7PA2 cells showed a partial recovery of COX IV levels, oxygen consumption rates and Δψm toward basal values, indicating that PARP-1 negatively affects the physiological bioenergetic set up of the cell.

These finding are in agreement with recent data showing that under oxidative stress conditions, PARP-1 silenced cells exhibited a phenotype with a markedly enhancement of cellular metabolism [31]. PARP-1^–/– mice showed higher mitochondrial content and oxidative metabolism, less fat accumulation and improved glucose tolerance [68]. Furthermore, PARP-1 inhibition produced an enhancement of mitochondrial activity in muscle and brown adipose tissue [30, 69] and enhanced fitness in mice by increasing the abundance of mitochondrial respiratory complexes and boosting mitochondrial respiratory capacity [70].

It is not yet clear how PARP-1 inhibition may help in restoring a proper energetic metabolism of mitochondrion. A direct action of PARP-1 on mitochondrial OxPhos proteins cannot be assessed since the existence of genuine PARP enzyme in the mitochondrial matrix is still a subject of debate [71 –73]. PARP-1 utilizes NAD⁺ as substrate, and thus asustained activation of the enzyme may deplete the intracellular pool of this key metabolite for cellular energy eliciting cell death [74]. Our results clearly indicate that Aβ leads to NAD⁺ depletion and energy deregulation via PARP-1 activation.

We hypothesize that inhibition of PARP-1 or poly(ADP-ribosylation) process may counteract Aβ insult, probably preserving NAD⁺ levels and in turn cellular respiration [38]. NAD⁺ depletion is effectively dangerous in organs that are metabolically active, such as heart, muscles and brain. Recent studies demonstrate that exogenous administration of NAD⁺ can attenuate oxidative DNA damage induced by Aβ in primary rat cortical neurons, suggesting the protective role of NAD⁺ in AD [75]. Extracellular NAD⁺ administration restores neuronal NAD⁺ levels after PARP-1 activation in astrocyte monocultures treated with NMDA, preventing glycolytic slow down, mitochondrial failure and neuronal death [76]. Furthermore, it has been demonstrated that mitochondrial respiratory function may be restored in APP(swe)/PS1(ΔE9) double transgenic (AD-Tg) mice by supplying nicotinamide mononucleotide (NMN), a NAD⁺ precursor, indicating this molecule as a promising agent in preserving mitochondrial function [77]. Therefore, NAD⁺-boosting strategies can be used to manage a wide spectrum of diseases from neurodegeneration to cancer [78]. In this scenario, PARP-1 inhibition makes NAD⁺ available to other NAD-dependent enzymes such as sirtuins. Noteworthy, SIRT1 in addition to its role in the modulation of energetic pathways depending on the type and the metabolic properties of tissues, has also a neuroprotective activity through suppression of Aβ amyloid production [79].

It is also of interest that ADPr can directly influence mitochondrial metabolism by inhibiting complex I of the electron transport chain [80] and its release from mitochondria, acts as molecular signaling. Consequently, PARP inhibition may downregulate ADPr production and thus maintain mitochondrial function [81]. The impairment of mitochondrial functions eventually leads to a deficit in ATP content that may be restored by glycolytic pathway. Therefore, we evaluated the glycolysis efficiency through the lactate measurement in SH-SY5Y cells treated with Aβ_25–35 and in 7PA2 incubated with or without PARP-1 inhibitors and in the presence or absence of oligomycin, an inhibitor of H+–ATP-synthase. Data showed an increase in lactate production in both cell types compared to control cells, as well as an enhancement of ATPgly/ATPOxPhos ratio. Although some assumptions of lactate method may becontested (i.e. the same rate of ATP consumption in presence or in absence of olygomicin), the overall data, herein presented, suggest that SH-SY5Y cells treated with Aβ_25–35 have a less active OxPhos than control cells and more active anaerobic metabolism to confront the ATP demand, as already reported by other authors for 7PA2 [82, 83]. The increase of the glycolytic flux seems therefore to represent a metabolic strategy in response to cellular Aβ insult. PARP-1 inhibitors were able to reduce lactate and ATP produced by glycolysis although this effect seems not to be direct but probably dependent on the restoration of mitochondrial potential and mitochondrial respiratory reserve capacity [84]. However, it is worthy underlining that glycolytic enzymes such as hexokinase and glyceraldehyde 3-phosphate dehydrogenase are the target of poly(ADP-ribosylation) by PARP-1 [85, 86]. The influence of PARP-1 on glycolysis may vary upon severity of injuries, and may also depend on the used cell types [87].

These findings on glycolytic flux variations prompted us to investigate, for the first time, the involvement of PKM2 in AD, monitoring its expression during the progression of the disease. PKM2 is highly upregulated in cancer cells, and the dynamic tuning of its activity causes the transition from aerobic respiration to glycolysis, a hallmark of the Warburg Effect [88]. Since an increased glycolytic flux is preferred after amyloid deposition, due to mitochondrial damage, we investigated the level of PKM2 in our mice and cellular models.

PKM2 expression in TgCRND8 mice was significantly increased at 1 month of age and decreased at 3 months of age relative to age-matched WT mice in both entorhinal cortex and hippocampal regions. Otherwise, PKM1 expression was increased in 3-month-old TgCRND8 mice compared with age-matched WT mice. The obtained results were confirmed by qRT-PCR experiments, indicating a time-dependent modulation of PKM2 and underlining the relevance of timing cellular response in the course of AD progression. The upregulation of PKM2 expression seems to indicate a critical link between early mitochondrial dysfunction and progression of the pathology. At 1 month of age, when plaques are not yet formed, but the mitochondrial impairment is already detectable, PKM2 increase is able to change and balance metabolic dysregulation. In contrast at 3 months of age, when the disease has almost reached the final state, the cells can counteract total mitochondrial failure only increasing the canonical glycolytic efflux increasing PKM1. Considering that PARP-1 activation is age dependent and that we reported an association between PARP-1 and PKM2 it is difficult to explain the data observed in Tg mice at 3 months of age where PKM2 dramatically decrease with respect to 1 month of age. These results suggest a complex mechanism involved in PKM2 expression in which PARP-1 is only one of the actors. However data reported seem to confirm a pivotal role of PARP-1 in PKM2 modulation at least at early stage of the disease.

In Aβ-treated SH-SY5Y and in 7PA2 cells, PKM2 increase is observed up to 48 h whereas PKM1 levels remain stable in both cell lines. The increase of PKM2 level in AD is unlikely to be ascribed to a control of PKM transcript splicing by c-myc phosphorylated [89] expression or Ras pathway activation which move the splicing toward production of the PKM2 transcript. PKM2 enhancement in both cell types and mice, despite their very different genetic background, strengthens the idea that PKM2 upregulation depends on basic physiologic processes common to both models such as bioenergetic metabolism [90]. In particular, the anaerobic metabolism observed in Aβ-induced state specific cells could be driven by PKM2 in a very early stage of the disease. The Warburg effect may take place in AD and may contribute to the resistance against Aβ toxicity, as reported by Newington recently [91]. Moreover, the use of PARP-1 inhibitors reversed PKM2 increase in SH-SY5Y cells treated with Aβ_25–35 and 7PA2 cells. We may speculate that a modulation of mitochondrial functions also occurs via PARP-1, through indirect downregulation of PKM2 expression even if a physical interaction between PARP-1 and PKM2 has been reported[92].

Finally, the outcome of these studies highlights that alteration of mitochondrial homeostasis participates in the earliest event of AD pathogenesis and that PARP-1 inhibitors are able to counteract bioenergetic impairment. Our data also point out that pyruvate kinase isoforms may reveal novel attractive markers for the progression of AD disease. The overall results may open new perspectives for the study of the molecular mechanisms that underlie the AD metabolic deficit, and indicate that PARP-1 inhibitors may represent an early therapeutic intervention.

Footnotes

ACKNOWLEDGMENTS

This paper was supported by funding from the EC 7th Framework Program (FP7/2007–2013) grant no. 278486 “DEVELAGE” and from Sapienza University of Roma, Scientific Research Programs 2013 and 2014. We are grateful to Prof. P. Sarti for kindly providing the 2k-Oxygraph, OROBOROS Instrument. English language has been revised by Ms. Jane Reynolds. This paper is dedicated to the memory of Dr. Giampiero Tempera.

Authors’ disclosures available online ().

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Bioenergetic Impairment in Animal and Cellular Models of Alzheimer’s Disease: PARP-1 Inhibition Rescues Metabolic Dysfunctions

Abstract

Keywords

INTRODUCTION

MATERIALS AND METHODS

Cell cultures

Treatments of cell culture

Animals

Mitochondria isolation

Citrate synthase activity

Western blotting

PARylated protein detection

COXIV, PKM1, and PKM2 detection

NAD+ measurement

Mitochondrial membrane potential assay

Oxygen consumption

Lactate and ATP measurement

Real time PCR

Statistical analysis

RESULTS

Amyloid-β peptide impairs bioenergetic metabolism in AD models

PARP-1 activity influences mitochondrial function and bioenergetic metabolism in cellular models of AD

PKM2 levels are affected by amyloid beta deposition in mice and in cellular models of AD

DISCUSSION

Footnotes

ACKNOWLEDGMENTS

References

NAD⁺ measurement