Role of endolysosome iron in amyloid-β protein-induced increases in reactive oxygen species,mitochondrial depolarization,and cytotoxicity

Abstract

Background

Alzheimer's disease (AD) is a neurodegenerative disease characterized pathologically by the presence of extracellular plaques containing amyloid-β (Aβ) protein, and intraneuronal accumulations of neurotoxic proteins including Aβ and hyperphosphorylated tau. Also implicated in AD pathophysiology are increased levels of reactive oxygen species (ROS) and iron dyshomeostasis. Ferrous iron (Fe²⁺) controls generation of ROS via Fenton-like reactions and Aβ₄₂ increases levels of intracellular ROS. Endosomes and lysosomes (endolysosomes) are acidic organelles that contain high levels of readily releasable stores of Fe²⁺ and lysosomotropic insults can trigger Fe²⁺ release from endolysosomes, which is sufficient to account for increased levels of cytoplasmic Fe²⁺ and ROS.

Objective

We tested the hypothesis that endolysosome stores of Fe²⁺ were sufficient to control Aβ₄₂-induced increases in mitochondrial Fe²⁺ and ROS, mitochondrial depolarization, and cell death.

Methods

Using SH-SY5Y human neuroblastoma cells, we investigated the effects of Aβ₄₂ (500 nM) and Aβ₄₀ (500 nM) peptides on endolysosome and mitochondrial stress responses.

Results

Aβ₄₂ but not Aβ₄₀ accumulated in and significantly decreased endolysosome Fe²⁺ levels. Further, Aβ₄₂ significantly (1) de-acidified endolysosomes, (2) caused endolysosome damage, (3) caused impaired autophagy, (4) increased levels of cytosolic Fe²⁺ and ROS, (5) increased levels of mitochondrial Fe²⁺ and ROS, (6) decreased mitochondrial membrane potentials, and (7) significantly increased cell death by apoptosis and ferroptosis. These neurotoxic effects were blocked by the endolysosome-specific iron chelator deferoxamine.

Conclusions

Endolysosome stores of Fe²⁺ appear to be important regulators of Aβ₄₂-induced cytotoxicity and targeting these iron stores may lead to new therapeutics against AD-like pathology.

Keywords

Alzheimer's disease amyloid-β cell death deferoxamine ferroptosis iron lysosomes mitochondria

Introduction

Alzheimer's disease (AD) is characterized clinically by age-dependent decreases in cognition, and pathologically by the extracellular deposition of amyloid-β (Aβ) proteins. Additionally, intraneuronal accumulation of neurotoxic Aβ proteins, hyperphosphorylated tau protein, and reactive oxygen species (ROS) have been implicated in AD pathophysiology.^1–4 Aβ peptides are processed from amyloid-β protein precursor (AβPP); a process catalyzed by the aspartyl protease β-site amyloid precursor protein cleaving enzyme 1 (BACE-1). BACE-1 is the rate-limiting enzyme that catalyzes the production of Aβ peptides of various lengths including Aβ₄₀ and Aβ₄₂^2,5–7; Aβ₄₀ is more abundant in brain, but Aβ₄₂ is more fibrillogenic, more toxic, and is more closely implicated in AD pathophysiology.^5,8–10

Endosomes and lysosomes (endolysosomes) are acidic organelles that regulate many physiologically important cellular functions and are of pathological relevance to many neurological disorders including AD.^11–13 At the onset of AD pathophysiology, endolysosomes change morphologically and functionally.^11,14,15 Like other organelles, endolysosomes exhibit stress responses; a process known as lysosomal stress responses (LSR).¹⁶ Insult-induced endolysosome de-acidification causes LSR, the release of Ca²⁺ and ferrous iron (Fe²⁺) from endolysosomes, and the accumulation of these divalent cations in the cytoplasm.^16–19 Endolysosomes are further linked to AD pathophysiology because BACE-1 is located mainly in endosomes and gamma-secretase, a second enzyme involved in amyloidogenesis, is localized in lysosomes.^13,20,21 Further, when endolysosomes are de-acidified, protein expression and activity levels of these amyloidogenic enzymes increase.^{20,21,22–24}

Endolysosomes are known as “master regulators of iron metabolism”; they contain readily releasable stores of Fe²⁺ that are mobilized by lysosomotropic insults that de-acidify endolysosomes.^17,19,25 Indeed, the levels of Fe²⁺ in endolysosomes are sufficient to account for insult-induced increases of Fe²⁺ and ROS in the cytosol and in mitochondria,^17,26,27 redox catastrophe and bioenergetic crisis in endolysosomes and mitochondria,²⁸ and increases in regulated cell death (RCD).^9,29,30

Aβ-induced RCD can occur by ferroptosis and apoptosis, although by distinct mechanisms.^31,32 Ferroptosis is an iron-based form of oxidative cell death that is regulated by erastin and cysteine while apoptosis is an energy-dependent form of cell death.^32,33 Common to both forms of RCD is mitochondrial dysfunction and inter-organellar iron dyshomeostasis.^32,34

Because disordered function of endolysosomes, mitochondria and inter-organellar signaling have been implicated in the pathophysiology of neurodegenerative disorders including AD,^12,35,36 we hypothesized that the endolysosome pool of readily releasable Fe²⁺ was sufficient to account for Aβ₄₂ peptide-induced increases in ROS and cell death. We showed here using SH-SY5Y human neuroblastoma cells that the highly fibrillogenic Aβ₄₂ but not Aβ₄₀ (1) trafficked into endolysosomes and de-acidified endolysosomes, (2) decreased endolysosome Fe²⁺ levels, (3) increased levels of cytosolic Fe²⁺ and ROS, (4) increased levels of mitochondrial Fe²⁺ and ROS (5) caused mitochondrial dysfunction, and (6) increased apoptotic and ferroptotic cell death; effects all blocked by chelating endolysosome Fe²⁺ stores with deferoxamine (DFO). Thus, endolysosome Fe²⁺ appears to play a key role in Aβ₄₂-induced neurotoxicity and might be targeted therapeutically to decrease AD neuropathogenesis.^37–44

Methods

Chemicals and biochemicals

DFO mesylate salt was purchased from Sigma Aldrich (cat. no. D9533-1G, St Louis, MO, USA). Unconjugated human Aβ₄₀ and Aβ₄₂ peptides (cat. nos. 1191 and 1428, respectively), ferostatin-1 (cat. no. 5180), ML-162 (cat. no. 7821) were sourced from Tocris, R&D Systems (Minneapolis, MN, USA). The human control Aβ_42-1 peptide (cat. no. ab120301) was purchased from Abcam (Waltham, MA, USA), and conjugated Aβ₄₂ (cat. no. AS-64161) was purchased from Anaspec Inc. (Fremont, CA, USA).

Cell cultures

SH-SY5Y human neuroblastoma cells were purchased from the American Type Culture Collection (cat. no. CRL-2266, Virginia, USA) and grown in Dulbecco's Modified Eagle Medium (Gibco, Life Technologies Corporation, NY, USA) with 10% heat-inactivated fetal bovine serum (cat. no. S10250, R&D Systems, Industry Way, GA, USA) and 1% penicillin/streptomycin (cat. no. 15140122, Invitrogen, Carlsbad, CA) in a humidified incubator maintained at 37°C with 5% CO₂. SH-SY5Y cells were grown in T75 flasks and sub-cultured in 35 mm² cell culture dishes (cat. no. P35GC-0-14-C, MatTek Corporation, Ashland, MA, USA), 100 mm² cell culture dishes (cat. no. 150466, Nunc, ThermoFisher Scientific, Roskilde, Denmark), and 24-well culture plates (Nunc, Thermo Fisher Scientific, Roskilde, Denmark) to about 80% confluency. Cells were passaged every 3 to 4 days by washing with pre-warmed 1× phosphate buffered saline (PBS) (Gibco, Life Technologies Limited, Paisley, UK) and cells were detached from plates using 0.05% trypsin (Invitrogen, Carlsbad, CA, USA). For flow cytometry experiments cells were seeded at a density of 5.0 × 10⁵ cells per well in 24-well cell culture dishes, for protein isolation 8.0 × 10⁶ cells were seeded in 100 mm² culture dishes, and for microscopy experiments 0.3 × 10⁶ cells were seeded in 35 mm² glass-bottom cell culture dishes; cells were treated the following day. Unless otherwise stated specifically, treatments were as follows; control (vehicle), DFO (25 M), and Aβ (500 nM) for 4 to 6 h. For DFO pre-treatments, cells were first treated with DFO for 2 h prior to addition of Aβ. We depended on the documented provenance of the cells provided by ATCC because SH-SY5Y cells are not listed as a commonly misidentified cell line by the International Cell Line Authentication Committee.

Amyloid-β staining and localization in endolysosomes

HiLyte Fluor-647-conjugated Aβ was used to examine Aβ endocytosis. Cells grown to 80% confluency in 35 mm² dishes were pretreated for 2 h with DFO (25 µM) or vehicle prior to the addition of HiLyte Fluor 647-labeled Aβ₄₂ (500 nM) for 4 to 6 h; cells were counter-stained with 50 nM of LysoTracker^TM Green DND-26 (Invitrogen, cat. no. L7526, Carlsbad, USA) for endolysosomes and 1 µg/ml of Hoechst-33342 (cat. no. 62249, ThermoFisher Scientific, Radford, IL, USA) for nuclei. After incubation for 5 min, cells were washed three-times with 1× PBS and Z-stack images were collected using our spinning disk confocal microscope (Oxford Instruments, Concord, MA, USA). To determine the specific endocytic mechanism(s) for the uptake of Aβ₄₂, we pre-treated cells for 2 h with the inhibitor of clathrin-dependent endocytosis dynasore (200 µM), the inhibitor of micropinocytosis methyl β–cyclodextrin (1 mM), the receptor associated protein (RAP) (2 µM) which is an antagonist of low-density lipoprotein receptor-related protein (LRP)-1, and also knocked down LRP-1 mRNA before incubating with HiLyte Fluor 647-labeled Aβ₄₂. LRP-1-deficient mutants were generated by transfecting seeded cells with LRP-1 shRNA plasmid solution (cat. No sc-108061, Santa Cruz Biotechnology, Inc, Dallas, TX, USA). For each set of experiments, the imaging settings remained the same for each treatment and for controls; at least 15 Hoechst-33342-positive cells per microscopic field were imaged without any intentional exclusion. Images were processed using ImageJ software (Version 1.54) to determine the mean fluorescence intensity (MFI) of Hilyte Fluor 647-conjugated Aβ₄₂ and a Pearson correlation test was used to determine the co-localization of Aβ₄₂ (Hilyte Fluor 647) in LysoTracker-positive endolysosomes.

Endolysosome pH

Endolysosome pH was determined using LysoSensor Yellow/Blue DND-160 (cat. no. L7545, ThermoFisher Scientific, Waltham, USA); a ratiometric dual-excitation dye used to measure pH in acidic organelles. SH-SY5Y cells cultured in 35 mm² dishes were exposed to the various treatments, incubated for 5 min at 37°C with media containing DND-160 (10 µM), and then imaged after replacement of DND-160-containing media with fresh media. Using a filter-based microscope imaging system (Zeiss, Germany), DND-160 fluorescence was captured every 30 s at an emission wavelength of 520 nm in response to excitation for 2 ms at wavelengths of 340 and 380 nm. Using a standard curve, we calculated the pH as previously described¹³ with the corresponding changes in proton concentrations within endolysosomes using the formula pH = −log[H⁺].

Endolysosome iron

Levels of Fe²⁺ in endolysosomes were measured using the fluorescence probe FeRhoNox-1 (cat. no. GC901, Goryo Chemicals, Darmstadt, Germany) using methods we and others have described previously.^25,45 The fluorescence of the dye has been shown to be unaffected by changes in pH in the range of 4.0 to 6.0.⁴⁵ Cells were seeded in 35 mm² dishes and grown to 80% confluency before being incubated with 10 µM FeRhoNox-1 at 37°C for 20 min, then addition of 50 nM LysoTracker^TM Green DND-26 (cat. no. L7526, Invitrogen, Carlsbad, USA) and 1 µg/ml Hoechst-33342 (cat. no. 62249, ThermoFisher) for 10 min. Cells were then washed three-times with 1× PBS and imaged with our spinning disk confocal microscope (Oxford Instruments, Concord, MA, USA). Imaging settings remained the same throughout each set of experiments; at least 15 Hoechst-33342-positive cells were imaged per microscopic field without any intentional exclusions. Images were processed using ImageJ software to determine the MFI of FeRhoNox-1. Cells seeded in 24-well dishes were used to confirm FeRhoNox-1 findings using flow cytometry. Following treatments, cells were washed three-times with 1× PBS to remove excess stain, re-suspended in 1× PBS, and data were acquired as we described previously.^19,25 A minimum of 10,000 cells were analyzed per treatment protocol.

Autophagy

Levels of autophagy were determined using Autophagy Assay Red (cat. no. 9156, ImmunoChemistry Technologies, LLC Bloomington, MN, USA) as per manufacturer's protocol. Cells were cultured in 24-well plates to 80% confluency and following the addition of various treatments for 6 h, cells were washed and re-suspended in 490 µL of 1× PBS. Following the addition of 10 µL of 50× autophagy probe, cells were incubated at 37°C for 25 min under dark conditions. Cells were then washed three-times by centrifuging at 200×g for 5 min at room temperature and then re-suspending in cellular assay buffer containing fixative. Data were acquired by flow cytometry at an excitation of 590 nm and emission at 620 nm. At least 10,000 cells were analyzed for each experimental treatment.

Cytosolic Fe²⁺ and ROS measurements

Cytosolic Fe²⁺ and ROS levels were measured using Phen Green^TM FL, diacetate (PGFL-DA) (cat. no P14313, Invitrogen, Carlsbad, USA) and CM-H2DCFDA (cat. no. C6827, Invitrogen, Carlsbad, USA), respectively. Cells were cultured in 24-well plates at 37°C and 5% CO₂ overnight before experimentation. For cytosolic Fe²⁺ detection, cells were incubated with 10 μM PGFL-DA in culture media for 20 min, preincubated for 1 h with DFO, then incubated with various experimental treatments. After incubations, cells were washed three times with PBS, resuspended in 500 μL PBS, and harvested into sterile tubes for flow cytometry. For cytosolic ROS, previously seeded and treated cells were stained with 10 μM CM-H2DCFDA in culture media for 20 min in a 37°C and 5% CO₂ incubator. After incubations, cells were washed three times with PBS, resuspended in 500 μL PBS, and harvested into sterile tubes for flow cytometry. Flow cytometry was performed using our Attune NxT flow cytometer (ThermoFisher, Waltham, USA) to measure the MFI using the respective probe manufacturer's recommended channels. Each sample was measured at least three times, and the average MFI from the measurements were used for analyses.

Mitochondrial iron and reactive oxygen species

Mitochondrial Fe²⁺ levels were measured using rhodamine B 4-[(2, 2′-bipyridin-4-yl) aminocarbonyl] benzyl ester (RDA) (cat. no. 952228-30-7, GuideChem, Milwaukee, USA). Cells grown to about 80% confluency in 24-well culture plates were treated with DFO (25 µM) for 2 h and/or Aβ (500 nM) for 4 to 6 h. After washing cells three-times with 1× PBS, cells were incubated with RDA (100 nM) for 20 min in a 37°C and 5% CO₂ incubator. Cells were then washed three-times with 1× PBS to remove extracellular dye and gently detached into tubes by repeated pipetting. A minimum of 10,000 cells were analyzed by flow cytometry (Attune NxT, ThermoFisher, Waltham, USA) using acquisition settings reported previously.¹⁹ To measure mitochondrial ROS, cultured cells were treated with DFO (25 µM) and/or Aβ (500 nM) for 6 h. Post-treatment the media was replaced, and cells were washed with 1× PBS before incubating with 2.5 µM of MitoSOX Red (cat. no. M36008, Invitrogen, Carlsbad, USA) for 20 min at 37°C. The cells were then rinsed three-times with 1× PBS, detached from the culture dishes by gentle repeated pipetting, and MFI was determined by flow cytometry (Attune NxT, ThermoFisher, Waltham, USA) at an excitation of 530 nm and an emission of 580 nm.

Mitochondrial membrane potential

SH-SY5Y cells were grown to about 80% confluency in 24-well culture dishes and treated with DFO (25 µM) for 2 h and/or Aβ₄₂ (500 nM) for 4 to 6 h in a 37°C and 5% CO₂ incubator. Cells were washed three-times with 1× PBS and then incubated with 100 nM MitoTracker™ Red CMXROS (Invitrogen, cat. no. M7512, Carlsbad, USA) for 15 min at 37°C. Cells were then washed three-times with 1× PBS, re-suspended in 500 µL 1× PBS, and detached from the culture plates by gentle repeated pipetting prior to data acquisition by flow cytometry (Attune NxT, ThermoFisher, Waltham, USA) with settings described previously.¹⁷ Similarly, after treatments with DFO (25 µM) and/or Aβ (500 nM), mitochondrial membrane depolarization was determined using MitoProbe JC-1 Assay Kit (cat. no. M34152, Life Technologies, ThermoFisher). After staining with MitoProbe JC-1 (2 µM, 20 min), cells were washed with 1× PBS, detached by gentle pipetting, and harvested into tubes for flow cytometry. The MFI of MitoProbe JC-1-stained cells was determined at an excitation of 520 nm and emission of 590 nm; data were acquired from at least 10,000 live cells per treatment condition.

Mitochondrial fragmentation

Mitochondrial fragmentation was determined using MitoTracker^TM Green FM (Invitrogen, cat. no. M7514, Carlsbad, CA). Following treatment of cells with DFO (25 µM) for 2 h and/or Aβ₄₂ (500 nM) for 4 to 6 h, cells were incubated with 100 nM MitoTracker^TM Green FM for 20 min in a 37°C and 5% CO₂ incubator. Cells were counter-stained with 1 μg/mL Hoechst 33342 for 10 min and washed three-times with 1× PBS. Z-stack images were acquired from Hoechst-positive cells using a spinning disk confocal microscope (Oxford Instruments, Concord, MA). Imaging settings remained consistent for control and experimental conditions and at least 15 Hoechst-33342-positive cell images were obtained for each field of view; there were no intentional exclusions of any cells from the fields of view. The images were processed using ImageJ software to determine morphological changes in MitoTracker-stained mitochondrial structures.

Cell death and ferroptosis assays

Cells seeded to about 80% confluency in 24-well culture plates were incubated with various treatments for 24 h. Post treatment, cells were harvested into sterile 1.5 ml Eppendorf tubes and centrifuged at 106×g (5418R, Eppendorf, Enfield, CT, USA) for 5 min. After centrifugation, cells were re-suspended in 2 µM staining solution of propidium iodide (PI) (cat. no. 195458, MP Biomedicals, Solon, Ohio, USA) and incubated in the dark for 15 min. The MFI of the PI-stained cell populations was determined by flow cytometry using acquisition settings described previously¹⁷; a minimum of 10,000 cells were analyzed for each treatment. Ferroptosis was determined by measuring protein levels of the anti-ferroptotic marker SLC7A11 as well as levels of lipid peroxidation using C11-BODIPY 581/591 (cat. no. D3861, Invitrogen, Carlsbad, USA) as described previously.^46,47 Cells were cultured to about 80% confluency in 24-well culture dishes before being treated with DFO (25 µM) for 2 h and/or Aβ_1-42 for 24 h. Cells were then incubated with 10 µM of C11-BODIPY 581/591 (cat. no. D3861, Invitrogen Carlsbad, CA) for 30 min at 37°C. Cells were washed with 1× PBS to remove excess stain, gently detached by repeated pipetting, and the MFI of C11-BODIPY 581/591 was acquired by flow cytometry. A minimum of 10,000 cells were analyzed for each treatment.

Western blots

Cells were lysed with 400 µl of 1× cell lysis buffer (cat. no. 9803, Cell Signaling Technology Inc., Danvers, MA, USA) supplemented with protease inhibitor cocktail (Halt Protease Inhibitor Single-Use Cocktail, EDTA-free, ThermoFisher Scientific, Rockford, IL, USA). 20 µg of protein from lysed cell pellets were separated on 4–20% SDS-PAGE, transferred to polyvinyl difluoride membranes (Invitrogen, ThermoFisher Scientific), blocked for 1 h, and incubated with primary antibodies overnight at 4°C. Membranes were washed and incubated with HRP-conjugated or Li-COR secondary antibodies. For HRP-tagged antibodies, membranes were incubated in SuperSignal West Pico PLUS Luminol (cat. no. 34577, ThermoFisher Scientific, USA). Membranes were analyzed using LI-COR Image Studio version 5.x software on a Li-COR Odyssey Fc Instrument. The primary antibodies were as follows; anti-beta actin (NBP1-47423; Novus Biologicals), anti-GAPDH antibody (cat. no. ab9485; Abcam), anti-Galectin-3 (cat. no. AF1197; Novus Biologicals), rabbit anti-p62 (cat. no. H00008878-M01; Novus Biologicals), anti-cytochrome-C (cat. no. sc-13156; Santa Cruz), anti-rabbit SLC7A11 (cat. no. NB300-317; Novus Biologicals), and rabbit anti-GPX4 (cat. no. NBP3-15362; Novus Biologicals).

Statistical analyses

Each experiment was repeated at least three independent times and the data shown are representative images or quantitative measurements represented as means ± S.D. All data values were imported into GraphPad Prism version 10.0.0 for Windows (GraphPad Software, Boston, MA, USA) and statistical analyses were performed using one-way ANOVA followed by Tukey's multiple comparisons test to determine significant differences due to treatment effects. A p value <0.05 was considered statistically significant.

Results

Aβ_42, but not Aβ₄₀, co-localized with LysoTracker-positive endolysosomes

Using Hilyte-647-conjugated Aβ₄₂ and Aβ₄₀ peptides, LysoTracker labelled endolysosomes, and Hoechst-33342 labelled nuclei, we found that Aβ₄₂, but not Aβ₄₀ colocalized with LysoTracker-positive endolysosomes (Figure 1A, B). The mean Pearson's correlation coefficient for co-localization between Aβ₄₂ in cells and LysoTracker-positive endolysosomes was greater than 0.7; the mean correlation coefficient was less than 0.2 for Aβ₄₀. Statistical comparisons between the two groups showed that Aβ₄₂ uptake into endolysosomes was significantly greater than Aβ₄₀. (Figure 1C).

Aβ₄₂ uptake into endolysosomes was LRP-1-mediated

Aβ₄₂ uptake was significantly decreased by 2 h pretreatment with receptor-associated protein (RAP) (2 µM), an antagonist of low-density lipoprotein receptor-related protein-1 (LRP-1). Other endocytosis mechanisms did not appear to be involved with Aβ₄₂ uptake; Aβ₄₂ uptake was not significantly affected by 2 h pretreatments of cells with the inhibitor of clathrin-dependent endocytosis dynasore or the inhibitor of caveolae-dependent pinocytosis methyl β-cyclodextrin (Figure 2A, B). LRP-1 mRNA was knocked down (Figure 2C, D) and this significantly decreased Aβ₄₂ uptake (Figure 2E, F).

DFO and Aβ₄₂ decreased levels of endolysosome Fe²⁺

Aβ₄₂, but not Aβ₄₀, concentration-dependently decreased endolysosome Fe²⁺ levels; 500 nM was the lowest concentration to significantly decrease levels of Fe²⁺ (Figure 3A, B). Aβ_42-1 at concentrations up to 1 μM did not significantly affect levels of endolysosome Fe²⁺ (data not shown). Accordingly, for all subsequent experiments we used 500 nM for Aβ₄₂ and the other Aβ peptides. Using semi-quantitative methods we described previously,^17,25 we found that DFO (25 µM), Aβ₄₂ (500 nM), and pretreatment with DFO (25 µM) significantly decreased endolysosome Fe²⁺ levels (Figure 3C, D). Quantitatively, DFO (25 µM), Aβ₄₂ (500 nM), and DFO plus Aβ₄₂, but not Aβ_42-1 (500 nM) and Aβ₄₀ (500 nM), significantly decreased levels of endolysosome Fe²⁺ (Figure 3E).

Aβ₄₂ deacidified endolysosomes

Aβ₄₂ (500 nM) significantly de-acidified endolysosomes (Figure 4A). DFO (25 µM) significantly acidified endolysosomes, and significantly blocked Aβ₄₂-induced endolysosome de-acidification (Figure 4A). Similarly, when the data were calculated as H⁺ concentration ([H⁺]), DFO significantly increased and Aβ₄₂ significantly decreased [H⁺] (Figure 4B). DFO significantly blocked Aβ₄₂-induced de-acidification (Figure 4A) and Aβ₄₂-induced decreases in endolysosome [H⁺] (Figure 4B).

Aβ₄₂ increased endolysosome damage and autophagy

Aβ₄₂ (500 nM) significantly increased levels of Galectin-3, an endolysosome damage-related protein (Figure 4C, D). DFO (25 µM) significantly decreased the levels of Galectin-3 protein and pretreatment with DFO blocked Aβ₄₂-induced increases in protein levels of Galectin-3 (Figure 4C, D). For the autophagy-related p62 protein, Aβ₄₂ (500 nM) significantly increased p62 protein levels, DFO (25 µM) decreased p62 protein levels, and pretreatment with DFO significantly blocked Aβ₄₂-induced increases in p62 protein levels (Figure 4E, F). Further, Aβ₄₂ (500 nM) increased Autophagy Red Probe MFI, DFO (25 µM) significantly decreased Autophagy Red Probe MFI, and pretreatment with DFO significantly blocked Aβ₄₂ -induced increases in Autophagy Red Probe MFI (Figure 4G).

DFO blocked Aβ₄₂-induced increases in levels of cytosolic Fe²⁺ and ROS

Aβ₄₂ (500 nM) significantly increased levels of Fe²⁺ (Figure 5A) and ROS (Figure 5B) in the cytosol. DFO (25 µM) significantly decreased cytosolic levels of Fe²⁺ and ROS, and pretreatment with DFO significantly blocked Aβ₄₂-induced increases in cytosolic Fe²⁺ and ROS levels (Figure 5A, B).

DFO blocked Aβ₄₂-induced increases in levels of mitochondrial Fe²⁺ and ROS

Aβ₄₂ (500 nM) significantly increased levels of Fe²⁺ (Figure 6A) and ROS in mitochondria (Figure 6B). DFO (25 µM) significantly decreased mitochondrial levels of Fe²⁺ and ROS, and pretreatment with DFO significantly blocked Aβ₄₂-induced increases in mitochondrial Fe²⁺ and ROS levels (Figure 6A, B). Aβ₄₀ (500 nM) and Aβ_42-1 (500 nM) did not significantly affect mitochondrial levels of Fe²⁺ and ROS.

DFO blocked Aβ₄₂-induced mitochondrial fragmentation and depolarization of mitochondrial membrane potentials

Mitochondria stained with 100 nM MitoTracker (Green) were long thread-like structures in control cells and these morphological features were even more pronounced following treatment with DFO (25 µM) (Figure 6C). Aβ₄₂ (500 nM) increased mitochondrial fragmentation; mitochondria were shortened and spherical and DFO blocked Aβ₄₂-induced mitochondrial shortening (Figure 6C). Treatment with Aβ₄₂ significantly decreased mitochondrial membrane potentials (MMP) as measured by MitoTracker CMXROS MFI (Figure 6D). DFO (25 µM) significantly increased MMP, and pretreatment with DFO significantly blocked Aβ₄₂-induced decreases in MMP. Aβ₄₀ (500 nM) and Aβ_42-1 reverse peptide (500 nM) did not significantly affect MMP_. Aβ₄₂ (500 nM) significantly depolarized mitochondrial membranes, DFO (25 µM) reduced mitochondrial membrane depolarization, and DFO blocked Aβ₄₂ (500 nM)-induced depolarization of mitochondrial membranes (Figure 6E).

DFO blocked Aβ₄₂-induced release of cytochrome c from mitochondria into the cytosol and Aβ₄₂-induced increases in cell death

Aβ₄₂ (500 nM) significantly increased, DFO (25 µM) significantly decreased, and pretreatment with DFO significantly blocked Aβ₄₂-induced increases in levels of cytochrome c in cytosol (Figure 6F, G). Aβ₄₀ (500 nM) and Aβ_42-1 (500 nM) did not significantly affect cytosolic levels of cytochrome c. Aβ₄₂ (500 nM) significantly increased, DFO (25 µM) significantly decreased, and pretreatment with DFO significantly blocked Aβ₄₂-induced increases in cell death. Aβ_42-1 (500 nM) and Aβ₄₀ (500 nM) did not significantly affect levels of cell death (Figure 6H).

DFO and FER-1 prevented Aβ₄₂-induced increases in lipid peroxidation and ferroptosis

BODIPY fluorescence staining intensity increases with increased lipid peroxidation^46,48 and Aβ₄₂ (500 nM) significantly increased MFI of BODIPY staining (Figure 7A-C). DFO (25 µM) and FER-1 (10 µM) alone did not significantly affect lipid peroxidation levels, but pretreatment with DFO and FER-1 significantly blocked Aβ₄₂-induced increases in lipid peroxidation (Figure 7A, B). Neither Aβ₄₀ (500 nM) nor Aβ_42-1 (500 nM) significantly affected levels of lipid peroxidation (Figure 7C). Aβ₄₂ (500 nM) increased, DFO (25 µM) but not FER-1 (10 µM) alone significantly decreased, and DFO (25 µM) and FER-1 (10 µM) both significantly blocked Aβ₄₂-induced increases in cell death (Figure 7D). Consistent with our confocal imaging data, we observed by flow cytometry that DFO (25 µM) and FER-1 (10 µM) did not significantly affect lipid peroxidation, but DFO and FER-1 significantly blocked Aβ₄₂-induced increases in lipid peroxidation (data not sown).

DFO and FER-1 blocked Aβ₄₂-induced decreases in levels of intracellular cystine and protein levels of SLC7A11

Aβ₄₂ (500 nM) and the known inducer of ferroptosis ML-162 (5 µM) significantly decreased levels of the anti-ferroptosis protein SLC7A11, and pre-treatment with DFO (25 µM) and FER-1 (10 µM) significantly blocked Aβ₄₂- and ML-162-induced decreases in SLC7A11 protein levels (Figure 8A,B). Because SLC7A11 regulates cellular uptake of cystine, an important precursor in the GPX-4-anti-ferroptotic pathway, we measured intracellular levels of cystine and found that DFO and FER-1 significantly blocked Aβ₄₂-induced decreases in levels of intracellular cystine; DFO and FER-1 alone had no significant effects (Figure 8C). ML-162, as a positive control, significantly decreased intracellular cystine levels and FER-1 significantly blocked the effects of ML-162 (Figure 8C).

Discussion

Endosomes and lysosomes (endolysosomes) are acidic organelles that help regulate important cellular processes including nutrient metabolism, autophagy, and clearance of misfolded proteins implicated in the pathogenesis of neurodegenerative disorders including AD.^12,16,49 AD is characterized pathologically by accumulation of extracellular plaques of misfolded Aβ proteins, intraneuronal highly fibrillogenic and neurotoxic Aβ proteins, hyperphosphorylated tau protein, and intracellular iron and ROS.^10,14,42,50 Aβ is generated in endolysosomes from AβPP through a series of reactions characterized by beta and gamma secretases, neither of which use iron as a co-factor; BACE-1 is the rate-limiting endosome-resident enzyme and gamma secretase is a lysosome-resident enzyme.^6,11,13 Endolysosome dysfunction appears to not only increase the onset of AD, but also contributes to AD persistence and age-related severity.^{11,12,14,51–53}

In AD pathophysiology, structural and functional abnormalities in endolysosomes are some of the earliest events recorded, and lysosome stress responses (LSR) are associated with AD pathophysiology.^11,12,14,50 LSR is characterized by morphological and functional changes including intraluminal deacidification, decreases in endolysosome Fe²⁺ levels, and changes in endolysosome numbers, sizes and positioning relative to the nucleus.^12,16 Insult-induced increases in LSR has been reported for silica nanoparticles, LDL cholesterol, proteins associated with neuro-HIV-1 and ALS as well as licit and illicit weak-base drugs.^{16,17,21–23}^,26,27 Because the pathophysiology of AD has been linked to endolysosome changes as well as increases in Fe²⁺ and ROS levels, we tested the hypothesis that readily releasable stores of endolysosome Fe²⁺ are sufficient to account for Aβ₄₂-induced neurotoxicity.

In AD, different Aβ species are present; Aβ₄₀ is present in higher amounts, but the highly fibrillogenic and neurotoxic Aβ₄₂ appears to be the most relevant to AD pathophysiology.^6,54 We found here that, Aβ₄₂ but not Aβ₄₀ is taken up into endolysosomes, de-acidified endolysosomes and caused inter-organellar iron dyshomeostasis leading to increases in levels of ROS and cell death. Aβ₄₂ uptake was mediated by low-density lipoprotein receptor-related protein-1 (LRP-1) and LRP-1 is abundantly expressed in primary and secondary neural cultures. Aβ₄₂ uptake was significantly reduced when LRP-1 mRNA levels were knocked-down or an antagonist of LRP-1 receptor-associated protein (RAP) was added and these observations confirm that LRP-1 plays an important role in Aβ clearance.^2,6,55,56

Intraluminal pH of endolysosomes ranges from 4.5-5.0,^57,58 and this can be altered by insults that target the vacuolar ATPase and other proton pumps. Similar to other lysosomotropic insults that we have previously reported on,^13,17,19,59 we found that Aβ₄₂ de-acidified endolysosomes. Studies have shown that the secondary structure of Aβ₄₂ is highly ionizable, and this property of Aβ₄₂ may have caused the increases observed in endolysosome pH.^6,60 Also, a recent study has shown that internalized Aβ binds to selected sub-units of the vacuolar-ATPase pump, which primarily maintains luminal pH of endolysosomes, and this interaction triggers dissociation of the proton pump leading to impaired v-ATPase activity.⁶¹ Because de-acidification of endolysosomes is central to LSR and appears to be upstream of mitochondrial dysfunction and neurotoxicity,^26–28^,35,62 it was important for us to examine processes downstream of LSR that may be important for Aβ₄₂-induced neurotoxicity.

Endolysosomes have large reservoirs of readily releasable divalent cations including iron and have been referred to as “master regulators of iron metabolism”.^{17,26,63–65} The two main oxidative states of iron are Fe²⁺ and Fe³⁺; Fe²⁺ is more physiologically important and is implicated in the pathophysiology of AD.^25,66–68 Also implicated in the pathophysiology of AD are increased levels of ROS as well as intraneuronal accumulation of Aβ proteins.^10,14,42,51 Because of the importance of Fe²⁺ in the formation of ROS through Fenton-like chemistry, we determined the role of endolysosome stores of Fe²⁺ in the neurotoxic effects of Aβ peptides. We found that Aβ₄₂ but not Aβ₄₀ decreased levels of endolysosome Fe²⁺ and increased Fe²⁺ levels in the cytosol and in mitochondria. DFO, an endocytosed iron chelator which also acidifies endolysosomes,^17,26 blocked Aβ₄₂-induced increases in endolysosome pH. Mechanistically, decreased levels of Fe²⁺ in endolysosomes might result from Aβ-induced net export of Fe²⁺ out of endolysosomes via various cation channels; effects which are associated with other lysosomotropic stressors.^{19,28,69–71} In the absence of DFO, Aβ₄₂ can hasten the production of Fe²⁺ in endolysosomes by dissociating ferritin-bound Fe³⁺ which favors the chemical reaction catalyzed by the endolysosome six transmembrane epithelial antigen of the prostrate (STEAP)-3 enzyme to produce Fe²⁺.^72,73 Further, decreased levels of endolysosome Fe²⁺ levels and Aβ₄₂-induced endolysosome deacidification may have contributed to endolysosome damage as evidenced by elevated galectin-3 and p-62 autophagy-related protein levels; effects blocked by DFO. These findings demonstrate that Aβ₄₂-induced build-up of autophagosomes and impaired autophagy in AD-like pathophysiology^74–77 may be linked to endolysosome iron dysregulation, similar to previous reports.^{17,19,26,44,78,79} Indeed, endolysosome-resident two pore channel inhibitors have been shown to block cytotoxic effects of lysosomotropic insults; findings consistent with the involvement of endolysosome iron.¹⁹

Aβ₄₂ -induced decreases in levels of endolysosome Fe²⁺ were sufficient to account for the resulting increases in cytosolic and mitochondrial Fe²⁺ levels. These changes in the levels of Fe²⁺ also resulted in Aβ₄₂-induced increases in cytosolic and mitochondrial ROS levels. Excessively high levels of ROS can overwhelm the antioxidant capacity of mitochondria and cause mitochondrial damage.^34,65,80 The depolarization of mitochondrial membranes can result in the release of mitochondrial proteins into the cytosol; a known cause of apoptosis.^81–83 Aβ₄₂ but not Aβ₄₀ significantly reduced mitochondrial membrane potentials and increased cytosolic levels of cytochrome c; effects blocked by DFO. Further, Aβ₄₂-induced mitochondrial membrane depolarization caused mitochondrial fragmentation. All these features are characteristic of insult-induced mitochondrial stress and mitochondrial damage.^17,82,84,85

Neuronal cell death can occur by apoptosis as well as ferroptosis. Ferroptosis is an oxidative process driven by iron-dependent phospholipid accumulation and as such is distinct from other forms of regulated cell death.^32,86 Ferroptosis is uniquely identified by a set of genes including glutathione peroxidase (GPX)-4, which catalyzes the reduction of lipid peroxides to prevent ferroptosis, and by the SLC7A11 antiporter that regulates cysteine metabolism; cysteine is a precursor in the production of GPX-4 substrates.^32,34,87 Aβ₄₂ decreased levels of SLC7A11 and cystine levels which may have reduced GPX-4 levels, and this suggests that Aβ₄₂ may induce ferroptosis by depleting cellular levels of the anti-ferroptotic proteins. Pretreating the cells with FER-1, a known ferroptosis inhibitor, or with DFO blocked Aβ₄₂-induced decreases in levels of SLC7A11, intracellular cystine levels and Aβ₄₂-induced increases in ferroptosis. Although the data here suggests that endolysosome iron stores are involved in ferroptosis via decreased expression of SLC7A11, the release of endolysosome iron can increase levels of ROS and the increased levels of ROS may cause feedback activation of endolysosome redox sensors including transient receptor potential mucolipin-1 (TRPML-1).²⁸ Such a vicious cycle between iron and reactive species suggests that both the release of endolysosome iron and the associated increases in ROS can potentially contribute to Aβ₄₂-induced decreases in expression levels of SLC7A11 and cell death by ferroptosis.^87,88

Endolysosomes dynamically participate in inter-organellar signaling with mitochondria, ER and cytosol,¹⁶ and evidence across multiple experimental models demonstrate that the Fe²⁺ stores in endolysosomes are sufficient to account for insult-induced intracellular iron dyshomeostasis, increases in intracellular ROS, and neurotoxicity. Here, our studies were limited to SH-SY5Y cells. However, we have consistently observed similar lysosomotropic insult-induced changes in cytoplasmic levels of Fe²⁺ and ROS in established cell lines as well as primary rodent and human neural cells.^{17,19,25–28} For example, although endolysosome Fe²⁺ levels were about 21% higher in SH-SY5Y human neuroblastoma (48.6 µM) and 54% higher in U87 astrocytoma cells (50.4 µM) compared to primary human (39.9 µM) and rodent (32.7 µM) neural cultures, ferrous ammonium citrate (FAC) significantly increased and DFO significantly decreased endolysosome Fe²⁺ levels in both primary neural cultures and established cell lines.^25,27 Further, HERV-K Env protein decreased similarly endolysosome Fe²⁺ levels and increased similarly cytosolic and mitochondrial Fe²⁺ and ROS levels in primary human neural cultures and established cell lines.²⁵ In SH-SY5Y and U87 cells, HIV-1 proteins gp120 and Tat decreased endolysosome Fe²⁺ levels and increased cytoplasmic Fe²⁺ levels.^17,19 These effects, including increases in cytoplasmic ROS, were blocked by DFO. Moreover, weak-base drugs decreased endolysosome Fe²⁺ levels and increased cytoplasmic Fe²⁺ and ROS levels in SH-SY5Y and U87 cells; effects blocked by DFO.²⁶ Thus, our results suggest strongly that lysosomotropic insult-induced changes in cytoplasmic levels of Fe²⁺ and ROS are not fundamentally different between primary and established neural cells.

Taken together, this work summarizes that Aβ₄₂ notably decreased endolysosome Fe²⁺ levels and caused downstream neurotoxic effects. Thus, targeting readily releasable stores of Fe²⁺ in endolysosomes may be useful in developing effective therapeutic strategies against AD (Figure 9).

Footnotes

Acknowledgements

The authors gratefully acknowledge present and previous members of the Geiger Laboratory.

ORCID iD

Darius N.K. Quansah

Ethical considerations

Not applicable.

Consent to participate

Not applicable.

Consent for publication

Not applicable.

Author contribution(s)

Darius N.K. Quansah: Conceptualization; Data curation; Formal analysis; Investigation; Methodology; Writing – original draft; Writing – review & editing.

Peter W. Halcrow: Formal analysis; Methodology; Writing – review & editing.

Nirmal Kumar: Methodology; Writing – review & editing.

Braelyn Liang: Methodology; Writing – review & editing.

Jonathan D. Geiger: Conceptualization; Funding acquisition; Methodology; Supervision; Writing – review & editing.

Funding

The authors disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This work in the Geiger laboratory was supported by the National Institute of General Medical Sciences under award number P20GM139759, the National Institute of Mental Health under award number R01MH119000, the National Institute of Neurological Diseases and Stroke under award number 2R01NS065957, and the National Institute of Drug Abuse under award number 2R01DA032444.

Declaration of conflicting interests

The authors declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Data availability statement

The data presented in this work as well as additional relatable information are available from the corresponding author upon reasonable request.

References

Zhao

Atagi

, et al. Apolipoprotein E lipoprotein particles inhibit amyloid-β uptake through cell surface heparan sulphate proteoglycan. Mol Neurodegener 2016; 11: 37.

Baranello

Bharani

Padmaraju

, et al. Amyloid-Beta protein clearance and degradation (ABCD) pathways and their role in Alzheimer’s disease. Curr Alzheimer Res 2015; 12: 32–46.

Kennedy

Stamford

Chen

, et al. The BACE1 inhibitor verubecestat (MK-8931) reduces CNS β-amyloid in animal models and in Alzheimer’s disease patients. Sci Transl Med 2016; 8: 363ra150–363ra150.

Bloom

. Amyloid-β and tau: the trigger and bullet in Alzheimer disease pathogenesis. JAMA Neurol 2014; 71: 505–508.

Canevari

Abramov

Duchen

. Toxicity of Amyloid β peptide: tales of calcium, mitochondria, and oxidative stress. Neurochem Res 2004; 29: 637–650.

Chen

Yan

, et al. Amyloid beta: structure, biology and structure-based therapeutic development. Acta Pharmacol Sin 2017; 38: 1205–1235.

. Advances in molecular biology and clinical study of amyloid precursor protein for Alzheimer’s disease. Zhongguo Yi Xue Ke Xue Yuan Xue Bao 2004; 26: 201–209.

Mayeux

Honig

Tang

M-X

, et al. Plasma Aβ40 and Aβ42 and Alzheimer’s disease: relation to age, mortality, and risk. Neurology 2003; 61: 1185–1190.

Butterfield

Boyd-Kimball

. Amyloid β-peptide(1-42) contributes to the oxidative stress and neurodegeneration found in Alzheimer disease brain. Brain Pathol 2006; 14: 426–432.

10.

Moore

Martin

de Mena

, et al. Short Aβ peptides attenuate Aβ42 toxicity in vivo. J Exp Med 2018; 215: 283–301.

11.

Nixon

. Amyloid precursor protein and endosomal–lysosomal dysfunction in Alzheimer’s disease: inseparable partners in a multifactorial disease. FASEB J 2017; 31: 2729–2743.

12.

Afghah

Chen

Geiger

. Role of endolysosomes and inter-organellar signaling in brain disease. Neurobiol Dis 2020; 134: 104670.

13.

Hui

Chen

Geiger

. Endolysosome involvement in LDL cholesterol-induced Alzheimer’s disease-like pathology in primary cultured neurons. Life Sci 2012; 91: 1159–1168.

14.

Cataldo

Peterhoff

Troncoso

, et al. Endocytic pathway abnormalities precede amyloid β deposition in sporadic Alzheimer’s disease and Down syndrome. Am J Pathol 2000; 157: 277–286.

15.

Rizzollo

Vangheluwe

, et al. The lysosome as a master regulator of iron metabolism. Trends Biochem Sci 2021; 46: 960–975.

16.

Lakpa

Khan

Afghah

, et al. Lysosomal stress response (LSR): physiological importance and pathological relevance. J Neuroimmune Pharmacol 2021; 16: 219–237.

17.

Halcrow

Kumar

Quansah

DNK

, et al. Endolysosome iron chelation inhibits HIV-1 protein-induced endolysosome De-acidification-induced increases in mitochondrial fragmentation, mitophagy, and cell death. Cells 2022; 11: 1811.

18.

Lakpa

Halcrow

Chen

, et al. Readily releasable stores of calcium in neuronal endolysosomes: physiological and pathophysiological relevance. Adv Exp Med Biol 2020; 1131: 681–697.

19.

Halcrow

Lakpa

Khan

, et al. HIV-1 gp120-Induced endolysosome de-acidification leads to efflux of endolysosome iron, and increases in mitochondrial iron and reactive oxygen Species. J Neuroimmune Pharmacol 2022; 17: 181–194.

20.

Chen

Hui

Geiger

. Role of endolysosomes and cholesterol in the pathogenesis of Alzheimer’s disease: insights into why statins might not provide clinical benefit. Austin J Pharmacol Ther 2014; 2: 1035.

21.

Hui

Soliman

Geiger

, et al. Acidifying endolysosomes prevented LDL-induced amyloidogenesis. J Alzheimers Dis JAD 2019; 67: 393–410.

22.

Hui

Lakpa

, et al. Effects of silica nanoparticles on endolysosome function in primary cultured neurons. Can J Physiol Pharmacol 2019; 97: 297–305.

23.

Hui

Soliman

, et al. Antiretroviral drugs promote amyloidogenesis by de-acidifying endolysosomes. J Neuroimmune Pharmacol Off J Soc NeuroImmune Pharmacol 2021; 16: 159–168.

24.

Chen

Hui

Geiger

. Role of LDL cholesterol and endolysosomes in amyloidogenesis and Alzheimer’s disease. J Neurol Neurophysiol 2014; 5: 236.

25.

Halcrow

Kumar

Afghah

, et al. Heterogeneity of ferrous iron-containing endolysosomes and effects of endolysosome iron on endolysosome numbers, sizes, and localization patterns. J Neurochem 2022; 161: 69–83.

26.

Halcrow

Quansah

DNK

Kumar

, et al. Weak base drug-induced endolysosome iron dyshomeostasis controls the generation of reactive oxygen species, mitochondrial depolarization, and cytotoxicity. NeuroImmune Pharmacol Ther 2024; 3: 33–46.

27.

Halcrow

Quansah

DNK

Kumar

, et al. HERV-K (HML-2) envelope protein induces mitochondrial depolarization and neurotoxicity via endolysosome iron dyshomeostasis. J Neurosci 2024; 44: e0826232024.

28.

Kumar

Halcrow

Quansah

DNK

, et al. Involvement of endolysosome iron in HIV-1 gp120-, morphine-, and iron supplementation-induced disruption of the reactive species interactome and induction of neurotoxicity. Redox Rep 2025; 30: 2546496.

29.

Wink

Nims

Saavedra

, et al. The Fenton oxidation mechanism: reactivities of biologically relevant substrates with two oxidizing intermediates differ from those predicted for the hydroxyl radical. Proc Natl Acad Sci U S A 1994; 91: 6604–6608.

30.

Fenton

HJH

. LXXIII.—oxidation of tartaric acid in presence of iron. J Chem Soc Trans 1894; 65: 899–910.

31.

Barber

Mead

Shaw

. Oxidative stress in ALS: a mechanism of neurodegeneration and a therapeutic target. Biochim Biophys Acta BBA - Mol Basis Dis 2006; 1762: 1051–1067.

32.

Dixon

Lemberg

Lamprecht

, et al. Ferroptosis: an iron-dependent form of non-apoptotic cell death. Cell 2012; 149: 1060–1072.

33.

Elmore

. Apoptosis: a review of programmed cell death. Toxicol Pathol 2007; 35: 495–516.

34.

Battaglia

Chirillo

Aversa

, et al. Ferroptosis and cancer: mitochondria meet the “iron maiden” cell death. Cells 2020; 9: 1505.

35.

Khan

Haughey

Nath

, et al. Involvement of organelles and inter-organellar signaling in the pathogenesis of HIV-1 associated neurocognitive disorder and Alzheimer’s disease. Brain Res 2019; 1722: 146389.

36.

Papadopoulos

Meyer

. Detection and clearance of damaged lysosomes by the endo-lysosomal damage response and lysophagy. Curr Biol 2017; 27: R1330–R1341.

37.

Guo

Wang

Zhong

M-L

, et al. Deferoxamine inhibits iron induced hippocampal tau phosphorylation in the Alzheimer transgenic mouse brain. Neurochem Int 2013; 62: 165–172.

38.

Guo

Wang

Zheng

, et al. Intranasal deferoxamine reverses iron-induced memory deficits and inhibits amyloidogenic APP processing in a transgenic mouse model of Alzheimer’s disease. Neurobiol Aging 2013; 34: 562–575.

39.

Fine

Baillargeon

Renner

, et al. Intranasal deferoxamine improves performance in radial arm water maze, stabilizes HIF-1α, and phosphorylates GSK3β in P301L tau transgenic mice. Exp Brain Res 2012; 219: 381–390.

40.

Zhang

Xie

, et al. Neuroprotection by iron chelator against proteasome inhibitor-induced nigral degeneration. Biochem Biophys Res Commun 2005; 333: 544–549.

41.

Xie

, et al. Neuroprotection of deferoxamine on rotenone-induced injury via accumulation of HIF-1α and induction of autophagy in SH-SY5Y cells. Neurochem Int 2010; 57: 198–205.

42.

Long

Holtzman

. Alzheimer disease: an update on pathobiology and treatment strategies. Cell 2019; 179: 312–339.

43.

Velasquez

Wray

. DeferoxamineStatPearls. Treasure Island (FL): StatPearls Publishing, 2023.

44.

Farr

Xiong

. Challenges and opportunities of deferoxamine delivery for treatment of Alzheimer’s disease, Parkinson’s disease, and intracerebral hemorrhage. Mol Pharm 2021; 18: 593–609.

45.

Hirayama

Okuda

Nagasawa

. A highly selective turn-on fluorescent probe for iron(II) to visualize labile iron in living cells. Chem Sci 2013; 4: 1250–1256.

46.

Von Krusenstiern

Robson

Qian

, et al. Identification of essential sites of lipid peroxidation in ferroptosis. Nat Chem Biol 2023; 19: 719–730.

47.

Swanda

, et al. Lysosomal cystine governs ferroptosis sensitivity in cancer via cysteine stress response. Mol Cell 2023; 83: 3347–3359.e9.

48.

Dai

Zhang

Zhou

, et al. Probing lipid peroxidation in ferroptosis: emphasizing the utilization of C11-BODIPY-based protocols. Methods Mol Biol Clifton NJ 2023; 2712: 61–72.

49.

Zhu

Yao

, et al. Lysosomal quality control of cell fate: a novel therapeutic target for human diseases. Cell Death Dis 2020; 11: 1–13.

50.

Cataldo

Petanceska

Terio

, et al. Abeta localization in abnormal endosomes: association with earliest Abeta elevations in AD and down syndrome. Neurobiol Aging 2004; 25: 1263–1272.

51.

Chu

Dodiya

Aebischer

, et al. Alterations in lysosomal and proteasomal markers in Parkinson’s disease: relationship to alpha-synuclein inclusions. Neurobiol Dis 2009; 35: 385–398.

52.

Sardiello

. Transcription factor EB: from master coordinator of lysosomal pathways to candidate therapeutic target in degenerative storage diseases. Ann N Y Acad Sci 2016; 1371: 3–14.

53.

Lee

J-H

Yang

D-S

Goulbourne

, et al. Faulty autolysosome acidification in Alzheimer’s disease mouse models induces autophagic build-up of aβ in neurons, yielding senile plaques. Nat Neurosci 2022; 25: 688–701.

54.

Dai

Chang

Liu

, et al. Aβ-40 Y10F increases βfibrils formation but attenuates the neurotoxicity of amyloid-β peptide. Int J Mol Sci 2012; 13: 5324–5337.

55.

Cam

Zerbinatti

. LRP In amyloid-β production and metabolism. Ann N Y Acad Sci 2006; 1086: 35–53.

56.

Kanekiyo

. The low-density lipoprotein receptor-related protein 1 and amyloid-β clearance in Alzheimer’s disease. Front Aging Neurosci 2014; 6: 93.

57.

Appelqvist

Wäster

Kågedal

, et al. The lysosome: from waste bag to potential therapeutic target. J Mol Cell Biol 2013; 5: 214–226.

58.

Huotari

Helenius

. Endosome maturation. EMBO J 2011; 30: 3481–3500.

59.

Chen

Hui

Soliman

, et al. Altered cholesterol intracellular trafficking and the development of pathological hallmarks of sporadic AD. J Park Dis Alzheimers Dis 2014; 1: 8.

60.

Zhang

Morozova-Roche

, et al. pH-Dependent conformational ensemble and polymorphism of amyloid-β core fragment. J Phys Chem B 2013; 117: 8392–8399.

61.

Kim

S-H

Cho

Y-S

Kim

, et al. Endolysosomal impairment by binding of amyloid beta or MAPT/tau to V-ATPase and rescue via the HYAL-CD44 axis in Alzheimer disease. Autophagy 2023; 19: 2318–2337.

62.

Tyagi

Naik

Tyagi

, et al. HIV infection, neurotoxicity, inflammation, premature aging, and therapeutic challenges to PLWH: an overview. Int J Mol Sci 2026; 27: 2192.

63.

Baringer

Lukacher

Palsa

, et al. Amyloid-β exposed astrocytes induce iron transport from endothelial cells at the blood–brain barrier by altering the ratio of apo- and holo-transferrin. J Neurochem 2023; 167: 248–261.

64.

Tian

, et al. The role of redox-mediated lysosomal dysfunction and therapeutic strategies. Biomed Pharmacother 2023; 165: 115121.

65.

Perluigi

Di Domenico

Butterfield

. Oxidative damage in neurodegeneration: roles in the pathogenesis and progression of Alzheimer disease. Physiol Rev 2024; 104: 103–197.

66.

Koskenkorva-Frank

Weiss

Koppenol

, et al. The complex interplay of iron metabolism, reactive oxygen species, and reactive nitrogen species: insights into the potential of various iron therapies to induce oxidative and nitrosative stress. Free Radic Biol Med 2013; 65: 1174–1194.

67.

Gleason

Bush

. Iron and ferroptosis as therapeutic targets in Alzheimer’s disease. Neurotherapeutics 2021; 18: 252–264.

68.

Lei

Ayton

Bush

. The essential elements of Alzheimer’s disease. J Biol Chem 2020; 296: 100105.

69.

Khan

Halcrow

Lakpa

, et al. Two-pore channels regulate tat endolysosome escape and tat-mediated HIV-1 LTR transactivation. FASEB J 2020; 34: 4147–4162.

70.

Lane

DJR

Ayton

Bush

. Iron and Alzheimer’s disease: an update on emerging mechanisms. J Alzheimers Dis JAD 2018; 64: S379–S395.

71.

Tahirbegi

Pardo

Alvira

, et al. Amyloid Aβ 42, a promoter of magnetite nanoparticle formation in Alzheimer’s disease. Nanotechnology 2016; 27: 465102.

72.

Balejcikova

Siposova

Kopcansky

, et al. Fe(II) formation after interaction of the amyloid β-peptide with iron-storage protein ferritin. J Biol Phys 2018; 44: 237–243.

73.

Everett

Brooks

Lermyte

, et al. Iron stored in ferritin is chemically reduced in the presence of aggregating Aβ(1–42). Sci Rep 2020; 10: 10332.

74.

Jia

Claude-Taupin

, et al. Galectin-3 coordinates a cellular system for lysosomal repair and removal. Dev Cell 2020; 52: 69–87.e8.

75.

Soares

Al-Dalahmah

Hillis

, et al. Novel galectin-3 roles in neurogenesis, inflammation and neurological diseases. Cells 2021; 10: 3047.

76.

Klionsky

Petroni

Amaravadi

, et al. Autophagy in major human diseases. EMBO J 2021; 40: e108863.

77.

Klionsky

. Regulation mechanisms and signaling pathways of autophagy. Annu Rev Genet 2009; 43: 67–93.

78.

Hatcher

Singh

Torti

, et al. Synthetic and natural iron chelators: therapeutic potential and clinical use. Future Med Chem 2009; 1: 1643–1670.

79.

Lloyd

Cable

Rice-Evans

. Evidence that desferrioxamine cannot enter cells by passive diffusion. Biochem Pharmacol 1991; 41: 1361–1363.

80.

Axelsen

Komatsu

Murray

IVJ

. Oxidative stress and cell membranes in the pathogenesis of Alzheimer’s disease. Physiology 2011; 26: 54–69.

81.

Barroso

Taylor

Morshedi

, et al. Mitochondrial membrane potential integrity and plasma membrane translocation of phosphatidylserine as early apoptotic markers: a comparison of two different sperm subpopulations. Fertil Steril 2006; 85: 149–154.

82.

Pieczenik

Neustadt

. Mitochondrial dysfunction and molecular pathways of disease. Exp Mol Pathol 2007; 83: 84–92.

83.

Zorova

Popkov

Plotnikov

, et al. Mitochondrial membrane potential. Anal Biochem 2018; 552: 50–59.

84.

Chu

Zheng

, et al. Mitochondrial mechanisms of apoptosis and necroptosis in liver diseases. Anal Cell Pathol (Amst) 2021; 2021: 8900122.

85.

Teodorof-Diedrich

Spector

. Human immunodeficiency virus type 1 gp120 and tat induce mitochondrial fragmentation and incomplete mitophagy in human neurons. J Virol 2018; 92: e00993–18.

86.

Zhu

Zhou

, et al. Characterization of ferroptosis signature to evaluate the predict prognosis and immunotherapy in glioblastoma. Aging 2021; 13: 17655–17672.

87.

Chen

Cao

Han

, et al. Patulin disrupts SLC7A11-cystine-cysteine-GSH antioxidant system and promotes renal cell ferroptosis both in vitro and in vivo. Food Chem Toxicol 2022; 166: 113255.

88.

Jyotsana

DelGiorno

. The role of cystine/glutamate antiporter SLC7A11/xCT in the pathophysiology of cancer. Front Oncol 2022; 12: 858462.

Role of endolysosome iron in amyloid-β protein-induced increases in reactive oxygen species,mitochondrial depolarization,and cytotoxicity

Abstract

Background

Objective

Methods

Results

Conclusions

Keywords

Introduction

Methods

Chemicals and biochemicals

Cell cultures

Amyloid-β staining and localization in endolysosomes

Endolysosome pH

Endolysosome iron

Autophagy

Cytosolic Fe2+ and ROS measurements

Mitochondrial iron and reactive oxygen species

Mitochondrial membrane potential

Mitochondrial fragmentation

Cell death and ferroptosis assays

Western blots

Statistical analyses

Results

Aβ42, but not Aβ40, co-localized with LysoTracker-positive endolysosomes

Aβ42 uptake into endolysosomes was LRP-1-mediated

DFO and Aβ42 decreased levels of endolysosome Fe2+

Aβ42 deacidified endolysosomes

Aβ42 increased endolysosome damage and autophagy

DFO blocked Aβ42-induced increases in levels of cytosolic Fe2+ and ROS

DFO blocked Aβ42-induced increases in levels of mitochondrial Fe2+ and ROS

DFO blocked Aβ42-induced mitochondrial fragmentation and depolarization of mitochondrial membrane potentials

DFO blocked Aβ42-induced release of cytochrome c from mitochondria into the cytosol and Aβ42-induced increases in cell death

DFO and FER-1 prevented Aβ42-induced increases in lipid peroxidation and ferroptosis

DFO and FER-1 blocked Aβ42-induced decreases in levels of intracellular cystine and protein levels of SLC7A11

Discussion

Footnotes

Acknowledgements

ORCID iD

Ethical considerations

Consent to participate

Consent for publication

Author contribution(s)

Funding

Declaration of conflicting interests

Data availability statement

References

Cytosolic Fe²⁺ and ROS measurements

Aβ_42, but not Aβ₄₀, co-localized with LysoTracker-positive endolysosomes

Aβ₄₂ uptake into endolysosomes was LRP-1-mediated

DFO and Aβ₄₂ decreased levels of endolysosome Fe²⁺

Aβ₄₂ deacidified endolysosomes

Aβ₄₂ increased endolysosome damage and autophagy

DFO blocked Aβ₄₂-induced increases in levels of cytosolic Fe²⁺ and ROS

DFO blocked Aβ₄₂-induced increases in levels of mitochondrial Fe²⁺ and ROS

DFO blocked Aβ₄₂-induced mitochondrial fragmentation and depolarization of mitochondrial membrane potentials

DFO blocked Aβ₄₂-induced release of cytochrome c from mitochondria into the cytosol and Aβ₄₂-induced increases in cell death

DFO and FER-1 prevented Aβ₄₂-induced increases in lipid peroxidation and ferroptosis

DFO and FER-1 blocked Aβ₄₂-induced decreases in levels of intracellular cystine and protein levels of SLC7A11