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Key Event Title
Increase, Mitochondrial Dysfunction
|Level of Biological Organization|
Key Event Components
Key Event Overview
AOPs Including This Key Event
|AOP Name||Role of event in AOP||Point of Contact||Author Status||OECD Status|
|Inhibition of Mt-ETC complexes leading to kidney toxicity||KeyEvent||Baki Sadi (send email)||Under development: Not open for comment. Do not cite|
|Kidney failure induced by inhibition of mitochondrial ETC||KeyEvent||Yann GUEGUEN (send email)||Under development: Not open for comment. Do not cite|
Key Event Description
Mitochondria are organelles found in all eukaryotic cells, crucial to the cellular consumption of oxygen, production of energy through the generation of ATP during oxidative phosphorylation, and regulation of cell death pathways (Alberts et al., 2014). The mitochondria are responsible for reduction of oxygen into water via the action of cytochrome c oxidase and other redox enzymes which transfer single electrons to oxygen and partially reduce it. The electron transfer is coupled with H+ ion transport across a membrane, producing the ion gradient that powers ATP synthesis (Alberts et al., 2014; Adiele et al., 2012). Under normal metabolic function, approximately 1-2% of the oxygen reduced by mitochondria converts into reactive oxygen species (ROS; such as superoxide, hydrogen peroxide, or hydroxyl radicals) at intermediate steps of the respiratory chain, as a result of electron transport (Kowaltowski and Vercesi, 1998; Volka et al., 2005; Li et al., 2003). This consistent and regular production of ROS and their signalling functionality at regulated levels contrasted with their harmful effects at high concentrations, justify the presence of antioxidant systems to regulate these processes.
Mitochondrial dysfunction, the loss of function or efficiency of oxidative phosphorylation, can be caused by a variety of factors and be apparent in a number of measurable ways. Some pathways of mitochondrial damage include: direct inhibition of mitochondrial proteins, indirect inhibition in upstream processes that affect mitochondrial metabolism, and indirect metabolic inhibition by ROS and physical damage to mitochondria. Dysfunction can be characterized through indicators of proton gradient loss, complex inhibition, or respiratory impairment such as mitochondrial permeability transition increase, mitochondrial membrane potential decrease, and ATP production (Shaki et al., 2013; Kruiderig et al., 1997). Any mitochondrial dysfunction impairs electron transfer and ATP production, which leads to deviation of electrons from their normal pathway in the electron transport chain (ETC), and increased ROS production. This, in turn, results in oxidative stress, mitochondrial permeability transition, and deregulation of cellular Ca2+ homeostasis (Nicholson, 2014; Shaki et al., 2013). Calcium, an imperative divalent cation to mitochondrial function, can be present at unsustainable levels due to increasing Ca2+ uptake, related to ROS generation and oxidative stress (reviewed Mei et al., 2013; Wang and Qin, 2010). Ca2+ accumulation and oxidative stress due additional ROS can trigger the opening of mitochondrial permeability transition pore (MPTP) by perturbing the osmolarity of mitochondria, disturbing Calcium homeostasis (Orrenius et al., 2015; Roos et al., 2012). The opening of the MPTP is a Ca2+-dependent process, that along with free proton movement collapses the mitochondrial membrane potential (MPP), halting ATP synthesis (Orrenius et al., 2015). ROS produced by the mitochondria can oxidize proteins and induce lipid peroxidation, compromising the barrier properties of the mitochondrial membrane (Orrenius et al., 2015) and therefore the proton gradient and ATP production. Respiration can also be impaired through mitochondrial DNA damage and increased permeability transition of the membrane as the mitochondrial inner membrane loses its impermeability to ions and other small molecules (up to a molecular weight of approximately 2kDa), this is loss of MPP and therefore proton gradient loss (Nicotera et al., 1998). Cytochrome c release is a major indicator of mitochondrial dysfunction as a combined result of a compromised mitochondrial membrane due to lipid peroxidation and the opening of the MPTP, and is commonly seen as an endpoint to mitochondrial toxicity (Chen et al., 2000). Mitochondrial damage can also be defined by loss of protein import and biosynthesis, as well as loss of mitochondrial motility as a result of failure to re-localize to sites with increased energy demands.
Metal-induced Mitochondrial Dysfunction
Mitochondria are an important site of Ca2+ regulation and storage, taking up Ca2+ ions electrophoretically from the cytosol through a Ca2+ uniporter, which can then accumulate in the mitochondria (Roos et al., 2012; Orrenius et al., 2015). Similarities between calcium and metals, such as cadmium and lead, makes the entrance and accumulation of these metals into the mitochondria via calcium metals possible by mode of molecular mimicry (Mathews et al., 2013; Adiele et al., 2012). The outer mitochondrial membrane also contains the divalent metal transporter (DMT1), which allows for mitochondrial uptake of divalent metals such as Fe and Mn. When cells are under heavy metal-induced stress, DMT has been shown to be overexpressed in the mitochondrial membrane, making the mitochondria targets of metal toxicity and accumulation.
Heavy metal exposure in aerobic organisms increases ROS formation through redox cycling, where metals with different valence states (Fe, Cu, Cr, etc.) directly produce ROS as they are reduced by cellular antioxidants and then react with oxygen (Shaki et al., 2012; Shaki et al., 2013; Pourahmad et al., 2006; Santos et al., 2007). The production of highly reactive hydroxyl radicals under mitochondrial oxidative stress and in the presence of transition metals occurs via the Fenton reaction or Haber-Weiss reaction (Hancock et al., 2001; Valko et al., 2005; Adam-Vizi et al., 2010). Metals and ROS are capable of damaging mitochondrial DNA as well as mechanisms of DNA repair and proliferation arrest (Valko et al., 2005). Metals and ROS have the potential to directly damage mitochondrial membranes and structure by binding to and oxidizing membrane lipids and proteins. This structural damage can collapse the MMP and lead to the opening of the MPTP (Orrenius et al., 2015; Roos et al., 2012; Pourahmad et al., 2006). Uranium and mercury, for example, have both been shown to directly inhibit the mitochondrial electron transport chain and interfere with ATP production (Shaki et al., 2012; Roos et al., 2012). Furthermore, as previously mentioned, metals have been shown to inhibit ROS-detoxifying enzymes. By binding to these enzymes, metals can inhibit their antioxidant functions, and cause an accumulation of ROS and increased synthesis of more antioxidant enzymes in order to combat the oxidative stress (Blajszczak and Bonini, 2017).
How It Is Measured or Detected
|Assay - What is being Measured||Description||Dose Range Studied||Assay Length / Ease of use, accuracy|
Rhodamine 123 Assay
Measuring Mitochondrial membrane potential (MMP) and its collapse
(Shaki et al., 2012)
Mitochondrial uptake of cationic fluorescent dye, rhodamine 123, is used for estimation of mitochondrial membrane potential. The fluorescence was monitored using Schimadzou RF-5000U fluorescence spectrophotometer at the excitation and emission wavelength of 490 nm and 535 nm, respectively.
|50, 100 and 500 μM of uranyl acetate||
Short / easy
TMRE fluorescence Assay
Measuring Mitochondrial permeability transition pore (MPTP) opening
(Huser et al., 1998)
|Laser scanning confocal microscopy in combination with the potentiometric fluorescence dye tetramethylrhodamine ethyl ester to monitor relative changes in membrane potential in single isolated cardiac mitochondria. The cationic dye distributes across the membrane in a voltage-dependent manner. Therefore, the large potential gradient across the inner mitochondrial membrane results in the accumulation of the fluorescent dye within the matrix compartment. Rapid depolarizations are caused by the opening of the transition pore.||1 µM cyclosporin A||
Short / easy
GSH / GSSG Determination Assay
Measuring cellular glutathione (GSH) status; ratio of GSH/GSSG
(Owen & Butterfield, 2010; Shaki et al., 2013)
|GSH and GSSG levels are determinted biochemically with DTNB (Ellman’s reagent). The developed yellow color was read at 412 nm on a spectrophotometer.||100 µM uranyl acetate||
Short / easy
Quantification of lipid peroxidation
(Yuan et al., 2016)
|MDA content, a product of lipid peroxidation, was measured using a thiobarbituric acid reactive substances (TBARS) assay. Briefly, the kidney cells were collected in 1 ml PBS buffer solution (pH 7.4) and sonicated. MDA reacts with thiobarbituric acid forming a colored product which can be measured at an absorbance of 532 nm.||200, 400, 800 µM uranyl acetate||
Medium / medium
Aequorin-based bioluminescence assay
Increase in mitochondrial Ca2+ influx
(Pozzan & Rudolf, 2009)
|Together with GFP, the aequorin moiety acts as Ca2+ sensor in vivo, which delivers emission energy to the GFP acceptor molecule in a BRET (Bioluminescence Resonance Energy Transfer) process; the Ca2+ can then be visualized with fluorescence microscopy.||
Short / easy
Western blot & immunostaining analyses
Measuring cytochrome c release(Chen et al., 2000)
|Examining the redistribution of Cyto c in cytosolic and mitochondrial cellular fractions. Cells are homogenized and centrifuged, then prepared for immunoblots. Cellular fractions were washed in PBS and lysed in 1% NP-40 buffer. Cellular proteins were separated by SDS–PAGE, transferred onto nitrocellulose membranes, probed using immunoblot analyses with antibodies specific to cyto c (6581A for Western and 65971A for immunostaining; Pharmingen)||
Short / easy
Quantikine Rat/Mouse Cytochrome c Immunoassay
Measuring cytochrome c release
(Shaki et al., 2012)
|Cytochrome C release was measured a monoclonal antibody specific for rat/mouse cytochrome c was precoated onto the microplate. Seventy-five microliter of conjugate (containing mono- clonal antibody specific for cytochrome c conjugated to horseradish peroxidase). After 2 h of incubation, the substrate solution (100 μl) was added to each well and incubated for 30 min. After 100 μl of the stop solution was added to each well; the optical density of each well was determined by the aforementioned microplate spectrophotometer set to 450 nm.||
Short / easy
Membrane potential and cell viability – Flow Cytometry
Measuring cytochrome c release
(Kruiderig et al., 1997)
|“Dc and viability were determined by analyzing the R123 and propidium iodide fluorescence intensity with a FACScan flow cytometer (Becton Dickinson, San Jose, CA) equipped with an argon laser, with the Lysis software program (Becton Dickinson). R123 is a cationic dye that accumulates in the negatively charged inner side of the mitochondria. When the potential drops, less R123 accumulates in the mitochondria, which results in a lower fluorescence signal. The potential was measured as follows: at the indicated times, a 500-ml sample of the cell suspension was taken and transferred to an Eppendorf minivial. To this sample, 100 ml of 6 mM R123 in buffer D was added. After incubation for 10 min at 37°C, the cell suspension was centrifuged for 5 min at 80 3 g. The cell pellet was resuspended in 200 ml of buffer D, containing 0.2 mM R123 and 10 mM propidium iodide, to prevent loss of R123 and to stain nonviable cells, respectively. The samples were transferred to FACScan tubes and analyzed immediately. Analysis was performed at a flow rate of 60 ml/min. R123 fluorescence was detected by the FL1 detector with an emission detection limit below 560 nm. Propidium iodide fluorescence was detected by the FL3 detector, with emission detection above 620 nm. Per sample 3,000 to 5,000 cells were counted (Van de Water et al., 1993)”||
Short / easy
Domain of Applicability
Mitochondrial dysfunction can occur in any eukaryotic cell.
Evidence for Perturbation by Stressor
Shaki et al. (2012) found that uranyl acetate (UA) exposure in isolated rat kidney mitochondria decreased the ATP production levels and ATP/ADP ratio in a concentration-dependent manner, through inhibition of complexes II and III of the ETC. Both of these levels were significantly changed at UA concentrations of 100 µM and 200 µM. In addition, a concentration-dependent decrease in activity of complex II with exposure to U was observed (Shaki et al., 2012). They also found that mitochondrial membrane potential damage and mitochondrial swelling both increased significantly time- and dose-dependently in the treated rat kidneys (Shaki et al., 2012). ATP/ADP ratios were also decreased significantly by treatment with 100 µM or more uranium (Shaki et al., 2012). Mitochondrial outer membrane damage was significantly decreased by treatment with 200 µM of uranium (Shaki et al., 2012).
Shaki et al. (2013) also investigated the effects of uranium on rat kidneys. They found that mitochondrial permeability transition was also impacted by uranium treatment, causing increased mitochondrial swelling and increased disruption of energy homeostasis (Shaki et al., 2013).
Hao et al. (2014) assessed the changes in mitochondrial potential in human kidney proximal tubular cells treated with uranium and found that the group treated with 500 µM of depleted uranium for 24 hours showed a significant decreased mitochondrial membrane potential.
In their study of the effects of depleted uranium treatment on human embryonic kidney cells, Hao et al. (2016) found that ETHE1, a mitochondrial protein involved in mitochondrial homeostasis and mitochondrial diseases, had significant dose- and time-dependant decreases in gene expression when treated with 125 µM or more depleted uranium (DU) for 2 hours or more.
Nanoparticles and Micrometer Particles
Karlsson et al. (2009) conducted experiments to examine the effects of micrometer and nanoparticle treatments of copper and iron on human alveolar type-II epithelial cells. Their results showed that copper oxide micrometer and nanoparticle treatments were able to cause dose-dependant mitochondrial depolarization with doses as low as 5 µg/cm2 (Karlsson et al., 2009). Iron(III) oxide nanoparticles and micrometer particles were both able to cause similar amounts of mitochondrial depolarization, along with iron (IV) oxide micrometer particles, however they were all much less toxic than copper oxide nanoparticles or micrometer particles (Karlsson et al., 2009).
The effects of gold nanoparticle (Au1.4MS) treatment on human cervical cancer cells were assessed by Pan et al. (2009), who found that the treated cells experienced a significant increase in permeability transition.
Huerta-García et al. (2014) studied the effects of titanium oxide nanoparticle treatment on glial tumor rat neuronal cells and cancerous human brain cells. Their results showed that in the treated rat and human cells there was a clear time-dependant increase in depolarization (Huerta-García et al., 2014). They also found that both the human and rat cells showed time-dependant decreases in mitochondrial membrane potential, with the TiO2 nanoparticles being more toxic to the human cells, which showed significant decrease as early as 2 hours post-treatment, while the rat cells did not show significant decrease until 6 hours post-treatment (Huerta-García et al., 2014).
Zhang et al. (2018) investigated the effects of copper nanoparticles on mitochondrial membrane potential in pig kidney cells and found that the treated cells showed a dose-dependant increase in the rate of mitochondrial membrane potential change from 40 µg/mL to 80 µg/mL when treated for 12 hours.
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