Reader TTS

ABSTRACT Heterozygous mutations of the lysosomal enzyme glucocerebrosidase (_GBA1_) represent the major genetic risk for Parkinson’s disease (PD), while homozygous _GBA1_ mutations cause

Gaucher disease, a lysosomal storage disorder, which may involve severe neurodegeneration. We have previously demonstrated impaired autophagy and proteasomal degradation pathways and

mitochondrial dysfunction in neurons from _GBA1_ knockout (_gba1__−/−_) mice. We now show that stimulation with physiological glutamate concentrations causes pathological [Ca2+]c responses

and delayed calcium deregulation, collapse of mitochondrial membrane potential and an irreversible fall in the ATP/ADP ratio. Mitochondrial Ca2+ uptake was reduced in _gba1_−/− cells as was

expression of the mitochondrial calcium uniporter. The rate of free radical generation was increased in _gba1_−/− neurons. Behavior of _gba1__+/−_ neurons was similar to _gba1__−/−_ in terms

of all variables, consistent with a contribution of these mechanisms to the pathogenesis of PD. These data signpost reduced bioenergetic capacity and [Ca2+]c dysregulation as mechanisms

driving neurodegeneration. SIMILAR CONTENT BEING VIEWED BY OTHERS MITOCHONDRIAL OXIDANT STRESS PROMOTES Α-SYNUCLEIN AGGREGATION AND SPREADING IN MICE WITH MUTATED GLUCOCEREBROSIDASE Article

Open access 11 December 2024 LYSOSOMAL LIPID ALTERATIONS CAUSED BY GLUCOCEREBROSIDASE DEFICIENCY PROMOTE LYSOSOMAL DYSFUNCTION, CHAPERONE-MEDIATED-AUTOPHAGY DEFICIENCY, AND ALPHA-SYNUCLEIN

PATHOLOGY Article Open access 06 October 2022 DYSREGULATION OF MITOCHONDRIA-LYSOSOME CONTACTS BY _GBA1_ DYSFUNCTION IN DOPAMINERGIC NEURONAL MODELS OF PARKINSON’S DISEASE Article Open access

22 March 2021 INTRODUCTION The _GBA1_ gene encodes for the lysosomal enzyme glucocerebrosidase (GBA1), which hydrolyzes the lipid glucosylceramide [1]. Homozygous mutations in _GBA1_ cause

an inherited lysosomal storage disease known as Gaucher disease (GD) [2], while heterozygous mutations in _GBA1_ are the major known genetic risk factor for Parkinson’s disease (PD) [3,4,5].

GD presents with a spectrum of symptoms and is classified into three ‘types’: [6] while type I shows limited or late CNS involvement, type II presents with a severe and progressive

neurodegeneration, and type III with less severe, chronic neurological symptoms. PD signs and symptoms include resting tremors, bradykinesia, and rigidity; anosmia, depression, and anxiety,

and at later stages, dementia. The links between specific GBA1 mutations and phenotype remain unclear [6, 7]. Both PD and lysosomal storage diseases such as GD are characterized by

dysfunction of the autophagy/lysosomal pathway and impaired mitochondrial function [8, 9], suggesting that deficiencies in each of these pathways can affect the other [10]. We have

previously shown that, in neurons from _gba1__−/−_ mice, autophagy is impaired upstream of the lysosome, associated with profoundly impaired mitochondrial function, decreased mitochondrial

membrane potential (Δψm), reduced mitochondrial respiration, and especially a dramatic decrease in uncoupled maximal respiratory capacity [11]. Interestingly, _gba1__+/−_ neurons showed a

modest reduction in Δψm, reflecting the absence of symptoms in the _gba1__+/−_ mice compared with _gba1__−/−_ [12]. The mechanism linking mitochondrial dysfunction to the underlying primary

defect in lysosomal biology remains unresolved but may be attributable to the accumulation of dysfunctional mitochondria due to impaired mitochondrial quality control pathways. This has also

been described in other lysosomal storage diseases (reviewed in [8]). Mitochondria show an intimate and complex relationship with calcium signaling, representing a major intersection

between cellular bioenergetics and cell signaling pathways [13,14,15]. Mitochondria take up Ca2+ from the cytosol, in a process mediated by the mitochondrial Ca2+ uniporter (MCU) complex

[16], its regulatory proteins [17,18,19], and balanced by the mitochondrial Ca2+ efflux pathway [20]. Dysregulation of [Ca2+]c signaling has been implicated widely in neurodegeneration and

has been identified as a key pathway in the selective degeneration of dopaminergic neurons in PD [21, 22]. We therefore explored the interplay between [Ca2+]c homeostasis and mitochondrial

bioenergetic capacity as potential contributors the neurodegeneration associated with _GBA1_ depletion, studying [Ca2+]c signaling in primary neuronal cultures from _gba1__−/−_, _gba1__+/−_,

and _gba1__+/+_ mice, stimulated with glutamate at physiological concentrations that are innocuous to control neurons (10 μM or below). As Ca2+-dependent mitochondrial dysfunction is

exacerbated by oxidative stress [15, 23], we also explored changes in free radical production associated with _GBA1_ depletion. We found that in both _gba1__−/−_ and _gba1__+/−_ neurons,

exposure to physiological glutamate concentrations caused delayed calcium deregulation (DCD), loss of Δψm and bioenergetic failure. Intriguingly, MCU protein expression was downregulated in

_gba1__−/−_, associated with a reduced mitochondrial Ca2+ buffering capacity. Very importantly, _gba1__+/−_ neurons, which model to a certain extent the heterozygous _GBA1_ mutations found

in PD patients, behaved similarly to _gba1__−/−_. These findings emphasize the fundamental importance of mitochondrial bioenergetic capacity in maintaining neuronal energy homeostasis in the

face of increased energetic demand associated with activity. Our data demonstrate the vulnerability of neurons in which mitochondrial function is perturbed to cellular [Ca2+]c overload,

suggesting that increased sensitivity to physiological glutamate concentrations may play an important role in neurodegeneration in GD and possibly also in GBA1-related PD. METHODS AND

MATERIALS MOUSE MODEL AND ANIMAL WELFARE Mice used for this work were described in Enquist et al. [12]. As K14-wt, K14-lnl/wt (lox/neomycin/lox), and K14-lnl/lnl and are herein referred as

_gba1__+/+_, _gba1__+/−_, and _gba1__−/−_. Mouse welfare was approved by the University College London Animal Welfare and Ethical Review Board (AWERB) and in accordance with personal

licenses granted by the UK Home Office and the Animal (Scientific Procedures) Act of 1986. The colony was maintained using breeding pairs heterozygous for GBA1 and pups were euthanized at P0

using cervical dislocation followed by decapitation for primary neuronal cultures preparation and liver extraction for genotyping. MOUSE GENOTYPING Genomic DNA was extracted from liver for

each pup (P0) using DNeasy Blood & Tissue Kit (Qiagen), genotyped by PCR using Q5 High Fidelity DNA Polymerase (NEB) and the following primers: GCex8-2 (Sigma, USA)

GTACGTTCATGGCATTGCTGTTCACT METex8-2 (Sigma, USA) ATTCCAGCTGTCCCTCGTCTCC NEO-AO2 (Sigma, USA) AAGACAGAATAAAACGCACGGGTGTTGG PCR was performed on T100 Thermal cycler (Biorad) using the

following PCR cycling conditions. Bands were visualized by gel electrophoresis. 98 °C 30 s ×15 98 °C 10 s 63 °C (0.5 °C touchdown) 30 s 72 °C 1.30 min 98 °C 10 s ×25 61 °C 30 s 72 °C 1.30 

min 72 °C 5 mins   10 °C Hold   NEURONAL AND ASTROCYTE PRIMARY CULTURE Mixed cultures of cortical neurons and astrocytes were obtained by dissecting brains from P0-P1 mice. Cortices from

each brain were isolated, kept in HBSS (H6648, Sigma) on ice separated from each other and genotyping was performed on liver extracted from each pup. Brain tissue was incubated in EBSS

(E2888, Sigma) and papain (LK003178, Worthington Biochemical Corp.) for 40 min at 37 °C and it was then dissociated by trituration in EBSS supplemented with DNAse (LK003172, Worthington

Biochemical Corp.) and papain inhibitor (LK003182, Worthington Biochemical Corp.). After spinning, the cell pellet was resuspended in Neurobasal (21103-049, Life Technologies), supplemented

with B27 (17504-044, Life Technologies), Glutamax (35050-038, Life Technologies), and 100 U/ml Penicillin–Streptomycin (1514-122, Life Technologies) counted and plated to appropriate

densities on coverslips (0.5·106 cells), 6-well plates (106 cells) or 96-well plates (30000 cells), coated with polylysine (P4707, Sigma). Half media changes were done every 4 or 7 days.

Cultures were maintained at 37 °C and 5% CO2 in humidified atmosphere and used between 10 and 15 days in vitro unless stated differently. CYTOSOLIC CALCIUM IMAGING Primary neurons and

astrocytes in mixed cultures were seeded onto 22-mm coverslips (or on 35mm FluoroDishes™ (Fisherscientific)) and stained with 5 μM FuraFF or Fura2 (F14181 and F1221, Thermo Fisher

Scientific) in recording buffer (150 mM NaCl, 4.25 mM KCl, 4 mM NaHCO3, 1.25 mM NaH2PO4, 1.2 mM CaCl2, 10 mM D-glucose, and 10 mM HEPES at pH 7.4) with pluronic acid 0.02%, at 37 °C and 5%

CO2 for 30 min. After washing, cells were imaged in recording buffer using a custom-made imaging widefield system built on an IX71 Olympus microscope equipped with a 20× water objective. A

Xenon arc lamp with a monochromator was used for excitation, exciting FuraFF or Fura2 fluorescence alternately at 340 nm ± 20 nm and 380 nm ± 20 nm and collecting emitted light through a

dichroic T510lpxru or a 79003-ET Fura2/TRITC (Chroma), and a band-pass filter 535/30 nm. Images were acquired using a Zyla CMOS camera (Andor) every 2–4 s and neurons were stimulated using

10 μM glutamate (G1626, Sigma). A total of 2 μM ionomycin was added at the end of each time course as a positive control. Electrical stimulation experiments were performed using a ‘myopacer’

(Ionoptix, Westwood USA) electrical stimulator and custom-made platinum electrodes (settings were: 40 V, 5 Hz 40 msec pulses and each stimulation period lasted 10 s). Activation of

metabotropic glutamate receptors was achieved by challenging neurons with 100 μM quisqualate (Q2128, Sigma), while inhibiting NMDA and AMPA/kainate receptors using 10 μM (2 

R)-amino-5-phosphonopentanoate (D-AP5) (0106/100, Bio Techne Ltd) and 20 μM CNQX (HB0205, HelloBio), respectively. Images were analyzed using ImageJ by selecting regions of interest (ROI) in

each cell and measuring average fluorescence intensity in the ROIs for each channel. After background subtraction, ratios between the signal excited at 380 nm and at 340 nm were calculated

at each time point and the resulting 340/380 ratioed traces representing cytosolic [Ca2+]c levels upon stimulation were plotted. Peak amplitude values were calculated for each cell using

Microsoft Excel and GraphPrism. RHODAMINE123 IMAGING Mixed cultures of primary neurons and astrocytes seeded on 22-mm coverslips were labeled with 10 μg/ml Rhodamine123 (R8004, Sigma) in

recording buffer, at 37 °C and 5% CO2 for 20 min. After washing, cells were imaged in recording buffer using a Zeiss 880 confocal microscope equipped with a 40× oil objective, exciting

neurons at 488 nm and collecting light longer than 505 nm. Images were acquired every 20 s with low laser power to avoid light-induced mitochondrial depolarization and photobleaching.

Neurons were stimulated using 10 μM glutamate (G1626, Sigma) and 1 μM Carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP) (C2920, Sigma) was added at the end of the time course to

evaluate the Rhod123 fluorescence intensity corresponding to 100% depolarization. Images were analyzed using ImageJ by selecting ROI in each field and measuring average fluorescence

intensity in the ROIs. For each trace, the signal was normalized between basal (the minimum value of intensity set to 0%) while the fully depolarized signal was set as the intensity of the

maximal data point, representing complete depolarization, at 100%. Cumulative frequency distribution analysis was also performed as for the measurements of [Ca2+]c. WESTERN BLOTTING Brains

tissue dissection was performed at P0-P1 and samples were snap frozen in liquid nitrogen and stored at −80 °C. To extract soluble proteins, brain tissue was homogenized in RIPA buffer (150 

mM NaCl, 0.5% Sodium deoxycholic acid, 0.1% SDS, 1% Triton X-100, 50 mM Tris pH 8.0, 1 mM PMSF (93482, Sigma)) and incubated 30 min on ice. After incubation, lysates were centrifuged and the

soluble fraction was collected to quantify protein concentration by Pierce BCA Protein Assay Kit (23225, Thermo Scientific). Sample buffer 4× with 2% beta-meracapto ethanol was added to

30–50 ug of total proteins, samples were boiled and loaded onto a Nu-Page gel (4–12% or 12%) (NP0335 and NP0341, Invitrogen) using MOPS or MEF buffer (NP001 or NP002, Invitrogen). Gels were

transferred to 0.45 μm PVDF membranes (IPVH00010, Millipore) in transfer buffer (NP0006, Invitrogen) with 20% methanol using a semi-dry system (Invitrogen). After blocking in 5% milk T-TBS

buffer (20 mM Tris, 150 mM NaCl pH 7.4, and 0.1% Tween 20), membranes were probed with primary antibodies (MCU, HPA016480 Sigma; EMRE, sc-86337 Santa Cruz; SOD1, sc-8637 Santa Cruz; SOD2,

sc-137254 Santa Cruz; grp75, sc-1058 Santa Cruz; β-actin, A2228 Sigma; MICU2, ab101465 Abcam; MCURI, ab86335, Abcam; GRIN2b, AGC-003 Alomone labs; GRIK2, AGC-009 Alomone labs;

Phospho-p40phox (Thr154), #4311 Cell Signaling Technology) overnight at 4 °C, washed and probed with an appropriate HRP-conjugated secondary antibody (anti-rabbit, 31463 Thermo Fisher

Scientific; anti-mouse, 31457 Thermo Fisher Scientific; anti-goat, A5420 Sigma) for 1 h at room temperature. Visualization was performed using Luminata Forte Western HRP substrate

(WBLUF0100, Millipore) and Chemidoc imaging system (Biorad). ATP:ADP MEASUREMENTS USING PERCEVALHR AND CONFOCAL IMAGING A total of 30,000–50,000 neurons were plated in 96-well plates with

clear bottom and transfected with the ATP:ADP sensor PercevalHR [24] at DIV8 using Lipofectamine LTX (15338-030, Life Technologies) in Optimem (31985-047, Life Technologies) for 1 h.

Conditioned media were kept and used to replace Optimem after transfection. After 48–72 h, imaging of single neurons expressing PercevalHR was performed in recording buffer exciting

PercevalHR at 405 nm and at 488 nm and collecting light at wavelengths longer than 510 nm, using a Zeiss 880 confocal microscope. Images were acquired every 10 s and neurons were stimulated

using 10 μM glutamate (G1626, Sigma) at 200 s. ATP:ADP ratios over time were obtained drawing ROIs in each neuron, measuring average fluorescence intensity in the ROIs for each channel in

ImageJ and calculating the 488 nm/405 nm ratio. MITOCHONDRIAL CALCIUM MEASUREMENT USING MITOCHONDRIAL-AEQUORIN A total of 30,000–50,000 mixed neurons and astrocytes were plated and grown

onto white 96-well plates, transduced with mtAequorin adenovirus [25] at DIV7–8 and incubated for 48–72 h. Afterward, media was replaced with 5 µM coelenterazine in Krebs Ringer Buffer (125 

mM NaCl, 5.5 mM D-Glucose, 5 mM KCl, 20 mM HEPES, 1 mM Na3PO4, 1 mM Glutamine, 100 mM Pyruvate, and 1.2 mM CaCl2). The plate was then incubated in the dark for 2 h at 37 °C. Luminescence

measurements were obtained using a plate reader (FluoStar Optima, BMG Labtech) every 1 s after 10 µM glutamate stimulation at 5 s. A total of 100 µM Digitonin was added at 25 s.

Mitochondrial Ca2+ concentrations were calculated as previously described [25]. MEASUREMENT OF RATES OF ROS GENERATION Mixed neurons and astrocytes plated on coverslips were imaged in

recording buffer using a Zeiss 510 confocal microscope, equipped with a UV laser (Coherent). 5 μM dihydroethidium (DHE, D1168 Thermo Fisher Scientific) was added to recording buffer after

starting the acquisition. Reduced DHE was excited at 351 nm and emitted light was recorded between 435 and 485 nm; oxidized DHE was excited at 543 nm and emitted light was collected using a

560-nm long-pass filter. Images were acquired over time every 8.93 s and 10 μM glutamate was added at 600 s to measure changes induced by glutamate stimulation in oxidation rates. Oxidation

rate curves were obtained calculating the ratio between reduced and oxidized DHE. Basal oxidation levels (basal slopes) were calculated considering the difference between ratio at 500 s and

the ratio at the beginning of the time course and dividing it by the time duration. Glutamate-induced oxidation rates were calculated as the difference between ratios at 1000 s and at 500 s

and dividing it by the time duration (glutamate slopes). MEASUREMENT OF GLUTATHIONE LEVELS USING MONOCHLOROBIMANE (MBC) Monocholorobimane (M1381MP, Invitrogen) imaging experiments were

performed further adapting a protocol set for neurons [26]. Briefly, neurons plated on coverslips as previously described and were incubated with 100 μM MCB in recording buffer until a

steady state was achieved. Images were acquired using a custom-made widefield imaging system built on an IX71 Olympus microscope equipped with a 20× water objective. A Xenon arc lamp with a

monochromator was used for excitation set at 380 nm and collecting emitted light through a dichroic mirror 79001-ET Fura2 (Chroma) and a band-pass emission filter at 525/36 nm, using a Zyla

CMOS camera (Andor). For image analysis, ROIs were then chosen and average MCB intensity for each neuron was calculated as an average of last three frames of the plateau and plotted in a

scatter plot. IMMUNOCYTOCHEMISTRY AND CONFOCAL IMAGING Neurons were plated on 96-well plates as previously described and grown until DIV12-14. Cells were then washed in PBS and fixed using

4% paraformaldehyde (P6148, Sigma) for 30 min at room temperature and then washed again. Blocking was performed by incubation with 3% Bovine Serum Albumin (A2153, Sigma) in PBS and staining

using a primary antibody against an extracellular epitope of Grin2b (AGC-003, Alomone labs) and a secondary anti-rabbit Alexa488 (Thermo Fisher Scientific). Nuclei were stained with Hoechst

33342 and images were then collected by exciting Alexa488 at 488 nm and Hoechst at 405 nm, using a 60× objective and a Zeiss 880 confocal microscope. MRNA EXTRACTION AND QPCR mRNA was

extracted from brain tissue using ReliaPrep™ RNA Cell Miniprep (Z6010, Promega); mRNA concentration was measured and 500 ng mRNA was used to obtain cDNA by means of the GoScript™ Reverse

Transcription Kit (A5000, Promega), following manufacturer instructions. The obtained cDNA was subjected to qPCR using SYBR® Green JumpStart Taq ReadyMix (S4438, Sigma) and CFX96 Real-Time

System (Biorad). Different pairs of primers where then used to quantify the mRNA expression levels of genes of interest: Cyclophilin A F 5′-CCCACCGTGTTCTTCGACA-3′ Cyclophilin A R

5′-CCAGTGCTCAGAGCTCGAAA-3′ GRIK2 F5′-TGTGGAATCTGGCCCTATGG-3′ GRIK2 R5′-TGAACTGTGTGAAGGACCGA-3′ Grin2b F 5′-CGCCCAGATCCTCGATTTCA-3′ Grin2b R5′-CTGGAAGAACATGGAGGACTCA-3′ MCU

F5′-GTCAGTTCACACTCAAGCCTAT-3′ MCU R 5′-TTGAAGCAGCAACGCGAACA-3′ MCURI F 5′-CTTCTGGGAGCAGGAAACTCTA-3′ MCURI R 5′-TGAGTAGCAAACCCATTGTC-3′ MICU1 F5′-GCTCCATAACGCCCAATGAG-3′ MICU1

R5′-GAAGGAGATGAGCCCACACT-3′ LIPID EXTRACTION AND LIPIDOMICS ANALYSIS Lipids were extracted from brains (dissections performed on newborn pups, _n_ = 3 per genotype) using Folch extraction

(chloroform:methanol = 2:1) and were analyzed, as previously described [27]. In this configuration, measurements of GBA1 substrate are not able to differentiate between the combined

glycosylceramides, and therefore include both glucosylceramides and galactosylceramides. STATISTICAL ANALYSIS Image quantification was performed using ImageJ and data were analyzed using

GraphPad Prism. When the number of data points (usually corresponding to cell number) was >150 (_n_ = 3 independent experiments), distribution analysis was performed (frequency

distribution or cumulative frequency distribution), while when the number of data points was lower mean ± SD or mean ± SEM were used, representing single data points on graphs to properly

account for data variability. Tests of normality were performed (Shapiro–Wilk test) to identify normal or non-normal populations. When normal, Student’s test or Anova tests (Bonferroni

post-test) were used as needed, otherwise the nonparametric Kruskal–Wallis test (Dunns post-test) was used. The chosen tests are clearly indicated in the figure legends; *_p_ < 0.05,

**_p_ < 0.01, and ***_p_ < 0.001. RESULTS _GBA1_ +/− AND _GBA1_ _−/−_ NEURONS SHOW DELAYED CALCIUM DEREGULATION IN RESPONSE TO 10 ΜM GLUTAMATE CONCENTRATIONS Mixed cultures of neurons

and astrocytes were loaded with the low-affinity [Ca2+]c indicator FuraFF-AM and images were acquired over time. Exposure to 10 μM glutamate (Fig. 1a) caused the expected rise in [Ca2+]c,

but responses clearly differed between _gba1__+/+_, _gba1__+/−_, and _gba1__−/−_ neurons. Both the early peak response, (ΔfuraFFearly) and responses at later time points differed

significantly between genotypes (Fig. 1bii and dii). Cumulative frequency distributions revealed a significant increase in the amplitude of the early glutamate response in both _gba1__−/−_

and _gba1__+/−_ neurons, compared with controls (Fig. 1bi, _n_ = 3 mice per genotype, _N_ = 30–60 cells per genotype per experiment). In the continued presence of glutamate, the initial

transient response was followed in the majority of _gba1__−/−_ cells by a delayed increase in [Ca2+]c, referred to as DCD, a characteristic response to toxic glutamate concentrations (100 μM

or higher) in control cells [28, 29]. At 10 μM glutamate, DCD was seen in very few control cells, but was a feature of the majority of _gba1__−/−_ cells, while _gba1__+/−_ cells showed an

intermediary response (Fig. 1a–c). The cumulative frequency distribution of the delayed response (ΔfuraFFDCD—Fig. 1di) and the scatter plot of the [Ca2+]c response at 400 s after glutamate

exposure (Fig. 1dii) indicated the percentage of neurons showing DCD (defined as ΔfuraFF > 0.1 ratio unit). In _gba1__+/−_ and in _gba1__−/−_ cultures, about 53 and 60% of the neurons

showed DCD, respectively, while this was seen in only 20% of neurons in _gba1__+/+_ cultures (Fig. 1d, _n_ = 3 mice per genotype, _N_ = 40–60 cells per genotype per experiment). Neuronal

responses to 1 μM glutamate, at the lower limit of the physiological range [30], also showed differences between genotypes consistent with these data (Supplementary Fig. 1A). [Ca2+]c

remained elevated 400 s after 1 μM glutamate in 10–15% of _gba1__+/−_ and _gba1__−/−_ neurons but only in 1% of _gba1__+/+_ neurons. In order to determine whether responses to Ca2+ signals

arising from different sources also showed deregulation, we explored responses to release ER Ca2+ by metabotropic glutamate receptors and to electrical pacing, promoting Ca2+ influx through

voltage-gated Ca2+ channels. Ionotropic receptors were inhibited using 10 μM D-AP5 and 20 μM CNQX and neurons challenged with 100 μM quisqualate, a group I metabotropic receptor agonist

[31]. Only very small responses were detectable using the low-affinity FuraFF (Supplementary Fig. 2A). Experiments with the higher affinity indicator Fura-2, revealed a small but significant

reduction in the [Ca2+]c peak response to quisqualate in _gba1__−/−_ neurons (Supplementary Fig. 2B-C). Responses to electrical pacing were undetectable using FuraFF, while measurements

with Fura-2 revealed no significant differences in peak responses between genotypes (Supplementary Fig. 2D). Altogether, these data show that DCD is a response only to Ca2+ influx through

ionotropic pathways, probably reflecting the much greater Ca2+ load that this represents. INCREASED SENSITIVITY OF _GBA1_ +/− AND _GBA1_ _−/−_ NEURONS TO GLUTAMATE IS NOT DUE TO DIFFERENT

EXPRESSION OF GLUTAMATE RECEPTORS The increased sensitivity to glutamate could reflect altered expression of ionotropic glutamate receptors. The ionotropic glutamate NMDA receptor subunit 2B

(_Grin2b_) and the ionotropic kainate receptor subunit 2 (_Grik2_) are both lifespan modifier genes in GWAS studies of mouse strains treated with the GBA1 inhibitor, Conduritol B Epoxide

(CBE) [32]. _Grin2b_ mRNA expression is higher in mouse strains in which lifespan is shortened following CBE treatment, suggesting that expression of the glutamate receptor is increased as a

secondary effect of GBA1 inhibition and sensitizes the cells. We therefore measured expression levels of mRNA for _Grin2b_ and for _Grik2_ in _gba1__+/+_, _gba1__+/−_, and _gba1__−/−_ mouse

brains by qPCR. mRNA levels of _Grik2_ were slightly higher for _gba1__+/−_ neurons compared with the other genotypes, while _Grin2b_ mRNA levels showed a small but significant decrease in

_gba1__+/−_ and _gba1__−/−_ compared with _gba1__+/+_ (Fig. 2a). However, western blots of brain lysates to quantify Grik2 and Grin2b protein levels (Fig. 2b) (_n_ = 4–5 per genotype)

revealed no significant difference among genotypes. Surface analysis for Grin2b by immunofluorescence [33] and confocal imaging also failed to reveal any significant differences between them

(Supplementary Fig. 3). Thus, the increased glutamate sensitivity and DCD is not a consequence of increased glutamate receptor expression. As DCD is a predictor of neuronal cell death [34,

35], these data show that _gba1__−/−_ and _gba1__+/−_ neurons are more vulnerable to glutamate-induced [Ca2+]c overload than the _gba1__+/+_. LIPID HOMEOSTASIS IS DIFFERENTIALLY ALTERED IN

THE BRAIN OF _GBA1_ _+/−_ AND _GBA1_ _−/−_ MICE The loss of GBA1 is expected to cause accumulation of its substrate glucosylceramide. Mass spectrometry analysis of lipids extracted from

_gba1__+/+_, _gba1__+/−_, and _gba1__−/−_ brains generates measurements of glycosylceramide, which corresponds to both glucosylceramide and galactosylceramide. The resulting data showed

accumulation of glycosylceramides only in _gba1__−/−_ but not in _gba1__+/−_ brains, suggesting that one copy of the gene may generate sufficient enzyme to minimize substrate accumulation

(Fig. 2b). However, it is important to emphasize that smaller increases in glucosylceramide levels in _gba1__+/−_ brains may have been masked when measured together with galactosylceramide,

because the latter are more abundant in subpopulations of cells in the brain. In fact, heterozygous GBA pure neurons may present glucosylceramide accumulation [36, 37]. Noteworthy, saturated

glycosylceramides (d18:0) were not elevated even in _gba1__−/−_ cells. Interestingly, other secondary substrates may also be implicated. For example, glycosphingosine accumulation occurs in

GD models and in patients with GD [38,39,40], and even if this does not seem to occur in _gba1__+/−_ neurons [40], we cannot exclude a contribution from changes in overall lipid metabolism.

Levels of ceramides, products of the GBA1 enzymatic activity, were not affected by the knockout, suggesting activation of compensatory pathways (Fig. 2e). Phosphatidylcholine (PC), lyso-PC,

and sphingomyelin were also unaffected (Fig. 2f–h), while phosphatidylethanolamine (PE) and phosphatidylserine (PS), which are important for mitochondrial function and for regulation of

autophagy [41,42,43,44], were both upregulated in _gba1__−/−_ brains compared with _gba1__+/+_ (Fig. 2i–l). PS was increased also in _gba1__+/−_ brains, further suggesting that this pathway

may contribute to the neuronal pathophysiology. LOW GLUTAMATE CONCENTRATIONS CAUSE LOSS OF MITOCHONDRIAL MEMBRANE POTENTIAL IN _GBA1_ +/− AND _GBA1_ _−/−_ NEURONS The [Ca2+]c increase in DCD

associated with glutamate excitotoxicity is closely coupled to collapse of Δψm and attributed to impaired [Ca2+]c homeostasis due to ATP depletion and the resultant failure of Ca2+

extrusion from the cytoplasm by Ca2+-H+ ATPases [29, 45], but this has not been demonstrated in a disease model. To further explore the relationship between mitochondrial (dys)function and

DCD, we used Rhodamine123, in ‘dequench mode’ [46], to study time-dependent changes in Δψm following exposure to glutamate. After an initial transient depolarization, coincident with the

initial [Ca2+]c response to glutamate, Δψm recovered almost to the baseline in the majority of _gba1__+/+_ neurons, while in _gba1__+/−_ and _gba1__−/−_ cells a large proportion of cells

showed a delayed collapse of Δψm (Fig. 3a). The secondary depolarization, quantified as the percentage change in the normalized Rhodamine123 signal at 400 s after glutamate stimulation (see

methods) was significantly different between genotypes (Fig. 3b): only 23% of the _gba1__+/+_ neurons presented at least 50% change of signal 400 s after 10 μM glutamate stimulation, while

an equivalent depolarization was seen in 42% of the _gba1__+/−_ and 43% of the _gba1__−/−_ neurons, closely resembling the distribution of the DCD responses. These measurements confirm the

close coupling between collapse of Δψm and DCD that we have previously described in other models [29]. INCREASED SENSITIVITY TO GLUTAMATE REFLECTS IMPAIRED CAPACITY TO MAINTAIN ATP

HOMEOSTASIS IN _GBA1_ +/− AND _GBA1_ _−/−_ NEURONS The appearance of DCD in response to low glutamate concentrations strongly resembles the glutamate response of neurons following inhibition

of oxidative phosphorylation by oligomycin [34, 45, 47] suggesting that glutamate-induced DCD reflects bioenergetic insufficiency in _gba1__+/−_ and _gba1__−/−_ neurons. We therefore

measured dynamic changes in neuronal ATP:ADP ratios in response to glutamate. Neurons were transfected with the ratiometric fluorescent probe PercevalHR, which reports changes in cytosolic

ATP:ADP [24]. Most neurons responded to glutamate with a rapid decrease in the ATP:ADP ratio (Fig. 4a), reported by the probe as a decrease in the ATP sensitive signal and an increase in the

ADP sensitive signal (Supplementary Fig. 4). The ATP:ADP ratio recovered over a few minutes in most control cells, but recovery was much slower or absent in the _gba1__−/−_ and to a lesser

extent in the _gba1__+/−_ neurons. Quantifying the ATP:ADP ratios before glutamate exposure, immediately after and 200 s after 10 µM glutamate stimulation (Fig. 4b–d), showed that there were

no significant differences between basal ATP:ADP ratios (before normalization) among the different genotypes (Fig. 4b), and the initial drop in ATP:ADP ratio was significantly higher in

_gba1__−/−_ neurons compared with wild-type (Fig. 4c). Moreover, recovery to the baseline was markedly impaired in both _gba1__+/−_ and _gba1__−/−_ cells (Fig. 4d). Interestingly, the same

experiments in control neurons in response to toxic concentrations of glutamate (100 μM) (Supplementary Fig. 5), showed an initial decrease in the ATP:ADP ratio, followed by a partial

recovery before undergoing a secondary decrease. This behavior was quite distinct from the responses seen in _gba1__+/−_ and _gba1__−/−_ neurons upon 10 μM glutamate stimulation. However,

when control neurons were treated first with 1 μM oligomycin, to inhibit oxidative phosphorylation, their responses to 10 μM glutamate resembled the responses of the _gba1__−/−_ neurons,

showing a decrease that failed to recover. These data further suggest that ATP depletion in _gba1__+/−_ and _gba1__−/−_ neurons in response to nontoxic glutamate concentrations is a

consequence of impaired mitochondrial function. These data suggest that even though energy homeostatic mechanisms maintain a normal ATP:ADP ratio at rest, the underlying loss of

mitochondrial bioenergetic capacity undermines the possibility to match ATP production to meet the increased energy demand following glutamate stimulation. MITOCHONDRIAL CALCIUM UPTAKE IS

REDUCED IN _GBA1_ _−/−_ AND IN _GBA1_ _+/−_ NEURONS The increased demand imposed on the cell by a [Ca2+]c signal may be matched by an increased energy supply driven by the upregulation of

the mitochondrial citric acid cycle in response to a rise in intramitochondrial Ca2+ concentration ([Ca2+]m) [48,49,50,51]. We therefore measured changes in [Ca2+]m in response to 10 μM

glutamate directly using mitochondria-targeted aequorin (Fig. 5a). Surprisingly, mitochondrial Ca2+ uptake was significantly reduced in both _gba1__−/−_ and _gba1__+/−_ neurons compared with

_gba1__+/+_. This is especially significant as the initial transient increase in cytosolic [Ca2+] in response to glutamate was increased in the _gba1__+/−_ and _gba1__−/−_ cells (see above,

Fig. 1b), consistent with impaired mitochondrial Ca2+ buffering. Resting cytosolic Ca2+ levels were not significantly different between populations, suggesting that the bioenergetic defect

was not severe enough to impair resting Ca2+ homeostasis (Supplementary Fig. 6). These findings may be attributable to the reduction in Δψm seen in _gba1__−/−_ and _gba1__+/−_ neurons [11].

However, we also explored the expression levels of the components of the mitochondrial Ca2+ uniporter complex, which may also contribute to altered mitochondrial Ca2+ uptake [17, 19,

52,53,54]. MCU, EMRE, MICU2, and MCUR1 protein levels were measured by western blot in brains from _gba1__+/+_, _gba1__+/−_, and _gba1__−/−_ mice (Fig. 5b). Quantification showed that MCU

expression was significantly reduced in the _gba1__+/−_ and _gba1__−/−_ cells, while expression levels of the associated regulatory proteins, EMRE, MICU2, and MCUR1 were not altered (_n_ = 

3–5 per genotype). Quantification of mRNA for MCU, EMRE, MICU1, and MCUR1 by qPCR (Supplementary Fig. 7) did not reveal any significant differences (_n_ = 3 per genotype), suggesting that

changes in MCU expression must be post transcriptionally regulated. RATES OF FREE RADICAL PRODUCTION ARE INCREASED IN _GBA1_ _−/−_ NEURONS Ca2+-dependent neuronal injury may be exacerbated

by the conjunction of raised [Ca2+]m with oxidative stress [23]. We therefore used dihydroethidium (DHE), a ratiometric fluorescent reporter sensitive to reactive oxygen species (ROS) to

determine whether basal rates of free radical production differ in _gba1__−/−_ cells (Fig. 6a) [55]. These data showed an increased basal rate of free radical production in _gba1__−/−_ cells

compared with the other genotypes (Fig. 6b). Exposure of cells to 10 μM glutamate caused a significant increase in the rate of ROS generation in each genotype, but the relative change was

not significantly different between _gba1__+/+_, _gba1__+/−_, and _gba1__−/−_ cultures (Fig. 6b) (_n_ = 3 independent experiments, _N_ = 10–20 cells per genotype per experiment). To address

the possibility that the increased rate of resting free radical production in _gba1__−/−_ was associated with impaired antioxidant defenses, we measured the expression of superoxide

dismutases (SODs) and glutathione (GSH) levels. Western blots of cytosolic SOD1 and mitochondrial SOD2 showed no difference between genotypes (_n_ = 5 per genotype) (Fig. 6c). GSH levels

were measured using monochlorobimane (MCB) as previously described [26]. MCB reacts with GSH, generating a fluorescent adduct, so a measure of steady-state MCB intensity gives a measure of

relative GSH content. MCB intensity in _gba1__+/+_, _gba1__+/−_, and _gba1__−/−_ showed no significant difference between genotypes (Fig. 6d—_n_ = 3 independent experiments, _N_ = 35–40

cells per genotype per experiment). DISCUSSION The goal of this study was to evaluate the functional consequences of mitochondrial dysfunction on cell physiology in neurons from the

transgenic _gba1__−/−_ mouse, a model for severe neuropathic GD, and from the _gba1__+/−_ mouse, which may illuminate mechanisms of neurodegeneration in GBA1-related PD. The functional

impact of impaired mitochondrial function is most dramatically revealed in neurons by an impaired capacity to respond to dynamic changes in metabolic demand, such as the increased energy

drain imposed by exposure to glutamate. Exposure of the cells to glutamate at concentrations that are innocuous for wild-type cells caused a profoundly dysregulated response in terms of

[Ca2+]c signaling and mitochondrial metabolism in the _gba1__−/−_ and, to a lesser extent, in the _gba1__+/−_ neurons. The _gba1__−/−_ mice suffer from an aggressive form of

neurodegeneration, and die only 2 weeks after birth [12], while _gba1__+/−_ mice do not show a disease-related phenotype. However, neurons from both _gba1__+/−_ and _gba1__−/−_ show a

significant decrease in Δψm and _gba1__−/−_ mixed cultures of neurons and astrocytes also show reduced basal respiratory activity and massively reduced maximal respiratory capacity [11]. We

found that neurons from both _gba1__−/−_ and _gba1__+/−_ mice showed abnormal responses to 10 μM glutamate, characterized as DCD, which is normally associated with ‘excitotoxicity’ in

response to much higher concentrations of glutamate. We attribute this vulnerability primarily to the decreased bioenergetic capacity, which is especially severe in the _gba1__−/−_ neurons

[11], and therefore to the failure to maintain ATP homeostasis in the face of increased energy demand imposed by glutamate. The decreased mitochondrial Ca2+ uptake that we have shown in both

_gba1__−/−_ and _gba1__+/−_ neurons, will likely contribute to this energetic failure, as it will limit the capacity of the mitochondria to increase ATP production in response to [Ca2+]c

signals. The observed reduction in mitochondrial Ca2+ uptake is attributable to the reduced Δψm but also to the reduced expression of the MCU protein, which seems to be associated with

changes in protein turnover rather than to transcriptional repression. The small reduction of _Grin2b_ mRNA in both _gba1__+/−_ and _gba1__−/−_ brains may represent another compensatory

mechanism, but was not reflected in changes in Grin2b protein expression, or in the localization of Grin2b at the plasma membrane. Thus, changes in Grin2b expression or localization are

unlikely to be responsible for DCD. Furthermore, responses of _gba1__−/−_ neurons to release of ER Ca2+ by metabotropic receptor activation showed a modest reduction, suggesting that the key

Ca2+ source that triggers DCD is delivered through ionotropic glutamate receptors. The response to the Ca2+ influx is likely compounded by an increased rate of ROS generation in _gba1__−/−_

neurons under basal conditions, as glutamate toxicity is exacerbated by oxidative stress [56]. Interestingly, we have found a marked cellular phenotype in _gba1__+/−_ neurons. We previously

showed, that Δψm is reduced in _gba1__+/−_ neurons [11], although defects in respiratory capacity were less severe than in homozygotes. These differences, together with the difference in

oxidative stress, that was increased only in _gba1__−/−_ neurons, may contribute to the difference of disease phenotype in the _gba1__−/−_ and _gba1__+/−_ mice. In agreement with our data, a

mouse model carrying the heterozygous _GBA1_ PD-associated mutation L444P was shown to have defective mitochondria, supporting a role of impaired bioenergetics in GBA1-associated PD [57].

Dopaminergic neurons at risk of neurodegeneration in PD are physiologically characterized by Ca2+-dependent pace-making activity, while intrinsic Ca2+ buffering capacity is reduced [22],

imposing a major energy demand, which will be amplified by mechanisms that compromise bioenergetic reserve, putting these cells especially at risk [58]. Since we have shown that partial

depletion of GBA1 in _gba1__+/−_ neurons sensitizes neurons to Ca2+ influx and show DCD in response to physiological glutamate concentrations, we suggest that the compromised mitochondrial

function in these cells may increase the risk of neurodegeneration in neurons that are already vulnerable because of their normal physiological activity. It is notable that both _gba1__−/−_

and _gba1__+/−_ neurons showed similar responses to glutamate and impaired [Ca2+]c handling, suggesting that it is unlikely that the metabolic and signaling defects are simply attributable

to the massive accumulation of the GBA1 substrate glucosylceramide, observed in _gba1__−/−_, which was evident despite the presence of galactosylceramides. However, subtle changes of

glucosylceramide levels in _gba1__+/−_ brains may have been masked by the higher levels of galactosylceramides in the mixed cells. Considering this and the deregulation observed in PE and PS

levels in _gba1__+/−_ and _gba1__−/−_ brains, broader evaluation of lipid homeostasis may help in understanding the pathophysiological mechanisms that couple reduced GBA1 to mitochondrial

dysfunction. Overall, our findings suggest that _gba1__−/−_ but also _gba1__+/−_ neurons are sensitive to dynamic changes in energy demand caused by an imposed workload, while under basal

conditions, both ATP and cytosolic Ca2+ levels in _gba1__−/−_ and _gba1__+/−_ neurons were not different from the control. This implies the activity of a vicious cycle in which every

mechanism enlisted to compensate a homeostatic Ca2+ stress, an increased energy demand and oxidative stress cause further deterioration of cell bioenergetic capacity, triggering a

pathological cascade. Our data highlight a general principle—that in any disease (from age-related neurodegenerative diseases to lysosomal storage disorders) in which mitochondrial

bioenergetic capacity is impaired, neurons will become more vulnerable to increased energy demand, which may be sufficient to initiate dysregulated calcium signaling and cell death. CHANGE

HISTORY * _ 09 MARCH 2020 A Correction to this paper has been published: https://doi.org/10.1038/s41418-020-0525-0 _ REFERENCES * Brady RO, Kanfer J, Shapiro D. The metabolism of

glucocerebrosides. I. Purification and properties of a glucocerebroside-cleaving enzyme from spleen tissue. J Biol Chem. 1965;240:39–43. Article  CAS  PubMed  Google Scholar  * Smith L,

Mullin S, Schapira AHV. Insights into the structural biology of Gaucher disease. Exp Neurol. 2017;298:180–90. Article  CAS  PubMed  Google Scholar  * Sidransky E, Nalls MA, Aasly JO,

Aharon-Peretz J, Annesi G, Barbosa ER, et al. Multicenter analysis of glucocerebrosidase mutations in Parkinson’s disease. N Engl J Med. 2009;361:1651–61. Article  CAS  PubMed  PubMed

Central  Google Scholar  * Westbroek W, Gustafson AM, Sidransky E. Exploring the link between glucocerebrosidase mutations and parkinsonism. Trends Mol Med. 2011;17:485–93. Article  CAS 

PubMed  PubMed Central  Google Scholar  * Lesage S, Anheim M, Condroyer C, Pollak P, Durif F, Dupuits C, et al. Large-scale screening of the Gaucher’s disease-related glucocerebrosidase gene

in Europeans with Parkinson’s disease. Hum Mol Genet. 2011;20:202–10. Article  CAS  PubMed  Google Scholar  * Grabowski GA. Phenotype, diagnosis, and treatment of Gaucher’s disease. Lancet.

2008;372:1263–71. Article  CAS  PubMed  Google Scholar  * Malini E, Grossi S, Deganuto M, Rosano C, Parini R, Dominisini S, et al. Functional analysis of 11 novel GBA alleles. Eur J Hum

Genet. 2014;22:511–6. Article  CAS  PubMed  Google Scholar  * Plotegher N, Duchen MR. Mitochondrial dysfunction and neurodegeneration in lysosomal storage disorders. Trends Mol Med.

2017;23:116–34. Article  CAS  PubMed  Google Scholar  * Plotegher N, Duchen MR. Crosstalk between lysosomes and mitochondria in Parkinson’s disease. Front Cell Dev Biol. 2017;5:2011–8.

Article  Google Scholar  * Raimundo N, Fernández-Mosquera L, Yambire KF, Diogo CV. Mechanisms of communication between mitochondria and lysosomes. Int J Biochem Cell Biol. 2016;79:345–9.

Article  CAS  PubMed  Google Scholar  * Osellame LD, Rahim Aa, Hargreaves IP, Gegg ME, Richard-londt A, Brandner S, et al. Mitochondria and quality control defects in a mouse model of

gaucher disease - links to Parkinson’s disease. Cell Metab. 2013;17:941–53. Article  CAS  PubMed  PubMed Central  Google Scholar  * Enquist IB, Lo Bianco C, Ooka A, Nilsson E, Månsson J-E,

Ehinger M, et al. Murine models of acute neuronopathic Gaucher disease. Proc Natl Acad Sci USA. 2007;104:17483–8. Article  CAS  PubMed  PubMed Central  Google Scholar  * Rizzuto R, De

Stefani D, Raffaello A, Mammucari C. Mitochondria as sensors and regulators of calcium signalling. Nat Rev Mol Cell Biol. 2012;13:566–78. Article  CAS  PubMed  Google Scholar  * Nicholls DG.

Mitochondrial function and dysfunction in the cell: Its relevance to aging and aging-related disease. Int J Biochem Cell Biol. 2002;34:1372–81. Article  CAS  PubMed  Google Scholar  *

Duchen MR. Mitochondria, calcium-dependent neuronal death and neurodegenerative disease. Pflügers Arch - Eur J Physiol. 2012;464:111–21. Article  CAS  Google Scholar  * De Stefani D,

Raffaello A, Teardo E, Szabo I, Rizzuto R. A forty-kilodalton protein of the inner membrane is the mitochondrial calcium uniporter. Nature. 2011;476:336–40. Article  PubMed  PubMed Central 

CAS  Google Scholar  * Kamer KJ, Mootha VK. MICU1 and MICU2 play nonredundant roles in the regulation of the mitochondrial calcium uniporter. EMBO Rep. 2014;15:299–307. Article  CAS  PubMed

  PubMed Central  Google Scholar  * Konig T, Troder SE, Bakka K, Korwitz A, Richter-Dennerlein R, Lampe PA, et al. The m-AAA protease associated with neurodegeneration limits MCU activity in

mitochondria. Mol Cell. 2016;64:148–62. Article  PubMed  CAS  Google Scholar  * Mallilankaraman K, Cardenas C, Doonan PJ, Chandramoorthy HC, Irrinki KM, Golenar T, et al. MCUR1 is an

essential component of mitochondrial Ca2+ uptake that regulates cellular metabolism. Nat Cell Biol. 2012;14:1336–43. Article  CAS  PubMed  PubMed Central  Google Scholar  * Palty R,

Silverman WF, Hershfinkel M, Caporale T, Sensi SL, Parnis J, et al. NCLX is an essential component of mitochondrial Na+/Ca2+exchange. Proc Natl Acad Sci. 2010;107:436–41. Article  CAS 

PubMed  Google Scholar  * Burbulla LF, Song P, Mazzulli JR, Zampese E, Wong YC, Jeon S, et al. Dopamine oxidation mediates mitochondrial and lysosomal dysfunction in Parkinson’s disease.

Science (80-). 2017;357:1255–61. Article  CAS  Google Scholar  * Surmeier DJ, Schumacker PT. Calcium, bioenergetics, and neuronal vulnerability in Parkinson’s disease. J Biol Chem.

2013;288:10736–41. Article  CAS  PubMed  Google Scholar  * Reyes RC, Brennan AM, Shen Y, Baldwin Y, Swanson RA. Activation of neuronal NMDA receptors induces superoxide-mediated oxidative

stress in neighboring neurons and astrocytes. J Neurosci. 2012;32:12973–8. Article  CAS  PubMed  PubMed Central  Google Scholar  * Tantama M, Martínez-François JR, Mongeon R, Yellen G.

Imaging energy status in live cells with a fluorescent biosensor of the intracellular ATP-to-ADP ratio. _Nat Commun_ 2013; 4. https://doi.org/10.1038/ncomms3550. * Bonora M, Giorgi C, Bononi

A, Marchi S, Patergnani S, Rimessi A, et al. Subcellular calcium measurements in mammalian cells using jellyfish photoprotein aequorin-based probes. Nat Protoc. 2013;8:2105–18. Article  CAS

  PubMed  Google Scholar  * Keelan J, Allen NJ, Antcliffe D, Pal S, Duchen MR. Quantitative imaging of glutathione in hippocampal neurons and glia in culture using monochlorobimane. J

Neurosci Res. 2001;66:873–84. Article  CAS  PubMed  Google Scholar  * Ferrazza R, Cogo S, Melrose H, Bubacco L, Greggio E, Guella G, et al. LRRK2 deficiency impacts ceramide metabolism in

brain. Biochem Biophys Res Commun. 2016;478:1141–6. Article  CAS  PubMed  Google Scholar  * Choi DW. Glutamate neurotoxicity in cortical cell culture is calcium dependent. Neurosci Lett.

1985;58:293–7. Article  CAS  PubMed  Google Scholar  * Abramov AY, Duchen MR. Mechanisms underlying the loss of mitochondrial membrane potential in glutamate excitotoxicity. Biochim Biophys

Acta Bioenerg. 2008;1777:953–64. Article  CAS  Google Scholar  * Moussawi K, Riegel A, Nair S, Kalivas PW. Extracellular glutamate: functional compartments operate in different concentration

ranges. Front Syst Neurosci. 2011;5:1–9. Article  CAS  Google Scholar  * Thio L, Clifford D, Zorumski C. Characterization of quisqualate receptor desensitization in cultured postnatal rat

hippocampal neurons. J Neurosci. 1991;11:3430–41. Article  CAS  PubMed  PubMed Central  Google Scholar  * Klein AD, Ferreira NS, Ben-Dor S, Duan J, Hardy J, Cox TM, et al. Identification of

modifier genes in a mouse model of gaucher disease. Cell Rep. 2016;16:2546–53. Article  CAS  PubMed  Google Scholar  * Pérez-Otaño I, Schulteis CT, Contractor A, Lipton SA, Trimmer JS,

Sucher NJ, et al. Assembly with the NR1 subunit is required for surface expression of NR3A-containing NMDA receptors. J Neurosci. 2001;21:1228–37. Article  PubMed  PubMed Central  Google

Scholar  * Vergun O, Keelan J, Khodorov BI, Duchen MR. Glutamate-induced mitochondrial depolarisation and perturbation of calcium homeostasis in cultured rat hippocampal neurones. J Physiol.

1999;519:451–66. Article  CAS  PubMed  PubMed Central  Google Scholar  * Tymianski M, Charlton MP, Carlen PL, Tator CH. Source specificity of early calcium neurotoxicity embryonic spinal

neurons in cultured. J Neurosci. 1993;13:2085–104. Article  CAS  PubMed  PubMed Central  Google Scholar  * Schöndorf DC, Aureli M, McAllister FE, Hindley CJ, Mayer F, Schmid B, et al.

iPSC-derived neurons from GBA1-associated Parkinson’s disease patients show autophagic defects and impaired calcium homeostasis. Nat Commun. 2014;5:1–17. Article  CAS  Google Scholar  *

Mazzulli JR, Xu YH, Sun Y, Knight AL, McLean PJ, Caldwell GA, et al. Gaucher disease glucocerebrosidase and α-synuclein form a bidirectional pathogenic loop in synucleinopathies. Cell.

2011;146:37–52. Article  CAS  PubMed  PubMed Central  Google Scholar  * Schöndorf DC, Ivanyuk D, Baden P, Sanchez-Martinez A, De Cicco S, Yu C, et al. The NAD+ precursor nicotinamide

riboside rescues mitochondrial defects and neuronal loss in iPSC and fly models of Parkinson’s disease. Cell Rep. 2018;23:2976–88. Article  PubMed  CAS  Google Scholar  * Dekker N, van

Dussen L, Hollak CEM, Overkleeft H, Scheij S, Ghauharali K, et al. Elevated plasma glucosylsphingosine in Gaucher disease: relation to phenotype, storage cell markers, and therapeutic

response. Blood. 2011;118:e118–e127. Article  PubMed  PubMed Central  Google Scholar  * Taguchi YV, Liu J, Ruan J, Pacheco J, Zhang X, Abbasi J et al. Glucosylsphingosine promotes

α-synuclein pathology in mutant GBA-associated Parkinson’s disease. J Neurosci. 2017;37:9617–31. * Van Der Veen JN, Lingrell S, Da Silva RP, Jacobs RL, Vance DE. The concentration of

phosphatidylethanolamine in mitochondria can modulate ATP production and glucose metabolism in mice. Diabetes. 2014;63:2620–30. Article  PubMed  CAS  Google Scholar  * Rockenfeller P, Koska

M, Pietrocola F, Minois N, Knittelfelder O, Sica V, et al. Phosphatidylethanolamine positively regulates autophagy and longevity. Cell Death Differ. 2015;22:499–508. Article  CAS  PubMed 

PubMed Central  Google Scholar  * Hsu P, Shi Y. Regulation of autophagy by mitochondrial phospholipids in health and diseases. Biochim Biophys Acta Mol Cell Biol Lipids. 2017;1862:114–29.

Article  CAS  PubMed  Google Scholar  * Steenbergen R, Nanowski TS, Beigneux A, Kulinski A, Young SG, Vance JE. Disruption of the phosphatidylserine decarboxylase gene in mice causes

embryonic lethality and mitochondrial defects. J Biol Chem. 2005;280:40032–40. Article  CAS  PubMed  Google Scholar  * Khodorov B, Pinelis V, Vergun O, Storozhevykh T, Vinskaya N.

Mitochondrial deenergization underlies neuronal calcium overload following a prolonged glutamate challenge. FEBS Lett. 1996;397:230–4. Article  CAS  PubMed  Google Scholar  * Duchen MR,

Surin A, Jacobson J. Imaging mitochondrial function in intact cells. Methods Enzymol. 2003;361:353–89. Article  CAS  PubMed  Google Scholar  * Abramov AY, Duchen MR. Impaired mitochondrial

bioenergetics determines glutamate-induced delayed calcium deregulation in neurons. Biochim Biophys Acta Gen Subj. 2010;1800:297–304. Article  CAS  Google Scholar  * Duchen MR.

Ca2+-dependent changes in the mitochondrial energetics in single dissociated mouse sensory neurons. Biochem J. 1992;283:41–50. Article  CAS  PubMed  PubMed Central  Google Scholar  * Pardo

B, Contreras L, Serrano A, Ramos M, Kobayashi K, Iijima M, et al. Essential role of aralar in the transduction of small Ca2+ signals to neuronal mitochondria. J Biol Chem. 2006;281:1039–47.

Article  CAS  PubMed  Google Scholar  * McCormack JG, Halestrap AP, Denton RM. Role of calcium ions in regulation of mammalian intramitochondrial metabolism. Physiol Rev. 1990;70:391–425.

Article  CAS  PubMed  Google Scholar  * Jouaville LS, Pinton P, Bastianutto C, Rutter GA, Rizzuto R. Regulation of mitochondrial ATP synthesis by calcium: evidence for a long-term metabolic

priming. Proc Natl Acad Sci USA. 1999;96:13807–12. Article  CAS  PubMed  PubMed Central  Google Scholar  * De Stefani D, Rizzuto R. Molecular control of mitochondrial calcium uptake. Biochem

Biophys Res Commun. 2014;449:373–6. Article  PubMed  CAS  Google Scholar  * Logan CV, Szabadkai G, Sharpe JA, Parry DA, Torelli S, Childs AM, et al. Loss-of-function mutations in MICU1

cause a brain and muscle disorder linked to primary alterations in mitochondrial calcium signaling. Nat Genet. 2014;46:188–93. Article  CAS  PubMed  Google Scholar  * Vais H, Mallilankaraman

K, Mak D-OD, Hoff H, Payne R, Tanis JE, et al. EMRE is a matrix Ca2+ sensor that governs gatekeeping of the mitochondrial Ca2+ uniporter. Cell Rep. 2016;14:403–10. Article  CAS  PubMed 

PubMed Central  Google Scholar  * Chouchani ET, Pell VR, Gaude E, Aksentijević D, Sundier SY, Robb EL, et al. Ischaemic accumulation of succinate controls reperfusion injury through

mitochondrial ROS. Nature. 2014;515:431–5. Article  CAS  PubMed  PubMed Central  Google Scholar  * Forder JP, Tymianski M. Postsynaptic mechanisms of excitotoxicity: Involvement of

postsynaptic density proteins, radicals, and oxidant molecules. Neuroscience. 2009;158:293–300. Article  CAS  PubMed  Google Scholar  * Yun SP, Kim D, Kim S, Kim S, Karuppagounder SS, Kwon

SH, et al. α-Synuclein accumulation and GBA deficiency due to L444P GBA mutation contributes to MPTP-induced parkinsonism. Mol Neurodegener. 2018;13:1–19. Article  CAS  PubMed  PubMed

Central  Google Scholar  * Rivero-Ríos P, Gómez-Suaga P, Fdez E, Hilfiker S. Upstream deregulation of calcium signaling in Parkinson’s disease. Front Mol Neurosci. 2014;7:53. PubMed  PubMed

Central  Google Scholar  Download references ACKNOWLEDGEMENTS We would like to thank Prof. Stefan Karlsson for providing us with the GBA1 knockout mouse model. NP was supported by Marie

Sklodowska-Curie Individual Fellowship (Horizon 2020 Grant No. 653434), FZ and MD were supported by Michael J. Fox Foundation Target Validation Grant (Grant ID 12159) and GB and MD by the

GSK/BBSRC CASE PhD studentship (BB/L502145/1). MD was also supported by BBSRC BB/P018726/1. AAR and SNW were part-funded by the UK Medical Research Council grants (G1000709 and MR/N026101/1)

and GM received support from the UK Gauchers Association. SNW received part funding from UK Medical Research Council grants MR/R015325/1, MR/P026494/1 and MR/N019075/1 and from SPARKS

(17UCL01). AAR is also funded by UK Medical Research Council Grants (MR/R015325/1, MR/S009434/1, MR/N026101/1 and MR/S036784/1), Action Medical Research (GN2485) and the Wellcome Trust

Institutional Strategic Support Fund/UCL Therapeutic Acceleration Support (TAS) Fund (ISSF3/H17RCO/TAS004). AUTHOR INFORMATION Author notes * Nicoletta Plotegher Present address: Department

of Biology, University of Padua, 35131, Padua, Italy AUTHORS AND AFFILIATIONS * Cell and Developmental Biology Department, University College London, London, WC1E6XA, UK Nicoletta Plotegher,

 Gauri Bhosale, Federico Zambon, Gyorgy Szabadkai & Michael R. Duchen * Institute for Women’s Health, University College London, London, WC1E6HU, UK Dany Perocheau & Simon N.

Waddington * Department of Physics, University of Trento, 38123, Povo (TN), Italy Ruggero Ferrazza & Graziano Guella * School of Pharmacy, University College London, London, WC1N1AX, UK

Giulia Massaro & Ahad A. Rahim * MRC Antiviral Gene Therapy Research Unit, Faculty of Health Sciences, University of the Witwatersrand, Johannesburg, South Africa Simon N. Waddington *

Department of Biomedical Sciences, University of Padua, 35131, Padua, Italy Gyorgy Szabadkai * The Francis Crick Institute, London, NW1 1AT, UK Gyorgy Szabadkai Authors * Nicoletta Plotegher

View author publications You can also search for this author inPubMed Google Scholar * Dany Perocheau View author publications You can also search for this author inPubMed Google Scholar *

Ruggero Ferrazza View author publications You can also search for this author inPubMed Google Scholar * Giulia Massaro View author publications You can also search for this author inPubMed 

Google Scholar * Gauri Bhosale View author publications You can also search for this author inPubMed Google Scholar * Federico Zambon View author publications You can also search for this

author inPubMed Google Scholar * Ahad A. Rahim View author publications You can also search for this author inPubMed Google Scholar * Graziano Guella View author publications You can also

search for this author inPubMed Google Scholar * Simon N. Waddington View author publications You can also search for this author inPubMed Google Scholar * Gyorgy Szabadkai View author

publications You can also search for this author inPubMed Google Scholar * Michael R. Duchen View author publications You can also search for this author inPubMed Google Scholar

CORRESPONDING AUTHOR Correspondence to Michael R. Duchen. ETHICS DECLARATIONS CONFLICT OF INTEREST The authors declare that they have no conflict of interest. ADDITIONAL INFORMATION

PUBLISHER’S NOTE Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. Edited by N. Bazan SUPPLEMENTARY INFORMATION AUTHOR

CONTRIBUTION TO THE ARTICLE RIGHTS AND PERMISSIONS This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation,

distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and

indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit

line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use,

you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/. Reprints and permissions ABOUT THIS

ARTICLE CITE THIS ARTICLE Plotegher, N., Perocheau, D., Ferrazza, R. _et al._ Impaired cellular bioenergetics caused by GBA1 depletion sensitizes neurons to calcium overload. _Cell Death

Differ_ 27, 1588–1603 (2020). https://doi.org/10.1038/s41418-019-0442-2 Download citation * Received: 05 February 2019 * Revised: 09 October 2019 * Accepted: 10 October 2019 * Published: 04

November 2019 * Issue Date: 01 May 2020 * DOI: https://doi.org/10.1038/s41418-019-0442-2 SHARE THIS ARTICLE Anyone you share the following link with will be able to read this content: Get

shareable link Sorry, a shareable link is not currently available for this article. Copy to clipboard Provided by the Springer Nature SharedIt content-sharing initiative