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Article

Pridopidine Protects ALS Patient-Derived Neural Progenitor Cells via Sigma-1 Receptor Activation

by
May Meltzer
1,*,
Maya Shefler Zamir
2,
Noam Tzuri
2,
Andrew M. Tan
1,3,
Michal Geva
1,
Michael R. Hayden
1,4,5 and
Rachel G. Lichtenstein
2,*
1
Prilenia Therapeutics B.V., 1411 DC Naarden, The Netherlands
2
Avram and Stella Goren-Goldstein Department of Biotechnology Engineering, Faculty of Engineering, Ben-Gurion University of the Negev, P.O. Box 653, Beer-Sheva 84105, Israel
3
Department of Neurology, Yale University School of Medicine, New Haven, CT 06510, USA
4
Regenerative Medicine and Stem Cell (RMSC) Research Center, Ben-Gurion University of the Negev, P.O. Box 653, Beer-Sheva 84105, Israel
5
Department of Medical Genetics, Centre for Molecular Medicine and Therapeutics, University of British Columbia, Vancouver, BC V5Z 4H4, Canada
*
Authors to whom correspondence should be addressed.
Int. J. Mol. Sci. 2026, 27(8), 3489; https://doi.org/10.3390/ijms27083489
Submission received: 4 February 2026 / Revised: 30 March 2026 / Accepted: 7 April 2026 / Published: 14 April 2026
(This article belongs to the Special Issue Sigma-1 Receptor: Signaling, Functions and Therapeutic Potential)
Browse Figures
Versions Notes

Abstract

The sigma-1 receptor (S1R) is an endoplasmic reticulum (ER)-resident protein enriched at the mitochondria-associated ER membranes (MAMs) that supports ER homeostasis, preserves mitochondrial function, and enhances cell survival under stress. Disruptions of MAM integrity and prolonged ER stress are well-recognized pathological features of amyotrophic lateral sclerosis (ALS), contributing to motor neuron dysfunction and degeneration. In this study, we evaluated the protective effects of pridopidine, a highly selective and potent S1R agonist currently in clinical development for Huntington’s disease (HD) and ALS, using neural progenitor cells (NPCs) derived from induced pluripotent stem cells (iPSCs) from a patient with sporadic ALS. Exposure of ALS NPCs to the ER stressor tunicamycin increased the ER stress markers binding immunoglobulin protein (BiP) and C/EBP homologous protein (CHOP), disrupted mitochondrial membrane potential, upregulated expression of the mitochondrial apoptotic marker, BAX, increased caspase-3 activation, and reduced cell viability. Pridopidine significantly attenuated tunicamycin-induced BiP and CHOP expression in a biphasic, dose-dependent manner (with maximal efficacy at 1 µM), consistent with the typical pharmacology of S1R agonists. Pridopidine restored mitochondrial membrane potential, reduced mitochondrial apoptotic signaling, shown by decreased BAX expression and caspase-3 activation, and improved survival of ALS-NPCs under ER stress. Co-treatment with the selective S1R antagonist, NE-100, attenuated these effects, supporting an S1R-mediated mechanism of action for pridopidine. Together, these results demonstrate that S1R activation by pridopidine mitigates ER-stress-induced mitochondrial dysfunction and cell loss in ALS-NPCs, resulting in enhanced survival of NPCs supporting the therapeutic potential of pridopidine in ALS.

1. Introduction

ALS is a progressive and fatal neurodegenerative disorder characterized by selective loss of motor neurons, leading to muscle atrophy, paralysis, and respiratory failure, with death often occurring within 3–5 years of symptom onset [1,2,3]. Current treatment options are either limited to a small and selective target population of SOD1 mutation carriers [4] or show limited benefits and do not alter disease progression [5,6]. Thus, there is a critical unmet need for effective disease-modifying therapies [7,8,9,10].
Sporadic and SOD1-mutant ALS types converge on shared cellular pathways that underline motor neuron degeneration. These forms of ALS share key cellular pathologies, including impaired ER–mitochondria crosstalk at the MAMs, prolonged ER stress, formation of toxic protein aggregates, mitochondrial dysfunction, and progressive loss of motor neurons in the spinal cord and cortex [3,11,12,13,14].
Among these processes associated with ALS, ER stress has emerged as one of the earliest pathogenic mechanisms [15,16,17,18]. Increased expression of ER stress markers, including the chaperone BiP/GRP78 and the pro-apoptotic transcription factor CHOP, has been consistently observed in ALS patient tissues and preclinical models [16,19,20,21]. Prolonged ER stress also contributes to neuronal loss in ALS by activating apoptotic signaling, including upregulation of the pro-apoptotic mediator, BAX, and downstream activation of caspase-3 [20,22].
Importantly, the S1R plays a central role in stabilizing MAMs integrity and regulating ER stress responses [19,23,24]. In healthy unstressed cells, the S1R is bound to BiP at the ER membrane in an inactive manner. Upon ER calcium depletion and elevated ER stress, or agonist binding, the S1R dissociates from BiP and interacts with other cellular proteins to reduce stress and enhance cell survival [25,26]. S1R interacts and stabilizes the inositol 1,4,5-trisphosphate receptors (IP3Rs), responsible for Ca2+ transfer from ER stores to the mitochondria, normalizing calcium homeostasis, reducing ER stress and enhancing mitochondrial function [27,28,29].
Genetic evidence supports the role of S1R in ALS. For example, complete loss-of-function mutations in SIGMAR1 cause juvenile ALS and distal hereditary motor neuropathies, while partial loss-of-function variants increase risk for adult-onset ALS. Consistent with these genetic findings, pharmacological S1R activation mitigates ER stress, enhances mitochondrial function, stabilizes calcium homeostasis, and promotes neuronal survival across multiple neurodegeneration models, including ALS, HD, Parkinson’s disease (PD), and Alzheimer’s disease (AD) [3,30,31,32,33,34].
In both in vitro and in vivo preclinical ALS models, pridopidine, via activation of the S1R, restores axonal transport, preserves neuromuscular junction integrity, reduces toxic mutant SOD1 aggregation, and improves motor neuron survival [35,36,37]. Beyond ALS, pridopidine stabilizes dendritic spines and enhances synaptic plasticity in association with improved cognitive or motor outcomes in HD, PD and AD models. These broad neuroprotective actions are consistent with S1R’s role as a key regulator of ER–mitochondria crosstalk, calcium homeostasis, mitochondrial function, and cellular stress responses [37,38,39,40,41].
A well-established fundamental property of S1R agonist activity is the biphasic dose–response curve, extensively demonstrated in preclinical models and in human studies [41,42]. Evidence for this dose–response characteristic of S1R agonists comes from various preclinical assays, including amelioration of cognitive impairment [43,44,45,46,47,48,49,50], electrophysiological measure of N-methyl-d-aspartic acid (NMDA)-induced neuronal activation in rat brains [51,52,53], potentiation of cellular calcium efflux [54,55], and anti-depressive effects in mice [56]. Pridopidine demonstrated a biphasic dose response in several nonclinical studies. Pridopidine restored the impaired homeostatic synaptic plasticity in HD cortical neurons with the strongest effect achieved at a concentration of 1 μM, while higher and lower doses showed diminished effects [57]. In the 6-OHDA PD mouse model, a low dose of pridopidine, but not a high dose, increased neuroprotection of dopaminergic neurons and restored behavioral abnormalities [58].
In this study, we investigated the cellular effects of pridopidine under ER stress conditions using NPCs derived from a sporadic ALS patient. Specifically, we evaluated whether pridopidine reduces ER stress by examining BiP and CHOP expression; restores mitochondrial membrane potential and reduces mitochondrial apoptotic signaling as measured by BAX expression and caspase-3 activation; and improves cell viability.

2. Results

2.1. Differentiation of Sporadic ALS Patient iPSCs into Neural Progenitor Cells

We used the iPSC-derived NPC model to evaluate pridopidine’s neuroprotective potential against ER stress in ALS. The sporadic CS14isALS-Tnxx iPSC line was differentiated into NPCs using published methods [59,60]. After 6 days in culture, either Basic fibroblast growth factor (bFGF) or Epidermal growth factor (EGF) was added to ensure progenitor self-renewal and maintain NPC phenotype [61] (Figure 1A).
Immunofluorescence further demonstrated characteristic NPC morphology, including elongated cell bodies, neurite-like projections, and dense intercellular networks—features clearly distinct from the compact, colony-like organization of undifferentiated iPSCs at day 0 (Figure 1B). These molecular and morphological features collectively support a consistent neuronal progenitor phenotype.
The NPCs kept their proliferation capacity; when they reached 90% confluence, the cells were dissociated and reseeded onto new plates. qPCR analysis confirmed a sustained proliferative capacity, comparable to that observed in pluripotent cells, as indicated by MKI67 expression (Figure 1C).
To validate that this treatment indeed preserved NPC identity, we assessed the expression of canonical progenitor markers. Flow cytometry, qPCR, and immunofluorescence confirmed sustained SOX2 and PAX6 expression, together with early neuronal marker TUJ1 (β-III tubulin), indicating retention of a stable NPC phenotype under these conditions (Figure 1D–F).
The neural induction protocol efficiently generated day-6 and day-13 NPCs (Figure 1E,F), with expression of neural markers PAX6 (52.97% ± 1.5% at day 6; 23.94% ± 0.7% at day 13), SOX2 (81.10% ± 2.3%), and TUJ1 (81.52% ± 2.6%). Together, these cytometric and morphological data demonstrate a robust and reproducible derivation of NPCs from the ALS iPSC line.

2.2. Pridopidine Reduces ER Stress Markers BiP and CHOP in ALS iPSC-Derived NPCs

Pridopidine was evaluated for its ability to attenuate ER stress in sporadic ALS patient-derived neural progenitor cells (ALS-NPCs). Cells were exposed to tunicamycin (1.25 µM, 16 h), and the expression of the ER stress markers BiP and CHOP was quantified by flow cytometry.
Tunicamycin exposure markedly increased the proportion of ALS-NPCs with elevated BiP expression by 47% compared with unstressed controls (p < 0.0001). Pridopidine significantly reduced the percentage of BiP-positive cells, with a clear biphasic dose response. The maximal effect of pridopidine was observed at 1 µM, with a 57% reduction in BiP-positive cells vs. tunicamycin-treated controls (p < 0.0001). Lower concentrations (0.1 and 0.5 µM) also produced significant decreases in BiP-positive cells (25%, p < 0.0001 and 42%, p < 0.000, respectively), whereas higher concentrations (5–50 µM) yielded progressively smaller effects (12–30% reduction), consistent with the characteristic biphasic response of S1R ligands (Figure 2A).
A similar pattern was observed for the effect of pridopidine on CHOP, a pro-apoptotic marker of prolonged ER stress. Tunicamycin increased CHOP-positive cells by 60% compared with unstressed controls (p < 0.0001). Pridopidine reduced CHOP expression in a biphasic manner, with the most pronounced effect at 1 µM pridopidine (50% reduction vs. tunicamycin-treated cells, p < 0.0001). Doses of 0.1 and 0.5 µM produced smaller but significant reductions (22%, p < 0.01 and 33% p < 0.001), while higher doses (5–50 µM) again showed progressively smaller effects (16–31% reduction), mirroring the S1R-dependent biphasic profile (Figure 2B).
Together, these findings demonstrate that pridopidine robustly attenuates tunicamycin-induced ER stress in ALS-NPCs, exhibiting the expected peak efficacy at 1 µM with reduced activity at lower and higher concentrations.

2.3. Pridopidine’s Effect on ER Stress Markers Is Mediated by Sigma-1 Receptor Activation

To determine whether the protective effects of pridopidine depend on S1R activation, ALS-NPCs were treated with tunicamycin (as before) (1.25 µM, 16 h), pridopidine (0.5 µM) and the selective S1R antagonist NE-100 (1 µM). BiP and CHOP expression levels were assessed by flow cytometry and immunocytochemistry as two complementary methodologies.
Tunicamycin markedly increased BiP expression by 73% compared with unstressed controls (p < 0.0001) (in line with the normal variability between independent experiments (47% in Figure 2). Pridopidine significantly reduced the proportion of BiP-positive cells by 42% (p < 0.001) relative to stressed, untreated cultures. Importantly, co-treatment with the selective S1R antagonist NE-100 robustly attenuated this effect, resulting in a non-significant and modest 17% reduction in BiP-positive cells (Figure 3A). Immunofluorescence confirmed these findings: tunicamycin induced a strong increase in BiP staining, which was markedly reduced by pridopidine (48% reduction, p < 0.001), and attenuated by NE-100 administration (non-significant, 12% reduction) (Figure 3C,D).
Pridopidine showed a parallel effect on CHOP. Tunicamycin increased CHOP-positive cells by 68% compared with unstressed controls (p < 0.0001) (Figure 3B). Pridopidine decreased CHOP-positive cells by 38% (p < 0.0001), an effect antagonized by NE-100 co-treatment, yielding a non-significant 8% reduction as compared with tunicamycin alone. Consistent with the flow cytometry results, immunocytochemistry demonstrated robust CHOP induction following tunicamycin exposure, whereas it was markedly reduced by pridopidine (41% reduction, p < 0.001) but blocked with NE-100 (6% reduction) (Figure 3E,F).
Together, these findings from two different ER stress markers show that pridopidine’s effects are mediated via S1R signaling.

2.4. Pridopidine Restores Mitochondrial Membrane Potential and Reduces Mitochondrial Apoptotic Signaling in ALS-NPCs

Given the central role of mitochondrial dysfunction in ALS pathology, we next assessed whether pridopidine could restore mitochondrial membrane potential (MMP) in sporadic ALS-NPCs subjected to tunicamycin-induced ER stress. Tunicamycin treatment caused a profound, 58.7% reduction in MMP compared to unstressed controls (p < 0.0001) (Figure 4A).
Based on ER stress results, showing the strongest effect with 1 µM pridopidine, we evaluated its effect on MMP at 0.1, 0.5 and 1µM. Pridopidine significantly restored MMP levels in stressed ALS-NPCs at all three concentrations, with the strongest effect seen at 1 µM (47%, p < 0.0001 increase in MMP relative to tunicamycin alone). Lower concentrations (0.1 and 0.5 µM) produced more modest but significant improvements (20%, p < 0.01; 30%, p < 0.0001, respectively).
Co-treatment with the S1R antagonist NE-100 (1 μM) markedly attenuated the effect of pridopidine, but not completely abolishing it, at the optimal dose of pridopidine. Pridopidine 1 μM alone restored MMP by 47%, and when co-treated with NE-100, a weaker increase in MMP was measured at 16% (p < 0.05).
Importantly, the S1R antagonist NE-100 alone did not affect the TMRE signal in tunicamycin-stressed ALS-NPCs, whereas NE-100 attenuated the effect of pridopidine, supporting the notion that pridopidine’s effect is mediated via S1R.
To further assess the effect of pridopidine on mitochondrial function, we assessed the downstream mitochondrial apoptotic response to ER stress. First, we performed qPCR to assess the expression of the pro-apoptotic mitochondrial marker BAX (Figure 4B). BAX promotes mitochondrial outer membrane permeabilization, linking mitochondrial dysfunction to activation of downstream apoptotic caspases [62]. Second, we assessed caspase-3 activation, a key executioner of the apoptotic pathway (Figure 4C,D).
As shown in Figure 4B, tunicamycin exposure significantly increased BAX transcription in ALS-NPCs as compared with untreated controls (p < 0.0001). In contrast, pridopidine treatment significantly reduced BAX levels compared with tunicamycin-treated cells, bringing levels close to baseline (p < 0.0001).
Along with the increase in BAX expression, tunicamycin treatment also increased caspase-3 activity as compared with untreated controls (100% vs. 33%, p < 0.0001) (Figure 4C,D). Here, pridopidine treatment reduced caspase-3 activation under ER stress conditions (28%, p < 0.0001 vs. tunicamycin), indicating suppression of downstream apoptotic execution.
Co-treatment with the S1R antagonist, NE-100, significantly reversed the anti-apoptotic effect of pridopidine, maintaining high caspase-3 activity, further supporting pridopidine activity via S1R.

2.5. Pridopidine Enhances Cell Viability Under ER Stress in Sporadic ALS NPCs

To assess whether pridopidine’s effect on attenuation of ER stress and enhanced mitochondrial function translates into improved cell survival, we quantified pridopidine’s effect on the viability of ALS-NPCs exposed to tunicamycin. Cell viability was measured by flow cytometry using a fixable viability dye (FVD; amine-reactive dead cell discriminator) and normalized to unstressed controls (set at 100%).
Pridopidine at 0.1, 0.5, and 1 μM was evaluated. Tunicamycin treatment caused a profound loss of cell viability, resulting in 79% cell death (corresponding to 21% cell viability) compared with unstressed controls (p< 0.0001).
Pridopidine treatment provided robust, dose-dependent protection against tunicamycin-induced cell death. Consistent with prior observations on ER stress and mitochondrial membrane potential, 1 μM pridopidine produced the strongest neuroprotection, increasing ALS-NPCs’ viability by 58% relative to tunicamycin-treated cells (p < 0.0001). Lower concentrations exhibited more modest effects, with viability increased by 19% at 0.1 μM and 37% at 0.5 μM (p < 0.01) (Figure 5).

3. Discussion

In this study, we demonstrate that pridopidine, a highly selective and potent S1R agonist, exerts neuroprotective effects in sporadic ALS-NPCs, showing attenuation of ER stress and restoration of MMP, and overall improved cell viability. Importantly, these effects were abolished or attenuated by the selective S1R antagonist NE-100, confirming that the observed benefits of pridopidine are primarily mediated through S1R activation.
Our sporadic ALS-NPC cultures are well-characterized, expressing PAX6 and SOX2, confirming neural progenitor identity, and TUJ1 (β-III tubulin), indicating early neuronal commitment. This system represents a mixed NPC/early neuron population transitioning toward a post-mitotic neuronal state. This developmental stage is relevant for capturing early pathogenic events, including ER stress and mitochondrial dysfunction, which are known to precede overt neurodegeneration in ALS. iPSC-derived NPCs are a well-established and widely used model for studying early cellular mechanisms in neurodegenerative diseases, including ALS (e.g., Lorenz et al., 2017 [63]; Zink et al., 2021 [64]). These cells retain the patient-specific genetic background while providing a controlled and reproducible experimental system.
Consistent with reports from ALS patient tissue and from ALS nonclinical models, tunicamycin exposure markedly induced ER stress, as measured by the increased number of ALS-NPCs expressing BiP and CHOP, impaired mitochondrial function and increased cell death [11,12,13,16,17,24]. Attractive therapeutic targets in ALS focus on reducing prolonged and maladaptive ER stress, the abnormal unfolded protein response, and mitochondrial impairment. These processes contribute directly to motor neuron vulnerability and degeneration [17,18,19,65]. The ability of pridopidine to reduce ER stress and enhance mitochondrial function and cell survival is in accordance with prior published studies demonstrating a key role for S1R in regulating normal ER and mitochondrial function. These findings are aligned with the central role of S1R at MAMs, where it regulates key cellular pathways including Ca2+ transfer from the ER into mitochondria, ATP production, and stress adaptation [12,13,14,28,29].
We have used tunicamycin in our model, which is an accepted and widely used pharmacological agent to induce ER stress through its inhibition of N-linked glycosylation with the understanding that it does not fully recapitulate the endogenous mechanisms underlying ER stress in ALS [22]. Nevertheless, prior studies have shown that tunicamycin induction of ER stress recapitulates disease-relevant mechanisms in ALS models, supporting its use as a relevant tool for studying ER stress-associated mechanisms in patient-derived systems [18,66].
Pridopidine exhibits a biphasic dose–response profile, similar to that reported for other S1R ligands [26,50,67]. Biphasic responses have been described for S1R ligands across numerous cellular and behavioral preclinical models, including calcium signaling, neuroprotection, cognition, and antidepressant-like activity [47,50,54,56,67]. Although the underlying mechanisms are not fully resolved, this biphasic profile is considered a hallmark of S1R pharmacology and may be driven by different mechanisms including ligand-dependent receptor conformational changes, receptor oligomerization, off-target activities, membrane microdomain distribution, and different downstream signaling pathways [26,29,34]. In the present study, pridopidine exerted a maximal reduction in BiP and CHOP expression, and the largest improvement in mitochondrial membrane potential and cell viability at a dose of 1 μM, with less robust responses at lower and higher dosages of the drug [17,50,66].
Pridopidine improved mitochondrial function, as measured by restoration of MMP, and attenuated the ER-stress-induced mitochondrial apoptotic signaling pathway, as measured by reduced BAX expression and decreased caspase-3 activation. Importantly, pridopidine’s effects were inhibited by the selective S1R antagonist, NE-100, indicating pridopidine’s protective effects are mediated via S1R activation.
In the HEALEY ALS platform trial, while pridopidine did not show significant benefit in the primary population (including early- and late-stage ALS patients) it showed notable clinical benefits in a homogenous subgroup of faster-progressing ALS participants [68]. In the faster-progressing group, pridopidine slowed disease progression, as measured by the ALSFRS-R scale, versus the placebo group, slowed the decline in respiratory and bulbar-related functions and produced meaningful improvements in speech [68,69]. Similarly, treatment benefits were observed in HD patients. In the recent Phase 3 PROOF-HD study, pridopidine showed clinically meaningful slowing of decline across measures of progression, function, cognition and motor abilities, in a subgroup of patients who were off antidopaminergic medications [70]. Importantly, pridopidine has a favorable safety profile and clinical benefits in ALS and HD patients [37,68].
Currently approved ALS treatments remain limited. For example, Riluzole provides modest survival benefit, and tofersen, recently approved, is limited only to patients with confirmed SOD1 mutations (~2% of ALS patients). Although edaravone was approved in the US, this drug was recently withdrawn in the EU following inconclusive efficacy data [9,10]. In addition, several emerging agents such as CNM-Au8 are still in development and show mixed clinical evidence [71]. Thus, broadly neuroprotective agents remain limited. S1R represents a compelling new therapeutic target due to its critical location and function within the central nervous system (CNS). S1R is highly expressed at MAMs, where it regulates ER–mitochondria crosstalk and cellular stress responses, pathways critical for neuronal health, function, and survival, that are disrupted in ALS. Activation of S1R with selective and potent agonists such as pridopidine represents a novel approach to therapeutic intervention [12,14].
In summary, this study provides the first demonstration that pridopidine attenuates ER stress, restores mitochondrial function, and improves viability in ALS patient-derived neural progenitor cells through S1R activation. The convergence of molecular, metabolic, and neuroprotective outcomes presented here provides important mechanistic support for pridopidine’s therapeutic potential. One limitation of our study is that using a single sporadic cell line may not represent the heterogeneity of ALS. The clinical benefits observed in the HEALEY Phase 2 trial [68] further support the ongoing Phase 3 trial assessing the therapeutic potential of pridopidine in patients with ALS (PREVAiLS; NCT07322003).

4. Methods and Materials

4.1. Experimental Model and Subject Details

All experiments were performed using the human iPSC line CS14isALS-Tnxx (CSBio; W14-C146) from the Ceders-Sinai Biomanufacturing Center, Los Angeles, CA, USA, originally derived by the CS iPSC Core Repository Project (https://biomanufacturing.cedars-sinai.org/ last accessed 18 December 2025) (detailed in Table 1).
Cells were cultured under serum-free, feeder-free conditions using growth factor-reduced Matrigel-coated plates (Corning, New York, NY, USA). Cultures were maintained in mTeSR Plus medium (STEMCELL Technologies) supplemented with 1% penicillin–streptomycin (GIBCO, Miami, FL, USA). Cells were incubated at 37 °C with 5% CO2 and passaged every 3–4 days using gentle dissociation with recombinant trypsin EDTA solution (SARTORIUS, Beit Haemek, Israel). The medium was changed daily. Routine assessments for mycoplasma contamination were performed using a commercial diagnostic kit (LiLiF, iNtRON Biotechnology, Inc., Seongnam-si, Republic of Korea), and the pluripotency of iPSC lines was regularly confirmed via qPCR and flow cytometry analysis of pluripotent marker OCT3/4 and SOX2.

4.2. Differentiation of iPSCs into NPCs

The iPSCs were differentiated into NPCs using a chemically defined protocol based on [59,60], with minor modifications. Briefly, dissociated iPSCs were seeded onto Matrigel-coated plates in mTeSR plus medium and cultured until they reached approximately 80% confluence. For early NPC differentiation, cells were transitioned to a medium consisting of DMEM/F12 supplemented with 1% N2 × 100 (Thermo Fisher Scientific, Waltham, MA, USA), 1 µM Dorsomorphin (TOCRIS, Bristol, UK), 1 mM L-glutamine (GIBCO, Miami, FL, USA), 1% penicillin–streptomycin, and 0.02% heparin (STEMCELL Technologies, Cambridge, MA, USA). After 72 h, retinoic acid (RA) (100 nM, Sigma-Aldrich, Darmstadt, Germany) was added to the medium for an additional 72 h to promote late-stage NPC differentiation. During the culture period, NPCs continued to proliferate. When they reached approximately 90% confluence, the cells were dissociated and reseeded onto new Matrigel-coated plates in differentiation medium supplemented with the ROCK inhibitor Y27632 (5 μM) (Cayman Chemical Company, Ann Arbor, MI, USA) as indicated above. The medium was changed daily. After 6 days, either bFGF or EGF (1.6 nM, Thermo Fisher Scientific) was added to the medium for an additional 6 days to promote long-term NPC maintenance.

4.3. Quantitative Real-Time PCR (qPCR)

RNA was extracted using the PureLink RNA Mini Kit (Invitrogen) according to the manufacturer’s instructions. For cDNA preparation, the high-capacity cDNA Reverse Transcription Kit (Applied Biosystems, Waltham, MA, USA) was used according to the manufacturer’s instructions. cDNA was prepared from 0.5 to 2 mg mRNA in RNase-free conditions. RNA purity and quantity were assessed using NanoDrop (Thermo Scientific NanoDrop™) (A260/A280 1.5–2 was considered suitable for further analysis). Gene expression analysis was performed using SYBR Green gene expression assays (Sigma-Merck). Reactions were run on a StepOnePlus applied detection system (Applied Biosystems), and PCR conditions consisted of 40 cycles of 95 °C for 10 s, followed by 60 °C for 30s. The housekeeping gene GAPDH was used as normalization control, where relative gene expression of target genes was calculated by the delta Ct method. Primer data are detailed in Table 2.

4.4. Induction of ER Stress and Treatment Conditions

Day-6 NPCs at 80% confluence were washed and incubated for 6 h at 37 °C with 5% CO2 in fresh NPC medium containing pridopidine (Cambrex or Recipharm) at 0.1–5 μM, NE100 (Cayman Chemical Company, Ann Arbor, MI, USA) at 0.1–5 μM, or a combination of NE100 and pridopidine at 0.1–1 μM. After 6 h, tunicamycin (Cayman Chemical Company, Ann Arbor, MI, USA or Cell Signaling Technology, Boston, MA, USA) at a final concentration of 1.25 μM was added, and cells were incubated overnight at 37 °C with 5% CO2. Optimal tunicamycin concentrations (0.13 µM to 12.3 µM) were determined via preliminary dose–response experiments. Control samples consisted of untreated NPCs, tunicamycin-only, pridopidine-only, and NE100-only treatment conditions. Untreated and treated cells were harvested to quantify BiP/GRP78 and CHOP in flow cytometry (FACS), confocal microscopy and apoptosis assessment by qPCR analysis of BAX expression. Quantification of differentiated NPCs in every experiment was performed by using PAX6 and SOX2 markers by flow cytometry and/or qPCR.

4.5. Detection of Mitochondrial Potential

Day-6 treated and untreated NPCs, as indicated above, at 80% confluence were washed and gently dissociated. For every treatment and control, cells were counted, and 5 × 104 cells were suspended per well in a 96-well plate. Following the instructions of the TMRE-Mitochondrial Membrane Potential Assay Kit (Abcam, Cambridge, UK), fluorescent dye was added to every well, and the plate was incubated at 37 °C for 30 min in the dark. Cells were washed twice in PBS and resuspended in 50 μL PBS, and fluorescence was measured using a plate reader at an excitation wavelength of 535 nm.

4.6. Immunostaining and Confocal Microscopy

Cells were fixed with 2% formaldehyde for 2 min and then with 4% paraformaldehyde (PFA) (Merck, Darmstadt, Germany) for 7 min, permeabilized with 0.2% (v/v) Triton-X 100 (Merck) for 5 min and blocked for 2 h in 3% BSA (Millipore, Burlington, MA, USA). Cells were then incubated overnight with primary antibodies PAX6 (Thermo Fisher Scientific; 42-6600), β-III tubulin (R&D Systems, Minneapolis, MN, USA; MAB1196), and SOX2 (GeneTex, Irvine, CA, USA; GTX627404) followed by 4 h incubation with the following secondary antibodies: donkey anti-mouse Alexa Fluor 647 and goat anti-rabbit Alexa Fluor 488 antibodies. Alexa Fluor 546 phalloidin (Thermo Fisher Scientific; A22283) was used for F-actin staining, Annexin-V (Cell Signaling Technology, 6592S) was used for detection of phosphatidylserine exposure, Hoechst was used for nuclear staining, and VECTASHIELD Mounting Medium (Vector Laboratories, Newark, CA, USA) was used for long-term preservation. Imaging was performed using a Nikon C1si laser scanning confocal microscope. For caspase-3 immunostaining, ALS-NPCs were seeded, treated and fixed as explained above. Following treatment and fixation, cells were incubated with an antibody against cleaved caspase-3 to assess apoptotic signaling. Nuclei were counterstained with Hoechst. Fluorescent images were acquired using confocal microscopy under identical acquisition settings across all treatment groups. Caspase-3 signal intensity was quantified using Fiji software (ImageJ, version 2.9.0) and expressed relative to the tunicamycin-treated condition.

4.7. Cell Viability and Flow Cytometry (FACS)

Cell viability was assessed using LIVE/DEAD Fixable Far Red Dead Cell Stain Kit (Thermo Fisher Scientific), which allows for the discrimination and quantification of live and dead cells in FACS. Cells were harvested, dissociated into single-cell suspensions and incubated for 30 min at 4 °C with the live/dead stain, and washed twice with FACS buffer containing PBS (Sartorius, Beit Haemek, Israel) and 10% FBS (Gibco, Miami, FL, USA). Following treatment, cells were permeabilized and fixed using the FOXP3/Transcription Staining Buffer Set (Invitrogen, Carlsbad, CA, USA). After fixation, intracellular staining was performed for ER stress marker analysis. Cells were stained with primary antibodies against BiP/GRP78 (Cell Signaling Technology; C50B12) and CHOP (Cell Signaling Technology; L63F7). Characterization of NPC differentiation included antibody staining for PAX6, SOX2 and β-III tubulin markers. Primary antibody staining was followed by incubation for 30 min with secondary antibodies: donkey anti-mouse Alexa Fluor 647 and goat anti-rabbit Alexa Fluor 488. A total of 1x105 cells were incubated with Annexin-V and 7-AAD solutions at 5 mg/mL for 5–30 min. Cells were analyzed using a FACS Canto flow cytometer (BD Biosciences, San Jose, CA, USA) equipped with CellQuest Pro software, Pro (version 5.2 BD, Biosciences, San Jose, CA, USA). Fluorescence intensities were recorded for each sample, and the normalized mean fluorescence intensity (nMFI) was calculated for each experimental condition. Results were reported as the percentage of NPCs positive for the measured proteins, with comparisons made between treated and untreated cells.

4.8. Statistical Analysis

All experiments were performed using at least three independent biological replicates. A biological replicate is defined as an independent differentiation of iPSCs into NPCs. Technical replicates represent repeated measurements within the same experiment (e.g., replicate wells, qPCR reactions, or flow cytometry measurements).
Statistical significance between groups was determined using a two-tailed Student t-test with Bonferroni post hoc correction, or a One-Way Analysis of Variance (ANOVA) followed by Tukey’s post hoc test for multiple comparisons. Results are presented as the mean ± SEM, with p < 0.05 considered statistically significant.

Author Contributions

M.M., M.G., R.G.L. and M.R.H. designed the research. M.M., M.S.Z., N.T. and R.G.L. performed the research and the analysis. M.M., M.G., A.M.T., R.G.L. and M.R.H. contributed to writing and editing the manuscript draft and final copy in the agreement with all authors. All authors have read and agreed to the published version of the manuscript.

Funding

This study was sponsored and funded by Prilenia Therapeutics B.V., which supported the study design, data collection, statistical analysis, and manuscript development.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The original contributions presented in this study are included in the article. Further inquiries can be directed to the corresponding authors.

Acknowledgments

We would like to thank Uzi Hadad for his help with the microscopy figure analyses.

Conflicts of Interest

Authors May Meltzer, Andrew M. Tan, and Michal Geva were employed by and hold stock options in Prilenia Therapeutics B.V. Author Michael R. Hayden is the founder and CEO of Prilenia Therapeutics B.V. and holds equity in the company. The remaining authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest. The authors declare that this study received funding from Prilenia Therapeutics B.V.

Abbreviations

IPSCsinduced pluripotent stem cells
NPCsneural progenitor cells
ALSamyotrophic lateral sclerosis
ERendoplasmic reticulum
S1RSigma-1 receptor
MAMmitochondria-associated membrane
BAX BCL2-associated X protein

References

  1. Masrori, P.; Van Damme, P. Amyotrophic Lateral Sclerosis: A Clinical Review. Eur. J. Neurol. 2020, 27, 1918–1929. [Google Scholar] [CrossRef]
  2. Riva, N.; Domi, T.; Pozzi, L.; Lunetta, C.; Schito, P.; Spinelli, E.G.; Cabras, S.; Matteoni, E.; Consonni, M.; Bella, E.D.; et al. Update on Recent Advances in Amyotrophic Lateral Sclerosis. J. Neurol. 2024, 271, 4693–4723. [Google Scholar] [CrossRef]
  3. Ionescu, A.; Gradus, T.; Altman, T.; Maimon, R.; Saraf Avraham, N.; Geva, M.; Hayden, M.; Perlson, E. Targeting the Sigma-1 Receptor via Pridopidine Ameliorates Central Features of ALS Pathology in a SOD1G93A Model. Cell Death Dis. 2019, 10, 210. [Google Scholar] [CrossRef]
  4. Blair, H.A. Tofersen: First Approval. Drugs 2023, 83, 1039–1043. [Google Scholar] [CrossRef]
  5. Bensimon, G.; Lacomblez, L.; Meininger, V. A Controlled Trial of Riluzole in Amyotrophic Lateral Sclerosis. N. Engl. J. Med. 1994, 330, 585–591. [Google Scholar] [CrossRef]
  6. Abe, K.; Aoki, M.; Tsuji, S.; Itoyama, Y.; Sobue, G.; Togo, M.; Hamada, C.; Tanaka, M.; Akimoto, M.; Nakamura, K.; et al. Safety and Efficacy of Edaravone in Well Defined Patients with Amyotrophic Lateral Sclerosis: A Randomised, Double-Blind, Placebo-Controlled Trial. Lancet Neurol. 2017, 16, 505–512. [Google Scholar] [CrossRef]
  7. Chen, J.J. Overview of Current and Emerging Therapies for Amytrophic Lateral Sclerosis. Am. J. Manag. Care 2020, 26, S191–S197. [Google Scholar] [CrossRef]
  8. Xu, X.; Shen, D.; Gao, Y.; Zhou, Q.; Ni, Y.; Meng, H.; Shi, H.; Le, W.; Chen, S.; Chen, S. A Perspective on Therapies for Amyotrophic Lateral Sclerosis: Can Disease Progression Be Curbed? Transl. Neurodegener. 2021, 10, 29. [Google Scholar] [CrossRef]
  9. Lu, L.; Deng, Y.; Xu, R. Current Potential Therapeutics of Amyotrophic Lateral Sclerosis. Front. Neurol. 2024, 15, 1402962. [Google Scholar] [CrossRef]
  10. Hoxhaj, P.; Hastings, N.; Kachhadia, M.P.; Gupta, R.; Sindhu, U.; Durve, S.A.; Azam, A.; Auz Vinueza, M.J.; Win, B.; Win, S.H.; et al. Exploring Advancements in the Treatment of Amyotrophic Lateral Sclerosis: A Comprehensive Review of Current Modalities and Future Prospects. Cureus 2023, 15, e45489. [Google Scholar] [CrossRef]
  11. Chen, J.; Bassot, A.; Giuliani, F.; Simmen, T. Amyotrophic Lateral Sclerosis (ALS): Stressed by Dysfunctional Mitochondria-Endoplasmic Reticulum Contacts (MERCs). Cells 2021, 10, 1789. [Google Scholar] [CrossRef]
  12. Candelise, N.; Salvatori, I.; Scaricamazza, S.; Nesci, V.; Zenuni, H.; Ferri, A.; Valle, C. Mechanistic Insights of Mitochondrial Dysfunction in Amyotrophic Lateral Sclerosis: An Update on a Lasting Relationship. Metabolites 2022, 12, 233. [Google Scholar] [CrossRef]
  13. Bernard-Marissal, N.; Chrast, R.; Schneider, B.L. Endoplasmic Reticulum and Mitochondria in Diseases of Motor and Sensory Neurons: A Broken Relationship? Cell Death Dis. 2018, 9, 333. [Google Scholar] [CrossRef]
  14. Johri, A.; Chandra, A. Connection Lost, MAM: Errors in ER–Mitochondria Connections in Neurodegenerative Diseases. Brain Sci. 2021, 11, 1437. [Google Scholar] [CrossRef]
  15. De Lorenzo, F.; Lüningschrör, P.; Nam, J.; Beckett, L.; Pilotto, F.; Galli, E.; Lindholm, P.; Rüdt von Collenberg, C.; Mungwa, S.T.; Jablonka, S.; et al. CDNF Rescues Motor Neurons in Models of Amyotrophic Lateral Sclerosis by Targeting Endoplasmic Reticulum Stress. Brain 2023, 146, 3783–3799. [Google Scholar] [CrossRef]
  16. Matus, S.; Valenzuela, V.; Medinas, D.B.; Hetz, C. ER Dysfunction and Protein Folding Stress in ALS. Int. J. Cell Biol. 2013, 2013, 674751. [Google Scholar] [CrossRef]
  17. Jeon, Y.-M.; Kwon, Y.; Lee, S.; Kim, H.-J. Potential Roles of the Endoplasmic Reticulum Stress Pathway in Amyotrophic Lateral Sclerosis. Front. Aging Neurosci. 2023, 15, 1047897. [Google Scholar] [CrossRef]
  18. Watts, M.E.; Giadone, R.M.; Ordureau, A.; Holton, K.M.; Harper, J.W.; Rubin, L.L. Analyzing the ER Stress Response in ALS Patient Derived Motor Neurons Identifies Druggable Neuroprotective Targets. Front. Cell. Neurosci. 2024, 17, 1327361. [Google Scholar] [CrossRef]
  19. Ghemrawi, R.; Khair, M. Endoplasmic Reticulum Stress and Unfolded Protein Response in Neurodegenerative Diseases. Int. J. Mol. Sci. 2020, 21, 6127. [Google Scholar] [CrossRef]
  20. Hetz, C.; Saxena, S. ER Stress and the Unfolded Protein Response in Neurodegeneration. Nat. Rev. Neurol. 2017, 13, 477–491. [Google Scholar] [CrossRef]
  21. Scheper, W.; Hoozemans, J.J.M. The Unfolded Protein Response in Neurodegenerative Diseases: A Neuropathological Perspective. Acta Neuropathol. 2015, 130, 315–331. [Google Scholar] [CrossRef]
  22. Almanza, A.; Carlesso, A.; Chintha, C.; Creedican, S.; Doultsinos, D.; Leuzzi, B.; Luís, A.; McCarthy, N.; Montibeller, L.; More, S.; et al. Endoplasmic Reticulum Stress Signalling—From Basic Mechanisms to Clinical Applications. FEBS J. 2019, 286, 241–278. [Google Scholar] [CrossRef]
  23. Walter, P.; Ron, D. The Unfolded Protein Response: From Stress Pathway to Homeostatic Regulation. Science 2011, 334, 1081–1086. [Google Scholar] [CrossRef]
  24. Chen, Q.; Thompson, J.; Hu, Y.; Lesnefsky, E.J. Tunicamycin-Induced Endoplasmic Reticulum Stress Damages Complex I in Cardiac Mitochondria. Life 2022, 12, 1209. [Google Scholar] [CrossRef]
  25. Hayashi, T.; Su, T.-P. Sigma-1 Receptor Chaperones at the ER-Mitochondrion Interface Regulate Ca2+ Signaling and Cell Survival. Cell 2007, 131, 596–610. [Google Scholar] [CrossRef]
  26. Su, T.-P.; Su, T.-C.; Nakamura, Y.; Tsai, S.-Y. The Sigma-1 Receptor as a Pluripotent Modulator in Living Systems. Trends Pharmacol. Sci. 2016, 37, 262–278. [Google Scholar] [CrossRef]
  27. Sharma, N.; Patel, C.; Shenkman, M.; Kessel, A.; Ben-Tal, N.; Lederkremer, G.Z. The Sigma-1 Receptor Is an ER-Localized Type II Membrane Protein. J. Biol. Chem. 2021, 297, 101299. [Google Scholar] [CrossRef]
  28. Ryskamp, D.A.; Korban, S.; Zhemkov, V.; Kraskovskaya, N.; Bezprozvanny, I. Neuronal Sigma-1 Receptors: Signaling Functions and Protective Roles in Neurodegenerative Diseases. Front. Neurosci. 2019, 13, 862. [Google Scholar] [CrossRef]
  29. Zhemkov, V.; Ditlev, J.A.; Lee, W.-R.; Wilson, M.; Liou, J.; Rosen, M.K.; Bezprozvanny, I. The Role of Sigma 1 Receptor in Organization of Endoplasmic Reticulum Signaling Microdomains. eLife 2021, 10, e65192. [Google Scholar] [CrossRef]
  30. Naia, L.; Ly, P.; Mota, S.I.; Lopes, C.; Maranga, C.; Coelho, P.; Gershoni-Emek, N.; Ankarcrona, M.; Geva, M.; Hayden, M.R.; et al. The Sigma-1 Receptor Mediates Pridopidine Rescue of Mitochondrial Function in Huntington Disease Models. Neurotherapeutics 2021, 18, 1017–1038. [Google Scholar] [CrossRef]
  31. Crouzier, L.; Denus, M.; Richard, E.M.; Tavernier, A.; Diez, C.; Cubedo, N.; Maurice, T.; Delprat, B. Sigma-1 Receptor Is Critical for Mitochondrial Activity and Unfolded Protein Response in Larval Zebrafish. Int. J. Mol. Sci. 2021, 22, 11049. [Google Scholar] [CrossRef]
  32. Crouzier, L.; Danese, A.; Yasui, Y.; Richard, E.M.; Liévens, J.-C.; Patergnani, S.; Couly, S.; Diez, C.; Denus, M.; Cubedo, N.; et al. Activation of the Sigma-1 Receptor Chaperone Alleviates Symptoms of Wolfram Syndrome in Preclinical Models. Sci. Transl. Med. 2022, 14, eabh3763. [Google Scholar] [CrossRef]
  33. Shenkman, M.; Geva, M.; Gershoni-Emek, N.; Hayden, M.R.; Lederkremer, G.Z. Pridopidine Reduces Mutant Huntingtin-induced Endoplasmic Reticulum Stress by Modulation of the Sigma-1 Receptor. J. Neurochem. 2021, 158, 467–481. [Google Scholar] [CrossRef]
  34. Zhemkov, V.; Geva, M.; Hayden, M.R.; Bezprozvanny, I. Sigma-1 Receptor (S1R) Interaction with Cholesterol: Mechanisms of S1R Activation and Its Role in Neurodegenerative Diseases. Int. J. Mol. Sci. 2021, 22, 4082. [Google Scholar] [CrossRef]
  35. Eddings, C.R.; Arbez, N.; Akimov, S.; Geva, M.; Hayden, M.R.; Ross, C.A. Pridopidine Protects Neurons from Mutant-Huntingtin Toxicity via the Sigma-1 Receptor. Neurobiol. Dis. 2019, 129, 118–129. [Google Scholar] [CrossRef]
  36. Estévez-Silva, H.M.; Mediavilla, T.; Giacobbo, B.L.; Liu, X.; Sultan, F.R.; Marcellino, D.J. Pridopidine Modifies Disease Phenotype in a SOD1 Mouse Model of Amyotrophic Lateral Sclerosis. Eur. J. Neurosci. 2022, 55, 1356–1372. [Google Scholar] [CrossRef]
  37. Goldberg, Y.P.; Navon-Perry, L.; Cruz-Herranz, A.; Chen, K.; Hecker-Barth, G.; Spiegel, K.; Cohen, Y.; Niethammer, M.; Tan, A.M.; Schuring, H.; et al. The Safety Profile of Pridopidine, a Novel Sigma-1 Receptor Agonist for the Treatment of Huntington’s Disease. CNS Drugs 2025, 39, 485–498. [Google Scholar] [CrossRef]
  38. Ryskamp, D.; Wu, J.; Geva, M.; Kusko, R.; Grossman, I.; Hayden, M.; Bezprozvanny, I. The Sigma-1 Receptor Mediates the Beneficial Effects of Pridopidine in a Mouse Model of Huntington Disease. Neurobiol. Dis. 2017, 97, 46–59. [Google Scholar] [CrossRef]
  39. Grachev, I.D.; Meyer, P.M.; Becker, G.A.; Bronzel, M.; Marsteller, D.; Pastino, G.; Voges, O.; Rabinovich, L.; Knebel, H.; Zientek, F.; et al. Sigma-1 and Dopamine D2/D3 Receptor Occupancy of Pridopidine in Healthy Volunteers and Patients with Huntington Disease: A [18F] Fluspidine and [18F] Fallypride PET Study. Eur. J. Nucl. Med. Mol. Imaging 2021, 48, 1103–1115. [Google Scholar] [CrossRef]
  40. Weng, T.-Y.; Tsai, S.-Y.A.; Su, T.-P. Roles of Sigma-1 Receptors on Mitochondrial Functions Relevant to Neurodegenerative Diseases. J. Biomed. Sci. 2017, 24, 74. [Google Scholar] [CrossRef]
  41. Malar, D.S.; Thitilertdecha, P.; Ruckvongacheep, K.S.; Brimson, S.; Tencomnao, T.; Brimson, J.M. Targeting Sigma Receptors for the Treatment of Neurodegenerative and Neurodevelopmental Disorders. CNS Drugs 2023, 37, 399–440. [Google Scholar] [CrossRef] [PubMed]
  42. Delprat, B.; Crouzier, L.; Su, T.-P.; Maurice, T. At the Crossing of ER Stress and MAMs: A Key Role of Sigma-1 Receptor? Adv Exp. Med. Biol. 2020, 1131, 699–718. [Google Scholar] [CrossRef] [PubMed]
  43. Maurice, T.; Goguadze, N. Sigma-1 (Σ1) Receptor in Memory and Neurodegenerative Diseases. Handb. Exp. Pharmacol. 2017, 244, 81–108. [Google Scholar] [CrossRef]
  44. Maurice, T.; Hiramatsu, M.; Itoh, J.; Kameyama, T.; Hasegawa, T.; Nabeshima, T. Behavioral Evidence for a Modulating Role of σ Ligands in Memory Processes. I. Attenuation of Dizocilpine (MK-801)-Induced Amnesia. Brain Res. 1994, 647, 44–56. [Google Scholar] [CrossRef]
  45. Maurice, T.; Lockhart, B.P. Neuroprotective and Anti-Amnesic Potentials of Sigma (σ) Receptor Ligands. Prog. Neuropsychopharmacol. Biol. Psychiatry 1997, 21, 69–102. [Google Scholar] [CrossRef]
  46. Maurice, T.; Urani, A.; Phan, V.L.; Romieu, P. The Interaction between Neuroactive Steroids and the Σ1 Receptor Function: Behavioral Consequences and Therapeutic Opportunities. Brain Res. Rev. 2001, 37, 116–132. [Google Scholar] [CrossRef]
  47. Monnet, F.P. Sigma-1 Receptor as Regulator of Neuronal Intracellular Ca2+: Clinical and Therapeutic Relevance. Biol. Cell 2005, 97, 873–883. [Google Scholar] [CrossRef]
  48. Matsuno, K.; Matsunaga, K.; Senda, T.; Mita, S. Increase in Extracellular Acetylcholine Level by Sigma Ligands in Rat Frontal Cortex. J. Pharmacol. Exp. Ther. 1993, 265, 851–859. [Google Scholar] [CrossRef]
  49. Matsuno, K.; Senda, T.; Matsunaga, K.; Mita, S. Ameliorating Effects of σ Receptor Ligands on the Impairment of Passive Avoidance Tasks in Mice: Involvement in the Central Acetylcholinergic System. Eur. J. Pharmacol. 1994, 261, 43–51. [Google Scholar] [CrossRef]
  50. Maurice, T.; Su, T.-P. The Pharmacology of Sigma-1 Receptors. Pharmacol. Ther. 2009, 124, 195–206. [Google Scholar] [CrossRef] [PubMed]
  51. Monnet, F.P.; Debonnel, G.; Junien, J.L.; De Montigny, C. N-Methyl-D-Aspartate-Induced Neuronal Activation Is Selectively Modulated by σ Receptors. Eur. J. Pharmacol. 1990, 179, 441–445. [Google Scholar] [CrossRef]
  52. Monnet, F.P.; Blier, P.; Debonnel, G.; de Montigny, C. Modulation by Sigma Ligands of N-Methyl-D-Aspartate-Induced [3H]Noradrenaline Release in the Rat Hippocampus: G-Protein Dependency. Naunyn. Schmiedebergs. Arch. Pharmacol. 1992, 346, 32–39. [Google Scholar] [CrossRef]
  53. Bermack, J.E.; Debonnel, G. Distinct Modulatory Roles of Sigma Receptor Subtypes on Glutamatergic Responses in the Dorsal Hippocampus. Synapse 2005, 55, 37–44. [Google Scholar] [CrossRef]
  54. Hayashi, T.; Maurice, T.; Su, T.P. Ca2+ Signaling via Σ1-Receptors: Novel Regulatory Mechanism Affecting Intracellular Ca2+ Concentration. J. Pharmacol. Exp. Ther. 2000, 293, 788–798. [Google Scholar] [CrossRef]
  55. Hong, W.; Nuwayhid, S.J.; Werling, L.L. Modulation of Bradykinin-induced Calcium Changes in SH-SY5Y Cells by Neurosteroids and Sigma Receptor Ligands via a Shared Mechanism. Synapse 2004, 54, 102–110. [Google Scholar] [CrossRef]
  56. Urani, A.; Romieu, P.; Roman, F.J.; Yamada, K.; Noda, Y.; Kamei, H.; Manh Tran, H.; Nagai, T.; Nabeshima, T.; Maurice, T. Enhanced Antidepressant Efficacy of Σ1 Receptor Agonists in Rats after Chronic Intracerebroventricular Infusion of β-Amyloid-(1–40) Protein. Eur. J. Pharmacol. 2004, 486, 151–161. [Google Scholar] [CrossRef]
  57. Smith-Dijak, A.I.; Nassrallah, W.B.; Zhang, L.Y.J.; Geva, M.; Hayden, M.R.; Raymond, L.A. Impairment and Restoration of Homeostatic Plasticity in Cultured Cortical Neurons From a Mouse Model of Huntington Disease. Front. Cell. Neurosci. 2019, 13, 209. [Google Scholar] [CrossRef]
  58. Francardo, V.; Geva, M.; Bez, F.; Denis, Q.; Steiner, L.; Hayden, M.R.; Cenci, M.A. Pridopidine Induces Functional Neurorestoration via the Sigma-1 Receptor in a Mouse Model of Parkinson’s Disease. Neurotherapeutics 2019, 16, 465–479. [Google Scholar] [CrossRef]
  59. Lee, G.; Chambers, S.M.; Tomishima, M.J.; Studer, L. Derivation of Neural Crest Cells from Human Pluripotent Stem Cells. Nat. Protoc. 2010, 5, 688–701. [Google Scholar] [CrossRef]
  60. Qu, Q.; Li, D.; Louis, K.R.; Li, X.; Yang, H.; Sun, Q.; Crandall, S.R.; Tsang, S.; Zhou, J.; Cox, C.L.; et al. High-Efficiency Motor Neuron Differentiation from Human Pluripotent Stem Cells and the Function of Islet-1. Nat. Commun. 2014, 5, 3449. [Google Scholar] [CrossRef]
  61. Uchida, N.; Buck, D.W.; He, D.; Reitsma, M.J.; Masek, M.; Phan, T.V.; Tsukamoto, A.S.; Gage, F.H.; Weissman, I.L. Direct Isolation of Human Central Nervous System Stem Cells. Proc. Natl. Acad. Sci. USA 2000, 97, 14720–14725. [Google Scholar] [CrossRef]
  62. McArthur, K.; Whitehead, L.W.; Heddleston, J.M.; Li, L.; Padman, B.S.; Oorschot, V.; Geoghegan, N.D.; Chappaz, S.; Davidson, S.; San Chin, H.; et al. BAK/BAX Macropores Facilitate Mitochondrial Herniation and MtDNA Efflux during Apoptosis. Science 2018, 359, eaao6047. [Google Scholar] [CrossRef]
  63. Lorenz, C.; Lesimple, P.; Bukowiecki, R.; Zink, A.; Inak, G.; Mlody, B.; Singh, M.; Semtner, M.; Mah, N.; Auré, K.; et al. Human IPSC-Derived Neural Progenitors Are an Effective Drug Discovery Model for Neurological MtDNA Disorders. Cell Stem Cell 2017, 20, 659–674.e9. [Google Scholar] [CrossRef]
  64. Zink, A.; Lisowski, P.; Prigione, A. Generation of Human IPSC-Derived Neural Progenitor Cells (NPCs) as Drug Discovery Model for Neurological and Mitochondrial Disorders. Bio-Protocol 2021, 11, e3939. [Google Scholar] [CrossRef]
  65. Bernard-Marissal, N.; Médard, J.-J.; Azzedine, H.; Chrast, R. Dysfunction in Endoplasmic Reticulum-Mitochondria Crosstalk Underlies SIGMAR1 Loss of Function Mediated Motor Neuron Degeneration. Brain 2015, 138, 875–890. [Google Scholar] [CrossRef] [PubMed]
  66. Lee, S.; Shang, Y.; Redmond, S.A.; Urisman, A.; Tang, A.A.; Li, K.H.; Burlingame, A.L.; Pak, R.A.; Jovičić, A.; Gitler, A.D.; et al. Activation of HIPK2 Promotes ER Stress-Mediated Neurodegeneration in Amyotrophic Lateral Sclerosis. Neuron 2016, 91, 41–55. [Google Scholar] [CrossRef]
  67. Maurice, T. Bi-Phasic Dose Response in the Preclinical and Clinical Developments of Sigma-1 Receptor Ligands for the Treatment of Neurodegenerative Disorders. Expert Opin. Drug Discov. 2021, 16, 373–389. [Google Scholar]
  68. Geva, M.; Goldberg, Y.P.; Leitner, M.L.; Chen, K.; Feng, W.; Emek, N.G.; Berry, J.D.; Paganoni, S.; Shefner, J.; Cudkowicz, M.E.; et al. Pridopidine Treatment Benefits in Definite, Probable, and Early (<18mo Onset) Subjects with ALS in the HEALEY ALS Platform Trial. Neurotherapeutics 2025, 22, e00675. [Google Scholar] [CrossRef]
  69. Shefner, J.M.; Oskarsson, B.; Macklin, E.A.; Chibnik, L.B.; Quintana, M.; Saville, B.R.; Detry, M.A.; Vestrucci, M.; Marion, J.; Mcglothlin, A.; et al. Pridopidine in Amyotrophic Lateral Sclerosis: The HEALEY ALS Platform Trial. JAMA 2025, 333, 1128–1137. [Google Scholar] [CrossRef]
  70. Reilmann, R.; Feigin, A.; Rosser, A.E.; Kostyk, S.K.; Saft, C.; Cohen, Y.; Schuring, H.; Hand, R.; Tan, A.M.; Chen, K.; et al. Pridopidine in Early-Stage Manifest Huntington’s Disease: A Phase 3 Trial. Nat. Med. 2025, 31, 3780–3789. [Google Scholar] [CrossRef]
  71. Hayden, D.; Lai, P.-Y.; Donahue, R.A.; Chen, H.-W.; Wang, J.; Mathai, N.; Lopes, G.; McCaffrey, A.; Scalia, J.; Luppino, S.; et al. CNM-Au8 in Amyotrophic Lateral Sclerosis. JAMA 2025, 333, 1138. [Google Scholar] [CrossRef]
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Figure 1. Directed differentiation of sporadic ALS patient-derived iPSCs into NPCs and early neurons. (A) Schematic representation of the stepwise neural induction protocol used to differentiate ALS patient- iPSCs into NPCs and neurons. Cells were cultured in mTeSR Plus medium until day 0, then sequentially transitioned through early NPC and NPC media containing DMEM/F12, N2, heparin, dorsomorphin, and retinoic acid, with final maturation in EGF/FGF-containing medium. (B) Representative immunofluorescence images show cytoskeletal organization during differentiation at day 0, day 3, and day 11 stained with Phalloidin (red, F-actin) and Hoechst (blue, nuclei). Scale bars represent 100/200 mm (n = 3 biological). (C) Quantification of the proliferation marker gene’s expression (MKI67) relative to sporadic ALS iPSCs (n = 3 technical replicates). (D) Representative immunofluorescence staining of NPCs at day 6 showing expression of PAX6 (magenta) and SOX2 (green), confirming acquisition of NPC identity. Nuclei were counterstained with Hoechst (blue). Scale bar: 5 µm (n = 3 biological replicates). (E) Flow cytometry analysis of day-6 NPCs and iPSCs showing the expression of neural progenitor markers PAX6 and SOX2. Quantification of normalized mean fluorescence intensity (nMFI) indicates strong upregulation of both markers upon differentiation (n ≥ 5 biological replicates, mean ± SEM) (F) Flow cytometry analysis of day-13 NPCs showing persistence PAX6 expression and induction of the early neuronal marker TUJ1, indicating progression toward neuronal lineage commitment. Quantification of nMFI demonstrates a significant increase in TUJ1 expression (n ≥ 3 technical replicates, mean ± SEM).
Figure 1. Directed differentiation of sporadic ALS patient-derived iPSCs into NPCs and early neurons. (A) Schematic representation of the stepwise neural induction protocol used to differentiate ALS patient- iPSCs into NPCs and neurons. Cells were cultured in mTeSR Plus medium until day 0, then sequentially transitioned through early NPC and NPC media containing DMEM/F12, N2, heparin, dorsomorphin, and retinoic acid, with final maturation in EGF/FGF-containing medium. (B) Representative immunofluorescence images show cytoskeletal organization during differentiation at day 0, day 3, and day 11 stained with Phalloidin (red, F-actin) and Hoechst (blue, nuclei). Scale bars represent 100/200 mm (n = 3 biological). (C) Quantification of the proliferation marker gene’s expression (MKI67) relative to sporadic ALS iPSCs (n = 3 technical replicates). (D) Representative immunofluorescence staining of NPCs at day 6 showing expression of PAX6 (magenta) and SOX2 (green), confirming acquisition of NPC identity. Nuclei were counterstained with Hoechst (blue). Scale bar: 5 µm (n = 3 biological replicates). (E) Flow cytometry analysis of day-6 NPCs and iPSCs showing the expression of neural progenitor markers PAX6 and SOX2. Quantification of normalized mean fluorescence intensity (nMFI) indicates strong upregulation of both markers upon differentiation (n ≥ 5 biological replicates, mean ± SEM) (F) Flow cytometry analysis of day-13 NPCs showing persistence PAX6 expression and induction of the early neuronal marker TUJ1, indicating progression toward neuronal lineage commitment. Quantification of nMFI demonstrates a significant increase in TUJ1 expression (n ≥ 3 technical replicates, mean ± SEM).
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Figure 2. Pridopidine reduces ER stress markers BiP and CHOP in sporadic ALS-NPCs in a biphasic manner. (A) Percentage of ALS-NPCs with elevated BiP expression following tunicamycin treatment (1.25 µM, 16 h) and co-treatment with increasing concentrations of pridopidine (0.1–50 µM). n = 3 biological replicates. (B) Percentage of ALS-NPCs with elevated CHOP expression under the same conditions. n = 3 biological replicates. Data are presented as mean ± SEM.; * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001 vs. tunicamycin-treated cells (two-way ANOVA with post hoc test).
Figure 2. Pridopidine reduces ER stress markers BiP and CHOP in sporadic ALS-NPCs in a biphasic manner. (A) Percentage of ALS-NPCs with elevated BiP expression following tunicamycin treatment (1.25 µM, 16 h) and co-treatment with increasing concentrations of pridopidine (0.1–50 µM). n = 3 biological replicates. (B) Percentage of ALS-NPCs with elevated CHOP expression under the same conditions. n = 3 biological replicates. Data are presented as mean ± SEM.; * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001 vs. tunicamycin-treated cells (two-way ANOVA with post hoc test).
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Figure 3. Reduction in ER stress markers BiP and CHOP by pridopidine is mediated via S1R activation. (A,B) Quantification of ER stress markers BiP (A) and CHOP (B) by flow cytometry in ALS-NPCs treated with tunicamycin (1.25 μM, 16 h) with or without pridopidine (0.5 μM) and the S1R antagonist NE-100 (1 μM). Tunicamycin markedly increased BiP and CHOP levels; pridopidine significantly reduced both markers, and NE-100 abolished this effect, indicating S1R dependence. Data represent mean ± SEM; n ≥ 5 biological replicates. Statistical analysis: **** p < 0.0001, *** p < 0.001, ns = not significant. (C,E) Representative immunofluorescence images of BiP (C) and CHOP (E). Tunicamycin induced strong BiP (green) and CHOP (red) accumulation, which was substantially reduced by pridopidine and restored by NE-100. Nuclei are counterstained with DAPI (blue). Scale bar: 5 μM. (D,F) Quantification of immunofluorescence signal for BiP (D) and CHOP (F). Pridopidine significantly decreased tunicamycin-induced BiP and CHOP intensities, whereas NE-100 attenuated this reduction. Data represent mean ± SEM; n = 3 biological replicates. Statistical analysis: **** p < 0.0001, *** p < 0.001.
Figure 3. Reduction in ER stress markers BiP and CHOP by pridopidine is mediated via S1R activation. (A,B) Quantification of ER stress markers BiP (A) and CHOP (B) by flow cytometry in ALS-NPCs treated with tunicamycin (1.25 μM, 16 h) with or without pridopidine (0.5 μM) and the S1R antagonist NE-100 (1 μM). Tunicamycin markedly increased BiP and CHOP levels; pridopidine significantly reduced both markers, and NE-100 abolished this effect, indicating S1R dependence. Data represent mean ± SEM; n ≥ 5 biological replicates. Statistical analysis: **** p < 0.0001, *** p < 0.001, ns = not significant. (C,E) Representative immunofluorescence images of BiP (C) and CHOP (E). Tunicamycin induced strong BiP (green) and CHOP (red) accumulation, which was substantially reduced by pridopidine and restored by NE-100. Nuclei are counterstained with DAPI (blue). Scale bar: 5 μM. (D,F) Quantification of immunofluorescence signal for BiP (D) and CHOP (F). Pridopidine significantly decreased tunicamycin-induced BiP and CHOP intensities, whereas NE-100 attenuated this reduction. Data represent mean ± SEM; n = 3 biological replicates. Statistical analysis: **** p < 0.0001, *** p < 0.001.
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Figure 4. Pridopidine restores mitochondrial membrane potential and attenuates mitochondrial apoptotic signaling, including BAX expression and caspase-3 activation, in sporadic ALS-NPCs. (A) MMP was measured in ALS-NPCs following tunicamycin exposure (1.25 µM, 16 h) in the presence or absence of pridopidine (0.1–1 µM) and the S1R antagonist NE-100 (1 µM). (B) Relative BAX gene expression as measured by qPCR under the same experimental conditions as in A. Tunicamycin increased BAX expression; pridopidine reduced BAX levels, consistent with pridopidine treatment reducing mitochondrial apoptotic signaling. n = 3 biological replicates. (C) Representative confocal images of caspase-3 immunostaining (green) and Hoechst nuclear staining (blue) in untreated cells, tunicamycin-treated cells, and cells co-treated with tunicamycin plus pridopidine, with or without NE-100. Scale bar, 100mm. (D) Quantification of relative caspase-3 activity under the indicated treatment conditions. Data are expressed as % of untreated control (mean ± SEM). * p < 0.05, ** p < 0.01, **** p < 0.0001 vs. tunicamycin alone (one-way ANOVA with post hoc test).
Figure 4. Pridopidine restores mitochondrial membrane potential and attenuates mitochondrial apoptotic signaling, including BAX expression and caspase-3 activation, in sporadic ALS-NPCs. (A) MMP was measured in ALS-NPCs following tunicamycin exposure (1.25 µM, 16 h) in the presence or absence of pridopidine (0.1–1 µM) and the S1R antagonist NE-100 (1 µM). (B) Relative BAX gene expression as measured by qPCR under the same experimental conditions as in A. Tunicamycin increased BAX expression; pridopidine reduced BAX levels, consistent with pridopidine treatment reducing mitochondrial apoptotic signaling. n = 3 biological replicates. (C) Representative confocal images of caspase-3 immunostaining (green) and Hoechst nuclear staining (blue) in untreated cells, tunicamycin-treated cells, and cells co-treated with tunicamycin plus pridopidine, with or without NE-100. Scale bar, 100mm. (D) Quantification of relative caspase-3 activity under the indicated treatment conditions. Data are expressed as % of untreated control (mean ± SEM). * p < 0.05, ** p < 0.01, **** p < 0.0001 vs. tunicamycin alone (one-way ANOVA with post hoc test).
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Figure 5. Pridopidine improves cell viability in sporadic ALS-NPCs under ER stress. Cell viability was assessed in ALS-NPCs treated with tunicamycin (1.25 µM, 16 h) with or without pridopidine (0.1–1 µM). Viability was measured using a fixable viability dye and quantified by flow cytometry, normalized to untreated controls (set to 100%). n = 3 biological replicates. Data represent mean ± SEM. ** p < 0.01, **** p < 0.0001 vs. tunicamycin alone (two-way ANOVA with post hoc test).
Figure 5. Pridopidine improves cell viability in sporadic ALS-NPCs under ER stress. Cell viability was assessed in ALS-NPCs treated with tunicamycin (1.25 µM, 16 h) with or without pridopidine (0.1–1 µM). Viability was measured using a fixable viability dye and quantified by flow cytometry, normalized to untreated controls (set to 100%). n = 3 biological replicates. Data represent mean ± SEM. ** p < 0.01, **** p < 0.0001 vs. tunicamycin alone (two-way ANOVA with post hoc test).
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Table 1. Summary of iPSC line.
Table 1. Summary of iPSC line.
iPSC Line NameGenderAgeEthnicityGenotype of LocusDiseasePrimary Tissue
CS14isALS-Tnxxmale42UnknownSporadicALSPBMC
Table 2. Primer data.
Table 2. Primer data.
Gene NameFull Gene NamePrimer Sequence
GAPDHGlyceraldehyde-3-phosphate dehydrogenaseForward 5′-CTTTTGCGTCGCCAG-3′
Backward 5′-TTGATGGCAACAATATCCAC-3′
MKI67Marker of proliferation Ki67Forward 5′-GACAGAGGTTCCTAAGAGAG-3′
Backward 5′-AACAATCAGATTTGCTTCCG-3′
BAXBcl2 Associated X,
Apoptosis Regulator		Forward 5′-AACTGGACAGTAACATGGAG-3′
Backward 5′-TTGCTGGCAAAGTAGAAAAG-3′
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MDPI and ACS Style

Meltzer, M.; Zamir, M.S.; Tzuri, N.; Tan, A.M.; Geva, M.; Hayden, M.R.; Lichtenstein, R.G. Pridopidine Protects ALS Patient-Derived Neural Progenitor Cells via Sigma-1 Receptor Activation. Int. J. Mol. Sci. 2026, 27, 3489. https://doi.org/10.3390/ijms27083489

AMA Style

Meltzer M, Zamir MS, Tzuri N, Tan AM, Geva M, Hayden MR, Lichtenstein RG. Pridopidine Protects ALS Patient-Derived Neural Progenitor Cells via Sigma-1 Receptor Activation. International Journal of Molecular Sciences. 2026; 27(8):3489. https://doi.org/10.3390/ijms27083489

Chicago/Turabian Style

Meltzer, May, Maya Shefler Zamir, Noam Tzuri, Andrew M. Tan, Michal Geva, Michael R. Hayden, and Rachel G. Lichtenstein. 2026. "Pridopidine Protects ALS Patient-Derived Neural Progenitor Cells via Sigma-1 Receptor Activation" International Journal of Molecular Sciences 27, no. 8: 3489. https://doi.org/10.3390/ijms27083489

APA Style

Meltzer, M., Zamir, M. S., Tzuri, N., Tan, A. M., Geva, M., Hayden, M. R., & Lichtenstein, R. G. (2026). Pridopidine Protects ALS Patient-Derived Neural Progenitor Cells via Sigma-1 Receptor Activation. International Journal of Molecular Sciences, 27(8), 3489. https://doi.org/10.3390/ijms27083489

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