Next Article in Journal
RETRACTED: Mancinelli et al. The Effects of Taurocholic Acid on Biliary Damage and Liver Fibrosis Are Mediated by Calcitonin-Gene-Related Peptide Signaling. Cells 2022, 11, 1591
Previous Article in Journal
The Spatial Signature of Glioblastoma: A Statistical Re-Assessment of Anatomical Distribution Based on Methylation Subtypes
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Cells
Volume 15
Issue 2
10.3390/cells15020179
cells-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Collapsin Response Mediator Protein 2 (CRMP2) Modulates Induction of the Mitochondrial Permeability Transition Pore in a Knock-In Mouse Model of Alzheimer’s Disease

by
Tatiana Brustovetsky
1,
Rajesh Khanna
2,3 and
Nickolay Brustovetsky
1,4,*
1
Department of Biochemistry, Molecular Biology, and Pharmacology, School of Medicine, Indiana University, Indianapolis, IN 46202, USA
2
Center for Advanced Pain Therapeutics and Research (CAPToR), College of Medicine University of Florida, Gainesville, FL 32610, USA
3
Department of Pharmacology & Therapeutics, and McKnight Brain Institute, College of Medicine University of Florida, Gainesville, FL 32610, USA
4
Stark Neurosciences Research Institute, Indiana University School of Medicine, Indianapolis, IN 46202, USA
*
Author to whom correspondence should be addressed.
Cells 2026, 15(2), 179; https://doi.org/10.3390/cells15020179
Submission received: 12 December 2025 / Revised: 10 January 2026 / Accepted: 15 January 2026 / Published: 19 January 2026
(This article belongs to the Section Mitochondria)
Browse Figures
Versions Notes

Abstract

Hyperphosphorylated collapsin response mediator protein 2 (CRMP2) is elevated in the cerebral cortex of an APP-SAA knock-in mouse model of Alzheimer’s disease and binds the adenine nucleotide translocase (ANT) in a phosphorylation-dependent manner. We propose that, in Alzheimer’s disease (AD) mitochondria, dissociation of hyperphosphorylated CRMP2 from ANT promotes opening of the permeability transition pore (PTP). We showed that purified ANT, when reconstituted into giant liposomes, forms large calcium-dependent channels resembling the PTP, which are effectively blocked by recombinant, unphosphorylated CRMP2. In synaptic mitochondria isolated from the cortices of APP-SAA knock-in mice and control B6J hAbeta mice, we observed an increased susceptibility to permeability transition pore (PTP) induction in AD mitochondria, accompanied by reduced viability of cultured cortical neurons. Pre-treatment of AD mice with the CRMP2-binding small molecule (S)-lacosamide ((S)-LCM), which prevents CRMP2 hyperphosphorylation and restores its interaction with ANT, attenuated PTP induction and improved neuronal viability. Interestingly, direct application of (S)-LCM to isolated mitochondria failed to suppress PTP induction, indicating that its protective effect requires upstream cellular mechanisms. These findings support a phosphorylation-dependent role for CRMP2 in regulating PTP induction in AD mitochondria and highlight (S)-LCM as a promising therapeutic candidate for mitigating mitochondrial dysfunction and enhancing neuronal viability in AD.

1. Introduction

Alzheimer’s disease (AD) is a severe neurodegenerative disorder with no effective cure. Despite decades of intensive research, its underlying mechanisms remain incompletely understood, in part due to the complexity of its pathological processes. While early breakthroughs—such as the identification of amyloid-β (Aβ) and tau—shaped the field over 25 years ago, progress since then has been incremental, underscoring the urgent need for novel insights into AD pathogenesis and therapeutic intervention [1].
The search for molecular drivers of AD has long centered on hallmark pathologies such as amyloid-β plaques and tau tangles. Yet, emerging evidence suggests that disruptions in neuronal signaling and cytoskeletal dynamics may play a pivotal role in disease onset and progression. Among the proteins implicated in these processes is Collapsin Response Mediator Protein 2 (CRMP2), a cytosolic phosphoprotein originally characterized for its role in axonal guidance and neurite outgrowth [2]. Increasing evidence implicates CRMP2 in the pathogenesis of AD [3,4,5,6], where dysregulated phosphorylation mediated by hyperactive kinases such as glycogen synthase kinase-3β (GSK-3β) and cyclin-dependent kinase 5 (Cdk5) contributes to disease-associated cellular abnormalities [7,8,9,10]. Notably, phosphorylation of CRMP2 at specific residues targeted by these kinases is markedly elevated in postmortem AD brains [11,12,13,14,15] and in transgenic mouse models of the disease [13,15,16]. Intriguingly, this hyperphosphorylation is detectable as early as two months of age in AD mice, preceding overt neuropathological symptoms and suggesting a potential initiating role in disease development [13]. Despite these associations, the functional consequences of CRMP2 hyperphosphorylation remain poorly understood. How this post-translational modification contributes to mitochondrial dysfunction and neuronal vulnerability in AD is an open question—one that may hold the key to novel therapeutic strategies.
Although traditionally viewed as a cytosolic protein, CRMP2 has recently been found to associate with mitochondria [17,18], with a subset localized to the intermembrane space between the inner and outer mitochondrial membranes [18]. Within this compartment, CRMP2 interacts with the adenine nucleotide translocase 1 (ANT) [17], a key regulator of mitochondrial bioenergetics [19]. Notably, this interaction is disrupted by phosphorylation of CRMP2—a modification mediated by kinases such as GSK-3β and Cdk5, both of which are hyperactive in AD. Elevated CRMP2 phosphorylation, observed in both human AD brains and transgenic mouse models [11,12,13,14,15,16], suggests that CRMP2 dissociation from the ANT may be an early event in AD pathogenesis. This may lead to reduced ANT activity [20]. However, other potential consequences of this dissociation remain unclear. It remains unclear whether preventing CRMP2 hyperphosphorylation and maintaining its association with the ANT can safeguard mitochondrial integrity and support neuronal survival in AD.
Mitochondria play a central role in buffering cytosolic calcium [21], especially during periods of neuronal activity. However, excessive mitochondrial calcium uptake can trigger an induction of the mitochondrial permeability transition pore (PTP)—a catastrophic event that leads to membrane depolarization, organelle swelling, impaired ATP synthesis, rupture of the outer mitochondrial membrane, and release of pro-apoptotic factors [21,22,23]. Notably, the propensity for PTP induction is elevated in Alzheimer’s disease (AD), potentially driven by interactions between amyloid-β (Aβ) and mitochondrial components such as cyclophilin D and the oligomycin sensitivity conferring protein (OSCP) of the F1F0-ATP synthase complex [24,25,26,27,28,29,30]. While these mechanisms have been implicated in AD-related mitochondrial dysfunction, the precise molecular events that facilitate PTP induction remain incompletely understood. This uncertainty leaves open the possibility that additional, yet unidentified, pathways contribute to mitochondrial vulnerability in AD.
In prior work, we identified bioenergetic deficits in cortical synaptic mitochondria from APP-SAA knock-in mice, a genetically faithful model of AD, which were linked to hyperphosphorylation of CRMP2 and its dissociation from the ANT [20], a key component of the mitochondrial PTP [31,32,33,34,35]. Pharmacological intervention with the small molecule (S)-LCM prevented CRMP2 hyperphosphorylation, preserved its interaction with the ANT, and restored mitochondrial oxidative metabolism [20]. Given the central role of the ANT in PTP regulation [31,32,33,34,35], these findings raised the possibility that CRMP2 phosphorylation status may influence PTP susceptibility in AD mitochondria. In this study, we directly tested this hypothesis and uncovered a previously unrecognized role for CRMP2 in modulating PTP induction, revealing a potential therapeutic axis for mitochondrial protection in AD.

2. Materials and Methods

2.1. Animals

All experimental procedures involving animals were conducted in accordance with the U.S. National Institutes of Health guidelines for the care and use of laboratory animals and were approved by the Institutional Animal Care and Use Committee at Indiana University School of Medicine (protocol #23156 MD/R/E). We employed APP-SAA knock-in (KI) (AD mice) mice [36] (Jackson Laboratory, Bar Harbor, ME, USA, strain #034711), which harbor a humanized Aβ sequence incorporating multiple familial Alzheimer’s disease mutations: R684H, F681Y, G676R in the Aβ region, KM670/671NL (Swedish) in exon 16, and E693G (Arctic) and T714I (Austrian) in exon 17 of the endogenous mouse App gene. These mice model AD pathology without the confounding effects of APP overexpression. Age-matched B6J hAbeta mice (Jackson Laboratory, strain #033013), which express a humanized Aβ1–42 region but lack pathogenic mutations, served as controls. Both male and female mice were included in all experimental groups. Breeding and housing were maintained at the Laboratory Animal Resource Center, Indiana University School of Medicine. The APP-SAA KI model was selected for its physiological relevance and reduced susceptibility to artifacts commonly observed in APP-overexpressing transgenic lines [36].

2.2. Mouse Oral Gavage

To administer compounds, mice received daily oral gavage for 7 consecutive days. (S)-LCM was delivered at a dose of 10 mg/kg body weight, dissolved in dimethyl sulfoxide (DMSO) and diluted in 0.2 mL sterile saline. Control animals received vehicle alone (10 μL DMSO in 0.2 mL saline). Gavage was performed using a precision 0.5 mL syringe fitted with a 20-gauge, ball-tipped feeding needle designed for murine oral delivery (FN-7910, Roboz Surgical Instrument Co., Gaithersburg, MD, USA). All procedures were conducted under standardized conditions to minimize stress and ensure consistent dosing.

2.3. Isolation and Purification of Brain Synaptic Mitochondria

Cortical synaptic mitochondria were isolated using a previously established protocol involving a discontinuous Percoll gradient (24%/40%), as described in detail elsewhere [18]. This method yields highly purified mitochondrial fractions enriched for synaptic populations. The integrity and purity of the resulting preparations have been validated in our prior work [18], confirming minimal contamination from non-mitochondrial compartments and suitability for downstream biochemical analyses.

2.4. Mitochondrial Swelling and Membrane Potential

Mitochondrial swelling was quantified by tracking changes in light scattering at 525 nm within a thermostated (37 °C), continuously stirred 0.3 mL optical chamber. The detection geometry was configured in a backscatter arrangement, with the photodetector positioned directly opposite the incident beam (180°), enhancing sensitivity to volumetric changes in the mitochondrial suspension. Experiments were conducted in a defined KCl-based buffer composed of 125 mM KCl, 0.5 mM MgCl2, 3 mM KH2PO4, 10 mM HEPES (pH 7.4), 10 μM EGTA, 3 mM pyruvate, and 1 mM malate. This medium supported simultaneous monitoring of mitochondrial swelling and membrane potential. Swelling was inferred from a reduction in light scattering, reflecting increased mitochondrial volume. Alamethicin (30 μg/mL), a pore-forming peptide [37], was used to induce maximal swelling, which served as the reference (100%) for normalization [38]. Calcium-induced swelling responses were expressed as a percentage of this alamethicin-defined maximum. To evaluate mitochondrial membrane potential, we employed a tetraphenylphosphonium (TPP+)-selective electrode, enabling real-time quantification of TPP+ distribution between the incubation medium and the mitochondrial matrix [39]. A rise in TPP+ concentration in the medium indicated membrane depolarization, whereas a decline signified polarization [38].

2.5. Mitochondrial Ca2+ Retention Capacity

Mitochondrial Ca2+ uptake was monitored in a 0.3 mL thermostated chamber maintained at 37 °C under continuous stirring, using a miniature Ca2+-selective electrode as previously described [38]. The assay was performed in a KCl-based buffer supplemented with 3 mM pyruvate, 1 mM malate, 0.1 mM ADP, and 1 μM oligomycin to support oxidative metabolism while inhibiting ATP synthase. In experiments with bongkrekic acid (BKA), ADP (and oligomycin) were omitted to avoid interference with the ANT. Calcium was introduced into the medium as sequential 10 μM CaCl2 pulses. Uptake was inferred from the progressive decline in free Ca2+ concentration in the incubation medium, reflecting mitochondrial sequestration. Quantification was expressed as calcium retention capacity, normalized to mitochondrial protein content (μmol Ca2+/mg protein) [38].

2.6. Immunoblotting

Brain tissues were lysed on ice using a buffer composed of 50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1% NP-40, 0.1% SDS, and 1 mM EDTA. Protease and phosphatase activities were inhibited by including commercial inhibitor cocktails (Roche, Indianapolis, IN, USA; Cat. #04906845001 and #04693124001). Homogenates were incubated on ice for 30 min and subsequently centrifuged at 100,000× g for 30 min at 4 °C. The resulting supernatant was collected for SDS-PAGE analysis. Protein samples (20 µg per lane) were resolved on 4–12% Bis-Tris polyacrylamide gels (Invitrogen (Carlsbad, CA, USA), Cat. #NP0335) and transferred to Hybond-ECL nitrocellulose membranes (Amersham Biosciences, (Piscataway, NJ, USA), Cat. #RPN78D). Membranes were blocked for 1 h at room temperature in phosphate-buffered saline (PBS, pH 7.2) containing 5% non-fat dry milk and 0.15% Triton X-100. Primary antibody incubations were performed overnight at 4 °C using rabbit anti-CRMP2 (Sigma, Cat. #C2993, 1:1000) or rabbit anti-ANT 1/2 (Proteintech, (Rosemont, IL, USA), Cat. #15997-1, 1:1000). After blocking, membranes were incubated with the following primary antibodies: sheep anti-CRMP2 pThr509/514 (Kinasource, Dundee, Scotland, UK; Cat. #PB-043; 1:1500), rabbit anti-CRMP2 pSer522 (ECM Biosciences, Versailles, KY, USA; Cat. #CP2191; 1:1500), rabbit anti-CRMP2 (Sigma-Aldrich, St. Louis, MO, USA; Cat. #C2993; 1:1000), rabbit anti-ANT1/2 (Proteintech, Rosemont, IL, USA; Cat. #15997-1; 1:1000), and mouse anti-GAPDH (Abcam, Cambridge, MA, USA; Cat. #ab9484; 1:2000). Blots were then incubated with horseradish peroxidase-conjugated secondary antibodies (goat anti-mouse or goat anti-rabbit IgG; Jackson ImmunoResearch Laboratories, West Grove, PA, USA, 1:25,000 or 1:20,000, respectively) for 1 h at room temperature. Signal detection was carried out using Supersignal West Pico chemiluminescent substrate (Pierce, (Rockford, IL, USA), Cat. #32106). Molecular weight markers were visualized using PageRuler Plus Prestained Protein Ladder (Thermo Fisher, (Waltham, MA, USA), Cat. #26619; 5 µL per lane). Immunoblot images were inverted, and band intensities were quantified by integrated density analysis following background subtraction using Adobe Photoshop (v22.2.0).

2.7. Co-Immunoprecipitation

Synaptic mitochondria were isolated from the cerebral cortex of 4-month-old APP-SAA knock-in mice using Percoll gradient fractionation. Animals were administered either (S)-LCM (10 mg/kg body weight, oral gavage for 7 days), vehicle (10 µL DMSO in 0.2 mL saline, oral gavage for 7 days), or received no treatment. Age-matched B6J hAβ mice were used as controls. Isolated mitochondria were lysed in buffer containing 125 mM KCl, 3 mM KH2PO4, 0.5 mM MgCl2, 10 mM HEPES (pH 7.4), 1% NP-40, 0.1% SDS, and protease inhibitors (Roche). Lysates were pre-cleared with Protein A/G agarose beads (Santa Cruz Biotechnology, Dallas, TX, USA, Cat # sc-2002) for 2 h at 4 °C, then incubated overnight at 4 °C with rabbit anti-CRMP2 (Sigma, Cat # C2993; 1:1000) or rabbit anti-ANT1/2 (Proteintech, Cat # 15997-1; 1:1000) under gentle agitation. Immunocomplexes were captured with Protein A/G beads for 2 h at 4 °C, washed three times with lysis buffer, and eluted by heating at 70 °C in SDS loading dye (Invitrogen). Proteins (20 µg per lane) were resolved on 3–8% Tris–acetate gels (Invitrogen, Cat # EA0375BOX) and immunoblotted as described previously [40]. Membranes were probed with anti-CRMP2 or anti-ANT1/2 (1:1000) and reprobed for bait protein controls to confirm immunoprecipitation efficiency. Input loading controls included anti-VDAC1 (Calbiochem; 1:1000) for CRMP2 and anti-Complex II 70 kDa subunit (Invitrogen; 1:1000) for ANT1/2. Immunoblot images were inverted, and band intensities quantified after background subtraction using Adobe Photoshop 22.2.0. All blots represent at least three independent experiments.

2.8. Mitochondrial Isolation and ANT Reconstitution in Proteoliposomes

Mitochondria were extracted from the brains of ten adult C57BL/6J mice using protocols previously established in our laboratory [18]. To purify the adenine nucleotide translocase (ANT), we adapted methodologies originally developed for yeast mitochondria [41], applying them to mouse brain-derived mitochondrial preparations. For reconstitution of the mouse brain ANT into proteoliposomes, we followed a strategy previously validated for yeast ANT [41]. A lipid mixture was prepared by dissolving 40 mg of phosphatidylcholine (Millipore-Sigma, (Burlington, MA, USA), Cat # 3356) and 1.5 mg of cardiolipin (Millipore-Sigma, Cat # C0563) in 0.45 mL of a detergent-rich buffer containing 11% (w/v) C12E8 (Millipore-Sigma, Cat # P8925), 87 mM Na2SO4, 1 mM EGTA, and 175 mM Tricine-OH (pH 8.0). This lipid solution was then combined with 1.5 mL of the ANT extract, yielding a final protein-to-phospholipid weight ratio of 0.015. The resulting mixture contained 2.5% (w/v) C12E8, 20 mM Na2SO4, 0.23 mM EGTA, 40.4 mM Tricine-OH (pH 8.0), and 9.6 mg/mL phospholipid. Proteoliposome formation was initiated by gradual detergent removal using Amberlite XAD-4 ion exchange beads (Millipore-Sigma, Cat # XAD4). To eliminate residual external components, the proteoliposomes were subsequently passed through a Sephadex G-75 column (30 × 1 cm; Millipore-Sigma, Cat # G75120) pre-equilibrated with a buffer containing 100 mM sucrose, 30 mM Na2SO4, 1 mM Tricine-OH (pH 7.5), and 1 mM EDTA.

2.9. Giant Liposomes

Giant liposomes were generated using a modified dehydration–rehydration technique based on protocols originally developed by Criado and Keller [42] and Schwarz et al. [43], with further refinements as described in our previous studies [32,33]. To ensure sterility and prevent microbial contamination, all solutions used during ANT reconstitution and liposome preparation were filtered through 0.2 μm Millipore membranes. Small ANT-containing proteoliposomes were diluted tenfold with azolectin vesicles, which were prepared from acetone-washed azolectin (type S-II, Sigma). Azolectin (2 mg) was first dissolved in 1 mL of chloroform, evaporated under a nitrogen stream, and rehydrated in 350 μL of buffer containing 100 mM KCl and 5 mM Hepes (pH 7.2). The resulting lipid suspension was extruded sequentially through polycarbonate membranes with pore sizes of 800 nm, 400 nm, and 200 nm using a LiposoFast™ extruder (Avestin Inc., Ottawa, ON, Canada), following established procedures [44]. To form giant liposomes, 5 μL of the vesicle suspension were placed on a glass slide and dehydrated under vacuum in a desiccator containing anhydrous CaCl2 for 5 min at room temperature. Rehydration was performed overnight at 4 °C using 10 μL of a buffer composed of 50 mM KCl and 2.5 mM Hepes (pH 7.2). This process yielded giant liposomes with diameters ranging from 20 to 60 μm, suitable for patch-clamp electrophysiological recordings.

2.10. Patch Clamping

Patch-clamp recordings in the excised patch configuration were conducted following previously established protocols [32,33]. Micropipettes were fabricated from 1.5 mm outer diameter borosilicate glass capillaries with an internal filament (Sutter Instruments (Novato, CA, USA) Cat # BF150-86-10) using a P-1000 micropipette puller (Sutter Instruments). The pipettes were filled with a standard bath solution composed of 100 mM KCl, 2 mM MgCl2, 0.5 mM CaCl2, 4 mM potassium gluconate, 5 mM MES, and 5 mM Tris, adjusted to pH 7.4. The resulting pipette resistance ranged between 3–5 MΩ. For patch formation, 5 μL of giant liposomes containing the ANT were introduced into a 3.0 mL recording chamber pre-filled with the same standard bath solution. Liposomes adhered to the chamber base were gently contacted with the pipette tip to initiate seal formation. A high-resistance seal was typically achieved either spontaneously upon contact or with minimal negative pressure applied via the pipette. Recordings were performed at ambient temperature (~22 °C) using a HEKA EPC-10 amplifier system. Data acquisition and analysis were carried out using PatchMaster v2x80 software (HEKA Elektronik, Holliston, MA, USA). Currents were low-pass filtered at 3.3 kHz and digitized at a sampling rate of 5 kHz. All voltages are reported relative to the pipette electrode potential.

2.11. Recombinant CRMP2 Protein Expression and Purification

Recombinant CRMP2 protein was produced using established bacterial expression protocols [45]. Sequence-verified pGex-Glu-CaV2.2-type channel constructs were introduced into Escherichia coli BL21 (DE3) cells for protein expression. The CRMP2-GST fusion plasmids were generously provided by Dr. Akihiro Kurimasa (Tottori, Japan). Protein induction was initiated with 1 mM isopropyl-β-D-thiogalactopyranoside (IPTG). Following induction, cultures were grown overnight at 16 °C and processed to yield CRMP2-enriched lysates prepared in Tris-based buffer containing 20 mM Tris (pH 7.5), 200 mM NaCl, 0.1 mM EDTA, 1 mM dithiothreitol (DTT), and a protease inhibitor cocktail. Cells were disrupted using an M-110L microfluidizer (Microfluidics Corp., Newton, MA, USA), and detergent extraction with 1% Triton X-100 followed by high-speed centrifugation removed insoluble material. The resulting supernatant was concentrated to 30 mg/mL, and CRMP2-containing preparations were flash-frozen and stored at −80 °C in 25 mM Tris-HCl, 100 mM glycine (pH 7.3), and 10% glycerol for subsequent biochemical analyses [46].

2.12. Statistics

Quantitative data are reported as the mean ± standard deviation (SD), with the number of independent experiments specified in the figure legends. Statistical comparisons were performed using either unpaired two-tailed t-tests or one-way analysis of variance (ANOVA), followed by Bonferroni’s multiple comparison post hoc test where appropriate. All analyses were conducted using GraphPad Prism® software (version 4.0; GraphPad Software Inc., La Jolla, CA, USA). Each dataset was derived from distinct preparations of isolated mitochondria or independently cultured neuronal samples.

3. Results

The adenine nucleotide translocase (ANT), a central component of the mitochondrial permeability transition pore (PTP) [31,32,33,34,35], is known to form large, Ca2+-activated ion channels in phospholipid membranes [32,33]. Given prior evidence of CRMP2 interaction with the ANT [20], we hypothesized that CRMP2 may modulate ANT channel activity and thereby influence the likelihood of PTP induction. To test this, we reconstituted purified ANT into giant liposomes. The ANT was isolated from brain lysates using hydroxyapatite chromatography, which yielded a strong enrichment of ANT protein (Figure 1A) while depleting CRMP2 below detectable levels (Figure 1B). This allowed us to assess ANT channel activity in a CRMP2-free environment. In our experiments, we generated and used giant ANT-reconstituted liposomes with diameters of 20–60 μm, which were suitable for electrophysiological patch-clamp studies (Figure 1C).
Patch-clamp recordings from ANT-containing liposomes revealed a prominent Ca2+-induced ion channel (Figure 2), consistent with previously described Ca2+-dependent ANT pores [32,33]. Channel activity was abolished by bongkrekic acid (BKA), a selective ANT inhibitor and known PTP suppressor [19,32,33,47]. The ANT-related ion currents revealed voltage gating behavior and underwent a closure at high positive voltage (Figure 2D). Notably, application of recombinant unphosphorylated CRMP2 (rCRMP2) led to rapid channel inactivation (Figure 2B,D), suggesting that CRMP2 directly interacts with the ANT pore to block ion conductance. These findings support a model in which CRMP2 binding to ANT suppresses pore formation and may thereby reduce the likelihood of PTP induction.
To evaluate this mechanism in a physiological context, we examined synaptic mitochondria isolated from APP-SAA KI mice (AD) and B6J hAbeta mice (Control, Ctrl). Upon Ca2+ challenge, AD mitochondria exhibited significantly greater depolarization and swelling, as measured by changes in membrane potential and light scattering at 525 nm [38] (Figure 3A,B,D). These responses are indicative of increased proclivity to PTP induction.
We next tested whether pharmacological modulation of CRMP2 phosphorylation could mitigate this vulnerability. Oral administration of (S)-LCM (10 mg/kg for 7 days) reduced CRMP2 phosphorylation, preserved CRMP2–ANT interaction (Supplementary Figures S1 and S2) [20], and protected AD mitochondria from Ca2+-induced PTP activation (Figure 3C,D). In contrast, direct application of (S)-LCM to isolated mitochondria failed to prevent PTP induction (Supplementary Figure S3), suggesting that its protective effect requires intact cellular signaling or upstream modulation of CRMP2 phosphorylation.
To further substantiate these findings, we measured mitochondrial Ca2+ retention capacity—a functional readout inversely correlated with PTP susceptibility [48]. AD mitochondria retained significantly less Ca2+ than Controls (Figure 4A,C), consistent with their heightened propensity to PTP induction. (S)-LCM treatment of AD mice restored Ca2+ retention capacity in AD mitochondria (Figure 4B,C), while (S)-LCM treatment of Control mice had negligible effects on mitochondrial Ca2+ retention capacity (Supplementary Figure S4). Direct application of (S)-LCM to isolated AD mitochondria had no effect (Supplementary Figure S5), again pointing to an indirect mechanism of action. As a positive control, cyclosporin A (CsA), a well-established PTP inhibitor [49,50], significantly increased Ca2+ retention capacity in AD (Figure 4B,C) and Control mitochondria (Supplementary Figure S4). Importantly, CsA failed to further enhance Ca2+ retention in mitochondria from (S)-LCM-treated AD mice (Figure 4B,C), suggesting that both interventions converge on the same pathway to suppress PTP induction.
To determine whether the Ca2+-induced PTP in Control and AD mitochondria was associated with the ANT, we employed bongkrekic acid (BKA), a specific ANT inhibitor and negative modulator of the PTP [19,32,33,47]. ADP and oligomycin were omitted from these experiments to prevent interference with the ANT. Under these conditions, mitochondria accumulated substantially less Ca2+ compared to those incubated with ADP and oligomycin (compare Figure 4 and Figure 5). Nevertheless, AD mitochondria retained only about half as much Ca2+ as control mitochondria (Figure 5). Treatment with BKA markedly increased and equalized Ca2+ retention capacity between the two groups. These results suggest that the Ca2+-induced PTP observed in our experiments is closely linked to the ANT.
Given the link between PTP induction and cell viability, we next asked whether increased PTP sensitivity in AD mitochondria correlates with neuronal vulnerability. In previous work, we showed that cortical neurons from double-transgenic APP/PS1 mice were more prone to cell death than wild-type neurons [51]. Extending this observation, we found that cultured cortical neurons from APP-SAA KI mice exhibited significantly higher cell death compared to neurons from B6J hAbeta controls (Figure 6). While these findings may not directly reflect in vivo neurodegeneration, they suggest that AD neurons are under greater stress in vitro. Notably, treatment with (S)-LCM (10 μM in culture medium for 7 days) conferred neuroprotection, significantly reducing cell death in AD neurons (Figure 6).
Together, our results support a model in which CRMP2 regulates ANT channel activity and PTP induction. Pharmacological preservation of CRMP2–ANT interaction via (S)-LCM confers mitochondrial and neuronal resilience in AD models, highlighting a potential therapeutic strategy for mitigating mitochondrial dysfunction and neurodegeneration.

4. Discussion

In our previous work, we identified CRMP2 hyperphosphorylation in APP-SAA knock-in (KI) mice, which coincided with its dissociation from the adenine nucleotide translocase (ANT) and a reduction in ANT-mediated ADP/ATP transport activity [20]. Beyond its canonical role in ADP/ATP exchange across the inner mitochondrial membrane, the ANT is also implicated in the formation of the mitochondrial permeability transition pore (PTP) [23,31,32,33,35,47,52,53,54,55,56,57,58], a pathological event that compromises mitochondrial integrity [59]. These findings led us to hypothesize that CRMP2 dissociation from the ANT may sensitize mitochondria to PTP induction in Alzheimer’s disease (AD). In the present study, we provide direct evidence that CRMP2 modulates PTP susceptibility in a phosphorylation-dependent manner. Specifically, cortical synaptic mitochondria from APP-SAA KI mice exhibited an increased propensity for PTP induction, which correlated with CRMP2 hyperphosphorylation and its disengagement from the ANT. Notably, pharmacological inhibition of CRMP2 hyperphosphorylation using (S)-LCM [20] mitigated PTP induction, revealing a previously unrecognized mechanism of PTP regulation in AD mitochondria.
PTP induction is known to be elevated in AD [24,25,26], potentially driven by interactions between amyloid-β (Aβ) and mitochondrial proteins such as cyclophilin D or the oligomycin sensitivity conferring protein (OSCP) of the F1F0-ATP synthase complex [24,25,26,27,28,29,30]. Activation of the PTP results in mitochondrial dysfunction, including mitochondrial depolarization, swelling, impaired ATP synthesis, rupture of the outer mitochondrial membrane, and release of pro-apoptotic proteins [21,22,23]. While these mechanisms are well-documented, the precise molecular triggers of PTP induction in AD remain incompletely understood. Given ANT’s established role in PTP formation under conditions of calcium overload [23,31,32,33,35,47,52,53,54,55,56,57,58], its contribution to mitochondrial dysfunction in AD warrants further investigation. Importantly, the potential involvement of CRMP2 in modulating ANT-dependent PTP induction—and its implications for neuronal survival—had not been previously explored.
CRMP2 is a cytosolic phosphoprotein that regulates the localization and activity of various intracellular targets [60]. It is phosphorylated by kinases such as GSK-3β and Cdk5 [12,61,62,63], both of which are hyperactive in AD [7,8,9,10]. Elevated CRMP2 phosphorylation has been observed in postmortem AD brains [11,12,13,14,15] and in transgenic mouse models including APP/PS1 and Tg2576 [13,15,16,64], suggesting its involvement in early AD events [13]. In vitro, brief exposure of cultured cortical neurons to the toxic Aβ25–35 peptide fragment rapidly increases CRMP2 phosphorylation [65], highlighting its sensitivity to Aβ-mediated stress. Our previous work revealed a correlation between CRMP2 hyperphosphorylation, dissociation of phosphorylated CRMP2 from the ANT, and a consequent reduction in ANT transport activity [20]. Yet, other potential functional consequences of CRMP2 hyperphosphorylation have remained elusive. Our study addresses this gap by focusing on CRMP2’s emerging role in regulation of mitochondrial functions and its relevance to AD pathogenesis [18,20,51].
Indeed, CRMP2 has been shown to associate with mitochondria [17,18], with a subset localized to the intermembrane space [18], where it interacts with the ANT1 [17], a central regulator of mitochondrial bioenergetics [19]. Hyperphosphorylation of CRMP2 disrupts this interaction [17], and our findings confirm that this dissociation occurs in AD models [20]. However, the downstream consequences of CRMP2 disengagement from the ANT—particularly in the context of PTP induction—were previously unknown. We now show that preserving CRMP2-ANT interaction by preventing CRMP2 hyperphosphorylation can attenuate PTP induction, suggesting a protective role for CRMP2 in maintaining mitochondrial stability.
Our previous work demonstrated that ANT transport activity is compromised in synaptic mitochondria from APP-SAA KI mice, coinciding with CRMP2 hyperphosphorylation and its dissociation from the ANT [20]. Treatment with (S)-LCM restored CRMP2-ANT interaction and improved ADP/ATP transport across the inner mitochondrial membrane, supporting a model in which CRMP2 positively regulates ANT function. This was further validated using recombinant CRMP2 (rCRMP2) in ANT-reconstituted proteoliposomes, where rCRMP2 enhanced ADP/ATP exchange [20]. Interestingly, CRMP2 may not be the only protein capable of modulating ANT activity. Previous studies have shown that Bcl-2 can stimulate ANT function, while Bax exerts an inhibitory effect [66], suggesting a broader regulatory network involving the ANT and its interacting partners.
Although ANT’s role in ADP/ATP exchange is well established, its involvement in PTP formation has been debated [31,32,33,34,35]. Early studies proposed the ANT as a core component of the PTP [31,32,33,34,35], but the lack of definitive genetic evidence delayed consensus. Genetic ablation studies provided critical insights: Wallace’s group generated mice lacking ANT1 and ANT2 in liver mitochondria and found that PTP induction required significantly higher Ca2+ loads in ANT-deficient mitochondria compared to wild-type [67]. While the authors concluded that ANT is not essential for PTP formation, this interpretation is contentious. For instance, skeletal muscle mitochondria lacking ANT require an eightfold increase in Ca2+ to trigger PTP [68], which strongly suggests that ANT contributes to PTP induction. Rather than negating ANT’s role, these findings imply the existence of compensatory or parallel mechanisms. Indeed, recent studies implicating F1F0-ATP synthase in PTP formation [69,70] support a multifactorial model of PTP regulation.
The PTP triggered in isolated mitochondria (Figure 3 and Figure 4) may involve either the ANT, or ATP synthase, or both. In our experiments, the pronounced effects of BKA—a specific ANT inhibitor and negative modulator of the PTP [19,32,33,47]—strongly indicate that ANT plays a central role in PTP induction. Indeed, we previously found no evidence of CRMP2 binding to ATP synthase [20]. In contrast, unphosphorylated CRMP2 directly interacts with the ANT [20] (Supplementary Figure S2) and inhibits ANT-mediated channel activity (Figure 2). The ANT-associated ion currents identified in our study display strong voltage dependence (Figure 2). These currents remain open at low membrane potentials but progressively close as the potential becomes more positive. Our results indicate that rCRMP2 enhances voltage gating at lower voltages, thereby potentially limiting conditions that favor PTP opening.
Collectively, our data indicate that the CRMP2-dependent modulation of the PTP observed here is mediated primarily through ANT rather than ATP synthase. However, we cannot entirely exclude a contribution of the ATP synthase-associated PTP to mitochondrial dysfunction in AD. Previous studies have demonstrated that induction of the PTP in ANT-deficient cells requires substantially higher Ca2+ concentrations [67,68]. Consequently, it is conceivable that in AD higher Ca2+ may trigger both the ANT-associated and the ATP synthase-related PTP.
The increased susceptibility to PTP induction observed in AD models may have deleterious consequences for neuronal health. In our experiments, cortical neurons from APP-SAA KI (AD) cultured for 21 days (21 DIV) had lower viability compared to cultured cortical neurons from B6J hAbeta (Control). The neuronal death became apparent in both groups, but was significantly more pronounced in neurons derived from AD mice. Although the precise mechanisms driving this increased cell death remain to be elucidated, our data suggest a protective role for CRMP2 modulation. Pre-treatment with the small molecule (S)-LCM (10 µM, administered in the growth medium for 7 days prior to analysis) substantially improved neuronal survival in AD cultures. This neuroprotection correlated with inhibition of CRMP2 hyperphosphorylation at Thr 509/514 and Ser 522, reduced PTP induction, and enhanced mitochondrial resilience. Thus, our findings reveal a mechanistic connection between CRMP2 phosphorylation, PTP regulation, and neuronal survival in Alzheimer’s disease. These results highlight the critical need to identify molecular modulators—such as CRMP2—that could serve as therapeutic targets to maintain mitochondrial integrity and promote neuronal viability in AD.

5. Conclusions

We demonstrate that CRMP2 regulates mitochondrial PTP induction in APP-SAA knock-in mice, a mouse model of Alzheimer’s disease, through phosphorylation-dependent binding to the ANT. Unphosphorylated CRMP2 binds the ANT and suppresses PTP induction, whereas hyperphosphorylated CRMP2 dissociates from the ANT, increasing propensity to PTP induction and correlating with enhanced cell death in APP-SAA cortical neuronal cultures. (S)-lacosamide binds CRMP2, preventing GSK-3β- and Cdk5-mediated hyperphosphorylation at Thr509/514 and Ser522, thereby preserving CRMP2-ANT interaction and impeding PTP induction in APP-SAA mitochondria. These findings establish CRMP2 as a therapeutic target for preventing mitochondrial dysfunction and neurodegeneration in Alzheimer’s disease.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/cells15020179/s1. Figure S1: CRMP2 phosphorylation is elevated in APP-SAA knock-in mice and reduced by (S)-LCM treatment, while total CRMP2 levels remain unchanged; Figure S2: CRMP2–ANT interaction in cortical synaptic mitochondria is disrupted in APP-SAA knock-in mice and restored by (S)-LCM treatment; Figure S3: (S)-lacosamide (10 μM) added directly to synaptic mitochondria isolated from cortices of APP-SAA KI mice (AD) failed to protect from Ca2+-induced mitochondrial swelling and depolarization, indicative of the permeability transition pore (PTP) induction; Figure S4: Ca2+ retention capacity of synaptic mitochondria isolated from cortices of 4-month old B6J hAbeta (Control, Ctrl) mice; Figure S5: (S)-lacosamide (10 μM) added directly to synaptic mitochondria isolated from cortices of APP-SAA KI mice (AD) failed to increase mitochondrial Ca2+ retention capacity.

Author Contributions

T.B. conducted experiments and carried out data analysis; R.K. contributed to data analysis, supplied essential reagents and tools, and participated in manuscript writing; N.B. conceived the study, performed experiments, analyzed data, and co-wrote the manuscript. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by the National Institute of Neurological Disorders and Stroke of the National Institutes of Health under award number R01 NS098772 (to N.B. and R.K.). N.B. additionally received support from the Indiana University Faculty Research Support Program—External Resubmission and from a Biomedical Research Grant from the Indiana University School of Medicine. The content is solely the responsibility of the authors and does not necessarily represent the official views of the Indiana University School of Medicine or the National Institutes of Health.

Institutional Review Board Statement

All animal procedures were approved by the Institutional Animal Care and Use Committee of the Indiana University School of Medicine (protocol #23156 MD/R/E, approved 26 January 2024) and were conducted in accordance with institutional and federal guidelines.

Informed Consent Statement

Not applicable.

Data Availability Statement

Data are contained within the article or Supplementary Materials.

Conflicts of Interest

The authors declare no conflicts of interest.

References

  1. Herrup, K. How Not to Study a Disease: The Stroty of Alzheimer’s; The MIT Press: Cambridge, MA, USA, 2021. [Google Scholar]
  2. Goshima, Y.; Nakamura, F.; Strittmatter, P.; Strittmatter, S.M. Collapsin-induced growth cone collapse mediated by an intracellular protein related to UNC-33. Nature 1995, 376, 509–514. [Google Scholar] [CrossRef] [PubMed]
  3. Hensley, K.; Venkova, K.; Christov, A.; Gunning, W.; Park, J. Collapsin response mediator protein-2: An emerging pathologic feature and therapeutic target for neurodisease indications. Mol. Neurobiol. 2011, 43, 180–191. [Google Scholar] [CrossRef] [PubMed]
  4. Hensley, K.; Kursula, P. Collapsin Response Mediator Protein-2 (CRMP2) is a Plausible Etiological Factor and Potential Therapeutic Target in Alzheimer’s Disease: Comparison and Contrast with Microtubule-Associated Protein Tau. J. Alzheimer’s Dis. 2016, 53, 1–14. [Google Scholar] [CrossRef] [PubMed]
  5. Quach, T.T.; Moutal, A.; Khanna, R.; Deems, N.P.; Duchemin, A.M.; Barrientos, R.M. Collapsin Response Mediator Proteins: Novel Targets for Alzheimer’s Disease. J. Alzheimer’s Dis. 2020, 77, 949–960. [Google Scholar] [CrossRef]
  6. Khanna, R.; Moutal, A.; Perez-Miller, S.; Chefdeville, A.; Boinon, L.; Patek, M. Druggability of CRMP2 for Neurodegenerative Diseases. ACS Chem. Neurosci. 2020, 11, 2492–2505. [Google Scholar] [CrossRef]
  7. Takashima, A. GSK-3 is essential in the pathogenesis of Alzheimer’s disease. J. Alzheimer’s Dis. 2006, 9, 309–317. [Google Scholar] [CrossRef]
  8. Cai, Z.; Zhao, Y.; Zhao, B. Roles of glycogen synthase kinase 3 in Alzheimer’s disease. Curr. Alzheimer Res. 2012, 9, 864–879. [Google Scholar] [CrossRef]
  9. Tsai, L.H.; Lee, M.S.; Cruz, J. Cdk5, a therapeutic target for Alzheimer’s disease? Biochim. Biophys. Acta 2004, 1697, 137–142. [Google Scholar] [CrossRef]
  10. Shukla, V.; Skuntz, S.; Pant, H.C. Deregulated Cdk5 activity is involved in inducing Alzheimer’s disease. Arch. Med. Res. 2012, 43, 655–662. [Google Scholar]
  11. Gu, Y.; Hamajima, N.; Ihara, Y. Neurofibrillary tangle-associated collapsin response mediator protein-2 (CRMP-2) is highly phosphorylated on Thr-509, Ser-518, and Ser-522. Biochemistry 2000, 39, 4267–4275. [Google Scholar] [CrossRef]
  12. Cole, A.R.; Knebel, A.; Morrice, N.A.; Robertson, L.A.; Irving, A.J.; Connolly, C.N.; Sutherland, C. GSK-3 phosphorylation of the Alzheimer epitope within collapsin response mediator proteins regulates axon elongation in primary neurons. J. Biol. Chem. 2004, 279, 50176–50180. [Google Scholar] [CrossRef]
  13. Cole, A.R.; Noble, W.; van Aalten, L.; Plattner, F.; Meimaridou, R.; Hogan, D.; Taylor, M.; LaFrancois, J.; Gunn-Moore, F.; Verkhratsky, A.; et al. Collapsin response mediator protein-2 hyperphosphorylation is an early event in Alzheimer’s disease progression. J. Neurochem. 2007, 103, 1132–1144. [Google Scholar] [CrossRef] [PubMed]
  14. Soutar, M.P.; Thornhill, P.; Cole, A.R.; Sutherland, C. Increased CRMP2 phosphorylation is observed in Alzheimer’s disease; does this tell us anything about disease development? Curr. Alzheimer Res. 2009, 6, 269–278. [Google Scholar] [CrossRef] [PubMed]
  15. Mokhtar, S.H.; Kim, M.J.; Magee, K.A.; Aui, P.M.; Thomas, S.; Bakhuraysah, M.M.; Alrehaili, A.A.; Lee, J.Y.; Steer, D.L.; Kenny, R.; et al. Amyloid-beta-dependent phosphorylation of collapsin response mediator protein-2 dissociates kinesin in Alzheimer’s disease. Neural Regen. Res. 2018, 13, 1066–1080. [Google Scholar] [PubMed]
  16. Petratos, S.; Li, Q.X.; George, A.J.; Hou, X.; Kerr, M.L.; Unabia, S.E.; Hatzinisiriou, I.; Maksel, D.; Aguilar, M.I.; Small, D.H. The beta-amyloid protein of Alzheimer’s disease increases neuronal CRMP-2 phosphorylation by a Rho-GTP mechanism. Brain 2008, 131, 90–108. [Google Scholar] [CrossRef]
  17. Rembutsu, M.; Soutar, M.P.; van Aalten, L.; Gourlay, R.; Hastie, C.J.; McLauchlan, H.; Morrice, N.A.; Cole, A.R.; Sutherland, C. Novel procedure to investigate the effect of phosphorylation on protein complex formation in vitro and in cells. Biochemistry 2008, 47, 2153–2161. [Google Scholar] [CrossRef]
  18. Brustovetsky, T.; Khanna, R.; Brustovetsky, N. CRMP2 Is Involved in Regulation of Mitochondrial Morphology and Motility in Neurons. Cells 2021, 10, 2781. [Google Scholar] [CrossRef]
  19. Klingenberg, M. The ADP and ATP transport in mitochondria and its carrier. Biochim. Biophys. Acta 2008, 1778, 1978–2021. [Google Scholar] [CrossRef]
  20. Brustovetsky, T.; Khanna, R.; Brustovetsky, N. Collapsin Response Mediator Protein 2 (CRMP2) Modulates Mitochondrial Oxidative Metabolism in Knock-In AD Mouse Model. Cells 2025, 14, 647. [Google Scholar] [CrossRef]
  21. Bernardi, P. Mitochondrial transport of cations: Channels, exchangers, and permeability transition. Physiol. Rev. 1999, 79, 1127–1155. [Google Scholar] [CrossRef]
  22. Bonora, M.; Patergnani, S.; Ramaccini, D.; Morciano, G.; Pedriali, G.; Kahsay, A.E.; Bouhamida, E.; Giorgi, C.; Wieckowski, M.R.; Pinton, P. Physiopathology of the Permeability Transition Pore: Molecular Mechanisms in Human Pathology. Biomolecules 2020, 10, 998. [Google Scholar] [CrossRef] [PubMed]
  23. Carraro, M.; Bernardi, P. The mitochondrial permeability transition pore in Ca2+ homeostasis. Cell Calcium 2023, 111, 102719. [Google Scholar] [PubMed]
  24. Du, H.; Guo, L.; Yan, S.; Sosunov, A.A.; McKhann, G.M.; Yan, S.S. Early deficits in synaptic mitochondria in an Alzheimer’s disease mouse model. Proc. Natl. Acad. Sci. USA 2010, 107, 18670–18675. [Google Scholar] [PubMed]
  25. Jia, K.; Du, H. Mitochondrial Permeability Transition: A Pore Intertwines Brain Aging and Alzheimer’s Disease. Cells 2021, 10, 649. [Google Scholar] [CrossRef]
  26. Samanta, S.; Akhter, F.; Roy, A.; Chen, D.; Turner, B.; Wang, Y.; Clemente, N.; Wang, C.; Swerdlow, R.H.; Battaile, K.P.; et al. New cyclophilin D inhibitor rescues mitochondrial and cognitive function in Alzheimer’s disease. Brain 2023, 147, 1710–1725. [Google Scholar] [CrossRef]
  27. Beck, S.J.; Guo, L.; Phensy, A.; Tian, J.; Wang, L.; Tandon, N.; Gauba, E.; Lu, L.; Pascual, J.M.; Kroener, S.; et al. Deregulation of mitochondrial F1FO-ATP synthase via OSCP in Alzheimer’s disease. Nat. Commun. 2016, 7, 11483. [Google Scholar]
  28. Gauba, E.; Chen, H.; Guo, L.; Du, H. Cyclophilin D deficiency attenuates mitochondrial F1Fo ATP synthase dysfunction via OSCP in Alzheimer’s disease. Neurobiol. Dis. 2019, 121, 138–147. [Google Scholar]
  29. Du, H.; Guo, L.; Fang, F.; Chen, D.; Sosunov, A.A.; McKhann, G.M.; Yan, Y.; Wang, C.; Zhang, H.; Molkentin, J.D.; et al. Cyclophilin D deficiency attenuates mitochondrial and neuronal perturbation and ameliorates learning and memory in Alzheimer’s disease. Nat. Med. 2008, 14, 1097–1105. [Google Scholar]
  30. Du, H.; Guo, L.; Zhang, W.; Rydzewska, M.; Yan, S. Cyclophilin D deficiency improves mitochondrial function and learning/memory in aging Alzheimer disease mouse model. Neurobiol. Aging 2011, 32, 398–406. [Google Scholar] [CrossRef]
  31. Woodfield, K.; Ruck, A.; Brdiczka, D.; Halestrap, A.P. Direct demonstration of a specific interaction between cyclophilin-D and the adenine nucleotide translocase confirms their role in the mitochondrial permeability transition. Biochem. J. 1998, 336, 287–290. [Google Scholar] [CrossRef]
  32. Brustovetsky, N.; Klingenberg, M. Mitochondrial ADP/ATP carrier can be reversibly converted into a large channel by Ca2+. Biochemistry 1996, 35, 8483–8488. [Google Scholar] [CrossRef]
  33. Brustovetsky, N.; Tropschug, M.; Heimpel, S.; Heidkamper, D.; Klingenberg, M. A large Ca2+-dependent channel formed by recombinant ADP/ATP carrier from Neurospora crassa resembles the mitochondrial permeability transition pore. Biochemistry 2002, 41, 11804–11811. [Google Scholar] [CrossRef] [PubMed]
  34. Neginskaya, M.A.; Morris, S.E.; Pavlov, E.V. Both ANT and ATPase are essential for mitochondrial permeability transition but not depolarization. iScience 2022, 25, 105447. [Google Scholar] [CrossRef] [PubMed]
  35. Karch, J.; Bround, M.J.; Khalil, H.; Sargent, M.A.; Latchman, N.; Terada, N.; Peixoto, P.M.; Molkentin, J.D. Inhibition of mitochondrial permeability transition by deletion of the ANT family and CypD. Sci. Adv. 2019, 5, eaaw4597. [Google Scholar] [CrossRef] [PubMed]
  36. Xia, D.; Lianoglou, S.; Sandmann, T.; Calvert, M.; Suh, J.H.; Thomsen, E.; Dugas, J.; Pizzo, M.E.; DeVos, S.L.; Earr, T.K.; et al. Novel App knock-in mouse model shows key features of amyloid pathology and reveals profound metabolic dysregulation of microglia. Mol. Neurodegener. 2022, 17, 41. [Google Scholar] [CrossRef]
  37. Bezrukov, S.M.; Vodyanoy, I. Probing alamethicin channels with water-soluble polymers. Effect on conductance of channel states. Biophys. J. 1993, 64, 16–25. [Google Scholar] [CrossRef]
  38. Hamilton, J.; Brustovetsky, T.; Rysted, J.E.; Lin, Z.; Usachev, Y.M.; Brustovetsky, N. Deletion of mitochondrial calcium uniporter incompletely inhibits calcium uptake and induction of the permeability transition pore in brain mitochondria. J. Biol. Chem. 2018, 293, 15652–15663. [Google Scholar] [CrossRef]
  39. Kamo, N.; Muratsugu, M.; Hongoh, R.; Kobatake, Y. Membrane potential of mitochondria measured with an electrode sensitive to tetraphenyl phosphonium and relationship between proton electrochemical potential and phosphorylation potential in steady state. J. Membr. Biol. 1979, 49, 105–121. [Google Scholar] [CrossRef]
  40. Brustovetsky, T.; Bolshakov, A.; Brustovetsky, N. Calpain activation and Na+/Ca2+ exchanger degradation occur downstream of calcium deregulation in hippocampal neurons exposed to excitotoxic glutamate. J. Neurosci. Res. 2010, 88, 1317–1328. [Google Scholar] [CrossRef]
  41. Gawaz, M.; Douglas, M.G.; Klingenberg, M. Structure-function studies of adenine nucleotide transport in mitochondria. II. Biochemical analysis of distinct AAC1 and AAC2 proteins in yeast. J. Biol. Chem. 1990, 265, 14202–14208. [Google Scholar] [CrossRef]
  42. Criado, M.; Keller, B.U. A membrane fusion strategy for single-channel recordings of membranes usually non-accessible to patch-clamp pipette electrodes. FEBS Lett. 1987, 224, 172–176. [Google Scholar] [CrossRef] [PubMed]
  43. Schwarz, M.; Gross, A.; Steinkamp, T.; Flugge, U.I.; Wagner, R. Ion channel properties of the reconstituted chloroplast triose phosphate/phosphate translocator. J. Biol. Chem. 1994, 269, 29481–29489. [Google Scholar] [CrossRef] [PubMed]
  44. MacDonald, R.C.; MacDonald, R.I.; Menco, B.P.; Takeshita, K.; Subbarao, N.K.; Hu, L.R. Small-volume extrusion apparatus for preparation of large, unilamellar vesicles. Biochim. Biophys. Acta 1991, 1061, 297–303. [Google Scholar] [CrossRef] [PubMed]
  45. Brittain, J.M.; Piekarz, A.D.; Wang, Y.; Kondo, T.; Cummins, T.R.; Khanna, R. An atypical role for collapsin response mediator protein 2 (CRMP-2) in neurotransmitter release via interaction with presynaptic voltage-gated calcium channels. J. Biol. Chem. 2009, 284, 31375–31390. [Google Scholar] [CrossRef]
  46. Dustrude, E.T.; Perez-Miller, S.; Francois-Moutal, L.; Moutal, A.; Khanna, M.; Khanna, R. A single structurally conserved SUMOylation site in CRMP2 controls NaV1.7 function. Channels 2017, 11, 316–328. [Google Scholar] [CrossRef]
  47. Hunter, D.R.; Haworth, R.A. The Ca2+-induced membrane transition in mitochondria. I. The protective mechanisms. Arch. Biochem. Biophys. 1979, 195, 453–459. [Google Scholar] [CrossRef]
  48. Schinzel, A.C.; Takeuchi, O.; Huang, Z.; Fisher, J.K.; Zhou, Z.; Rubens, J.; Hetz, C.; Danial, N.N.; Moskowitz, M.A.; Korsmeyer, S.J. Cyclophilin D is a component of mitochondrial permeability transition and mediates neuronal cell death after focal cerebral ischemia. Proc. Natl. Acad. Sci. USA 2005, 102, 12005–12010. [Google Scholar] [CrossRef]
  49. Fournier, N.; Ducet, G.; Crevat, A. Action of cyclosporine on mitochondrial calcium fluxes. J. Bioenerg. Biomembr. 1987, 19, 297–303. [Google Scholar] [CrossRef]
  50. Crompton, M.; Ellinger, H.; Costi, A. Inhibition by cyclosporin A of a Ca2+-dependent pore in heart mitochondria activated by inorganic phosphate and oxidative stress. Biochem. J. 1988, 255, 357–360. [Google Scholar]
  51. Brustovetsky, T.; Khanna, R.; Brustovetsky, N. CRMP2 participates in regulating mitochondrial morphology and motility in Alzheimer’s disease. Cells 2023, 12, 1287. [Google Scholar] [CrossRef]
  52. Ruck, A.; Dolder, M.; Wallimann, T.; Brdiczka, D. Reconstituted adenine nucleotide translocase forms a channel for small molecules comparable to the mitochondrial permeability transition pore. FEBS Lett. 1998, 426, 97–101. [Google Scholar] [CrossRef] [PubMed]
  53. Lee, J.; Schriner, S.E.; Wallace, D.C. Adenine nucleotide translocator 1 deficiency increases resistance of mouse brain and neurons to excitotoxic insults. Biochim. Biophys. Acta 2009, 1787, 364–370. [Google Scholar] [CrossRef] [PubMed]
  54. Carraro, M.; Carrer, A.; Urbani, A.; Bernardi, P. Molecular nature and regulation of the mitochondrial permeability transition pore(s), drug target(s) in cardioprotection. J. Mol. Cell Cardiol. 2020, 144, 76–86. [Google Scholar] [CrossRef] [PubMed]
  55. Bernardi, P. Mechanisms for Ca2+-dependent permeability transition in mitochondria. Proc. Natl. Acad. Sci. USA 2020, 117, 2743–2744. [Google Scholar] [CrossRef]
  56. Bround, M.J.; Bers, D.M.; Molkentin, J.D. A 20/20 view of ANT function in mitochondrial biology and necrotic cell death. J. Mol. Cell Cardiol. 2020, 144, A3–A13. [Google Scholar] [CrossRef]
  57. Neginskaya, M.A.; Solesio, M.E.; Berezhnaya, E.V.; Amodeo, G.F.; Mnatsakanyan, N.; Jonas, E.A.; Pavlov, E.V. ATP Synthase C-Subunit-Deficient Mitochondria Have a Small Cyclosporine A-Sensitive Channel, but Lack the Permeability Transition Pore. Cell Rep. 2019, 26, 11–17. [Google Scholar] [CrossRef]
  58. Bernardi, P.; Carraro, M.; Lippe, G. The mitochondrial permeability transition: Recent progress and open questions. FEBS J. 2022, 289, 7051–7074. [Google Scholar] [CrossRef]
  59. Brustovetsky, N. The Role of Adenine Nucleotide Translocase in the Mitochondrial Permeability Transition. Cells 2020, 9, 2686. [Google Scholar] [CrossRef]
  60. Khanna, R.; Wilson, S.M.; Brittain, J.M.; Weimer, J.; Sultana, R.; Butterfield, A.; Hensley, K. Opening Pandora’s jar: A primer on the putative roles of CRMP2 in a panoply of neurodegenerative, sensory and motor neuron, and central disorders. Future Neurol. 2012, 7, 749–771. [Google Scholar]
  61. Yoshimura, T.; Kawano, Y.; Arimura, N.; Kawabata, S.; Kikuchi, A.; Kaibuchi, K. GSK-3beta regulates phosphorylation of CRMP-2 and neuronal polarity. Cell 2005, 120, 137–149. [Google Scholar] [CrossRef]
  62. Uchida, Y.; Ohshima, T.; Sasaki, Y.; Suzuki, H.; Yanai, S.; Yamashita, N.; Nakamura, F.; Takei, K.; Ihara, Y.; Mikoshiba, K.; et al. Semaphorin3A signalling is mediated via sequential Cdk5 and GSK3beta phosphorylation of CRMP2: Implication of common phosphorylating mechanism underlying axon guidance and Alzheimer’s disease. Genes Cells 2005, 10, 165–179. [Google Scholar] [CrossRef]
  63. Li, T.; Hawkes, C.; Qureshi, H.Y.; Kar, S.; Paudel, H.K. Cyclin-dependent protein kinase 5 primes microtubule-associated protein tau site-specifically for glycogen synthase kinase 3beta. Biochemistry 2006, 45, 3134–3145. [Google Scholar] [CrossRef]
  64. Toba, J.; Nikkuni, M.; Ishizeki, M.; Yoshii, A.; Watamura, N.; Inoue, T.; Ohshima, T. PPARgamma agonist pioglitazone improves cerebellar dysfunction at pre-Abeta deposition stage in APPswe/PS1dE9 Alzheimer’s disease model mice. Biochem. Biophys. Res. Commun. 2016, 473, 1039–1044. [Google Scholar] [CrossRef]
  65. Yang, Z.; Kuboyama, T.; Tohda, C. A Systematic Strategy for Discovering a Therapeutic Drug for Alzheimer’s Disease and Its Target Molecule. Front. Pharmacol. 2017, 8, 340. [Google Scholar] [CrossRef] [PubMed]
  66. Belzacq, A.S.; Vieira, H.L.; Kroemer, G.; Brenner, C. The adenine nucleotide translocator in apoptosis. Biochimie 2002, 84, 167–176. [Google Scholar] [CrossRef] [PubMed]
  67. Kokoszka, J.E.; Waymire, K.G.; Levy, S.E.; Sligh, J.E.; Cai, J.; Jones, D.P.; MacGregor, G.R.; Wallace, D.C. The ADP/ATP translocator is not essential for the mitochondrial permeability transition pore. Nature 2004, 427, 461–465. [Google Scholar] [CrossRef] [PubMed]
  68. Panov, A.V.; Andreeva, L.; Greenamyre, J.T. Quantitative evaluation of the effects of mitochondrial permeability transition pore modifiers on accumulation of calcium phosphate: Comparison of rat liver and brain mitochondria. Arch. Biochem. Biophys. 2004, 424, 44–52. [Google Scholar] [CrossRef]
  69. Giorgio, V.; von Stockum, S.; Antoniel, M.; Fabbro, A.; Fogolari, F.; Forte, M.; Glick, G.D.; Petronilli, V.; Zoratti, M.; Szabo, I.; et al. Dimers of mitochondrial ATP synthase form the permeability transition pore. Proc. Natl. Acad. Sci. USA 2013, 110, 5887–5892. [Google Scholar] [CrossRef]
  70. Alavian, K.N.; Beutner, G.; Lazrove, E.; Sacchetti, S.; Park, H.A.; Licznerski, P.; Li, H.; Nabili, P.; Hockensmith, K.; Graham, M.; et al. An uncoupling channel within the c-subunit ring of the F1FO ATP synthase is the mitochondrial permeability transition pore. Proc. Natl. Acad. Sci. USA 2014, 111, 10580–10585. [Google Scholar] [CrossRef]
Cells 15 00179 g001
Figure 1. ANT purification with hydroxyapatite chromatography (A) and removal of CRMP2 (B). Giant liposomes used in our patch-clamp experiments (C). In (A,B), typical immunoblots demonstrating ANT enrichment in the brain tissue lysates from B6J hAbeta mice following purification with hydroxyapatite (HA) chromatography ((A), after HA) and removal of CRMP2 from this solution ((B), after HA). Mtc, mitochondria. In (C), the giant liposomes with reconstituted ANT were prepared as we described previously [32,33] and measured approximately 40–60 μm in diameter.
Figure 1. ANT purification with hydroxyapatite chromatography (A) and removal of CRMP2 (B). Giant liposomes used in our patch-clamp experiments (C). In (A,B), typical immunoblots demonstrating ANT enrichment in the brain tissue lysates from B6J hAbeta mice following purification with hydroxyapatite (HA) chromatography ((A), after HA) and removal of CRMP2 from this solution ((B), after HA). Mtc, mitochondria. In (C), the giant liposomes with reconstituted ANT were prepared as we described previously [32,33] and measured approximately 40–60 μm in diameter.
Cells 15 00179 g001
Cells 15 00179 g002
Figure 2. Electrophysiological characterization of ANT channel activity and modulation by recombinant CRMP2. (A,B) Representative ion current traces recorded from excised patches of giant liposomes reconstituted with ANT, under a step voltage protocol at a holding potential (Vhold) of +140 mV. Currents were recorded in the presence of 0.5 mM CaCl2 (black traces), with 1 µM bongkrekic acid (Millipore-Sigma, Cat # B6179) (BKA; green traces), with 10 µg/mL recombinant CRMP2 (rCRMP2; blue traces), added to both bath and pipette solutions, and in the absence of Ca2+ in the bath solution (red traces). (C,D) Representative ion currents recorded using a voltage ramp protocol ranging from −200 mV to +200 mV over 10 s, under the same experimental conditions as in panels (A,B). These recordings illustrate the modulatory effects of rCRMP2 and BKA on ANT channel activity and the calcium dependence of the observed currents.
Figure 2. Electrophysiological characterization of ANT channel activity and modulation by recombinant CRMP2. (A,B) Representative ion current traces recorded from excised patches of giant liposomes reconstituted with ANT, under a step voltage protocol at a holding potential (Vhold) of +140 mV. Currents were recorded in the presence of 0.5 mM CaCl2 (black traces), with 1 µM bongkrekic acid (Millipore-Sigma, Cat # B6179) (BKA; green traces), with 10 µg/mL recombinant CRMP2 (rCRMP2; blue traces), added to both bath and pipette solutions, and in the absence of Ca2+ in the bath solution (red traces). (C,D) Representative ion currents recorded using a voltage ramp protocol ranging from −200 mV to +200 mV over 10 s, under the same experimental conditions as in panels (A,B). These recordings illustrate the modulatory effects of rCRMP2 and BKA on ANT channel activity and the calcium dependence of the observed currents.
Cells 15 00179 g002
Cells 15 00179 g003
Figure 3. Induction of the PTP in synaptic mitochondria isolated from cortices of 4-month old APP-SAA KI (AD) and B6J hAbeta (Control, Ctrl) mice. Mitochondria were isolated and PTP induction was assessed as we described previously [38]. Here and in Figure 4, mitochondria were incubated at 37 °C in KCl-based medium supplemented with 1 mM malate plus 3 mM pyruvate [38]. In (A), mitochondria from Ctrl mice. AL, alamethicin a pore-forming agent [37], causing maximal (100%) swelling [38]. In (B) and (C), AD mice were pretreated with either a vehicle (Veh, 10 µL of DMSO in 0.2 mL saline) or 10 mg (S)-LCM/kg, delivered by oral gavage for 7 days prior to experiment, respectively. In (D) and (E), statistical summary of changes in external TPP+ concentration ([TPP+]ext) and light scattering, respectively. Where indicated, 50 μM Ca2+ was applied. Mitochondrial swelling was monitored by following light scattering of mitochondrial suspension (red traces) at 525 nm; mitochondrial membrane potential was followed with a tetraphenylphosphonium (TPP+)-sensitive electrode (black traces) [38]. A decrease in light scattering is indicative of mitochondrial swelling, whereas an increase in [TPP+]ext is indicative of mitochondrial depolarization. Mitochondria were isolated from 4-month old AD and Ctrl mice of both sexes. Data are mean ± SD, N = 5 separate experiments, ** p < 0.01, *** p < 0.001.
Figure 3. Induction of the PTP in synaptic mitochondria isolated from cortices of 4-month old APP-SAA KI (AD) and B6J hAbeta (Control, Ctrl) mice. Mitochondria were isolated and PTP induction was assessed as we described previously [38]. Here and in Figure 4, mitochondria were incubated at 37 °C in KCl-based medium supplemented with 1 mM malate plus 3 mM pyruvate [38]. In (A), mitochondria from Ctrl mice. AL, alamethicin a pore-forming agent [37], causing maximal (100%) swelling [38]. In (B) and (C), AD mice were pretreated with either a vehicle (Veh, 10 µL of DMSO in 0.2 mL saline) or 10 mg (S)-LCM/kg, delivered by oral gavage for 7 days prior to experiment, respectively. In (D) and (E), statistical summary of changes in external TPP+ concentration ([TPP+]ext) and light scattering, respectively. Where indicated, 50 μM Ca2+ was applied. Mitochondrial swelling was monitored by following light scattering of mitochondrial suspension (red traces) at 525 nm; mitochondrial membrane potential was followed with a tetraphenylphosphonium (TPP+)-sensitive electrode (black traces) [38]. A decrease in light scattering is indicative of mitochondrial swelling, whereas an increase in [TPP+]ext is indicative of mitochondrial depolarization. Mitochondria were isolated from 4-month old AD and Ctrl mice of both sexes. Data are mean ± SD, N = 5 separate experiments, ** p < 0.01, *** p < 0.001.
Cells 15 00179 g003
Cells 15 00179 g004
Figure 4. Ca2+ retention capacity of synaptic mitochondria isolated from cortices of 4-month old APP-SAA KI (AD) and B6J hAbeta (Control, Ctrl) mice. The Ca2+ retention capacity was evaluated as we described previously in the KCl-based incubation medium supplemented with 1 mM malate plus 3 mM pyruvate [38]. In (A), synaptic mitochondria from AD and Ctrl mice. In (B), where indicated, mitochondria were isolated from AD mice pretreated with either a vehicle (Veh, 10 µL of DMSO in 0.2 mL saline) or 10 mg/kg (S)-LCM delivered by oral gavage for 7 days prior to mitochondrial isolation, or 1 μM cyclosporin A (CsA, positive control) was added to isolated mitochondria during the experiment. Ca2+ uptake and Ca2+ retention by mitochondria were monitored by following Ca2+ concentration in the medium ([Ca2+]ext) with a Ca2+-selective electrode [38]. To assess mitochondrial Ca2+ retention capacity, multiple pulses of 10 µM Ca2+ were applied to mitochondria until a failure to take Ca2+ up and the release of accumulated Ca2+. In (C), statistical summary. Data are mean ± SD, ** p < 0.01, *** p < 0.001, N = 5 separate experiments.
Figure 4. Ca2+ retention capacity of synaptic mitochondria isolated from cortices of 4-month old APP-SAA KI (AD) and B6J hAbeta (Control, Ctrl) mice. The Ca2+ retention capacity was evaluated as we described previously in the KCl-based incubation medium supplemented with 1 mM malate plus 3 mM pyruvate [38]. In (A), synaptic mitochondria from AD and Ctrl mice. In (B), where indicated, mitochondria were isolated from AD mice pretreated with either a vehicle (Veh, 10 µL of DMSO in 0.2 mL saline) or 10 mg/kg (S)-LCM delivered by oral gavage for 7 days prior to mitochondrial isolation, or 1 μM cyclosporin A (CsA, positive control) was added to isolated mitochondria during the experiment. Ca2+ uptake and Ca2+ retention by mitochondria were monitored by following Ca2+ concentration in the medium ([Ca2+]ext) with a Ca2+-selective electrode [38]. To assess mitochondrial Ca2+ retention capacity, multiple pulses of 10 µM Ca2+ were applied to mitochondria until a failure to take Ca2+ up and the release of accumulated Ca2+. In (C), statistical summary. Data are mean ± SD, ** p < 0.01, *** p < 0.001, N = 5 separate experiments.
Cells 15 00179 g004
Cells 15 00179 g005
Figure 5. Ca2+ retention capacity of synaptic mitochondria from cortical tissue of 4-month-old APP-SAA KI (AD) and B6J hAβ (Control) mice: the effect of bongkrekic acid (BKA). Synaptic mitochondria were incubated in a KCl-based medium supplemented with 1 mM malate and 3 mM pyruvate, without ADP or oligomycin, as previously described [38]. Ca2+ uptake and retention were monitored by measuring extracellular Ca2+ concentration ([Ca2+]ext) using a Ca2+-selective electrode [38]. To determine retention capacity, sequential 10 μM Ca2+ pulses were applied until mitochondria failed to take up additional Ca2+ and released accumulated Ca2+. Panels (A) and (B) show representative traces from synaptic mitochondria of control and AD mice, respectively. Where indicated, mitochondria were treated with 5 μM BKA. Panel (C) summarizes Ca2+ retention capacity across groups. Data are presented as mean ± SD; ** p < 0.01, *** p < 0.001; N = 5 independent experiments.
Figure 5. Ca2+ retention capacity of synaptic mitochondria from cortical tissue of 4-month-old APP-SAA KI (AD) and B6J hAβ (Control) mice: the effect of bongkrekic acid (BKA). Synaptic mitochondria were incubated in a KCl-based medium supplemented with 1 mM malate and 3 mM pyruvate, without ADP or oligomycin, as previously described [38]. Ca2+ uptake and retention were monitored by measuring extracellular Ca2+ concentration ([Ca2+]ext) using a Ca2+-selective electrode [38]. To determine retention capacity, sequential 10 μM Ca2+ pulses were applied until mitochondria failed to take up additional Ca2+ and released accumulated Ca2+. Panels (A) and (B) show representative traces from synaptic mitochondria of control and AD mice, respectively. Where indicated, mitochondria were treated with 5 μM BKA. Panel (C) summarizes Ca2+ retention capacity across groups. Data are presented as mean ± SD; ** p < 0.01, *** p < 0.001; N = 5 independent experiments.
Cells 15 00179 g005
Cells 15 00179 g006
Figure 6. Differential apoptotic and necrotic cell death in cortical neurons from APP-SAA KI (AD model) and B6J hAbeta (Control) mice. Primary cortical neurons were cultured for 21 days and stained with Hoechst 33342 (blue, nuclear marker), YO-PRO-1 (green, apoptotic marker), and propidium iodide (red, necrotic marker) to assess cell death phenotypes. (A,B), representative bright field and fluorescence micrographs of neurons derived from Control mice. (C,D), neurons from APP-SAA KI mice treated with vehicle (Veh; 0.01% DMSO) for 7 days prior to analysis show increased apoptotic and necrotic markers. (E,F), neurons from APP-SAA KI mice incubated with 10 µM (S)-LCM for 7 days exhibit reduced cell death, indicating neuroprotective effect. (G,H), quantitative analysis of apoptotic and necrotic cell populations across experimental conditions. Data represent mean ± SD from five independent experiments using neurons from separate platings. Statistical significance was determined by one-way ANOVA followed by post hoc testing; *** p < 0.001.
Figure 6. Differential apoptotic and necrotic cell death in cortical neurons from APP-SAA KI (AD model) and B6J hAbeta (Control) mice. Primary cortical neurons were cultured for 21 days and stained with Hoechst 33342 (blue, nuclear marker), YO-PRO-1 (green, apoptotic marker), and propidium iodide (red, necrotic marker) to assess cell death phenotypes. (A,B), representative bright field and fluorescence micrographs of neurons derived from Control mice. (C,D), neurons from APP-SAA KI mice treated with vehicle (Veh; 0.01% DMSO) for 7 days prior to analysis show increased apoptotic and necrotic markers. (E,F), neurons from APP-SAA KI mice incubated with 10 µM (S)-LCM for 7 days exhibit reduced cell death, indicating neuroprotective effect. (G,H), quantitative analysis of apoptotic and necrotic cell populations across experimental conditions. Data represent mean ± SD from five independent experiments using neurons from separate platings. Statistical significance was determined by one-way ANOVA followed by post hoc testing; *** p < 0.001.
Cells 15 00179 g006
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Brustovetsky, T.; Khanna, R.; Brustovetsky, N. Collapsin Response Mediator Protein 2 (CRMP2) Modulates Induction of the Mitochondrial Permeability Transition Pore in a Knock-In Mouse Model of Alzheimer’s Disease. Cells 2026, 15, 179. https://doi.org/10.3390/cells15020179

AMA Style

Brustovetsky T, Khanna R, Brustovetsky N. Collapsin Response Mediator Protein 2 (CRMP2) Modulates Induction of the Mitochondrial Permeability Transition Pore in a Knock-In Mouse Model of Alzheimer’s Disease. Cells. 2026; 15(2):179. https://doi.org/10.3390/cells15020179

Chicago/Turabian Style

Brustovetsky, Tatiana, Rajesh Khanna, and Nickolay Brustovetsky. 2026. "Collapsin Response Mediator Protein 2 (CRMP2) Modulates Induction of the Mitochondrial Permeability Transition Pore in a Knock-In Mouse Model of Alzheimer’s Disease" Cells 15, no. 2: 179. https://doi.org/10.3390/cells15020179

APA Style

Brustovetsky, T., Khanna, R., & Brustovetsky, N. (2026). Collapsin Response Mediator Protein 2 (CRMP2) Modulates Induction of the Mitochondrial Permeability Transition Pore in a Knock-In Mouse Model of Alzheimer’s Disease. Cells, 15(2), 179. https://doi.org/10.3390/cells15020179

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop