Elevated mitochondrial ROS and dysfunction

Within the inner mitochondrial membrane, the electron transport chain (ETC) functions to couple the transfer of electrons from NADH and FADH₂ to molecular oxygen with the translocation of protons, ultimately driving ATP synthesis. During this process, partial reduction of oxygen can lead to the generation of ROS, primarily superoxide (O₂•⁻), which can subsequently be converted to other reactive species such as hydrogen peroxide (H₂O₂) and hydroxyl radicals (•OH) (Brand et al. 2010; Brand et al. 2016). Complex I (NADH:ubiquinone oxidoreductase) (Cocheme et al. 2008; Sharma et al. 2009) and complex III (ubiquinol:cytochrome c oxidoreductase) (Castello et al. 2007) are recognized as major sites of ROS production under both physiological and pathophysiological conditions. 

Fig. 1. Reactive oxygen species. Two molecules of superoxide can react to generate hydrogen peroxide (H2O2) in a reaction known as dismutation, which is accelerated by the enzyme superoxide dismutase (SOD). In the presence of iron, superoxide and H2O2 react to generate hydroxyl radicals. In addition to superoxide, H2O2 and hydroxyl radicals, other reactive oxygen species (ROS) occur in biological systems., which can be generated from singlet oxygen by antibody molecules. The colour coding indicates the reactivity of individual molecules (yellow, limited reactivity; orange, moderate reactivity; red, high reactivity and non-specificity) (Adapted by permission from Macmillan Publishers Ltd, Lambeth, 2004, copyright (2004)). 

Complex I and ROS Formation. Complex I is the largest and most intricate component of the ETC, responsible for the transfer of electrons from NADH to ubiquinone (CoQ) (Sharma et al. 2009). This multistep process involves the passage of electrons through a flavin mononucleotide (FMN) and a series of iron–sulfur (Fe–S) clusters before reaching the CoQ-binding site. ROS formation at complex I primarily occurs through two distinct mechanisms: the "forward" and "reverse" electron transport (RET) modes. In the forward mode, electrons flow from NADH through the FMN and Fe–S centers to ubiquinone, which is reduced to ubiquinol. During this process, a small percentage of electrons may leak prematurely to molecular oxygen, generating superoxide, primarily at the FMN site (Cadenas et al. 1977; Turrens et al. 1980; Turrens et al. 1985). This leakage is relatively limited under normal physiological conditions but can increase when the NADH/NAD⁺ ratio is high or when electron flow downstream is inhibited (Wong et al. 2017; Quinlan et al. 2014; Lambert et al. 2004). The RET mechanism is a potent source of ROS, especially under conditions of high proton motive force and elevated levels of reduced CoQ (Miwa et al. 2003). In this scenario, electrons can flow in reverse from ubiquinol back to complex I’s FMN site, leading to substantial ROS generation. This is particularly evident during conditions such as ischemia-reperfusion, when succinate accumulation during ischemia leads to its rapid oxidation and massive ROS production upon reperfusion. 

Complex III and ROS Formation. Complex III transfers electrons from ubiquinol to cytochrome c via the Q-cycle, a mechanism involving two distinct ubiquinone-binding sites: the Qo site (outer) and the Qi site (inner). ROS production by complex III is primarily associated with the Qo site, where a semiquinone intermediate can reduce oxygen to superoxide (Dröse et al. 2011; Dröse et al. 2008; Dröse 2009; Votyakova et al. 2001). This occurs on the outer surface of the inner mitochondrial membrane, allowing superoxide to access both the mitochondrial matrix and the intermembrane space. 

Under normal conditions, the ROS produced by complex III are relatively low and are efficiently detoxified by mitochondrial antioxidant systems such as superoxide dismutases (SODs) and glutathione peroxidases. However, when electron flow through complex III is perturbed — for instance, by inhibition of the Qo site (e.g., by antimycin A) — the lifetime of the semiquinone intermediate is prolonged, significantly enhancing ROS production.(Quinlan et al. 2011; Sarewicz et al. 2010; Erecinska et al. 1976; Dröse et al. 2008; Zhang et al. 1998; Quinlan et al. 2012). 

Physiological vs. Pathophysiological ROS Production. Under physiological conditions, mitochondrial ROS serve as important signaling molecules that modulate processes including hypoxic response, cellular proliferation, and apoptosis (Ullrich et al. 2014; Handy et al. 2012). Low-level ROS production is tightly regulated and balanced by antioxidant defenses, ensuring redox homeostasis. In contrast, pathophysiological conditions such as ischemia-reperfusion injury, neurodegenerative diseases, and metabolic disorders are characterized by dysregulation of ETC components and antioxidant systems (Lin et al. 2000). This leads to excessive and sustained ROS production, causing oxidative damage to mitochondrial DNA, lipids, and proteins, further impairing mitochondrial function and contributing to disease progression (Kowaltoski et al. 2009). 

Oxidative Modifications of Mitochondrial Proteins. Mitochondrial proteins, particularly those involved in oxidative phosphorylation (OXPHOS), are highly susceptible to ROS-induced oxPTMs due to their proximity to ROS-generating sites within the electron transport chain (ETC). Cysteine thiol groups are especially reactive and can undergo a range of oxidative modifications, including sulfenylation (–SOH), disulfide bond formation, S-glutathionylation, sulfinylation (–SO₂H), and irreversible sulfonylation (–SO₃H). Additionally, methionine residues may be oxidized to methionine sulfoxide, and tyrosine residues can form dityrosine cross-links or undergo nitration in the presence of peroxynitrite (ONOO⁻). These oxPTMs alter protein conformation, catalytic activity, and protein–protein interactions. For example, oxidation of critical thiols in ATP synthase or adenine nucleotide translocase (ANT) impairs ATP production and disrupts nucleotide exchange across the inner mitochondrial membrane. Similarly, oxidative inactivation of mitochondrial aconitase, a key tricarboxylic acid (TCA) cycle enzyme containing an iron–sulfur cluster, leads to metabolic dysfunction. Proteins of the mitochondrial quality control system, including chaperones and proteases, can also be oxidatively modified, further exacerbating proteotoxic stress and contributing to mitochondrial dysfunction Chung et al. 2013; Mu et al. 2024; Mailloux et al. 2013; Chinta et al. 2011; Brown et al. 2004; Clementi et al. 1998; Nakagawa et al. 2007; Gardner et al. 2002; Tortora et al. 2007; Castro et al. 1994; Aulak et al. 2004; Tretter et al. 1999; Tretter et al. 2000; Yang et al. 2008). 

Oxidative Damage to Mitochondrial DNA. Mitochondrial DNA, located in close proximity to the inner mitochondrial membrane where the ETC resides, is highly vulnerable to ROS-induced damage due to its lack of protective histones and limited DNA repair capacity. Oxidative lesions in mtDNA include strand breaks, base modifications such as 8-oxo-2'-deoxyguanosine (8-oxo-dG), and abasic sites. Among these, 8-oxo-dG is a prominent biomarker of oxidative DNA damage and can result in G→T transversions during replication. ROS-induced damage to mtDNA leads to impaired transcription and translation of mitochondrial-encoded subunits of respiratory complexes, which further compromises OXPHOS efficiency and creates a vicious cycle of increasing ROS production. mtDNA mutations and deletions accumulate over time, contributing to aging and degenerative disease phenotypes. Additionally, oxidized mtDNA can be released into the cytosol, where it acts as a damage-associated molecular pattern (DAMP), triggering inflammatory responses via activation of the cGAS–STING pathway or inflammasomes (Cooke et al. 2003; Evans et al. 2004; Floyd et al. 1986; Chen et al. 2010; Sanders et al. 2017; Bender et al. 2006; Sharma et al. 2019; Druzhyna et al. 2008; Hahm et al. 2022; Shokolenko et al. 2014). 

Lipid Peroxidation in Mitochondrial Membranes. The mitochondrial inner membrane is rich in polyunsaturated fatty acids, particularly cardiolipin, which is essential for the structural organization and function of respiratory supercomplexes. ROS, especially hydroxyl radicals, initiate lipid peroxidation through the abstraction of hydrogen atoms from lipid acyl chains, generating lipid peroxyl radicals and hydroperoxides. These reactive lipid species propagate oxidative damage across the membrane and compromise its integrity and fluidity. Peroxidation of cardiolipin disrupts the optimal assembly of ETC complexes and impairs cytochrome c anchoring. Oxidized cardiolipin also promotes the release of cytochrome c into the cytosol, a key step in the initiation of intrinsic apoptosis. Moreover, lipid peroxidation products such as 4-hydroxynonenal (4-HNE) and malondialdehyde (MDA) form covalent adducts with mitochondrial proteins, further impairing enzymatic function and contributing to oxidative stress signaling and cell death pathways (Schlame et al. 1993; Chu et al. 2013; Skulachev et al. 2010; Bayir et al. 2006; Milazzo et al. 2017; Xiao et al. 2017; Chen et al. 1994; Mohammadi-Bardbori et al. 2008; Satoh et al. 1997; Chen et al. 2024; Bernardi et al. 2023; Riojas-Hernandez et al. 2015). 

Consequences for Mitochondrial Function. Collectively, oxidative modifications of mitochondrial proteins, DNA, and lipids disrupt mitochondrial bioenergetics, including impaired electron transport, ATP depletion, altered redox balance, and dysregulated calcium handling. Mitochondrial membrane potential (Δψm) collapses under severe oxidative stress, leading to the opening of the mitochondrial permeability transition pore (mPTP), mitochondrial swelling, and eventual release of pro-apoptotic factors. These events mark the progression from mitochondrial dysfunction to cell death via apoptosis or necrosis. Furthermore, persistent oxidative damage impairs mitophagy, the selective removal of damaged mitochondria, allowing dysfunctional organelles to accumulate and propagate ROS signaling. In chronic disease states, this contributes to a feed-forward loop of mitochondrial damage, inflammation, and tissue degeneration (Frank et al. 2012). 

Amplex Red Ultra assay. The Amplex® Red Ultra assay has emerged as a sensitive and specific technique for the quantitative measurement of hydrogen peroxide (H₂O₂), a primary and relatively stable ROS generated by mitochondria (Grivennikova et al. 2018). The assay is based on the horseradish peroxidase (HRP)-catalyzed reaction between H₂O₂ and the Amplex Red Ultra reagent (10-acetyl-3,7-dihydroxyphenoxazine), producing the highly fluorescent compound resorufin. The fluorescence intensity of resorufin is directly proportional to the amount of H₂O₂ present, allowing real-time monitoring of mitochondrial ROS production. In a mitochondrial context, ROS primarily originate as superoxide anions (O₂•⁻), predominantly produced at complexes I and III of the electron transport chain. These anions are rapidly dismutated to H₂O₂ by mitochondrial superoxide dismutase (SOD2) in the matrix and SOD1 in the intermembrane space. The resultant H₂O₂ diffuses through membranes and accumulates in the surrounding medium, where it can be quantitatively detected using the Amplex Red Ultra assay (Munro et al. 2019; Quinlan et al. 2013). To perform the assay, isolated mitochondria or permeabilized cells are incubated in a reaction buffer containing Amplex Red Ultra, HRP, and appropriate mitochondrial substrates and inhibitors. The choice of substrate (e.g., glutamate/malate for complex I or succinate for complex II) and respiratory conditions (e.g., state 2, 3, or uncoupled respiration) allows the assessment of ROS production under defined bioenergetic states. Inhibitors such as rotenone (complex I), antimycin A (complex III), or myxothiazol can be used to modulate electron flow and enhance ROS formation at specific complexes, enabling mechanistic dissection of ROS sources (Murphy et al. 2022). Importantly, care must be taken to avoid artifacts from exogenous ROS or enzyme autoxidation, and all reactions should be controlled for background fluorescence (Starkov et al. 2010; Tretter et al. 2014). The improved formulation of Amplex Red Ultra offers enhanced resistance to autoxidation and greater photostability compared to conventional Amplex Red, making it particularly suitable for high-sensitivity assays and extended kinetic measurements. The assay is typically performed using a microplate reader set to excitation and emission wavelengths of ~530 nm and ~590 nm, respectively. The high sensitivity of the Amplex Red Ultra assay allows detection of H₂O₂ in the picomolar to low nanomolar range, making it well-suited for studies requiring precise quantification of mitochondrial ROS output. 

MitoSOX. MitoSOX™ dyes are a family of fluorogenic probes specifically designed for the selective detection of mitochondrial superoxide in live cells. The most commonly used derivative, MitoSOX™ Red, is a mitochondrial-targeted derivative of hydroethidine (HE), conjugated to a triphenylphosphonium (TPP⁺) moiety that drives its accumulation into the mitochondrial matrix due to the negative membrane potential (Δψm) (Zielonka et al. 2010; Mukhopadhyay et al. 2007). Upon entry into the mitochondria, MitoSOX Red is oxidized specifically by superoxide to form a highly fluorescent ethidium-like product that binds to mitochondrial DNA, emitting red fluorescence upon excitation. The reaction between superoxide and MitoSOX is rapid and relatively selective, although some degree of nonspecific oxidation can occur, particularly in the presence of other oxidants such as hydrogen peroxide, peroxynitrite, or hydroxyl radicals (Roelofs et al. 2015). Therefore, appropriate experimental controls are essential to validate superoxide specificity, including the use of superoxide dismutase mimetics or mitochondrial uncouplers that reduce Δψm and consequently MitoSOX uptake. In practical applications, live cells or isolated mitochondria are incubated with MitoSOX Red under controlled conditions, typically at concentrations of 2–5 μM for 10–30 minutes at 37°C. The cells are then washed to remove excess dye, and fluorescence is detected using fluorescence microscopy, flow cytometry, or plate-based fluorometry. The fluorescence excitation and emission maxima for the oxidized product are approximately 510 nm and 580 nm, respectively (Kauffman et al. 2010). Quantitative analysis requires normalization to mitochondrial content or membrane potential, often using additional stains such as MitoTracker Green or TMRE. MitoSOX Red is particularly valuable in experiments aiming to identify the specific sites or triggers of mitochondrial superoxide production. For instance, mitochondrial ROS generation can be stimulated by respiratory chain inhibitors such as rotenone (complex I) or antimycin A (complex III), redox cyclers which enhance electron leakage and superoxide formation. Similarly, metabolic substrates can modulate ROS production by altering electron flow and redox status within the ETC. In these contexts, MitoSOX allows for real-time detection of dynamic changes in ROS generation in response to pharmacological or genetic perturbations. 

Electron Paramagnetic Resonance (EPR). Electron paramagnetic resonance (EPR) spectroscopy, also known as electron spin resonance (ESR), is a powerful and direct analytical technique for the detection of unpaired electrons in paramagnetic species, including free radicals such as reactive oxygen species (ROS). In the context of mitochondrial research, EPR provides a highly specific and quantitative method for detecting and characterizing ROS generated within the mitochondrial respiratory chain, especially under conditions of oxidative stress or during the evaluation of mitochondrial-targeted therapies (Chen et al. 2014). Unlike fluorescence-based probes, which infer ROS presence through secondary reactions, EPR directly detects the presence of short-lived radical species or their stabilized spin-adducts. Native ROS such as superoxide (O₂•⁻), hydroxyl radicals (•OH), and nitric oxide (NO•) are typically too reactive and short-lived to be measured directly by EPR. Therefore, EPR detection of mitochondrial ROS relies primarily on the use of spin-trapping agents (Griendling et al. 2016). These compounds react with transient radicals to form more stable radical adducts that can be detected by EPR. Among the most widely used spin traps for mitochondrial studies is 5,5-dimethyl-1-pyrroline N-oxide (DMPO), which forms stable adducts with O₂•⁻ and •OH. Alternatively, cyclic nitrones such as DEPMPO and EMPO offer enhanced specificity and longer half-lives of the spin-adducts, improving sensitivity in complex biological systems (Hardy et al. 2007; Hardy et al. 2014).  

To measure mitochondrial ROS formation using EPR, isolated mitochondria or permeabilized cells are incubated with suitable substrates and respiratory chain inhibitors in the presence of the spin trap. For example, in the presence of succinate and antimycin A, mitochondria generate high levels of ROS at complex III, leading to enhanced spin-adduct formation detectable by EPR. Similarly, reverse electron transport from succinate oxidation can be induced to stimulate superoxide production at complex I, allowing site-specific investigation of ROS sources. EPR spectra provide detailed information on the chemical environment of the radical species, including hyperfine splitting constants and g-values, which help identify the specific type of ROS involved. Quantification is achieved by integrating the signal intensity of the spin-adduct, which is proportional to the radical concentration. To improve signal specificity and reduce background noise, mitochondria-targeted spin traps such as mito-TEMPO or mito-CP (cyclic nitroxides conjugated to lipophilic cations like triphenylphosphonium) can be employed. These agents accumulate within mitochondria due to the membrane potential and provide localized detection of ROS, particularly superoxide. One of the key advantages of EPR is its ability to distinguish between different radical species, a capability that is not matched by conventional fluorescence or chemiluminescence assays. Furthermore, EPR enables real-time kinetic monitoring of radical generation and decay under tightly controlled experimental conditions. However, the technique also presents challenges: it requires specialized instrumentation, careful sample preparation under anaerobic or low-temperature conditions to preserve radical species, and relatively high concentrations of spin traps, which may perturb native redox equilibria. 

Coelenterazine chemiluminescence is a sensitive and non-invasive analytical method for detecting elevated mitochondrial reactive oxygen species (ROS), particularly superoxide (O₂•⁻) and hydrogen peroxide (H₂O₂), in intact cells and isolated mitochondria (Lucas et al. 1992; Teranishi et al. 1997). Coelenterazine, a luminescent substrate originally derived from marine organisms such as Renilla and Aequorea, undergoes oxidative decarboxylation upon reaction with ROS to form coelenteramide, emitting photons in the blue spectral range (typically 480–500 nm). The intensity of this light emission is directly proportional to the amount of ROS present, enabling real-time quantification of oxidative stress under physiological and pathophysiological conditions. The utility of coelenterazine for mitochondrial ROS detection lies in its cell-permeable and lipophilic nature, which allows it to diffuse across membranes and accumulate in intracellular compartments, including the mitochondrial matrix. Several coelenterazine derivatives have been developed to enhance specificity and mitochondrial targeting, such as coelenterazine-h and mito-coelenterazine, which exhibit improved sensitivity for mitochondrial superoxide and reduced background luminescence. These derivatives preferentially localize to mitochondria due to their increased hydrophobicity or conjugation with lipophilic cationic moieties, facilitating the detection of ROS specifically originating from the mitochondrial electron transport chain. In experimental applications, cells or mitochondria are incubated with coelenterazine under controlled metabolic conditions. Substrates and inhibitors of the electron transport chain can be used to modulate ROS production at specific sites. For instance, the use of succinate in combination with antimycin A enhances ROS generation at complex III, while reverse electron transport induced by succinate oxidation in the presence of rotenone highlights superoxide production at complex I. The luminescence signal is recorded using highly sensitive luminometers or photon-counting devices, providing kinetic data on ROS production in real time. Coelenterazine chemiluminescence is particularly suited for high-throughput screening and longitudinal studies due to its non-destructive nature and compatibility with live-cell imaging platforms (Tarpey et al. 1999; Daiber et al. 2004). Unlike fluorescence-based ROS probes, which often require excitation light that may introduce phototoxicity or interfere with mitochondrial function, chemiluminescence avoids exogenous illumination and thereby minimizes assay artifacts. Furthermore, the rapid and continuous signal output of coelenterazine enables temporal resolution of ROS dynamics in response to metabolic shifts, pharmacological interventions, or environmental stressors. 

Redox cycling is a universal event occurring in any cells of any species as well as in bacteria and yeast (Cocheme and Murphy, 2008). Mitochondrial dysfunction is a universal event occurring in cells of any species (Farooqui and Farooqui, 2012). Many invertebrate species (e.g. D. melanogaster and C. elegans) are considered as potential models to study mitochondrial functionality. Data on marine invertebrates, such as molluscs and crustaceans and non-Drosophila species, are emerging (Martinez-Cruz et al., 2012). Mitochondrial dysfunction can be measured in animal models used for toxicity testing (Waerzeggers et al., 2010; Winklhofer and Haass, 2010) as well as in humans (Winklhofer and Haass, 2010). brain region-specific mitochondrial membrane potential and susceptibility towards dysfunction of mitochondrial oxidative phosphorylation (OXPHOS) was also observed by Pickrell et al. (2011). Here the striatum was found to be especially sensitive towards disturbance of OXPHOS due to the high striatal mitochondrial OXPHOS and membrane potential, which is prone to collapse when OXPHOS activity is reduced. This instance becomes important when studies on compound effects on isolated mitochondria are not of the correct origin, which would– for studying Parkinsonism– be the brain, and here the nigrostriatal area. In addition to mitochondrial differences between organs and intra-organ regions, species-specific mitochondrial activity was also measured.  

Aulak KS, Koeck T, Crabb JW, Stuehr DJ. Dynamics of protein nitration in cells and mitochondria. Am J Physiol Heart Circ Physiol. 2004 Jan;286(1):H30-8. doi: 10.1152/ajpheart.00743.2003. PMID: 14684358. 

Bayir H, Fadeel B, Palladino MJ, Witasp E, Kurnikov IV, Tyurina YY, Tyurin VA, Amoscato AA, Jiang J, Kochanek PM, DeKosky ST, Greenberger JS, Shvedova AA, Kagan VE. Apoptotic interactions of cytochrome c: redox flirting with anionic phospholipids within and outside of mitochondria. Biochim Biophys Acta. 2006 May-Jun;1757(5-6):648-59. doi: 10.1016/j.bbabio.2006.03.002. Epub 2006 Mar 31. PMID: 16740248.

Bender A, Krishnan KJ, Morris CM, Taylor GA, Reeve AK, Perry RH, Jaros E, Hersheson JS, Betts J, Klopstock T, Taylor RW, Turnbull DM. High levels of mitochondrial DNA deletions in substantia nigra neurons in aging and Parkinson disease. Nat Genet. 2006 May;38(5):515-7. doi: 10.1038/ng1769. Epub 2006 Apr 9. PMID: 16604074. 

Bernardi P, Gerle C, Halestrap AP, Jonas EA, Karch J, Mnatsakanyan N, Pavlov E, Sheu SS, Soukas AA. Identity, structure, and function of the mitochondrial permeability transition pore: controversies, consensus, recent advances, and future directions. Cell Death Differ. 2023 Aug;30(8):1869-1885. doi: 10.1038/s41418-023-01187-0. Epub 2023 Jul 17. PMID: 37460667; PMCID: PMC10406888. 

Brand MD. Mitochondrial generation of superoxide and hydrogen peroxide as the source of mitochondrial redox signaling. Free Radic Biol Med. 2016 Nov;100:14-31. doi: 10.1016/j.freeradbiomed.2016.04.001. Epub 2016 Apr 13. PMID: 27085844. 

Brand MD. The sites and topology of mitochondrial superoxide production. Exp Gerontol. 2010 Aug;45(7-8):466-72. doi: 10.1016/j.exger.2010.01.003. Epub 2010 Jan 11. PMID: 20064600; PMCID: PMC2879443. 

Brown GC, Borutaite V. Inhibition of mitochondrial respiratory complex I by nitric oxide, peroxynitrite and S-nitrosothiols. Biochim Biophys Acta. 2004 Jul 23;1658(1-2):44-9. doi: 10.1016/j.bbabio.2004.03.016. PMID: 15282173. 

Cadenas E, Boveris A, Ragan CI, Stoppani AO. Production of superoxide radicals and hydrogen peroxide by NADH-ubiquinone reductase and ubiquinol-cytochrome c reductase from beef-heart mitochondria. Arch Biochem Biophys. 1977 Apr 30;180(2):248-57. doi: 10.1016/0003-9861(77)90035-2. PMID: 195520. 

Castello PR, Drechsel DA, Patel M. Mitochondria are a major source of paraquat-induced reactive oxygen species production in the brain. J Biol Chem. 2007 May 11;282(19):14186-93. doi: 10.1074/jbc.M700827200. Epub 2007 Mar 27. PMID: 17389593; PMCID: PMC3088512. 

Castro L, Rodriguez M, Radi R. Aconitase is readily inactivated by peroxynitrite, but not by its precursor, nitric oxide. J Biol Chem. 1994 Nov 25;269(47):29409-15. PMID: 7961920. 

Chen JJ, Yu BP. Alterations in mitochondrial membrane fluidity by lipid peroxidation products. Free Radic Biol Med. 1994 Nov;17(5):411-8. doi: 10.1016/0891-5849(94)90167-8. PMID: 7835747. 

Chen Q, Niu Y, Zhang R, Guo H, Gao Y, Li Y, Liu R. The toxic influence of paraquat on hippocampus of mice: involvement of oxidative stress. Neurotoxicology. 2010 Jun;31(3):310-6. doi: 10.1016/j.neuro.2010.02.006. Epub 2010 Mar 6. PMID: 20211647. 

Chen S, Li Q, Shi H, Li F, Duan Y, Guo Q. New insights into the role of mitochondrial dynamics in oxidative stress-induced diseases. Biomed Pharmacother. 2024 Sep;178:117084. doi: 10.1016/j.biopha.2024.117084. Epub 2024 Aug 1. PMID: 39088967. 

Chinta SJ, Andersen JK. Nitrosylation and nitration of mitochondrial complex I in Parkinson's disease. Free Radic Res. 2011 Jan;45(1):53-8. doi: 10.3109/10715762.2010.509398. Epub 2010 Sep 6. PMID: 20815786. 

Chu CT, Ji J, Dagda RK, Jiang JF, Tyurina YY, Kapralov AA, Tyurin VA, Yanamala N, Shrivastava IH, Mohammadyani D, Wang KZQ, Zhu J, Klein-Seetharaman J, Balasubramanian K, Amoscato AA, Borisenko G, Huang Z, Gusdon AM, Cheikhi A, Steer EK, Wang R, Baty C, Watkins S, Bahar I, Bayir H, Kagan VE. Cardiolipin externalization to the outer mitochondrial membrane acts as an elimination signal for mitophagy in neuronal cells. Nat Cell Biol. 2013 Oct;15(10):1197-1205. doi: 10.1038/ncb2837. Epub 2013 Sep 15. PMID: 24036476; PMCID: PMC3806088. 

Chung HS, Wang SB, Venkatraman V, Murray CI, Van Eyk JE. Cysteine oxidative posttranslational modifications: emerging regulation in the cardiovascular system. Circ Res. 2013 Jan 18;112(2):382-92. doi: 10.1161/CIRCRESAHA.112.268680. PMID: 23329793; PMCID: PMC4340704. 

Clementi E, Brown GC, Feelisch M, Moncada S. Persistent inhibition of cell respiration by nitric oxide: crucial role of S-nitrosylation of mitochondrial complex I and protective action of glutathione. Proc Natl Acad Sci U S A. 1998 Jun 23;95(13):7631-6. doi: 10.1073/pnas.95.13.7631. PMID: 9636201; PMCID: PMC22706. 

Cochemé HM, Murphy MP. Complex I is the major site of mitochondrial superoxide production by paraquat. J Biol Chem. 2008 Jan 25;283(4):1786-98. doi: 10.1074/jbc.M708597200. Epub 2007 Nov 26. PMID: 18039652. 

Cooke MS, Evans MD, Dizdaroglu M, Lunec J. Oxidative DNA damage: mechanisms, mutation, and disease. FASEB J. 2003 Jul;17(10):1195-214. doi: 10.1096/fj.02-0752rev. PMID: 12832285. 

Dröse S, Bleier L, Brandt U. A common mechanism links differently acting complex II inhibitors to cardioprotection: modulation of mitochondrial reactive oxygen species production. Mol Pharmacol. 2011 May;79(5):814-22. doi: 10.1124/mol.110.070342. Epub 2011 Jan 28. PMID: 21278232. 

Dröse S, Brandt U. The mechanism of mitochondrial superoxide production by the cytochrome bc1 complex. J Biol Chem. 2008 Aug 1;283(31):21649-54. doi: 10.1074/jbc.M803236200. Epub 2008 Jun 3. PMID: 18522938. 

Dröse S, Hanley PJ, Brandt U. Ambivalent effects of diazoxide on mitochondrial ROS production at respiratory chain complexes I and III. Biochim Biophys Acta. 2009 Jun;1790(6):558-65. doi: 10.1016/j.bbagen.2009.01.011. Epub 2009 Feb 6. PMID: 19364480. 

Druzhyna NM, Wilson GL, LeDoux SP. Mitochondrial DNA repair in aging and disease. Mech Ageing Dev. 2008 Jul-Aug;129(7-8):383-90. doi: 10.1016/j.mad.2008.03.002. Epub 2008 Mar 13. PMID: 18417187; PMCID: PMC2666190. 

Erecińska M, Wilson DF. The effect of antimycin A on cytochromes b561, b566, and their relationship to ubiquinone and the iron-sulfer centers S-1 (+N-2) and S-3. Arch Biochem Biophys. 1976 May;174(1):143-57. doi: 10.1016/0003-9861(76)90333-7. PMID: 180891. 

Evans MD, Dizdaroglu M, Cooke MS. Oxidative DNA damage and disease: induction, repair and significance. Mutat Res. 2004 Sep;567(1):1-61. doi: 10.1016/j.mrrev.2003.11.001. PMID: 15341901.

Farooqui T, Farooqui AA, 2012. Oxidative stress in Vertebrates and Invertebrate: Molecular Aspects of Cell Signalling. Wiley-Blackwell, Chapter 27, pp. 377–385.

Floyd RA, Watson JJ, Wong PK, Altmiller DH, Rickard RC. Hydroxyl free radical adduct of deoxyguanosine: sensitive detection and mechanisms of formation. Free Radic Res Commun. 1986;1(3):163-72. doi: 10.3109/10715768609083148. PMID: 2577733.  

Frank M, Duvezin-Caubet S, Koob S, Occhipinti A, Jagasia R, Petcherski A, Ruonala MO, Priault M, Salin B, Reichert AS. Mitophagy is triggered by mild oxidative stress in a mitochondrial fission dependent manner. Biochim Biophys Acta. 2012 Dec;1823(12):2297-310. doi: 10.1016/j.bbamcr.2012.08.007. Epub 2012 Aug 16. PMID: 22917578. 

Gardner PR. Aconitase: sensitive target and measure of superoxide. Methods Enzymol. 2002;349:9-23. doi: 10.1016/s0076-6879(02)49317-2. PMID: 11912933. 

Hahm JY, Park J, Jang ES, Chi SW. 8-Oxoguanine: from oxidative damage to epigenetic and epitranscriptional modification. Exp Mol Med. 2022 Oct;54(10):1626-1642. doi: 10.1038/s12276-022-00822-z. Epub 2022 Oct 21. PMID: 36266447; PMCID: PMC9636213. 

Handy DE, Loscalzo J. Redox regulation of mitochondrial function. Antioxid Redox Signal. 2012 Jun 1;16(11):1323-67. doi: 10.1089/ars.2011.4123. Epub 2012 Feb 3. PMID: 22146081; PMCID: PMC3324814. 

Kowaltowski AJ, de Souza-Pinto NC, Castilho RF, Vercesi AE. Mitochondria and reactive oxygen species. Free Radic Biol Med. 2009 Aug 15;47(4):333-43. doi: 10.1016/j.freeradbiomed.2009.05.004. Epub 2009 May 8. PMID: 19427899. 

Lambert AJ, Brand MD. Inhibitors of the quinone-binding site allow rapid superoxide production from mitochondrial NADH:ubiquinone oxidoreductase (complex I). J Biol Chem. 2004 Sep 17;279(38):39414-20. doi: 10.1074/jbc.M406576200. Epub 2004 Jul 15. PMID: 15262965. 

Lin MT, Beal MF. Mitochondrial dysfunction and oxidative stress in neurodegenerative diseases. Nature. 2006 Oct 19;443(7113):787-95. doi: 10.1038/nature05292. PMID: 17051205. 

Mailloux RJ, Jin X, Willmore WG. Redox regulation of mitochondrial function with emphasis on cysteine oxidation reactions. Redox Biol. 2013 Dec 19;2:123-39. doi: 10.1016/j.redox.2013.12.011. PMID: 24455476; PMCID: PMC3895620. 

Martinez-Cruz O, Sanchez-Paz A, Garcia-Carre~ no F, Jimenez-Gutierrez L, Navarrete del Toro Mde L and Muhlia-Almazan A, 2012. Invertebrates Mitochondrial Function and Energetic Challenges (www.intechopen.com). In: Clark K (ed.). Bioenergetics, ISBN 978-953-51-0090-4, InTech, pp. 181–218. 

Milazzo L, Tognaccini L, Howes BD, Sinibaldi F, Piro MC, Fittipaldi M, Baratto MC, Pogni R, Santucci R, Smulevich G. Unravelling the Non-Native Low-Spin State of the Cytochrome c-Cardiolipin Complex: Evidence of the Formation of a His-Ligated Species Only. Biochemistry. 2017 Apr 4;56(13):1887-1898. doi: 10.1021/acs.biochem.6b01281. Epub 2017 Mar 20. PMID: 28277678. 

Miwa S, St-Pierre J, Partridge L, Brand MD. Superoxide and hydrogen peroxide production by Drosophila mitochondria. Free Radic Biol Med. 2003 Oct 15;35(8):938-48. doi: 10.1016/s0891-5849(03)00464-7. PMID: 14556858. 

Mohammadi-Bardbori A, Ghazi-Khansari M. Alternative electron acceptors: Proposed mechanism of paraquat mitochondrial toxicity. Environ Toxicol Pharmacol. 2008 Jul;26(1):1-5. doi: 10.1016/j.etap.2008.02.009. Epub 2008 Feb 29. PMID: 21783880. 

Mu B, Zeng Y, Luo L, Wang K. Oxidative stress-mediated protein sulfenylation in human diseases: Past, present, and future. Redox Biol. 2024 Oct;76:103332. doi: 10.1016/j.redox.2024.103332. Epub 2024 Aug 30. PMID: 39217848; PMCID: PMC11402764. 

Nakagawa H, Komai N, Takusagawa M, Miura Y, Toda T, Miyata N, Ozawa T, Ikota N. Nitration of specific tyrosine residues of cytochrome C is associated with caspase-cascade inactivation. Biol Pharm Bull. 2007 Jan;30(1):15-20. doi: 10.1248/bpb.30.15. PMID: 17202652. 

Pickrell AM, Fukui H, Wang X, Pinto M, Moraes CT, 2011. The striatum is highly susceptible to mitochondrial oxidative phosphorylation dysfunctions. Journal of Neuroscience, 31, 9895–9904. 

Quinlan CL, Gerencser AA, Treberg JR, Brand MD. The mechanism of superoxide production by the antimycin-inhibited mitochondrial Q-cycle. J Biol Chem. 2011 Sep 9;286(36):31361-72. doi: 10.1074/jbc.M111.267898. Epub 2011 Jun 27. PMID: 21708945; PMCID: PMC3173136. 

Quinlan CL, Goncalves RL, Hey-Mogensen M, Yadava N, Bunik VI, Brand MD. The 2-oxoacid dehydrogenase complexes in mitochondria can produce superoxide/hydrogen peroxide at much higher rates than complex I. J Biol Chem. 2014 Mar 21;289(12):8312-25. doi: 10.1074/jbc.M113.545301. Epub 2014 Feb 10. PMID: 24515115; PMCID: PMC3961658. 

Quinlan CL, Treberg JR, Perevoshchikova IV, Orr AL, Brand MD. Native rates of superoxide production from multiple sites in isolated mitochondria measured using endogenous reporters. Free Radic Biol Med. 2012 Nov 1;53(9):1807-17. doi: 10.1016/j.freeradbiomed.2012.08.015. Epub 2012 Aug 17. PMID: 22940066; PMCID: PMC3472107. 

Riojas-Hernández A, Bernal-Ramírez J, Rodríguez-Mier D, Morales-Marroquín FE, Domínguez-Barragán EM, Borja-Villa C, Rivera-Álvarez I, García-Rivas G, Altamirano J, García N. Enhanced oxidative stress sensitizes the mitochondrial permeability transition pore to opening in heart from Zucker Fa/fa rats with type 2 diabetes. Life Sci. 2015 Nov 15;141:32-43. doi: 10.1016/j.lfs.2015.09.018. Epub 2015 Sep 25. PMID: 26407476. 

Sanders LH, Paul KC, Howlett EH, Lawal H, Boppana S, Bronstein JM, Ritz B, Greenamyre JT. Editor's Highlight: Base Excision Repair Variants and Pesticide Exposure Increase Parkinson's Disease Risk. Toxicol Sci. 2017 Jul 1;158(1):188-198. doi: 10.1093/toxsci/kfx086. PMID: 28460087; PMCID: PMC6075191. 

Sarewicz M, Borek A, Cieluch E, Swierczek M, Osyczka A. Discrimination between two possible reaction sequences that create potential risk of generation of deleterious radicals by cytochrome bc₁. Implications for the mechanism of superoxide production. Biochim Biophys Acta. 2010 Nov;1797(11):1820-7. doi: 10.1016/j.bbabio.2010.07.005. Epub 2010 Jul 15. PMID: 20637719; PMCID: PMC3057645. 

Satoh T, Enokido Y, Aoshima H, Uchiyama Y, Hatanaka H. Changes in mitochondrial membrane potential during oxidative stress-induced apoptosis in PC12 cells. J Neurosci Res. 1997 Nov 1;50(3):413-20. doi: 10.1002/(SICI)1097-4547(19971101)50:3<413::AID-JNR7>3.0.CO;2-L. PMID: 9364326. 

Schlame M, Haldar D. Cardiolipin is synthesized on the matrix side of the inner membrane in rat liver mitochondria. J Biol Chem. 1993 Jan 5;268(1):74-9. PMID: 8380172. 

Sharma P, Sampath H. Mitochondrial DNA Integrity: Role in Health and Disease. Cells. 2019 Jan 29;8(2):100. doi: 10.3390/cells8020100. PMID: 30700008; PMCID: PMC6406942. 

Sharma LK, Lu J, Bai Y. Mitochondrial respiratory complex I: structure, function and implication in human diseases. Curr Med Chem. 2009;16(10):1266-77. doi: 10.2174/092986709787846578. PMID: 19355884; PMCID: PMC4706149. 

Shokolenko IN, Wilson GL, Alexeyev MF. Aging: A mitochondrial DNA perspective, critical analysis and an update. World J Exp Med. 2014 Nov 20;4(4):46-57. doi: 10.5493/wjem.v4.i4.46. PMID: 25414817; PMCID: PMC4237642. 

Skulachev VP, Antonenko YN, Cherepanov DA, Chernyak BV, Izyumov DS, Khailova LS, Klishin SS, Korshunova GA, Lyamzaev KG, Pletjushkina OY, Roginsky VA, Rokitskaya TI, Severin FF, Severina II, Simonyan RA, Skulachev MV, Sumbatyan NV, Sukhanova EI, Tashlitsky VN, Trendeleva TA, Vyssokikh MY, Zvyagilskaya RA. Prevention of cardiolipin oxidation and fatty acid cycling as two antioxidant mechanisms of cationic derivatives of plastoquinone (SkQs). Biochim Biophys Acta. 2010 Jun-Jul;1797(6-7):878-89. doi: 10.1016/j.bbabio.2010.03.015. Epub 2010 Mar 20. PMID: 20307489. 

Tórtora V, Quijano C, Freeman B, Radi R, Castro L. Mitochondrial aconitase reaction with nitric oxide, S-nitrosoglutathione, and peroxynitrite: mechanisms and relative contributions to aconitase inactivation. Free Radic Biol Med. 2007 Apr 1;42(7):1075-88. doi: 10.1016/j.freeradbiomed.2007.01.007. Epub 2007 Jan 8. PMID: 17349934. 

Tretter L, Adam-Vizi V. Inhibition of alpha-ketoglutarate dehydrogenase due to H2O2-induced oxidative stress in nerve terminals. Ann N Y Acad Sci. 1999;893:412-6. doi: 10.1111/j.1749-6632.1999.tb07867.x. PMID: 10672279

Tretter L, Adam-Vizi V. Inhibition of Krebs cycle enzymes by hydrogen peroxide: A key role of [alpha]-ketoglutarate dehydrogenase in limiting NADH production under oxidative stress. J Neurosci. 2000 Dec 15;20(24):8972-9. doi: 10.1523/JNEUROSCI.20-24-08972.2000. PMID: 11124972; PMCID: PMC6773008. 

Turrens JF, Alexandre A, Lehninger AL. Ubisemiquinone is the electron donor for superoxide formation by complex III of heart mitochondria. Arch Biochem Biophys. 1985 Mar;237(2):408-14. doi: 10.1016/0003-9861(85)90293-0. PMID: 2983613. 

Turrens JF, Boveris A. Generation of superoxide anion by the NADH dehydrogenase of bovine heart mitochondria. Biochem J. 1980 Nov 1;191(2):421-7. doi: 10.1042/bj1910421. PMID: 6263247; PMCID: PMC1162232. 

Ullrich V, Schildknecht S. Sensing hypoxia by mitochondria: a unifying hypothesis involving S-nitrosation. Antioxid Redox Signal. 2014 Jan 10;20(2):325-38. doi: 10.1089/ars.2012.4788. Epub 2012 Sep 11. PMID: 22793377. 

Votyakova TV, Reynolds IJ. DeltaPsi(m)-Dependent and -independent production of reactive oxygen species by rat brain mitochondria. J Neurochem. 2001 Oct;79(2):266-77. doi: 10.1046/j.1471-4159.2001.00548.x. PMID: 11677254. 

Waerzeggers, Yannic Monfared, Parisa Viel, Thomas Winkeler, Alexandra Jacobs, Andreas H, 2010. Mouse models in neurological disorders: applications of non-invasive imaging. Biochimica et Biophysica Acta (BBA) – Molecular Basis of Disease, 1802, 819–839. 

Winklhofer K, Haass C, 2010. Mitochondrial dysfunction in Parkinson’s disease. Biochimica et Biophysica Acta (BBA)- Molecular Basis of Disease, 1802, 29–44. 

Wong HS, Dighe PA, Mezera V, Monternier PA, Brand MD. Production of superoxide and hydrogen peroxide from specific mitochondrial sites under different bioenergetic conditions. J Biol Chem. 2017 Oct 13;292(41):16804-16809. doi: 10.1074/jbc.R117.789271. Epub 2017 Aug 24. PMID: 28842493; PMCID: PMC5641882. 

Xiao M, Zhong H, Xia L, Tao Y, Yin H. Pathophysiology of mitochondrial lipid oxidation: Role of 4-hydroxynonenal (4-HNE) and other bioactive lipids in mitochondria. Free Radic Biol Med. 2017 Oct;111:316-327. doi: 10.1016/j.freeradbiomed.2017.04.363. Epub 2017 Apr 27. PMID: 28456642. 

Yang ES, Lee JH, Park JW. Ethanol induces peroxynitrite-mediated toxicity through inactivation of NADP+-dependent isocitrate dehydrogenase and superoxide dismutase. Biochimie. 2008 Sep;90(9):1316-24. doi: 10.1016/j.biochi.2008.03.001. Epub 2008 Mar 19. PMID: 18405671. 

Zhang L, Yu L, Yu CA. Generation of superoxide anion by succinate-cytochrome c reductase from bovine heart mitochondria. J Biol Chem. 1998 Dec 18;273(51):33972-6. doi: 10.1074/jbc.273.51.33972. PMID: 9852050.