Increase, Mitochondrial complex III antagonism

1397-94-0

Antimycin A

Antipiricullin Fintrol Virosin

DTXSID9032325

1951-25-3

IYIKLHRQXLHMJQ-UHFFFAOYSA-N

Amiodarone

Methanone, (2-butyl-3-benzofuranyl)[4-[2-(diethylamino)ethoxy]-3,5-diiodophenyl]- 2-Butyl-3-[3,5-diiodo-4-(2-diethylaminoethoxy)benzoyl]benzofuran 2-Butyl-3-benzofuranyl p-[(2-diethylamino)ethoxy]-m,m-diiodophenyl ketone 2-n-Butyl-3',5'-diiodo-4'-N-diethylaminoethoxy-3-benzoylbenzofuran Amidorone Amiodaron amiodarona Ancaron Ketone, 2-butyl-3-benzofuranyl 4-[2-(diethylamino)ethoxy]-3,5-diiodophenyl Sedacoron Sedacorone

DTXSID7022592

3562-84-3

WHQCHUCQKNIQEC-UHFFFAOYSA-N

Benzbromarone

DTXSID4022652

151-50-8

NNFCIKHAZHQZJG-UHFFFAOYSA-N

Potassium cyanide

K(CN) Potassium cyanide, cianuro de potasio Cyanure de potassium Hydrocyanic acid, potassium salt Kaliumcyanid Potassiuim cyanide UN 1680 UN3413

DTXSID0024268

83366-66-9

VRBKIVRKKCLPHA-UHFFFAOYSA-N

Nefazodone

3H-1,2,4-Triazol-3-one, 2-[3-[4-(3-chlorophenyl)-1-piperazinyl]propyl]-5-ethyl-2,4-dihydro-4-(2-phenoxyethyl)-

DTXSID2023357

657-24-9

XZWYZXLIPXDOLR-UHFFFAOYSA-N

Metformin

Imidodicarbonimidic diamide, N,N-dimethyl- 1,1-Dimethylbiguanide Biguanide, 1,1-dimethyl- Cidophage Dimethylbiguanide Emiphage Ficonax Fluamine Flumamine Gliguanid Haurymelin metformina metformine Metiguanide Metphage N,N-Dimethylbiguanide N,N-Dimethyldiguanide N,N-Dimethylimidodicarbonimidic diamide N,N-Dimethyl-imidodicarbonimidic diamide N1,N1-Dimethylbiguanide N'-Dimethylguanylguanidine

DTXSID2023270

130929-57-6

JRURYQJSLYLRLN-BJMVGYQFSA-N

Entacapone

2-Propenamide, 2-cyano-3-(3,4-dihydroxy-5-nitrophenyl)-N,N-diethyl-, (2E)-

DTXSID5046439

1404-19-9

MNULEGDCPYONBU-WMBHJXFZSA-N

Oligomycin

oligomicina oligomycine

DTXSID7040570

15307-79-6

KPHWPUGNDIVLNH-UHFFFAOYSA-M

Diclofenac sodium

Benzeneacetic acid, 2-[(2,6-dichlorophenyl)amino]-, monosodium salt [2-[(2,6-dichlorophenyl)amino]phenyl]acetate de sodium [2-[(2,6-diclorofenil)amino]fenil]acetato de sodio [o-(2,6-Dichloroanilino)phenyl]acetic acid sodium salt {2-[(2,6-Dichlorophenyl)amino]phenyl}acetate de sodium 2-(2,6-Dichloroanilino)phenylacetic acid sodium salt 2-[(2,6-Dichlorophenyl)amino]benzene acetic acid monosodium salt Acetic acid, [o-(2,6-dichloroanilino)phenyl]-, monosodium salt Allvoran Assaren Benfofen Benzeneacetic acid, 2-[(2,6-dichlorophenyl)amino]-, sodium salt (1:1) Cataflam Delphimix Diacron Dichronic Diclobene Diclobenin Diclodyn Diclofen SR 100 Diclofenac sodium salt Diclofenac-Na Emulgel Diclofenacsodium Emulgel Diclokalium Diclophenac sodium Diclo-Phlogont Diclo-Puren Diclord Diclorep Dicloreum Diklovit Dolobasan Duravolten Dyloject Effekton Evofenac Feloran Fortfen Hyanalgese D Inflaban Kriplex N-(2,6-Dichlorophenyl)-o-aminophenylacetic acid sodium salt Natrium-[2-[(2,6-dichlorphenyl)amino]phenyl]acetat Neriodin Novapirina Orthofen Orthophen Primofenac Profenac Prophenatin Rhumalgan sodium [2-[(2,6-dichlorophenyl)amino]phenyl]acetate Sodium [o-(2,6-dichloroanilino)phenyl]acetate Sodium 2-(2,6-dichloroanilino)-phenyl-acetate Sodium diclofenac Sorelmon Tsudohmin Valetan Voltaren Voltaren Ophtha Voltaren Ophtha CD Voltarol Voveran

DTXSID3037208

6659-45-6

DTFARBHXORYQBF-HBGVWJBISA-N

1',2'-Dihydrorotenone

DTXSID5041227

134098-61-6

YYJNOYZRYGDPNH-MFKUBSTISA-N

Fenpyroximate

DTXSID7032557

96489-71-3

DWFZBUWUXWZWKD-UHFFFAOYSA-N

Pyridaben

3(2H)-Pyridazinone, 4-chloro-2-(1,1-dimethylethyl)-5-(((4-(1,1-dimethylethyl)phenyl)methyl)thio)-

DTXSID5032573

7440-61-1

JFALSRSLKYAFGM-UHFFFAOYSA-N

Uranium

Uranium, isotope of mass 238 238U Element UN 2979 (DOT) Uranium I

DTXSID1042522

7440-43-9

BDOSMKKIYDKNTQ-UHFFFAOYSA-N

Cadmium

Cadimium CADMIUM BLUE CADMIUM, IN PLATTEN, STANGEN, BROCKEN,KOERNER

DTXSID1023940

PR:000031316

PR NADH-ubiquinone oxidoreductase chain 1

CHEBI:36080

CHEBI protein

GO:0005739

GO mitochondrion

GO:0006120

GO mitochondrial electron transport, NADH to ubiquinone

GO:0008137

GO NADH dehydrogenase (ubiquinone) activity

D057165

MESH Proteostasis deficiencies

GO:0008219

GO cell death

WIKI decreased

WIKI abnormal

WIKI increased

WIKI functional change

Antimycin A

2018-12-19T09:50:21

Amiodarone

2016-11-29T18:42:27

Benzbromarone

2019-03-07T05:10:21

KCN

2018-12-19T09:51:10

Nefazodone

2019-03-07T05:13:43

Metformin

2019-03-07T05:14:41

Entacapone

2019-03-07T05:14:55

Oligomycin

2019-03-07T05:17:22

Aurovertin

2019-03-07T05:19:56

2019-03-07T05:20:13

Diclofenac sodium

2016-11-29T18:42:09

1',2'-dihydrorotenone

2016-11-29T18:42:27

Fenpyroximate

2019-03-06T10:45:28

Pyridaben

2019-03-06T10:45:48

Uranium

2021-08-05T14:28:50

Nanoparticles and Micrometer Particles

2022-02-04T13:43:43

Cadmium

2017-10-25T08:33:12

WCS_9606

common toxicological species human

10116

NCBI rat

10090

NCBI mouse

10118

NCBI Rattus sp.

WikiUser_25

Wikiuser: Cyauk human and other cells in culture

10116

NCBI Rattus norvegicus

Increase, Mitochondrial complex III antagonism

Molecular

The mitochondrial complex III (mitochondrial cytochrome bc1 complex) is an enzyme complex located in the inner membrane of mitochondria. It is the third out of 5 complexes that together form the mitochondrial respiratory chain. It consists out of multiple subunits, including cytochrome b/b6, cytochrome c1 and an 2Fe-2S cluster. The combination of these subunits catalyze following process: <ol> <li> The cytochrome c1 is involved in the process of oxidizing ubiquinol (coming directly from complex I, from complex I via complex II or from complex II) to a semiquinone radical and back to quinone. A process that results in two free electrons that are transferred via cytochrome c to next complex. </li> </ol>   The electron transfer in this process mediates the translocation of protons from the mitochondrial matrix through the inner membrane to the intermembrane space. The created proton gradient will be used to catalyze the reaction in which ADP is converted into ATP.   The initiation event is the reversibly or irreversibly interaction to any of the subunits in the mitochondrial complex III, leading to an perturbation of the electron flow and an absence of proton transport via this complex.

Complex inhibition assays specifically for complex III It is important to release that the activity of complex II depends on the input from complex II. So screening of effects at complex II is advised. The assay exist in multiple forms. Some assay can be performed on whole cells and other needed complex isolation based on antibody interactions. However, they all rely on the following detection of complex II/III activity: The reduction of cytochrome c, which has an absorbance at 550nm. Mitochondrial Membrane potential using fluorescent dyes. Positive charged molecules will accumulated in the mitochondria in an inverse proportion as the membrane potential. More polarised mitochondria will accumulate more dye (Rhodamine123, TMRE, TMRM) - leading to a higher fluorescent signal - and the absence of membrane potential leads to an absence of fluorescent signal. An exception is the dye JC1, because this dye has green fluorescence when present in low concentrations (depolarisation) and red fluorescence when accumulated (hyperpolarized)   Measurements Complex inhibition assays <ol> <li> Cayman MitoCheck MitoCheck® Complex II/III Activity Assay Kit (Item No. 700950) </li> <li> Abcam MitoTox™ Complex II + III OXPHOS Activity Assay Kit (ab109905) </li> <li> BioVision Mitochondrial Complex III Activity Assay Kit (K520) </li> </ol> Membrane potential dyes <ol start="4"> <li> Perry, 2011, mitochondrial membrane potential probes and proton gradient </li> <li> Mitra, 2010, analysis of mitochondrial dynamics and functions using imaging approaches </li> </ol>

AOPWiki 2018-12-19T09:35:55

2020-12-04T17:24:45

Mitochondrial Complex IV inhibition

Molecular

The mitochondrial complex IV (Cytochrome c oxidase) is an enzyme complex located in the inner membrane of mitochondria. It is the forth out of 5 complexes that together form the mitochondrial respiratory chain. It consists out of multiple subunits of cytochrome c oxidase, including cytochrome a and cytochrome a3. The combination of these subunits catalyze following process: <ol> <li> The cytochrome c oxidase is involved in the process of oxidizing reduced cytochrome c to its oxidized form. A process that results in a free electron. </li> </ol> <ol> <li> 4 oxidation rounds results in enough electron to reduce 1 molecule oxygen to 2 molecules water. </li> </ol>   The electron transfer in this process mediates the translocation of protons from the mitochondrial matrix through the inner membrane to the intermembrane space. The created proton gradient will be used to catalyze the reaction in which ADP is converted into ATP.   The initiation event is the reversibly or irreversibly interaction to any of the subunits in the mitochondrial complex IV, leading to an perturbation of the electron flow and an absence of proton transport via this complex.

<ol> <li> Complex inhibition assays specifically for complex IV The activity of complex IV is always tested in the bovine heart mitochondria provided within the kit. The kit relies on the following detection of complex IV activity: The reduction of cytochrome c, which leads to a reduction in absorbance at 550nm can be measured. </li> <li> Mitochondrial Membrane potential using fluorescent dyes. Positive charged molecules will accumulated in the mitochondria in an inverse proportion as the membrane potential. More polarised mitochondria will accumulate more dye (Rhodamine123, TMRE, TMRM) - leading to a higher fluorescent signal - and the absence of membrane potential leads to an absence of fluorescent signal. An exception is the dye JC1, because this dye has green fluorescence when present in low concentrations (depolarisation) and red fluorescence when accumulated (hyperpolarized </li> </ol>   Measurements Complex inhibition assays <ol> <li> Cayman MitoCheck MitoCheck® Complex IV Activity Assay Kit (Item No. 700990) </li> <li> Abcam MitoTox™ Complex IV OXPHOS Activity Assay Kit (ab109906) Complex IV Human Enzyme Activity Microplate Assay Kit (ab109909) </li> <li> BioVision Cytochrome Oxidase Activity Colorimetric Assay Kit (K287) </li> </ol> Membrane potential dyes <ol start="4"> <li> Perry, 2011, mitochondrial membrane potential probes and proton gradient </li> <li> Mitra, 2010, analysis of mitochondrial dynamics and functions using imaging approaches </li> </ol>

AOPWiki 2018-12-19T09:36:11

2019-03-07T05:15:41

Mitochondrial Complex V inhibition

Molecular

The mitochondrial complex V (ATPase) is an enzyme complex located in the inner membrane of mitochondria. It is the fifth out of 5 complexes that together form the mitochondrial respiratory chain. <ol> <li> The ATPase catalysed the production of ATP from ADP and Pi using the proton gradient created by complex I,III and IV. </li> </ol>   The initiation event is the reversibly or irreversibly interaction to any of the subunits in the mitochondrial complex V, leading to an perturbation of ATP production.

<ol> <li> Complex inhibition assays specifically for complex V The activity of complex IV is always tested in the bovine heart mitochondria provided within the kit or using freshly isolated mitochondria. The kit relies on the following detection of complex V activity: ATP is converted by ATPase to ADP. The ADP is used by pyruvate kinase to convert Phosphoenolpyruvate to pyruvate. The pyruvate is converted to lactate by lactate dehydrogenase, while oxidising NADH to NAD+. NADH oxidation is measured at 340nm. </li> <li> Mitochondrial Membrane potential using fluorescent dyes. Positive charged molecules will accumulated in the mitochondria in an inverse proportion as the membrane potential. More polarised mitochondria will accumulate more dye (Rhodamine123, TMRE, TMRM) - leading to a higher fluorescent signal - and the absence of membrane potential leads to an absence of fluorescent signal. An exception is the dye JC1, because this dye has green fluorescence when present in low concentrations (depolarisation) and red fluorescence when accumulated (hyperpolarized) </li> </ol>   Measurements Complex inhibition assays <ol> <li> Cayman MitoCheck MitoCheck® Complex V Activity Assay Kit (Item No. 701000) </li> <li> Abcam MitoTox™ Complex V OXPHOS Activity Assay Kit (ab109907) </li> </ol> Membrane potential dyes <ol start="3"> <li> Perry, 2011, mitochondrial membrane potential probes and proton gradient </li> <li> Mitra, 2010, analysis of mitochondrial dynamics and functions using imaging approaches </li> </ol>

AOPWiki 2018-12-19T09:36:25

2019-03-07T05:21:52

Decrease in mitochondrial oxidative phosphorylation

Cellular

AOPWiki 2018-12-19T09:36:59

2018-12-19T09:36:59

Increased reactive oxygen species (in the mitochondria)

Cellular

AOPWiki 2018-12-19T09:37:12

2018-12-19T09:37:12

Mitochondrial Injury

Cellular

AOPWiki 2018-12-19T09:37:40

2018-12-19T09:37:40

Necrotic Tissue

Tissue

Taxa = necrotic tissue in the liver During the process of towards necrotic tissue, too many cells die. In practice it is difficult to distinguish separate forms of cell death (apoptosis, necrosis and necroptosis), especially in vivo.

Histopathology: performed on tissue sections · Hematoxylin (nuclei) and eosin staining: determine morphology · TUNEL staining: cell death https://medical-dictionary.thefreedictionary.com/Necrotic+tissue https://livertox.nih.gov/Phenotypes_ahn.html (Weng et al. 2015) (Guicciardi et al. 2013)

Guicciardi, M.E. et al., 2013. Apoptosis and Necrosis in the Liver. Comprehensive Physiology, 3(2), pp.977–1010. Available at: http://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=3867948&tool=pmcentrez&rendertype=abstract. Weng, H. et al., 2015. Two sides of one coin: Massive hepatic necrosis and progenitor cell-mediated regeneration in acute liver failure. Frontiers in Physiology, 6(MAY), pp.1–12. AOPWiki 2018-12-19T09:40:01

2019-03-07T04:19:14

Liver Injury

Organ

Hepatic Injury after apoptosis.(Landesmann, 2016) Liver injury is the altered state of the liver wherein the normal homeostasis of all process in the liver are perturbed.   4 types of liver injury are distinguished in patients:   Hepatocellular (= acute hepatitis) Characteristics = elevation of serum transaminases (ALT+AST)   Choleostatic (= obstruction of the bile duct (bile cannot flow from liver to duodenum)) Characteristics = a) elevation in serum alkaline phosphatase (ALP) with normal or mild elevations in serum transaminases (ALT+AST) b) elevated bilirubin levels   Infiltrative (= sarcoidosis, tuberculosis, liver abscess, metastatic malignancy) Characteristics = a) elevation in serum alkaline phosphatase with normal or mild elevations (less than five times normal) in serum transaminases b) no effects at bilirubin levels   Autoimmune (= autoimmune disease against liver components) Characteristics = can present itself as hepatocellular when hepatocytes are target (= autoimmune hepatitis) or cholestatic when biliary ducts is the target (primary biliary cirrhosis) Drug induced liver injury mostly manifest itself as hepatocellular injury, cholestasis or a mixture of both. In a mixture hepatitis the amount of hepatocellular and cholestatic features vary per case. Biopsy results with mixed hepatitis is a combination of: Hepatocellular = liver cell necrosis, inflammation Choleostatic = bile stasis, portal inflammation, injury of bile ducts Patient with any kind of mixed hepatitis demonstrates the following symptoms: First symptoms = Fatigue, anorexia and nausea Later symptoms = jaundice (=skin and eye white become yellows/greenish) dark urine and pruritus (= sensitization of itch)

Indicators of liver injury include: Levels of: ALT, AST, ALP, bilirubin, GGT, NTP, Ceruloplasmin, AFP (Guicciardi et al. 2013)   Test in patients or in vivo(Gowda et al. 2009) (Musana et al. 2004): · Biochemistry assays · Levels of: ALT, AST, ALP, bilirubin, GGT, NTP, Ceruloplasmin, AFP, · Imaging scans · Ultrasound · CT · MRI · Biopsy

Landesmann, B. (2016). Adverse Outcome Pathway on Protein Alkylation Leading to Liver Fibrosis, (2). Guicciardi ME, Malhi H, Mott JL, Gores GJ (2013) Apoptosis and Necrosis in the Liver Maria. Compr Physiol 3:977–1010 . doi: 10.1002/cphy.c120020.Apoptosis Musana, K.A., Yale, S.H. & Abdulkarim, A.S., 2004. Tests of liver injury. Clin Med Res, 2(2), pp.129–131. Gowda, S. et al., 2009. A review on laboratory liver function tests. The Pan African medical journal, 3(November), p.17. Available at: http://www.ncbi.nlm.nih.gov/pubmed/21532726%5Cnhttp://www.pubmedcentral.nih.gov/articlerender.fcgi?artid=PMC2984286. AOPWiki 2018-12-19T09:40:21

2019-03-07T04:23:21

Binding of inhibitor, NADH-ubiquinone oxidoreductase (complex I)

Molecular

Electron transport through the mitochondrial respiratory chain (oxidative phosphorylation) is mediated by five multimeric complexes (I–V) that are embedded in the mitochondrial inner membrane (Fig. 1). NADH-ubiquinone oxidoreductase is the Complex I (CI) of electron transport chain (ETC). It is a large assembly of proteins that spans the inner mitochondrial membrane. In mammals, it is composed of about 45-47 protein subunits (human 45) of which 7 are encoded by the mitochondrial genome (ND1, ND2, ND3, ND4, ND4L, ND5, and ND6) and the remainder by the nuclear genome (Greenamyre, 2001). CI oxidizes NADH elevating the NAD+/NADH ratio by transferring electrons via a flavin mononucleotide (FMN) cofactor and several iron-sulfur centers to ubiquinone (Friedrich et al., 1994) (Fig. 1). Binding of an inhibitor to CI inhibits the NADH–ubiquinone oxido-reductase activity, i.e. blocks the electron transfer. Recent studies suggest that a wide variety of CI inhibitors share a common binding domain at or close to the ubiquinone reduction site (Ino et al., 2003). Furthermore, the structural factors required for inhibitory actions have been characterized on the basis of structure-activity relationships (Miyoshi, 1998, Hideto, 1998). Based on molecular docking simulations, in silico models mimicking the binding of chemicals to the pocket of NADH ubiquinone oxidoreductase have been created according to the crystal structure of mitochondrial CI. To investigate the ability of chemicals to bind to the active pocket, around 100 individual docking simulations have been performed. These confirmed the possible site of interaction between the chemical and the pocket of CI. In particular, Miao YJ and coworkers recently investigated the IC50 values of 24 chemicals (annonaceous acetogenins) for inhibition of mitochondrial CI (Miao et al., 2014). Based on their binding sites, CI inhibitors are classified as follows (Degli Esposti, 1998) (Fig. 2): (i) type A inhibitors are antagonists of fully oxidized ubiquinone binding; (ii) type B inhibitors displace the partially reduced ubisemiquinone intermediate; (iii) type C inhibitors are antagonists of the fully reduced ubiquinol product. The affinity of the different types of CI inhibitors to their diverse CI binding sites is described in the paragraph Evidence for Chemical Initiation of this Molecular Initiating Event (see below) in the context of a specific type of inhibitor. <a class="image" href="/wiki/index.php/File:AOP-003-Figure1-smaller.JPG"><img alt="AOP-003-Figure1-smaller.JPG" src="/wiki/images/b/b9/AOP-003-Figure1-smaller.JPG" style="height:461px; width:593px" /></a> Fig. 1. The electron transport chain in the mitochondrion. CI (NADH-coenzyme Q reductase or NADH dehydrogenase) accepts electrons from NADH and serves as the link between glycolysis, the citric acid cycle, fatty acid oxidation and the electron transport chain. Complex II also known as succinate-coenzyme Q reductase or succinate dehydrogenase, includes succinate dehydrogenase and serves as a direct link between the citric acid cycle and the electron transport chain. The coenzyme Q reductase or Complex III transfers the electrons from CoQH2 to reduce cytochrome c which is the substrate for Complex IV (cytochrome c reductase). Complex IV transfers the electrons from cytochrome c to reduce molecular oxygen into water. Finally, this gradient is used by the ATP synthase complex (Complex V) to make ATP via oxidative phosphorylation. mtDNA: mitochondrial DNA; nDNA: nuclear DNA. <a class="image" href="/wiki/index.php/File:MIE_Fig._2.jpg"><img alt="MIE Fig. 2.jpg" src="/wiki/images/6/63/MIE_Fig._2.jpg" style="height:301px; width:436px" /></a> Fig. 2. Schematic representation of CI and proposed inhibition binding sites by inhibitors of class A, B and C. Nicotinamide adenine dinucleotide (NADH, reduced and NAD, oxidized), flavin mononucleotide (FMN) and Ubiquinone (Q) (taken from Haefeli, 2012).

Two different types of approaches have been used. The first is to measure binding as such, and the corresponding assays are described below; the second is to infer binding indirectly from assays that quantify e.g. CI activity and to assume that the activity can only be altered upon binding. The second type of approach is dealt with in the chapter entitled KE1: Inhibition of NADH ubiquinone oxidoreductase (complex I). However, it has to be noted here that indirect assays can lead to wrong conclusions. For instance, some compounds may trigger oxidative stress without actually binding to CI. Such compounds, by triggering the generation of reactive oxygen species (ROS), may damage CI protein components, thus causing a reduction of CI activity. Measurement of binding by quantitative autoradiography To assess binding of an inhibitor at the rotenone binding site of CI in tissues (e.g. in the substantia nigra or in the striatum), the standard approach is to quantify the displacement of a radioactively labelled ligand of this binding site by the toxicant under evaluation. Most commonly, binding of [3H]-labeled dihydrorotenone (DHR) is measured and compared in control tissue and treated tissue. Binding of this rotenone-derivative is detected by autoradiography. Unselective binding is determined by measurement of [3H]-DHR binding in the presence of an excess of unlabeled rotenone. Since a rotenone-derivative is used for the assay, only CI inhibitors that bind to the rotenone-binding site in CI are detected. This was observed for e.g., meperdine, amobarbital, or MPP+. This method allows a spatial resolution of CI expression and the mapping of the binding of a competitive inhibitor on CI. The method can be used for (a) in vitro measurements and for (b) ex vivo measurements: a) In vitro measurements. Tissues are embedded in a matrix for cutting by a cryostat. The tissue slices are then mounted onto slides. For the binding experiment, they are incubated with the test compound in the presence of labeled [3H]-DHR. Then the tissue slices are washed and prepared for autoradiographic detection (Greenamyre et al. 1992; Higgins and Greenamyre, 1996). b) Ex vivo measurements. As rotenone can pass the blood brain barrier, the in vitro method was further extended for in vivo labeling of CI in the brains of living animals, and detection of binding after preparation of the tissue from such animals. Animals are exposed to test compounds and [3H]-DHR is applied intraventricularly for 2-6 h before the brain is dissected and arranged for the preparation of tissue slices (Talpade et al. 2000). In untreated animals, this method allows a precise spatial resolution of the expression pattern of CI. In animals with impaired CI activity, either as a result of CI deficiencies, or upon treatment with CI inhibitors, the assay allows an assessment of the degree of CI inhibition. Complex I Enzyme Activity (Colorimetric) The analysis of mitochondrial OXPHOS CI enzyme activity can be performed using human, rat, mouse and bovine cell and tissue extracts (abcam: <a class="external free" href="http://www.abcam.com/complex-i-enzyme-activity-microplate-assay-kit-colorimetric-ab109721" rel="nofollow" target="_blank">http://www.abcam.com/complex-i-enzyme-activity-microplate-assay-kit-colorimetric-ab109721</a>). Capture antibodies specific for CI subunits are pre-coated in the microplate wells. Samples are added to the microplate wells which have been pre-coated with a specific capture antibody. After the target has been immobilized in the well, CI activity is determined by following the oxidation of NADH to NAD+ and the simultaneous reduction of a dye which leads to increased absorbance at OD=450 nm. By analyzing the enzyme's activity in an isolated context, outside of the cell and free from any other variables, an accurate measurement of the enzyme's functional state can be evaluated.

CI has a highly conserved subunit composition across species, from lower organisms to mammals (Cardol, 2011). Fourteen subunits are considered to be the minimal structural requirement for physiological functionality of the enzyme. These units are well conserved among bacterial (E. coli), human (H. sapiens), and Bovine (B. taurus) (Vogel et al., 2007b; Ferguson, 1994). However, the complete structure of CI is reported to contain between 40 to 46 subunits and the number of subunits differs, depending on the species (Gabaldon 2005; Choi et al., 2008). In vertebrates CI consists of at least 46 subunits (Hassinen, 2007), particularly, in humans 45 subunits have been described (Vogel et al, 2007b). Moreover, enzymatic and immunochemical evidence indicate a high degree of similarity between mammalian and fungal counterparts (Lummen, 1998). Mammalian CI structure and activity have been characterized in detail (Vogel et al., 2007a; Vogel et al., 2007b), referring to different human organs including the brain. There is also a substantial amount of studies describing CI in human muscles, brain, liver, as well as bovine heart (Janssen et al., 2006; Mimaki et al. 2012) (Okun et al., 1999).

CL:0000255

CL eukaryotic cell

High High High

Betarbet R, Sherer TB, MacKenzie G, Garcia-Osuna M, Panov AV, Greenamyre JT. (2000). Chronic systemic pesticide exposure reproduces features of Parkinson's disease. Nat Neurosci 3:1301-1306. Cardol, P., (2011) Mitochondrial NADH:ubiquinone oxidoreductase (complex I) in eukaryotes: A highly conserved subunit composition highlighted by mining of protein databases Biochimica et Biophysica Acta 1807, 1390–1397. Choi WS., Kruse S.E., Palmiter R, Xia Z., (2008) Mitochondrial complex I inhibition is not required for dopaminergic neuron death induced by rotenone, MPP, or paraquat. PNAS, 105, 39, 15136-15141. Degli Esposti (1998) Inhibitors of NADH-ubiquinone reductase: an overview Biochimica et Biophysica Acta 1364-222-235. Desplats P, Patel P, Kosberg K, Mante M, Patrick C, Rockenstein E, Fujita M, Hashimoto M, Masliah E. (2012). Combined exposure to Maneb and Paraquat alters transcriptional regulation of neurogenesis-related genes in mice models of Parkinson’s disease. Mol Neurodegener 7:49. <a href="https://www.ncbi.nlm.nih.gov/pubmed/?term=Fendel%20U%5BAuthor%5D&cauthor=true&cauthor_uid=18486594">Fendel U</a>, <a href="https://www.ncbi.nlm.nih.gov/pubmed/?term=Tocilescu%20MA%5BAuthor%5D&cauthor=true&cauthor_uid=18486594">Tocilescu MA</a>, <a href="https://www.ncbi.nlm.nih.gov/pubmed/?term=Kerscher%20S%5BAuthor%5D&cauthor=true&cauthor_uid=18486594">Kerscher S</a>, <a href="https://www.ncbi.nlm.nih.gov/pubmed/?term=Brandt%20U%5BAuthor%5D&cauthor=true&cauthor_uid=18486594">Brandt U</a>. (2008) Exploring the inhibitor binding pocket of respiratory complex I.<a href="https://www.ncbi.nlm.nih.gov/pubmed/18486594" title="Biochimica et biophysica acta.">Biochim Biophys Acta.</a> 2008, 1777(7-8):660-5. Ferguson SJ. Similarities between mitochondrial and bacterial electron transport with particular reference to the action of inhibitors. Biochem Soc Trans. 1994 Feb;22(1):181-3. Friedrich T, van Heek P, Leif H, Ohnishi T, Forche E, Kunze B, Jansen R, TrowitzschKienast W, Hofle G & Reichenbach H (1994) Two binding sites of inhibitors in NADH: ubiquinone oxidoreductase (complex I). Relationship of one site with the ubiquinone-binding site of bacterial glucose:ubiquinone oxidoreductase. Eur J Biochem 219(1–2): 691–698. Gabaldon, T., Rainey, D., Huynen, M.A. (2005) Tracing the evolution of a large protein complex in the eukaryotes, NADH:ubiquinone oxidoreductase (Complex I), J. Mol. Biol. 348; 857–870. Greenamyre, J T., Sherer, T.B., Betarbet, R., and Panov A.V. (2001) Critical Review Complex I and Parkinson’s Disease Life, 52: 135–141. Greenamyre JT, Higgins DS, Eller RV (1992) Quantitative autoradiography of dihydrorotenone binding to complex I of the electron transport chain. J Neurochem. 59(2):746-9. Grivennikova, V.G., Maklashina, E.O., E.V. Gavrikova, A.D. Vinogradov (1997) Interaction of the mitochondrial NADH-ubiquinone reductase with rotenone as related to the enzyme active/inactive transition Biochim. Biophys. Acta, 1319 (1997), pp. 223–232. Haefeli, RH (2012) Molecular Effects of Idebenone. Doctoral thesis <a class="external free" href="http://edoc.unibas.ch/19016/1/Molecular_Effects_of_Idebenone_Roman_Haefeli.pdf" rel="nofollow" target="_blank">http://edoc.unibas.ch/19016/1/Molecular_Effects_of_Idebenone_Roman_Haefeli.pdf</a>. Hassinen I (2007) Regulation of Mitochondrial Respiration in Heart Muscle. In Mitochondria – The Dynamic Organelle Edited by Schaffer & Suleiman. Springer ISBN-13: 978-0-387-69944-8. Hideto M. Structure–activity relationships of some complex I inhibitors. Biochimica et Biophysica Acta 1364 _1998. 236–244. Higgins DS Jr1, Greenamyre JT. (1996). [3H]dihydrorotenone binding to NADH: ubiquinone reductase (complex I) of the electron transport chain: an autoradiographic study. J Neurosci. 1996 Jun 15;16(12):3807-16. Ichimaru, N., Murai, M., Kakutani, N., Kako, J., Ishihara, A., Nakagawa, Y., … Miyoshi, H. (2008). Synthesis and Characterization of New Piperazine-Type Inhibitors for Mitochondrial NADH-Ubiquinone Oxidoreductase (Complex I). Biochemistry, 47(40), 10816–10826. Ino T, Takaaki N, Hideto M. Characterization of inhibitor binding sites of mitochondrial complex I using fluorescent inhibitor. Biochimica et Biophysica Acta 1605 (2003) 15– 20. Janssen RJ, Nijtmans LG, van den Heuvel LP, Smeitink JA. Mitochondrial complex I: structure, function and pathology. J Inherit Metab Dis. 2006 Aug;29(4):499-515. Keeney PM, Xie J,Capaldi RA,Bennett JP Jr. (2006) Parkinson's disease brain mitochondrial complex I has oxidatively damaged subunits and is functionally impaired and misassembled. J Neurosci. 10;26(19):5256-64. Lin CJ, Lee CC, Shih YL, Lin CH, Wang SH, Chen TH, Shih CM. (2012). Inhibition of Mitochondria- and Endoplasmic Reticulum Stress-Mediated Autophagy Augments Temozolomide-Induced Apoptosis in Glioma Cells. PLoS ONE 7:e38706. Lümmen, P., (1998) Complex I inhibitors as insecticides and acaricides1, Biochimica et Biophysica Acta (BBA) - Bioenergetics, Volume 1364, Issue 2, Pages 287-296. Majander A, Finel M, Wikstrom M. (1994) Diphenyleneiodonium inhibits reduction of iron–sulfur clusters in the mitochondrial NADH–ubiquinone oxidoreductase (complex I) J Biol Chem. 269:21037–21042. Miao YJ, Xu XF, Xu F, Chen Y, Chen JW, Li X. (2014) The structure-activity relationships of mono-THF ACGs on mitochondrial complex I with a molecular modelling study. Nat Prod Res.28(21):1929-35. Mimaki M, Wang X, McKenzie M, Thorburn DR, Ryan MT. Understanding mitochondrial complex I assembly in health and disease. Biochim Biophys Acta. 2012 Jun;1817(6):851-62. doi: 10.1016/j.bbabio.2011.08.010. Miyoshi H. Structure-activity relationships of some complex I inhibitors. Biochim Biophys Acta. 1998, 6:236-244. Okun, J.G, Lümmen, P and Brandt U., (1999) Three Classes of Inhibitors Share a Common Binding Domain in Mitochondrial Complex I (NADH:Ubiquinone Oxidoreductase) J. Biol. Chem. 274: 2625-2630. doi:10.1074/jbc.274.5.2625. Ramsay R., Krueger MJ., Youngster SK., Gluck MR., Casida J.E. and Singer T.P. Interaction of 1-Methyl-4-Phenylpyridinium Ion (MPP+) and Its Analogs with the Rotenone/Piericidin Binding Site of NADH Dehydrogenase. Journal of Neurochemistry, 1991, 56: 4, 1184–1190. Sava V, Velasquez A, Song S, Sanchez-Ramos J. (2007). Dieldrin elicits a widespread DNA repair and antioxidative response in mouse brain. J Biochem Mol Toxicol 21:125-135. <pre> Schuler, F. and Casida, JE. (2001) Functional coupling of PSST and ND1 subunits in NADH:ubiquinone oxidoreductase established by photoaffinity labeling, Biochimica et Biophysica Acta (BBA) - Bioenergetics, Volume 1506, Issue 1, 2 July 2001, Pages 79-87, ISSN 0005-2728, https://doi.org/10.1016/S0005-2728(01)00183-9.</pre> Talpade DJ, Greene JG, Higgins DS Jr, Greenamyre JT (2000) In vivo labeling of mitochondrial complex I (NADH:ubiquinone oxidoreductase) in rat brain using [(3)H]dihydrorotenone. J Neurochem. 75(6):2611-21. Vogel R.O., van den Brand M.A., Rodenburg R.J., van den Heuvel L.P., Tsuneoka M., Smeitink J.A., Nijtmans L.G. (2007a). Investigation of the complex I assembly chaperones B17.2L and NDUFAF1 in a cohort of CI deficient patients. Mol. Genet. Metab. 91:176–182. Vogel, R.O. Smeitink, J.A. Nijtmans L.G. (2007b) Human mitochondrial complex I assembly: a dynamic and versatile process Biochim. Biophys. Acta, 1767-. 1215–1227. Zharova, TV, and Vinogradov, A.(1997) A competitive inhibition of the mitochondrial NADH-ubiquinone oxidoreductase (Complex I) by ADP-ribose. Biochimica et Biophysica Acta, 1320:256-64. AOPWiki 2016-11-29T18:41:27

2018-03-28T04:51:46

Inhibition, NADH-ubiquinone oxidoreductase (complex I)

Cellular

Under physiological conditions complex I (CI) couples the oxidation of NADH to NAD+ by reducing flavin mononucleotide (FMN) to FMNH2. FMNH2 is then oxidized through a semiquinone intermediate. Each electron moves from the FMNH2 to Fe-S clusters, and from the Fe-S clusters to ubiquinone (Q). Transfer of the first electron results in the formation of the free-radical (semiquinone) form of Q, and transfer of the second electron reduces the semiquinone form to the ubiquinol form (CoQH2). Altogether, four protons are translocated from the mitochondrial matrix to the inter-membrane space for each molecule of NADH oxidized at CI. This leads to the establishment of the electrochemical potential difference (proton-motive force) that may be used to produce ATP (Garrett and Grisham, 2010). Binding of an inhibitor attenuates or completely blocks the activity of CI, i.e. the oxidation of NADH is impaired and protons are not moved. This causes two major consequences: first, electrons are channelled toward oxygen instead Q. This impairs normal oxygen reduction into water at complex IV and leads to the formation of the ROS superoxide at other sites of the respiratory chain. Superoxide may cause damage of proteins, lipid and DNA of the cell, or damage components of the mitochondria after transformation into e.g. hydrogen peroxide. These processes result in mitochondrial dysfunction (Voet and Voet., 2008). The second consequence is the increase of the NADH/NAD+ ratio in mitochondria. This affects the function of key dehydrogenase enzymes in the citric acid cycle and can lead to its block, resulting in an inhibition of mitochondrial ATP production and mitochondrial respiration. Prolonged treatment with an inhibitor results in a severe, progressive and irreversible inhibition of complex I, most likely by indirect mechanisms involving oxidative damage (Cleeter et al., 1992). The functional consequences of CI inhibition have been titrated in a time- and dose-dependent manner (Barrientos and Moraes, 1999), with mitochondrial dysfunction measured by a range of different assays (Barrientos and Moraes, 1999; Greenamyre et al., 2001). These included quantification of ROS derived from mitochondria, and of cellular respiration (see KE2: Mitochondrial dysfunction).

As CI has an enzymatic function as such, but also contributes to the overall function of oxidative phosphorylation, there are two fundamental approaches to assess CI inhibition. The first approach measures the enzymatic activity of the complex itself; the second one assesses the overall activity of oxidative phosphorylation of entire mitochondria, and indirectly infers from this a potential dysfunction of CI. I. Direct detection of complex I activity. This type of assay is always performed in homogenates of cells or tissues, and requires at least a partial purification of mitochondria or respiratory chain components. In order to focus on CI activity, the activities of Complexes III (e.g. antimycin A) and complex IV (e.g. cyanide) need to be blocked by pharmacological inhibitors in these setups. 1. Forward Electron Transfer. Submitochondrial particles or intact isolated mitochondria are incubated with NADH as electron donor and with an electron acceptor to measure the flow of electrons from NADH, through CI to the acceptor. As readout, either the consumption of NADH, or the reduction of the electron acceptor is followed photometrically or fluorometrically (Lenaz et al. 2004; Spinazzi et al. 2012; Long et al. 2009; Kirby et al. 2007). The physiological electron acceptor of CI is Coenzyme Q10 (CoQ10). Due to its hydrophobicity, it is not suitable for use in an experimental in vitro setup. Short-chain analogs of CoQ10, such as CoQ1 or decylubiquinone (DB) with a 10 carbon-atom linear saturated side chain are hence applied as alternatives. With these non-physiological electron acceptors, it is important to consider that the activity of CI can easily be underestimated. As water-soluble electron acceptors, either ferricyanide or 2,6-dichlorophenolidophenol (DCIP) are used. However the reduction of such compounds is not strictly coupled to the transduction of energy. To identify the portion of rotenone-inhibitable CI activity, all samples investigated are assayed in parallel following treatment with rotenone. In contrast to the autoradiography assays, direct CI activity detection allows the identification also of CI inhibitors that bind to sites of CI different from the rotenone binding site. 2. Reverse Electron Transfer. An alternative setup for the direct measurement of CI activity with minimal interference by the activities of complex III and complex IV make use of the observation of a general reversibility of oxidative phosphorylation and electron flow across the mitochondrial respiratory chain (Ernster et al. 1967). With this method, electrons enter the respiratory chain via complex II. Based on the reverse flux, this method allows the complete circumvention of complexes III and IV. As electron donor, succinate is applied, together with NAD+ as electron acceptor. Formation of NADH from NAD+ can be determined photometrically. The succinate-linked NAD+ reduction can be performed either with intact isolated mitochondria or with submitochondrial particles. For the direct assessment of CI activity, submitochondrial particles are used. For assays with intact mitochondria, the succinate-linked reduction of NAD+ is performed in the presence of ATP as energy source. Potassium cyanide (KCN) is added for inhibition of forward electron transport towards complex IV. 3. Complex I activity dipstick assay. To assess CI activity and its inhibition in cell or tissue homogenates without interference by other components of the respiratory chain, CI-selective antibodies attached to a matrix (e.g. multiwell plates) are used (Willis et al., 2009). Homogenized tissue can directly be added for capturing of CI, the unbound supernatant is washed away and leaves a complex of the antibody and mitochondrial CI. For activity determination, NADH as electron donor and nitroblue tetrazolium (NBT) as acceptor are added. Reduced NBT forms a colored precipitate, its signal intensity is proportional to the amount of CI bound to the antibody. CI inhibitors can directly be added for an assessment of their inhibitory potential. This method, when applied in e.g. 96-well or 384-well plates, allows screening of large sets of potential CI inhibitors without any interference by other elements of the mitochondrial respiratory chain. II. Indirect measurements of complex I activity. Such assays mostly require / allow the use of live cells. 1. Oxygen consumption. Electrons, fed into the mitochondrial respiratory chain either by CI or complex II, ultimately reduce molecular oxygen to water at complex IV. In a closed system, this consumption of oxygen leads to a drop of the overall O2 concentration, and this can serve as parameter for mitochondrial respiratory activity. Measurements are traditionally done with a Clark electrode, or with more sophisticated optical methods. At the cathode of a Clark electrode, oxygen is electrolytically reduced, which initiates a current in the electrode, causing a potential difference that is ultimately recorded. Clark electrodes however have the disadvantage that oxygen is consumed. Furthermore, interferences with nitrogen oxides, ozone, or chlorine are observed (Stetter et al., 2008). To circumvent these limitations, optical sensors have been developed that have the advantage that no oxygen is consumed, combined with a high accuracy and reversibility. Optical oxygen sensors work according to the principle of dynamic fluorescence quenching. The response of the respective fluorescence dye is proportional to the amount of oxygen in the sample investigated (Wang and Wolfbeis, 2014). In a model of isolated mitochondria in the absence of complex II substrates, oxygen consumption can serve as surrogate readout for the assessment of the degree of CI inhibition. It is however essential to realize that also complex III and complex IV activities are involved and their inhibition also results in a decline in O2 consumption. In addition to that, CI inhibitors can lead to a one-electron reduction of molecular oxygen at the site of CI to yield superoxide. The amount of superoxide formed hence contributes to the consumption of oxygen, but this must not be interpreted as oxygen consumption as a result of controlled and coupled electron flux through the complexes of the mitochondrial respiratory chain. A modern convenient method to measure oxygen consumption is provided by the Seahorse technology of extracellular flux (XF) analysis, in which cells are kept in a very small volume, so that changes of oxygen levels can be detected very sensitively by an oxygen sensor. To allow manipulation of the mitochondria in cells, the cell membrane can be permeabilized with saponin (SAP), digitonin (DIG) or recombinant perfringolysin O (rPFO) (XF-plasma membrane permeabilizer (PMP) reagent), to allow addition of specific substrates to measure activity of different respiratory chain complexes, including CI. (Salabei et al., 2014). 2. Intracellular ATP levels. Intracellular ATP levels originate both from mitochondria and from glycolysis. If glycolytic ATP production is impaired or inhibited, the cellular production of ATP is a measure of mitochondrial function. If it is assumed that the ATP consumption remains constant, then the steady state ATP levels can serve as indirect readout for mitochondrial activity, and the latter depends on the functioning of CI. Inhibitors of CI reduce cellular ATP levels, but it has to be remembered that intracellular ATP levels are also affected by inhibitors of other parts of the respiratory chain, of the citric acid cycle or of the transport of energy substrates. For a proper interpretation of assay results, it has to be ascertained in each particular test system, that ATP production from other sources is excluded and that the cellular ATP consumption remains constant. ATP levels can be easily measured from lysates of in vitro cell cultures or from tissues by a luminometric luciferase/luciferin assay. The amount of light emitted is proportional to the amount of ATP in the sample (Nguyen et al. 1988, Leist et al., 1997). 3. Other approaches. As mitochondrial activity is coupled to many cellular functions, there is a multitude of other indirect assays that are sensitive to inhibitors of CI. Some of these tests may indeed be very sensitive, while they have a low specificity. Thus, their application requires usually a good control of the experimental system and care with the interpretation of the data. One exemplary approach is the measurement of NADH/NAD+ ratios in mitochondria by imaging methods. This provides resolution on the level of individual mitochondria within a living cell (van Vliet et al., 2014).

The CI is well-conserved across species from lower organisms to mammals. The central subunits of CI harboring the bioenergetic core functions are conserved from bacteria to humans. CI from bacteria and from mitochondria of Yarrowia lipolytica, a yeast genetic model for the study of eukaryotic CI (Kerscher et al., 2002) was analyzed by x-ray crystallography (Zickermann et al., 2015, Hofhaus et al., 1991; Baradaran et al., 2013). The CI of the mitochondria of eukaryotes and in the plasma membranes of purple photosynthetic bacteria are closely related to respiratory bacteria and the close homology of sequences, function, and prosthetic groups shows a common ancestry (Friedrich et al., 1995).

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Baradaran R., John M. Berrisford, Gurdeep S. Minhas , Leonid A. Sazanov. Crystal structure of the entire respiratory complex I. Nature , 2013,| 494,443–448. Barrientos A., and Moraes C.T. (1999) Titrating the Effects of Mitochondrial Complex I Impairment in the Cell Physiology. Vol. 274, No. 23, pp. 16188–16197. Cleeter MW, Cooper JM, Schapira AH. Irreversible inhibition of mitochondrial complex I by 1-methyl-4-phenylpyridinium: evidence for free radical involvement. J Neurochem. 1992 Feb;58(2):786-9. Degli Esposti (1998) Inhibitors of NADH-ubiquinone reductase: an overview Biochimica et Biophysica Acta 1364-222-235. Ernster L, Lee C (1967) Energy-linked reduction of NAD+ by succinate. Methods Enzym. 10:729-738. Friedrich, T., Steinmüller, K. & Weiss, H. (1995) The proton-pumping respiratory complex I of bacteria and mitochondria and is homologue of chloroplasts. FEBS Lett. (Minireview), 367, 107-111. Garrett and Grisham, Biochemistry, Brooks/Cole, 2010, pp 598-611. Greenamyre, J T., Sherer, T.B., Betarbet, R., and Panov A.V. (2001) Critical Review Complex I and Parkinson’s Disease Life, 52: 135–141. Hofhaus, G., Weiss, H. and Leonard, K. (1991): Electron microscopic analysis of the peripheral and the membrane parts of mitochondrial NADH dehydrogenase (Complex I). J. Mol. Biol. 221, 1027-1043. Kerscher, S. Dröse, K. Zwicker, V. Zickermann, U. Brandt Yarrowia lipolytica, a yeast genetic system to study mitochondrial complex I. Biochim. Biophys. Acta 1555, 83–91 (2002). Kirby DM, Thorburn DR, Turnbull DM, Taylor RW (2007) Biochemical assays of respiratory chain complex activity. Methods Cell Biol. 80:93-119. Leist M, Single B, Castoldi AF, Kühnle S, Nicotera P (1997) Intracellular adenosine triphosphate (ATP) concentration: a switch in the decision between apoptosis and necrosis. J Exp Med. 185:1481-6. Leist M. Current approaches and future role of high content imaging in safety sciences and drug discovery. ALTEX. 2014;31(4):479-93. Lenaz G, Fato R, Baracca A, Genova ML (2004) Mitochondrial quinone reductases: complex I. Methods Enzymol. 382:3-20. Long J, Ma J, Luo C, Mo X, Sun L, Zang W, Liu J (2009) Comparison of two methods for assaying complex I activity in mitochondria isolated from rat liver, brain and heart. Life Sci. 85(7-8):276-80. Nguyen VT, Morange M, Bensaude O. (1988) Firefly luciferase luminescence assays using scintillation counters for quantitation in transfected mammalian cells. Anal Biochem. 171(2):404-8. van Vliet E, Daneshian M, Beilmann M, Davies A, Fava E, Fleck R, Julé Y, Kansy M, Kustermann S, Macko P, Mundy WR, Roth A, Shah I, Uteng M, van de Water B, Hartung T, Spinazzi M, Casarin A, Pertegato V, Salviati L, Angelini C (2012) Assessment of mitochondrial respiratory chain enzymatic activities on tissues and cultured cells. Nat Protoc. 7(6):1235-46. Salabei J.K., Gibb A.A. and Hill BG. (2014) Comprehensive measurement of respiratory activity in permeabilized cells using extracellular flux analysis. Nature Protocols, 9, 421–438. Stetter JR, Li J (2008) Amperometric gas sensors--a review. Chem Rev. 108(2):352-66. Wang XD, Wolfbeis OS (2014) Optical methods for sensing and imaging oxygen: materials, spectroscopies and applications. Chem Soc Rev. 43(10):3666-761. Voet DJ and Voet JG; Pratt CW (2008). Chapter 18, Mitochondrial ATP synthesis. Principles of Biochemistry, 3rd Edition. Wiley. p. 608. <a class="internal mw-magiclink-isbn" href="/wiki/index.php/Special:BookSources/9780470233962">ISBN 978-0-470-23396-2</a>. Willis JH, Capaldi RA, Huigsloot M, Rodenburg RJ, Smeitink J, Marusich MF (2009) Isolated deficiencies of OXPHOS complexes I and IV are identified accurately and quickly by simple enzyme activity immunocapture assays. Biochim Biophys Acta. 1787(5):533-8. Zickermann V., Christophe Wirth, Hamid Nasiri, Karin Siegmund, Harald Schwalbe, Carola Hunte, Ulrich Brandt. Mechanistic insight from the crystal structure of mitochondrial complex I. Science 2 January 2015: Vol. 347 no. 6217 pp. 44-49. AOPWiki 2016-11-29T18:41:27

2018-03-12T11:03:29

Impaired, Proteostasis

Cellular

The concept of proteostasis refers to the homeostasis of proteins in space and time, i.e. the correct balance between protein synthesis, modification, transport and degradation. Disturbance of proteostasis results in pathological changes either by loss of function events (lack of a pivotal protein/protein function) or by a gain of undesired functions (aggregation of a protein leading to the formation of inclusions and new structures in cells and disturbing turnover of many unrelated proteins). Proteostasis regulation is the main defence mechanism against toxic proteins, whose accumulation could greatly compromise normal cellular function and viability. Therefore, the chaperone and degradation systems assuring the removal of misfolded and aggregated proteins, as well as damaged, dysfunctional cellular organelles (e.g. defective mitochondria) play a key role in cellular homeostasis (Lee et al., 2012). The two major degradation systems are the ubiquitin–proteasome system (UPS) and the autophagy–lysosome pathway (ALP) (Korolchuk et al., 2010; Kroemer et al., 2010; Ravikumar et al., 2010). The UPS works through the attachment of multiple ubiquitin molecules to a protein substrate, followed by the subsequent degradation of the tagged polyubiquitinated protein by the proteasome (Ciechanover, 1998; Ciechanover and Brundin, 2003). A compromised function of the UPS leads to the accumulation of ubiquitylated proteins, such as α-synuclein (Ii et al. 1997; Spillantini et al. 1997; Sulzer and Zecca 2000). The accumulation of polyubiquitinated proteins, as a consequence of a dysfunctional proteasome activity, is observed in some pathologies, and experimental inhibition of the proteasome has been shown to trigger parkinsonian neurodegeneration (McNaught and Jenner 2001; Hardy et al., 2001). ALP involves the engulfment of cytoplasmic materials into autophagosomes, which are degraded by lysosomal enzymes after fusion of autophagosomes with lysosomes (Kuma et al., 2004) or direct import of proteins into lysosomes (Cuervo, 2004; Mizushima et al., 2008). Autophagy also plays an essential role for the removal of damaged organelles, such as mitochondria. Both, excessive autophagy or reduced autophagic flux can compromise cell survival (Rothermel and Hill, 2007), and several genetic forms of PD are linked to the autophagy-related genes Pink1, Parkin or Uchl1. Autophagy enables cell survival during mitochondrial stress by clearing the damaged organelles (Lee et al., 2012). One of the main aggregated proteins found to accumulate in nigrostriatal cells during Parkinson's disease is α-synuclein. Aggregation of α-synuclein can obstruct normal cellular transport, leading to impaired intracellular trafficking and/or trapping of cellular organelles in inappropriate locations, this resulting in synaptic and cell dysfunctions (Bartels et al., 2011) (Bellucci A., et al., 2012; Cookson MR., 2005; Games D., et al., 2013; Hunn BH., et al., 2015). Importantly, accumulation of α-synuclein affects mitochondrial trafficking. The polarity and correct function of different types of cells depend on an efficient transport of mitochondria to areas of high energy consumption (Sheng, 2014). Therefore, the correct distribution of mitochondria to various parts of a cell is essential to preserve cell function (Schwarz, 2013; Zhu et al., 2012).

1. Evaluation of UPS function. General turnover assays: quantitative evaluation can be based on the detection of increased ubiquitin or ubiquinated proteins, as well as proteasomal subunits, either by immunocyto/histochemistry or by western blotting (Rideout et al., 2001; Ortega and Lucas, 2014). UPS activity can be continuously monitored by quantitating (by mean of flow cytometry or microscopy) the level of e.g. EGFP-degron fusion proteins (green fluorescent protein) that are selectively degraded by the proteasome (Bence et al., 2001). Proteasome activity assay. Various fluorogenic substrates (e.g., Suc-Leu-Leu-Val-Tyr-AMC for the chymotrypsin-like activity) can be used for the determination of proteasomal activity in in vivo or in vitro applications. These substrates may be applied to tissue or cell homogenates, but specific measurements require partial purification of the proteasome (Kisselev and Goldberg, 2005). Detection of α-synuclein (AS) aggregates. The most common methods to detect AS aggregates use immunostaining for AS (in cells or in tissues). In cell culture, AS may also be epitope-tagged or coupled to GFP to allow an indirect detection. The detection of small, not microscopically-visible AS aggregates is indicative of protease-resistance. Tissue slices may be exposed to proteases before immunostaining for AS. Alternatively, small or large aggregates may be biochemically enriched by differential centrifugation and proteolytic treatment, and then analyzed, e.g., by western blot, mass spectrometry or ELISA-like immunoquantification. 2. Evaluation of ALP function. Quantification of lysosomes or autophagosomes. Disturbances of ALP often result in counter-regulations that can be visualized by staining of lysosomes or parts of the autophagy system. Several weakly basic dyes can be used to stain acidic organelles (lysosomes) in live cells. For example, the dye LysoTracker Red stains lysosomes and can be used to monitor autophagy (Klionsky et al., 2007; Klionsky et al., 2008). The autofluorescent drug monodansylcadaverine (MDC) has also been used as autophago-lysosome marker (Munafó and Colombo, 2002). A convenient way to stain lysosomes in tissue or fixed cells is the use of antibodies against the Lysosomal-Associated Membrane Protein 1 (LAMP-1) (Rajapakshe et al., 2015) or against cathepsins (Foghsgaard et al., 2001). For qualitative or semiquantitative estimates of lysosomes and related organelles, transmission electron microscopy has been frequently used (Barth et al., 2010). Monitoring of autophagy-related molecules. The amount and the localization of autophagy-related proteins can change during disturbance of the ALP. Especially in cell culture, but also in transgenic mice, various techniques have been used to monitor autophagy by mean of fluorescence-tags or other substrates, e.g., ATG, autophagy-related protein or autophagy substrates, to monitor their fate in cells and thus provide information on disturbed ALP, or the over-expression of GFP–LC3, in which GFP (green fluorescent protein) is expressed as a fusion protein at the amino terminus of LC3 (microtubule-associated protein 1A/1B-light chain 3), which is the a mammalian homologue of S. cerevisiae ATG8 (Kadowaki and Karim, 2009). Monitoring autophagic flux. The lysosomal degradation of the autophagic cargo constitutes the autophagic flux, which can be measured by assessing the rate of turnover of long-lived proteins that are normally turned over by autophagy (Bauvy et al., 2009). This is performed by labelling intracellular proteins with either [14C]-leucine or [14C]-valine, followed by a long culture period in standard medium. The release of radioactive leucin or valin into the culture medium corresponds to the protein degradation rate in cells, and it may be measured by liquid scintillation counting. Monitoring the conversion of LC3-I to LC3-II. The progression of autophagy (autophagic flux) can be studied by the conversion of LC3-I into LC3-II (i.e. a post-translational modification specific for autophagy) by mean of Western blot analysis. The amount of LC3-II correlates with the number of autophagosomes. Conversion of LC3 can be used to examine autophagic activity in the presence or absence of lysosomal activity (Klionsky et al., 2007; Klionsky et al., 2008). The technology can also be used in vivo, e.g. by the use of transgenic mice that overexpress GFP–LC3 (Kuma et al., 2004). 3. Evaluation of intracellular transport of mitochondria and other organelles. A range of technologies has been used to visualize mitochondrial dynamics in live cells (Jakobs, 2006; Grafstein and Forman, 1980). They usually employ a combination of mitochondrial labelling with fluorescent dyes (e.g. DiOC6 (3, 3′-Dihexyloxacarbocyanine iodide), JC-1 (5,5′,6,6′-Tetrachloro-1,1′,3,3′ tetraethylbenzimida-zolylcarbo-cyanine iodide), MitoTracker, MitoFluor probes, etc.), followed by video- or confocal microscopy for live cell imaging (Schwarz, 2013; Pool et al., 2006). Most frequently, mitochondrial mobility is observed along neurites, and measurable endpoints may be mitochondrial speed and direction with regard to the cell soma (Schildknecht et al. 2013). Additionally, also mitochondrial fusion and fission have been monitored by such methods (Exner et al., 2012). The transport of other organelles along neurites may be monitored using similar methods, and the microtubule structures that serve as transport scaffold may be co-stained.

The ubiquitin proteasome system is highly conserved in eukaryotes, from yeast to human. Ubiquitin is a small (8.5 kDa) regulatory protein that has been found in almost all tissues of eukaryotic organisms. For instance, drosophila has been used as PD model to study the role of ubiquitin in α-synuclein induced-toxicity (Lee et al., 2009). Human and yeast ubiquitin share 96% sequence identity. Neither ubiquitin nor the ubiquitination machinery are known to exist in prokaryotes. Autophagy is ubiquitous in eukaryotic cells and is the major mechanism involved in the clearance of oxidatively or otherwise damaged/worn-out macromolecules and organelles (Esteves et al., 2011). Due to the high degree of conservation, most of the knowledge on autophagy proteins in vertebrates is derived from studies in yeast (Klionsky et al., 2007). Autophagy is seen in all eukaryotic systems, including fungi, plants, slime mold, nematodes, fruit flies and insects, rodents (i.e., laboratory mice and rats), and humans. It is a fundamental and phylogenetically conserved self-degradation process that is characterized by the formation of double-layered vesicles (autophagosomes) around intracellular cargo for delivery to lysosomes and proteolytic degradation.

CL:0000255

CL eukaryotic cell

High High Moderate

Barth S., Danielle Glick, and Kay F Macleod, Autophagy: assays and artifacts. J Pathol. 2010 Jun; 221(2): 117–124. Bartels T, Choi JG, Selkoe DJ (Sep 2011). "α-Synuclein occurs physiologically as a helically folded tetramer that resists aggregation". Nature 477 (7362): 107–10. Bauvy C, Meijer AJ, Codogno P. Assaying of autophagic protein degradation. Methods Enzymol. 2009;452:47–61. Bellucci A., M. Zaltieri, L. Navarria, J. Grigoletto, C. Missale, and P. Spano, “From α-synuclein to synaptic dysfunctions: new insights into the pathophysiology of Parkinson’s disease,” Brain Research, vol. 1476, pp. 183–202, 2012. Bence NF, Sampat RM, Kopito RR. Impairment of the ubiquitin–proteasome system by protein aggregation. Science 2001;292:1552–5. Ciechanover A. (1998) The ubiquitin-proteasome pathway: on protein death and cell life. EMBO J. 17, 7151±7160. Ciechanover A., and Brundin P., 2003, The Ubiquitin Proteasome System in Neurodegenerative Diseases: Sometimes the Chicken, Sometimes the Egg. Neuron, 427–446 Cookson MR., “The biochemistry of Parkinson’s disease,” Annual Review of Biochemistry, vol. 74, pp. 29–52, 2005. Cuervo A.M., “Autophagy: many paths to the same end,” Molecular and Cellular Biochemistry, vol. 263, no. 1, pp. 55–72, 2004. Exner N, Lutz AK, Haass C, Winklhofer KF. Mitochondrial dysfunction in Parkinson's disease: molecular mechanisms and pathophysiological consequences. EMBO J. 2012 Jun 26;31(14):3038-62. Esteves AR, Arduíno DM, Silva DF, Oliveira CR, Cardoso SM. 2011. Mitochondrial Dysfunction: The Road to Alpha-Synuclein Oligomerization in PD. Parkinsons Dis. 2011:693761. Foghsgaard L, Wissing D, Mauch D, Lademann U, Bastholm L, Boes M, Elling F, Leist M, Jäättelä M. Cathepsin B acts as a dominant execution protease in tumor cell apoptosis induced by tumor necrosis factor. J Cell Biol. 2001 May 28;153(5):999-1010. Games D., P. Seubert, E. Rockenstein et al., “Axonopathy in an α-synuclein transgenic model of Lewy body disease is associated with extensive accumulation of c-terminal-truncated α-synuclein,” American Journal of Pathology, vol. 182, no. 3, pp. 940–953, 2013. Grafstein B., and Forman DS. Intracellular transport in neurons. Physiological Reviews Published 1 October 1980 Vol. 60 no. 4. Hardy J. Rideout, Kristin E. Larsen, David Sulzer and Leonidas Stefanis, Proteasomal inhibition leads to formation of ubiquitin/a-synuclein-immunoreactive inclusions in PC12 cells. Journal of Neurochemistry, 2001, 78, 899±908 Hunn BH., S. J. Cragg, J. P. Bolam, M. G. Spillantini, and R. Wade-Martins, “Impaired intracellular trafficking defines early Parkinson’s disease,” Trends in Neurosciences, vol. 38, no. 3, pp.178–188, 2015. Ii K., Ito H., Tanaka K. and Hirano A. (1997) Immunocytochemical co-localization of the proteasome in ubiquitinated structures in neurodegenerative diseases and the elderly. J. Neuropathol. Exp. Neurol. 56, 125-131. Jakobs S., High resolution imaging of live mitochondria, 2006, Biochimica et Biophysica Acta (BBA) - Molecular Cell Research. 1763, Issues 5–6 Pages 561–575 Kadowaki M, Karim MR. Cytosolic LC3 ratio as a quantitative index of macroautophagy. Methods Enzymol. 2009;452:199–213. [PubMed] Kisselev AF, Goldberg AL. Monitoring activity and inhibition of 26S proteasomes with fluorogenic peptide substrates. Methods Enzymol. 2005;398:364–378. Klionsky DJ., Ana Maria Cuervo & Per O. Seglen. Methods for Monitoring Autophagy from Yeast to Human. Autophagy 2007, 3:3, 181-206; Klionsky D.J., Abeliovich H., Agostinis P., Agrawal D.K., Aliev G., Askew D.S., Baba M., Baehrecke E.H., Bahr B.A., Ballabio A., et al Guidelines for the use and interpretation of assays for monitoring autophagy in higher eukaryotes. Autophagy 2008;4:151-175. Korolchuk VI, Menzies FM, Rubinsztein DC (2010) Mechanisms of cross-talk between the ubiquitin–proteasome and autophagy–lysosome systems. FEBS Lett 584:1393–1398 Kroemer G, Mariño G, Levine B (2010) Autophagy and the integrated stress response. J. Molecular cell 40:280–293. Kuma A, Hatano M, Matsui M, Yamamoto A, Nakaya H, Yoshimori T, et al. The role of autophagy during the early neonatal starvation period. Nature. 2004;432:1032–1036. Lee J, Giordano S, Zhang J; Giordano; Zhang (January 2012). "Autophagy, mitochondria and oxidative stress: cross-talk and redox signalling". Biochem. J. 441 (2): 523–40. Lee FK, Wong AK, Lee YW, Wan OW, Chan HY, Chung KK. The role of ubiquitin linkages on alpha-synuclein induced-toxicity in a Drosophila model of Parkinson's disease. J Neurochem. 2009 Jul;110(1):208-19 McNaught K. S. and Jenner P. (2001) Proteasomal function is impaired in substantia nigra in Parkinson's disease. Neurosci. Lett. 297, 191-194. Mizushima N. et al., 2008. Autophagy fights disease through cellular self-digestion. Nature. 451(7182):1069-75. Review. Munafó DB, Colombo MI. Induction of autophagy causes dramatic changes in the subcellular distribution of GFP-Rab24. Traffic. 2002 Jul;3(7):472-82. Ortega Z. and Lucas J.J. (2014) Ubiquitin–proteasome system involvement in Huntington’s disease Front Mol Neurosci. 2014; 7: 77. Pool M., Rippstein P., Mcbride H. Kothary R., 2006 Trafficking of Macromolecules and Organelles in Cultured Dystonia musculorum Sensory Neurons Is Normal. J. Comparative Neurology 494:549–558 (2006) Rajapakshe AR, Podyma-Inoue KA, Terasawa K, Hasegawa K, Namba T, Kumei Y, Yanagishita M, Hara-Yokoyama M. Lysosome-associated membrane proteins (LAMPs) regulate intracellular positioning of mitochondria in MC3T3-E1 cells. Exp Cell Res. 2015 Feb 1;331(1):211-22. doi: 10.1016/j.yexcr.2014.09.014. Ravikumar B, Sarkar S, Davies JE et al (2010) Regulation of mammalian autophagy in physiology and pathophysiology. Physiol Rev 90:1383–1435. doi:10.1152/physrev.00030.2009 Rothermel BA, Hill JA (2007) Myocyte autophagy in heart disease: friend or foe? Autophagy 3:632–634. Rideout HJ, Larsen KE, Sulzer D, Stefanis L. 2001.Proteasomal inhibition leads to formation of ubiquitin/a-synuclein-immunoreactive inclusions in PC12 cells. Journal of Neurochemistry. 78, 899-908. Schildknecht S, Karreman C, Pöltl D, Efrémova L, Kullmann C, Gutbier S, Krug A, Scholz D, Gerding HR, Leist M. Generation of genetically-modified human differentiated cells for toxicological tests and the study of neurodegenerative diseases. ALTEX. 2013;30(4):427-44. Schwarz TL.Mitochondrial trafficking in neurons. Cold Spring Harb Perspect Biol. 2013 Jun 1;5(6). pii: a011304. Sheng ZH., Mitochondrial trafficking and anchoring in neurons: new insight and implications. J of Cell Biology, vol. 204. No.7 pp. 1087-1098, 2014. Spillantini M. G., Schmidt M. L., Lee V. M., Trojanowski J. Q., Jakes R. and Goedert M. (1997) Alpha-synuclein in Lewy bodies.Nature 388, 839-840. Sulzer D. and Zecca L. (2000) Intraneuronal dopamine-quinonem synthesis: a review. Neurotoxicity Res. 1, 181-195. Zhu XH, Qiao H, Du F, Xiong Q, Liu X, Zhang X, Ugurbil K, Chen W. Quantitative imaging of energy expenditure in human brain. Neuroimage. 2012;60(4):2107-17) AOPWiki 2016-11-29T18:41:27

2018-03-15T12:46:08

Cell injury/death

Cellular

Two types of cell death can be distinguished by morphological features, although it is likely that these are two ends of a spectrum with possible intermediate forms. Apoptosis involves shrinkage, nuclear disassembly, and fragmentation of the cell into discrete bodies with intact plasma membranes. These are rapidly phagocytosed by neighbouring cells. An important feature of apoptosis is the requirement for adenosine triphosphate (ATP) to initiate the execution phase. In contrast, necrotic cell death is characterized by cell swelling and lysis. This is usually a consequence of profound loss of mitochondrial function and resultant ATP depletion, leading to loss of ion homeostasis, including volume regulation, and increased intracellular Ca2+. The latter activates a number of nonspecific hydrolases (i.e., proteases, nucleases, and phospholipases) as well as calcium dependent kinases. Activation of calpain I, the Ca2+-dependent cysteine protease cleaves the death-promoting Bcl-2 family members Bid and Bax which translocate to mitochondrial membranes, resulting in release of truncated apoptosis-inducing factor (tAIF), cytochrome c and endonuclease in the case of Bid and cytocrome c in the case of Bax. tAIF translocates to cell nuclei, and together with cyclophilin A and phosphorylated histone H2AX (γH2AX) is responsible for DNA cleavage, a feature of programmed necrosis. Activated calpain I has also been shown to cleave the plasma membrane Na+–Ca2+ exchanger, which leads to build-up of intracellular Ca2+, which is the source of additional increased intracellular Ca2+. Cytochrome c in cellular apoptosis is a component of the apoptosome. DNA damage activates nuclear poly(ADP-ribose) polymerase-1(PARP-1), a DNA repair enzyme. PARP-1 forms poly(ADP-ribose) polymers, to repair DNA, but when DNA damage is extensive, PAR accumulates, exits cell nuclei and travels to mitochondrial membranes, where it, like calpain I, is involved in AIF release from mitochondria. A fundamental distinction between necrosis and apoptosis is the loss of plasma membrane integrity; this is integral to the former but not the latter. As a consequence, lytic release of cellular constituents promotes a local inflammatory reaction, whereas the rapid removal of apoptotic bodies minimizes such a reaction. The distinction between the two modes of death is easily accomplished in vitro but not in vivo. Thus, although claims that certain drugs induce apoptosis have been made, these are relatively unconvincing. DNA fragmentation can occur in necrosis, leading to positive TUNEL staining (see explanation below). Conversely, when apoptosis is massive, it can exceed the capacity for rapid phagocytosis, resulting in the eventual appearance of secondary necrosis. Two alternative pathways - either extrinsic (receptor-mediated) or intrinsic (mitochondria-mediated) - lead to apoptotic cell death. The initiation of cell death begins either at the plasma membrane with the binding of TNF or FasL to their cognate receptors or within the cell. The latter is due to the occurrence of intracellular stress in the form of biochemical events such as oxidative stress, redox changes, covalent binding, lipid peroxidation, and consequent functional effects on mitochondria, endoplasmic reticulum, microtubules, cytoskeleton, or DNA. The intrinsic mitochondrial pathway involves the initiator, caspase-9, which, when activated, forms an “apoptosome” in the cytosol, together with cytochrome c, which translocates from mitochondria, Apaf-1 and dATP. The apoptosome activates caspase-3, the central effector caspase, which in turn activates downstream factors that are responsible for the apoptotic death of a cell (Fujikawa, 2015). Intracellular stress either directly affects mitochondria or can lead to effects on other organelles, which then send signals to the mitochondria to recruit participation in the death process (Fujikawa, 2015; Malhi et al., 2010). Constitutively expressed nitric oxide synthase (nNOS) is a Ca2+-dependent cytosolic enzyme that forms nitric oxide (NO) from L-arginine, and NO reacts with the free radical such as superoxide (O2−) to form the very toxic free radical peroxynitrite (ONOO−). Free radicals such as ONOO−, O2 − and hydroxyl radical (OH−) damage cellular membranes and intracellular proteins, enzymes and DNA (Fujikawa, 2015; Malhi et al., 2010; Kaplowitz, 2002; Kroemer et al., 2009).

Necrosis: Lactate dehydrogenase (LDH) is a soluble cytoplasmic enzyme that is present in almost all cells and is released into extracellular space when the plasma membrane is damaged. To detect the leakage of LDH into cell culture medium, a tetrazolium salt is used in this assay. In the first step, LDH produces reduced nicotinamide adenine dinucleotide (NADH) when it catalyzes the oxidation of lactate to pyruvate. In the second step, a tetrazolium salt is converted to a colored formazan product using newly synthesized NADH in the presence of an electron acceptor. The amount of formazan product can be colorimetrically quantified by standard spectroscopy. Because of the linearity of the assay, it can be used to enumerate the percentage of necrotic cells in a sample (Chan et al., 2013).  The MTT assay is a colorimetric assay for assessing cell viability. NAD(P)H-dependent cellular oxidoreductase enzymes may reflect the number of viable cells present. These enzymes are capable of reducing the tetrazolium dye MTT 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide to its insoluble formazan, which has a purple color. Other closely related tetrazolium dyes include XTT, MTS and the WSTs. Tetrazolium dye assays can also be used to measure cytotoxicity (loss of viable cells) or cytostatic activity (shift from proliferation to quiescence) of potential medicinal agents and toxic materials. MTT assays are usually done in the dark since the MTT reagent is sensitive to light (Berridgeet al.,2005). Propidium iodide (PI) is an intercalating agent and a fluorescent molecule used to stain necrotic cells. It is cell membrane impermeant so it stains only those cells where the cell membrane is destroyed. When PI is bound to nucleic acids, the fluorescence excitation maximum is 535 nm and the emission maximum is 617 nm (Moore et al.,1998) Alamar Blue (resazurin) is a fluorescent dye. The oxidized blue non fluorescent Alamar blue is reduced to a pink fluorescent dye in the medium by cell activity (O'Brien et al., 2000) (12). Neutral red uptake, which is based on the ability of viable cells to incorporate and bind the supravital dye neutral red in lysosomes (Repetto et al., 2008)(13). Moreover, quantification of ATP, signaling the presence of metabolically active cells, can be performed (CellTiter-Glo; Promega). ATP assay: Quantification of ATP, signaling the presence of metabolically active cells (CellTiter-Glo; Promega). Apoptosis: TUNEL is a common method for detecting DNA fragmentation that results from apoptotic signalling cascades. The assay relies on the presence of nicks in the DNA which can be identified by terminal deoxynucleotidyl transferase or TdT, an enzyme that will catalyze the addition of dUTPs that are secondarily labeled with a marker. It may also label cells that have suffered severe DNA damage. Caspase activity assays measured by fluorescence. During apoptosis, mainly caspase-3 and -7 cleave PARP to yield an 85 kDa and a 25 kDa fragment. PARP cleavage is considered to be one of the classical characteristics of apoptosis. Antibodies to the 85 kDa fragment of cleaved PARP or to caspase-3 both serve as markers for apoptotic cells that can be monitored using immunofluorescence (Li, Peng et al., 2004). Hoechst 33342 staining: Hoechst dyes are cell-permeable and bind to DNA in live or fixed cells. Therefore, these stains are often called supravital, which means that cells survive a treatment with these compounds. The stained, condensed or fragmented DNA is a marker of apoptosis (Loo, 2002; Kubbies and Rabinovitch, 1983).  Acridine Orange/Ethidium Bromide staining is used to visualize nuclear changes and apoptotic body formation that are characteristic of apoptosis. Cells are viewed under a fluorescence microscope and counted to quantify apoptosis.

Cell death is an universal event occurring in cells of any species (Fink and Cookson,2005).

CL:0000255

CL eukaryotic cell

Not Specified Unspecific

Not Specified

All life stages

High High High High

<ul> <li>Fujikawa, D.G. (2015), The role of excitotoxic programmed necrosis in acute brain injury, Comput Struct Biotechnol J, vol. 13, pp. 212-221.</li> <li>Malhi, H. et al. (2010), Hepatocyte death: a clear and present danger, Physiol Rev, vol. 90, no. 3, pp. 1165-1194.</li> <li>Kaplowitz, N. (2002), Biochemical and Cellular Mechanisms of Toxic Liver Injury, Semin Liver Dis, vol. 22, no. 2, <a class="external free" href="http://www.medscape.com/viewarticle/433631" rel="nofollow" target="_blank">http://www.medscape.com/viewarticle/433631</a> (accessed on 20 January 2016).</li> <li>Kroemer, G. et al., (2009), Classification of cell death: recommendations of the Nomenclature Committee on Cell Death, Cell Death Differ, vol. 16, no. 1, pp. 3-11.</li> <li>Chan, F.K., K. Moriwaki and M.J. De Rosa (2013), Detection of necrosis by release of lactate dehydrogenase (LDH) activity, Methods Mol Biol, vol. 979, pp. 65–70.</li> <li>Berridge, M.V., P.M. Herst and A.S. Tan (2005), Tetrazolium dyes as tools in cell biology: new insights into their cellular reduction. Biotechnology Annual Review, vol. 11, pp 127-152.</li> <li>Moore, A, et al.(1998), Simultaneous measurement of cell cycle and apoptotic cell death,Methods Cell Biol, vol. 57, pp. 265–278.</li> <li>Li, Peng et al. (2004), Mitochondrial activation of apoptosis, Cell, vol. 116, no. 2 Suppl,pp. S57-59, 2 p following S59.</li> <li>Loo, D.T. (2002), TUNEL Assay an overview of techniques, Methods in Molecular Biology, vol. 203: In Situ Detection of DNA Damage, chapter 2, Didenko VV (ed.), Humana Press Inc.</li> <li>Kubbies, M. and P.S. Rabinovitch (1983), Flow cytometric analysis of factors which influence the BrdUrd-Hoechst quenching effect in cultivated human fibroblasts and lymphocytes, Cytometry, vol. 3, no. 4, pp. 276–281.</li> <li>Fink, S.L. and B.T. Cookson (2005), Apoptosis, pyroptosis, and necrosis: mechanistic description of dead and dying eukaryotic cells, Infect Immun, vol. 73, no. 4, pp.1907-1916.</li> <li>O'Brien J, Wilson I, Orton T, Pognan F. 2000. Investigation of the Alamar Blue (resazurin) fluorescent dye for the assessment of mammalian cell cytotoxicity. European journal of biochemistry / FEBS 267(17): 5421-5426.</li> <li>Repetto G, del Peso A, Zurita JL. 2008. Neutral red uptake assay for the estimation of cell viability/cytotoxicity. Nature protocols 3(7): 1125-1131.</li> </ul> AOPWiki 2016-11-29T18:41:22

2022-07-15T09:46:25

Binding of inhibitor to mitochondrial complex III

Binding of inhibitor, mitochondrial complex III

Molecular

AOPWiki 2019-06-07T09:31:02

2019-06-07T09:31:02

Binding of inhibitor to mitochondrial complex IV

Binding of inhibitor, mitochondrial complex IV

Molecular

AOPWiki 2019-06-07T09:31:21

2019-06-07T09:31:21

Binding of inhibitor to mitochondrial complex V

Binding of inhibitor, mitochondrial complex V

Molecular

AOPWiki 2019-06-07T09:31:37

2019-06-07T09:31:37

N/A, Mitochondrial dysfunction 1

Cellular

Mitochondrial dysfunction is a consequence of inhibition of the respiratory chain leading to oxidative stress. Mitochondria can be found in all cells and are considered the most important cellular consumers of oxygen. Furthermore, mitochondria possess numerous redox enzymes capable of transferring single electrons to oxygen, generating the superoxide (O2-). Some mitochondrial enzymes that are involved in reactive oxygen species (ROS) generation include the electron-transport chain (ETC) complexes I, II and III; pyruvate dehydrogenase (PDH) and glycerol-3-phosphate dehydrogenase (GPDH). The transfer of electrons to oxygen, generating superoxide, happens mainly when these redox carriers are charged enough with electrons and the potential energy for transfer is elevated, like in the case of high mitochondrial membrane potential. In contrast, ROS generation is decreased if there are not enough electrons and the potential energy for the transfer is not sufficient (reviewed in Lin and Beal, 2006). Cells are also able to detoxify the generated ROS due to an extensive antioxidant defence system that includes superoxide dismutases, glutathione peroxidases, catalase, thioredoxins, and peroxiredoxins in various cell organelles (reviewed in Lin and Beal, 2006). It is worth mentioning that, as in the case of ROS generation, antioxidant defences are also closely related to the redox and energetic status of mitochondria. If mitochondria are structurally and functionally healthy, an antioxidant defence mechanism balances ROS generation, and there is not much available ROS production. However, in case of mitochondrial damage, the antioxidant defence capacity drops and ROS generation takes over. Once this happens, a vicious cycle starts and ROS can further damage mitochondria, leading to more free-radical generation and further loss of antioxidant capacity. During mitochondrial dysfunction the availability of ATP also decreases, which is considered necessary for repair mechanisms after ROS generation. A number of proteins bound to the mitochondria or endoplasmic reticulum (ER), especially in the mitochondria-associated ER membrane (MAM), are playing an important role of communicators between these two organelles (reviewed Mei et al., 2013). ER stress induces mitochondrial dysfunction through regulation of Ca2+ signaling and ROS production (reviewed Mei et al., 2013). Prolonged ER stress leads to release of Ca2+ at the MAM and increased Ca2+ uptake into the mitochondrial matrix, which induces Ca2+-dependent mitochondrial outer membrane permeabilization and apoptosis. At the same, ROS are produced by proteins in the ER oxidoreductin 1 (ERO1) family. ER stress activates ERO1 and leads to excessive production of ROS, which, in turn, inactivates SERCA and activates inositol-1,4,5- trisphosphate receptors (IP3R) via oxidation, resulting in elevated levels of cytosolic Ca2+, increased mitochondrial uptake of Ca2+, and ultimately mitochondrial dysfunction. Just as ER stress can lead to mitochondrial dysfunction, mitochondrial dysfunction also induces ER Stress (reviewed Mei et al., 2013). For example, nitric oxide disrupts the mitochondrial respiratory chain and causes changes in mitochondrial Ca2+ flux which induce ER stress. Increased Ca2+ flux triggers loss of mitochondrial membrane potential (MMP), opening of mitochondrial permeability transition pore (mPTP), release of cytochrome c and apoptosis inducing factor (AIF), decreasing ATP synthesis and rendering the cells more vulnerable to both apoptosis and necrosis (Wang and Qin, 2010). Summing up: Mitochondria play a pivotal role in cell survival and cell death because they are regulators of both energy metabolism and apoptotic/necrotic pathways (Fiskum, 2000; Wieloch, 2001; Friberg and Wieloch, 2002). The production of ATP via oxidative phosphorylation is a vital mitochondrial function (Kann and Kovács, 2007; Nunnari and Suomalainen, 2012). The ATP is continuously required for signalling processes (e.g. Ca2+ signalling), maintenance of ionic gradients across membranes, and biosynthetic processes (e.g. protein synthesis, heme synthesis or lipid and phospholipid metabolism) (Kang and Pervaiz, 2012), and (Green, 1998; McBride et al., 2006). Inhibition of mitochondrial respiration contributes to various cellular stress responses, such as deregulation of cellular Ca2+ homeostasis (Graier et al., 2007) and ROS production (Nunnari and Suomalainen, 2012; reviewed Mei et al., 2013).). It is well established in the existing literature that mitochondrial dysfunction may result in: (a) an increased ROS production and a decreased ATP level, (b) the loss of mitochondrial protein import and protein biosynthesis, (c) the reduced activities of enzymes of the mitochondrial respiratory chain and the Krebs cycle, (d) the loss of the mitochondrial membrane potential, (e) the loss of mitochondrial motility, causing a failure to re-localize to the sites with increased energy demands (f) the destruction of the mitochondrial network, and (g) increased mitochondrial Ca2+ uptake, causing Ca2+ overload (reviewed in Lin and Beal, 2006; Graier et al., 2007), (h) the rupture of the mitochondrial inner and outer membranes, leading to (i) the release of mitochondrial pro-death factors, including cytochrome c (Cyt. c), apoptosis-inducing factor, or endonuclease G (Braun, 2012; Martin, 2011; Correia et al., 2012; Cozzolino et al., 2013), which eventually leads to apoptotic, necrotic or autophagic cell death (Wang and Qin, 2010). Due to their structural and functional complexity, mitochondria present multiple targets for various compounds.

Mitochondrial dysfunction can be detected using isolated mitochondria, intact cells or cells in culture as well as in vivo studies. Such assessment can be performed with a large range of methods (revised by Brand and Nicholls, 2011) for which some important examples are given. All approaches to assess mitochondrial dysfunction fall into two main categories: the first assesses the consequences of a loss-of-function, i.e. impaired functioning of the respiratory chain and processes linked to it. Some assay to assess this have been described for KE1, with the limitation that they are not specific for complex I. In the context of overall mitochondrial dysfunction, the same assays provide useful information, when performed under slightly different assay conditions (e.g. without addition of complex III and IV inhibitors). The second approach assesses a ‘non-desirable gain-of-function’, i.e. processes that are usually only present to a very small degree in healthy cells, and that are triggered in a cell, in which mitochondria fail. I. Mitochondrial dysfunction assays assessing a loss-of function. 1. Cellular oxygen consumption. See KE1 for details of oxygen consumption assays. The oxygen consumption parameter can be combined with other endpoints to derive more specific information on the efficacy of mitochondrial function. One approach measures the ADP-to-O ratio (the number of ADP molecules phosphorylated per oxygen atom reduced (Hinkle, 1995 and Hafner et al., 1990). The related P/O ratio is calculated from the amount of ADP added, divided by the amount of O2 consumed while phosphorylating the added ADP (Ciapaite et al., 2005; Diepart et al., 2010; Hynes et al., 2006; James et al., 1995; von Heimburg et al., 2005). 2. Mitochondrial membrane potential (Δψm ). The mitochondrial membrane potential (Δψm) is the electric potential difference across the inner mitochondrial membrane. It requires a functioning respiratory chain in the absence of mechanisms that dissipate the proton gradient without coupling it to ATP production. The classical, and still most quantitative method uses a tetraphenylphosphonium ion (TPP+)-sensitive electrode on suspensions of isolated mitochondria. The Δψm can also be measured in live cells by fluorimetric methods. These are based on dyes which accumulate in mitochochondria because of Δψm. Frequently used are tetramethylrhodamineethylester (TMRE), tetramethylrhodaminemethyl ester (TMRM) (Petronilli et al., 1999) or 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazole carbocyanide iodide (JC-1). Mitochondria with intact membrane potential concentrate JC-1, so that it forms red fluorescent aggregates, whereas de-energized mitochondria cannot concentrate JC-1 and the dilute dye fluoresces green (Barrientos et al., 1999). Assays using TMRE or TMRM measure only at one wavelength (red fluorescence), and depending on the assay setup, de-energized mitochondria become either less fluorescent (loss of the dye) or more fluorescent (attenuated dye quenching). 3. Enzymatic activity of the electron transport system (ETS). Determination of ETS activity can be dene following Owens and King's assay (1975). The technique is based on a cell-free homogenate that is incubated with NADH to saturate the mitochondrial ETS and an artificial electron acceptor [l - (4 -iodophenyl) -3 - (4 -nitrophenyl) -5-phenylte trazolium chloride (INT)] to register the electron transmission rate. The oxygen consumption rate is calculated from the molar production rate of INT-formazan which is determined spectrophotometrically (Cammen et al., 1990). 4. ATP content. For the evaluation of ATP levels, various commercially-available ATP assay kits are offered  based on luciferin and luciferase activity. For isolated mitochondria various methods are available to continuously measure ATP with electrodes (Laudet 2005), with luminometric methods, or for obtaining more information on different nucleotide phosphate pools (e.g. Ciapaite et al., (2005). II. Mitochondrial dysfunction assays assessing a gain-of function. 1. Mitochondrial permeability transition pore opening (PTP). The opening of the PTP is associated with a permeabilization of mitochondrial membranes, so that different compounds and cellular constituents can change intracellular localization. This can be measured by assessment of the translocation of cytochrome c, adenylate kinase or AIF from mitochondria to the cytosol or nucleus. The translocation can be assessed biochemically in cell fractions, by imaging approaches in fixed cells or tissues or by life-cell imaging of GFP fusion proteins (Single 1998; Modjtahedi 2006). An alternative approach is to measure the accessibility of cobalt to the mitochondrial matrix in a calcein fluorescence quenching assay in live permeabilized cells (Petronilli et al., 1999). 2. mtDNA damage as a biomarker of mitochondrial dysfunction. Various quantitative polymerase chain reaction (QPCR)-based assays have been developed to detect changes of DNA structure and sequence in the mitochondrial genome. mtDNA damage can be detected in blood after low-level rotenone exposure, and the damage persists even after CI activity has returned to normal. With a more sustained rotenone exposure, mtDNA damage is also detected in skeletal muscle. These data support the idea that mtDNA damage in peripheral tissues in the rotenone model may provide a biomarker of past or ongoing mitochondrial toxin exposure (Sanders et al., 2014a and 2014b). 3. Generation of ROS and resultant oxidative stress. a. General approach. Electrons from the mitochondrial ETS may be transferred ‘erroneously’ to molecular oxygen to form superoxide anions. This type of side reaction can be strongly enhanced upon mitochondrial damage. As superoxide may form hydrogen peroxide, hydroxyl radicals or other reactive oxygen species, a large number of direct ROS assays and assays assessing the effects of ROS (indirect ROS assays) are available (Adam-Vizi, 2005; Fan and Li 2014). Direct assays are based on the chemical modification of fluorescent or luminescent reporters by ROS species. Indirect assays assess cellular metabolites, the concentration of which is changed in the presence of ROS (e.g. glutathione, malonaldehyde, isoprostanes,etc.) At the animal level the effects of oxidative stress are measured from biomarkers in the blood or urine. b. Measurement of the cellular glutathione (GSH) status. GSH is regenerated from its oxidized form (GSSH) by the action of an NADPH dependent reductase (GSSH + NADPH + H+ à 2 GSH + NADP+). The ratio of GSH/GSSG is therefore a good indicator for the cellular NADH+/NADPH ratio (i.e. the redox potential). GSH and GSSH levels can be determined by HPLC, capillary electrophoresis, or biochemically with DTNB (Ellman’s reagent). As excess GSSG is rapidly exported from most cells to maintain a constant GSH/GSSG ratio, a reduction of total glutathione (GSH/GSSG) is often a good surrogate measure for oxidative stress. c. Quantification of lipid peroxidation. Measurement of lipid peroxidation has historically relied on the detection of thiobarbituric acid (TBA)-reactive compounds such as malondialdehyde generated from the decomposition of cellular membrane lipid under oxidative stress (Pryor et al., 1976). This method is quite sensitive, but not highly specific. A number of commercial assay kits are available for this assay using absorbance or fluorescence detection technologies. The formation of F2-like prostanoid derivatives of arachidonic acid, termed F2-isoprostanes (IsoP) has been shown to be more specific for lipid peroxidation. A number of commercial ELISA kits have been developed for IsoPs, but interfering agents in samples requires partial purification before analysis. Alternatively, GC/MS may be used, as robust (specific) and sensitive method. d. Detection of superoxide production. Generation of superoxide by inhibition of complex I and the methods for its detection are described by Grivennikova and Vinogradov (2014). A range of different methods is also described by BioTek (<a class="external free" href="http://www.biotek.com/resources/articles/reactive-oxygen-species.html" rel="nofollow" target="_blank">http://www.biotek.com/resources/articles/reactive-oxygen-species.html</a>). The reduction of ferricytochrome c to ferrocytochrome c may be used to assess the rate of superoxide formation (McCord, 1968). Like in other superoxide assays, specificity can only be obtained by measurements in the absence and presence of superoxide dismutase. Chemiluminescent reactions have been used for their increased sensitivity. The most widely used chemiluminescent substrate is lucigenin. Coelenterazine has also been used as a chemiluminescent substrate. Hydrocyanine dyes are fluorogenic sensors for superoxide and hydroxyl radical, and they become membrane impermeable after oxidation (trapping at site of formation). The best characterized of these probes are Hydro-Cy3 and Hydro-Cy5. generation of superoxide in mitochondria can be visualized using fluorescence microscopy with MitoSOX™ Red reagent (Life Technologies). MitoSOX™ Red reagent is a cationic derivative of dihydroethidium that permeates live cells and accumulates in mitochondria. e. Detection of hydrogen peroxide (H2O2) production. There are a number of fluorogenic substrates, which serve as hydrogen donors that have been used in conjunction with horseradish peroxidase (HRP) enzyme to produce intensely fluorescent products in the presence of hydrogen peroxide (Zhou et al., 1997: Ruch et al., 1983). The more commonly used substrates include diacetyldichloro-fluorescein, homovanillic acid, and Amplex® Red. In these examples, increasing amounts of H2O2 form increasing amounts of fluorescent product (Tarpley et al., 2004). Summing up, mitochondrial dysfunction can be measured by: • ROS production: superoxide (O2-), and hydroxyl radicals (OH−) • Nitrosative radical formation such as ONOO− or directly by: • Loss of mitochondrial membrane potential (MMP) • Opening of mitochondrial permeability transition pores (mPTP) • ATP synthesis • Increase in mitochondrial Ca2+ • Cytochrome c release • AIF (apoptosis inducing factor) release from mitochondria • Mitochondrial Complexes enzyme activity • Measurements of mitochondrial oxygen consumption • Ultrastructure of mitochondria using electron microscope and mitochondrial fragmentation measured by labelling with DsRed-Mito expression (Knott et al, 2008) Mitochondrial dysfunction-induced oxidative stress can be measured by: • Reactive carbonyls formations (proteins oxidation) • Increased 8-oxo-dG immunoreactivity (DNA oxidation) • Lipid peroxidation (formation of malondialdehyde (MDA) and 4- hydroxynonenal (HNE) • 3-nitrotyrosine (3-NT) formation, marker of protein nitration • Translocation of Bid and Bax to mitochondria • Measurement of intracellular free calcium concentration ([Ca2+]i): Cells are loaded with 4 μM fura-2/AM). • Ratio between reduced and oxidized form of glutathione (GSH depletion) (Promega assay, TB369; Radkowsky et al., 1986) • Neuronal nitric oxide synthase (nNOS) activation that is Ca2+-dependent. All above measurements can be performed as the assays for each readout are well established in the existing literature (e.g. Bal-Price and Brown, 2000; Bal-Price et al., 2002; Fujikawa, 2015; Walker et al., 1995). See also KE <a href="/wiki/index.php/Event:209" title="Event:209"> Oxidative Stress, Increase</a> <table border="1" cellpadding="1" cellspacing="1"> <tbody> <tr> <td> Assay Type & Measured Content </td> <td>Description</td> <td>Dose Range Studied</td> <td> Assay Characteristics (Length/Ease of use/Accuracy) </td> </tr> <tr> <td> Rhodamine 123 Assay Measuring Mitochondrial membrane potential (MMP) and its collapse  (Shaki et al., 2012) </td> <td> 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. </td> <td>50, 100 and 500 μM of uranyl acetate</td> <td> Short / easy Medium accurancy </td> </tr> <tr> <td> TMRE fluorescence Assay Measuring Mitochondrial permeability transition pore (mPTP) opening (Huser et al., 1998) </td> <td>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.</td> <td>1 µM cyclosporin A</td> <td> Short / easy Low accurancy </td> </tr> <tr> <td> GSH / GSSG Determination Assay Measuring  cellular glutathione (GSH) status; ratio of GSH/GSSG (Owen & Butterfield, 2010; Shaki et al., 2013) </td> <td>GSH and GSSG levels are determinted biochemically with DTNB (Ellman’s reagent). The developed yellow color was read at 412 nm on a spectrophotometer.</td> <td>100 µM uranyl acetate</td> <td> Short / easy Low accurancy </td> </tr> <tr> <td> TBARS Assay Quantification of lipid peroxidation (Yuan et al., 2016) </td> <td>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.</td> <td>200, 400, 800 µM uranyl acetate</td> <td> Medium / medium High accurancy </td> </tr> <tr> <td> Aequorin-based bioluminescence assay Increase in mitochondrial Ca2+ influx (Pozzan & Rudolf, 2009) </td> <td>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.</td> <td> </td> <td> Short / easy Low accurancy </td> </tr> <tr> <td> Western blot & immunostaining analyses Measuring cytochrome c release (Chen et al., 2000)</td> <td>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)</td> <td> </td> <td> Short / easy Medium accurancy </td> </tr> <tr> <td> Quantikine Rat/Mouse Cytochrome c Immunoassay Measuring cytochrome c release (Shaki et al., 2012) </td> <td>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.</td> <td> </td> <td> Short / easy Low accurancy </td> </tr> <tr> <td> Membrane potential and cell viability – Flow Cytometry Measuring cytochrome c release (Kruidering et al., 1997) </td> <td>“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)”</td> <td> </td> <td> Short / easy Medium accurancy </td> </tr> </tbody> </table>

Mitochondrial dysfunction is a universal event occurring in cells of any species (Farooqui and Farooqui, 2012). Many invertebrate species (drosophila, C, elegans) are considered as potential models to study mitochondrial function. New 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 (Winklhofer and Haass, 2010; Waerzeggers et al., 2010) as well as in humans (Winklhofer and Haass, 2010).

CL:0000255

CL eukaryotic cell

Not Specified Unspecific

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All life stages

High High High

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Am J Physiol Cell Physiol 292:C641-576. Karlsson, H. L., Gustafsson, J., Cronholm, P., & Möller, L. (2009). Size-dependent toxicity of metal oxide particles—A comparison between nano- and micrometer size. Toxicology Letters, 188(2), 112-118. doi:<a href="https://doi.org/10.1016/j.toxlet.2009.03.014" target="_blank">10.1016/j.toxlet.2009.03.014</a> Knott Andrew B., Guy Perkins, Robert Schwarzenbacher & Ella Bossy-Wetzel. Mitochondrial fragmentation in neurodegeneration. Nature Reviews Neuroscience, 2008, 229: 505-518. Kruidering, M., Van De Water, B., De Heer, E., Mulder, G. J., & Nagelkerke, J. F. (1997). Cisplatin-induced nephrotoxicity in porcine proximal tubular cells: Mitochondrial dysfunction by inhibition of complexes I to IV of the respiratory chain. The Journal of Pharmacology and Experimental Therapeutics, 280(2), 638-649. Llaudet E, Hatz S, Droniou M, Dale N. Microelectrode biosensor for real-time measurement of ATP in biological tissue. Anal Chem. 2005, 77(10):3267-73. Lee HC, Wei YH. (2012). Mitochondria and aging. Adv Exp Med Biol 942:311-327. Li N, Ragheb K, Lawler G, Sturgis J, Rajwa B, et al. Mitochondrial complex I inhibitor rotenone induces apoptosis through enhancing mitochondrial reactive oxygen species production. J Biol Chem.2003;278:8516–8525. Lin MT, Beal MF. Mitochondrial dysfunction and oxidative stress in neurodegenerative diseases. Nature 2006. 443:787-795. Martin LJ. (2011). Mitochondrial pathobiology in ALS. J Bioenerg Biomembr 43:569 – 579. Martinez-Cruz, Oliviert Sanchez-Paz, Arturo Garcia-Carreño, Fernando Jimenez-Gutierrez, Laura Ma. de los Angeles Navarrete del Toro and Adriana Muhlia-Almazan. Invertebrates Mitochondrial Function and Energetic Challenges (www.intechopen.com), Bioenergetics, Edited by Dr Kevin Clark, <a class="internal mw-magiclink-isbn" href="/wiki/index.php/Special:BookSources/9789535100904">ISBN 978-953-51-0090-4</a>, Publisher InTech, 2012, 181-218. McBride HM, Neuspiel M, Wasiak S. (2006). Mitochondria: more than just a powerhouse. Curr Biol 16:R551–560. McCord, J.M. and I. Fidovich (1968) The Reduction of Cytochrome C by Milk Xanthine Oxidase. J. Biol. Chem. 243:5733-5760. Mei Y, Thompson MD, Cohen RA, Tong X. (2013) Endoplasmic Reticulum Stress and Related Pathological Processes. J Pharmacol Biomed Anal.. 1:100-107. Miccadei, S., & Floridi, A. (1993). Sites of inhibition of mitochondrial electron transport by cadmium. Elsevier Scientific Publishers Ireland Ltd., 89, 159-167.Xu, X. M., & Møller, S. G. (2010). ROS removal by DJ-1: Arabidopsis as a new model to understand Parkinson's Disease. Plant signaling & behavior, 5(8), 1034–1036. doi:10.4161/psb.5.8.12298 Modjtahedi N, Giordanetto F, Madeo F, Kroemer G. Apoptosis-inducing factor: vital and lethal. Trends Cell Biol. 2006 May;16(5):264-72. Nunnari J, Suomalainen A. (2012). Mitochondria: in sickness and in health. Cell 148:1145–1159. Hajnóczky G, Csordás G, Das S, Garcia-Perez C, Saotome M, Sinha Roy S, Yi M. (2006). Mitochondrial calcium signalling and cell death: approaches for assessing the role of mitochondrial Ca2+ uptake in apoptosis. Cell Calcium 40:553-560. Oliviert Martinez-Cruz, Arturo Sanchez-Paz, Fernando Garcia-Carreño, Laura Jimenez-Gutierrez, Ma. de los Angeles Navarrete del Toro and Adriana Muhlia-Almazan. Invertebrates Mitochondrial Function and Energetic Challenges (www.intechopen.com), Bioenergetics, Edited by Dr Kevin Clark, <a href="/wiki/index.php/Special:BookSources/9789535100904">ISBN 978-953-51-0090-4</a>, Publisher InTech, 2012, 181-218. Owen, J. B., & Butterfield, D. A. (2010). Measurement of oxidized/reduced glutathione ratio. Methods in Molecular Biology (Clifton, N.J.), 648, 269-277. doi:10.1007/978-1-60761-756-3_18 [doi] Owens R.G. and King F.D. The measurement of respiratory lectron-transport system activity in marine zooplankton. Mar. Biol. 1975, 30:27-36. Pan, Y., Leifer, A., Ruau, D., Neuss, S., Bonrnemann, J., Schmid, G., . . . Jahnen-Dechent, W. (2009). Gold nanoparticles of diameter 1.4 nm trigger necrosis by oxidative stress and mitochondrial damage. Small, 5(8), 2067-2076. doi:10.1002/smll.200900466 Petronilli V, Miotto G, Canton M, Brini M, Colonna R, Bernardi P, Di Lisa F: Transient and long-lasting openings of the mitochondrial permeability transition pore can be monitored directly in intact cells by changes in mitochondrial calcein fluorescence. Biophys J 1999, 76:725-734. Pozzan, T., & Rudolf, R. (2009). Measurements of mitochondrial calcium in vivo. Biochimica Et Biophysica Acta (BBA) - Bioenergetics, 1787(11), 1317-1323. doi:<a href="https://doi.org/10.1016/j.bbabio.2008.11.012" target="_blank">https://doi.org/10.1016/j.bbabio.2008.11.012</a> Promega GSH-Glo Glutathione Assay Technical Bulletin, TB369, Promega Corporation, Madison, WI. Pryor, W.A., J.P. Stanley, and E. Blair. (1976) Autoxidation of polyunsaturated fatty acids: II. 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Nano copper induces apoptosis in PK-15 cells via a mitochondria-mediated pathway. Biological Trace Element Research, 181(1), 62-70. doi:10.1007/s12011-017-1024-0 Zhou, M., Z.Diwu, Panchuk-Voloshina, N. and R.P. Haughland (1997), A Stable nonfluorescent derivative of resorufin for the fluorometric determination of trace hydrogen peroxide: application in detecting the activity of phagocyte NADPH oxidase and other oxidases. Anal. Biochem 253:162-168. AOPWiki 2016-11-29T18:41:23

2022-03-07T07:12:30

<upstream-id>f85263db-9bde-4c2f-91d3-e2447439a6a1</upstream-id> <downstream-id>10998181-3576-4b5b-8bcb-548ebf9dff23</downstream-id> The initiation event is the reversibly or irreversibly interaction to any of the subunits in the mitochondrial complex I, III or IV. Inhibition of complex I, III or IV leads to a reduction in proton gradient and therefore impair or completely prevent the production of mitochondrial membrane dependent ATP production. Threshold relationship = The cell can cope with inhibition of the various mitochondrial complexes and the following decreased OXPHOS depending on the number of mitochondria affected. The more mitochondria within a cell are perturbed, the lower the concentration of available ATP will become. The cell can influence the threshold at which these low ATP levels will become lethal, for example by upregulating the number of available mitochondria (biogenesis).

It is broadly accepted that complex inhibition leads to decreased mitochondrial oxidative phosphorylation. Deficiencies in complex I in patient leads to decreased activity of the complex

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430ae0ee80> AOPWiki 2018-12-19T09:41:24

2019-03-07T08:37:17

<upstream-id>a8e1ed8c-fea7-4f3f-ba6c-c0013875bbb6</upstream-id> <downstream-id>10998181-3576-4b5b-8bcb-548ebf9dff23</downstream-id> The initiation event is the reversibly or irreversibly interaction to any of the subunits in the mitochondrial complex I, III or IV. Inhibition of complex I, III or IV leads to a reduction in proton gradient and therefore impair or completely prevent the production of mitochondrial membrane dependent ATP production. Threshold relationship = The cell can cope with inhibition of the various mitochondrial complexes and the following decreased OXPHOS depending on the number of mitochondria affected. The more mitochondria within a cell are perturbed, the lower the concentration of available ATP will become. The cell can influence the threshold at which these low ATP levels will become lethal, for example by upregulating the number of available mitochondria (biogenesis).

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430ae31688> AOPWiki 2018-12-19T09:41:36

2019-03-07T08:38:03

<upstream-id>b60e8426-2921-4e24-82ae-06929d67d457</upstream-id> <downstream-id>10998181-3576-4b5b-8bcb-548ebf9dff23</downstream-id> The initiation event is the reversibly or irreversibly interaction to any of the subunits in the mitochondrial complex V. Inhibition of complex IV directly prevents the production of ATP in the mitochondria , because the catalyzing element isn’t functional anymore. Threshold relationship = The cell can cope with inhibition of the various mitochondrial complexes and the following decreased OXPHOS depending on the number of mitochondria affected. The more mitochondria within a cell are perturbed, the lower the concentration of available ATP will become. The cell can influence the threshold at which these low ATP levels will become lethal, for example by upregulating the number of available mitochondria (biogenesis).

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430ae78740> AOPWiki 2018-12-19T09:41:48

2019-03-07T08:38:54

<upstream-id>990f1b78-619a-4c89-800b-e785f12290b3</upstream-id> <downstream-id>ff2781e4-7276-4600-8093-1011e3cd310b</downstream-id> Logically when liver tissue is degraded, the liver will fail to execute its function. Again, this can be tissue type dependent(Krishna 2017)

Necrotic tissue is frequently observed in liver disease, e.g. a failing liver. Reviews: (Guicciardi et al. 2013): Apoptosis and necrosis in the liver. (Luedde et al. 2014): Cell death in liver disease. (Krishna 2017): Patterns of Necrosis in Liver disease. (Malhi et al. 2006): Apoptosis and necrosis in the liver (including TNF signaling (NFkB and Caspase8) part)

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430aebe3f8> AOPWiki 2018-12-19T09:45:26

2019-03-07T06:07:46

<upstream-id>62efeca6-1197-47a7-b325-c5b962eeee68</upstream-id> <downstream-id>0e9327f7-7d9f-4be1-8be2-1d766cbdae6f</downstream-id> It is well documented that binding of an inhibitor to CI inhibits its activity (see MIE). Naturally occurring and synthetic CI inhibitors have been shown to inhibit the catalytic activity of CI, leading to partial or total inhibition of its activity in a dose response manner (Degli Esposti and Ghelli, 1994; Ichimaru et al. 2008; Barrientos and Moraes, 1999; Betarbet et al., 2000). Indeed, binding of inhibitors stops the electron flow from CI to ubiquinone. Therefore, the Fe-S clusters of CI become highly reduced and no further electrons can be transferred from NADH to CI. This leads to the inhibition of the NADH oxido-reductase function, i.e. CI inhibition.

The weight of evidence supporting the relationship between binding of an inhibitor to NADH-ubiquinone oxidoreducatse and its inhibition is strong. Biological Plausibility There is an extensive understanding of the functional relationship between binding of an inhibitor to NADH-ubiquinone oxidoreductase (CI) and its inhibition. As the first entry complex of mitochondrial respiratory chain, CI oxidizes NADH and transfers electrons via a flavin mononucleotide cofactor and several Fe-S complexes to ubiquinone. The electron flow is coupled to the translocation of protons from the matrix to the intermembrane space. This helps to establish the electrochemical gradient that is used to fuel ATP synthesis (Greenamyre et al., 2001). If an inhibitor binds to CI, the electron transfer is blocked. This compromises ATP synthesis and maintenance of Δψm, leading to mitochondrial dysfunction. As CI exerts a higher control over oxidative phosphorylation in synaptic mitochondria than in non-synaptic mitochondria in the brain (Davey and Clark, 1996), specific functional defects observed in PD may be explained. It is well documented that CI inhibition is one of the main sites at which electron leakage to oxygen occurs. This results in a production of ROS, such as superoxide (Efremov and Sazanow, 2011) and hydrogen peroxide, which are main contributors to oxidative stress (Greenamyre et al., 2001).

A variety of studies show a significant correlation between binding of an inhibitor to CI and its inhibition, usually measured by the decreased mitochondrial respiration. Different classes of CI inhibitors, such as rotenone, MPP+, piericidin A, acetogenins, pyridaben, and various piperazin derivatives (Ichimaru et al. 2008) have been shown to bind to the ubiquitin site of CI, leading to a partial or total inhibition of oxidoreductase activity in a dose response manner (Grivennikova et al., 1997; Barrientos and Moraes, 1999; Betarbet et al., 2000). The reduction of CI activity is well documented in a variety of studies using isolated mitochondria or cells, as well as in in vivo experiments and in human post mortem PD brains. Usually it is measured by assays described in 2nd Key Event Relationship (KER): Inhibition of complex I leads to mitochondrial dysfunction. It has been shown that binding of rotenone to CI (e.g. Betarbet et al., 2000, Greenamyre et al., 2001) or MPP+ (e.g. Krug et al., 2014; Langston, 1996) can reproduce the anatomical, neurochemical, behavioural and neuropathological features of PD. Therefore, the empirical support for this KER will be mainly based on the experiments performed after exposure to rotenone or MPP+. <ul> <li>The binding of rotenone to CI resulted in time- and dose-dependent inhibition of CI activity measured in sub-mitochondrial particles. The kinetics of the active CI inhibition was determined after exposure to rotenone at 20, 30 and 40 nM at different times of exposure (30 sec, 1 min or 2 min) (Grivennikova et al., 1997). This study suggests that two rotenone binding sites exist in CI: one affecting NADH oxidation by ubiquinone and the other one operating in ubiquinol-NAD+ reductase action.</li> </ul> <ul> <li>Partial inhibition of CI produces a mild, late-onset mitochondrial damage. The threshold effect seen in brain mitochondria (25–50% decrease in activity) may not directly impact ATP levels or Δψm but could have long-term deleterious effects triggered by oxidative stress, as it has been shown that an electron leak upstream of the rotenone binding site in CI leads to ROS production (Greenamyre et al., 2001).</li> </ul> <ul> <li>Exposure of rats to rotenone (2 days, 2 mg/kg) produced free brain rotenone concentration of 20–30 nM and resulted in 73% inhibition of specific binding to CI of [3H] dihydrorotenone (Betarbet et al., 2000). However, oximetry analysis indicated that in brain mitochondria (but not liver mitochondria) this rotenone concentration (30 nM maximum) was insufficient to inhibit glutamate (CI substrate)-supported respiration (Betarbet et al., 2000) suggesting that this rotenone concentration did not alter mitochondrial oxygen consumption in isolated brain mitochondria.</li> </ul> <ul> <li>Rotenone has been reported to be a specific and potent mitochondrial CI inhibitor with IC50 values from 0.1 nM to 100 nM depending on the system and methods used (Lambert and Brand, 2004; Ichimaru et al., 2008; Chinopoulos and Vizi, 2001; Beretta et al., 2006).</li> <li>Mesencephalic cultures prepared from C57/BL6 mice and treated with 5, or 10 nM rotenone for 24 h inhibited CI activity by 11% or 33%, respectively (Choi et al., 2008).</li> </ul> <ul> <li>The inhibition of CI was studied in the human osteosarcoma-derived cell line (143B) after the exposure to rotenone or using a genetic model (40% loss of CI activity in human xenomitochondrial cybrids (HXC) lines). Different degrees of CI inhibition were quantitatively correlated with levels of decreased cellular respiration (Barrientos and Moraes, 1999). Only when CI was inhibited by 35-40% (< 5 nM rotenone), cell respiration decreased linearly until 30% of the normal rate. Increasing concentrations of rotenone produced further but slower decrease in CI activity and cell respiration (Fig. 1). Cells with the complete rotenone-induced CI inhibition still maintain a cell respiration rate of approximately 20% because of an electron flow through complex II. At high concentrations (5–6-fold higher than the concentration necessary for 100% CI inhibition), rotenone showed a secondary, toxic effect at the level of microtubule assembly (Barrientos and Moraes 1999).</li> </ul> <ul> <li>Bovine sub-mitochondrial particles were used to test rotenone affinity binding at 20 nM. This concentration of rotenone reduced the NADH oxidation rate by approximately 50% (Okun et al., 1999.</li> </ul> <ul> <li>MPP+ (an active metabolite of MPTP) is an inhibitor of CI (Nicklas et al., 1987; Mizuno et al, 1989; Sayre et al., 1986). Inhibition of the mitochondrial CI by MPP+ supresses aerobic glycolysis and ATP production (Book chapter in Cheville 1994).</li> </ul> <ul> <li>MPP+ binds loosely to CI and causes reversible inhibition of its activity: approximately 40% inhibition was observed at 10 mM concentration within 15 min of incubation. However, prolonged incubation (> 15min) produces up to 78% of irreversible inhibition of CI (Cleeter et al., 1992).</li> <li> MPP+, interrupts mitochondrial electron transfer at the NADH dehydrogenase-ubiquinone junction, as do the respiratory chain inhibitors rotenone, piericidin A and barbiturates. The 4'-alkyl derivatives of MPP+ inhibit NADH oxidation in submitochondrial particles at much lower concentrations than does MPP+ itself. The MPP+ analogues with short alkyl chains prevent the binding of 14C-labelled piericidin A to the membrane and thus must act at the same site, but analogues with alkyl chains longer than heptyl do not prevent binding of [14C]piericidin (Ramsay et al., 1991). </li> <li> 13 complex I inhibitors  were found to decrease labelling at the PSST site without effect on ND1 labelling.  The results suggest that the common action of MPP(+) and stigmatellin on the functional coupling of the PSST and ND1 subunits is initiated by binding at a semiquinone binding site in complex I (Schuler and Casida, 1991). </li> </ul> Human studies <ul> <li>There are many studies that show impaired catalytic activity of CI in multiple PD post-mortem brain tissues. For example (Parker and Swerdlow, 1998), five PD brains were used to measure activities of complexes I, III, IV, and of complexes I/III together (NADH: cytochrome c reductase). These measurements were performed in purified frontal cortex mitochondria and revealed a significant loss of CI activity in these PD samples as compared to controls.</li> </ul> <ul> <li>Human data indicate that impairment of CI activity may contribute to the pathogenic processes of PD (for example, Greenamyre et al., 2001; Schapira et al., 1989; Shults, 2004).</li> </ul>

It is not clear the number of subunits constituting CI in mammals, as according to the existing literature different numbers are cited (between 41-46) (Vogel et al., 2007a; Hassinen, 2007). The majority of data claims that mammalian CI is composed of 46 (Greenamyre et al., 2001; Hassinen, 2007) or 45 subunits (Vogel et al., 2007a). It is not sure whether there may exist tissue-specific subunits of CI isoforms (Fearnley et al., 2001). It is unclear, which subunit(s) bind rotenone or other inhibitors of CI. Additionally, it is not clear whether CI has other uncharacterized functions, taking into consideration its size and complexity (43-46 subunits vs. 11 subunits of complex III or 13 subunits of complex IV) (Greenamyre et al., 2001). There is no strict linear relationship between inhibitor binding and reduced mitochondrial function. Low doses of rotenone that inhibit CI activity partially do not alter mitochondrial oxygen consumption. Therefore, bioenergetic defects can not account alone for rotenone-induced neurodegeneration. Instead, under such conditions, rotenone neurotoxicity may result from oxidative stress (Betarbet et al., 2000). Few studies used human brain cells/human brain mitochondria. Therefore, full quantitative data for humans are not available.

The kinetics of binding and CI inhibition by rotenone has been quantitatively evaluated in a dose-dependent manner using the sub-mitochondrial particles (Grivennikova et al., 1997). The consequences of CI inhibition were quantitatively measured by a variety of assays that are used to study mitochondrial dysfunction (see Key Event Relationship (KER): Inhibition of Complex I leads to mitochondrial dysfunction). There are also many in vitro and in vivo studies combining the quantification of CI inhibition and DA cell death (e.g. Choi et al., 2008, Betarbet et al., 2000, see KER Mitochondrial dysfunction induces degeneration of nigrostriatal pathway). The binding of different classes of inhibitors (e.g., pesticides, drugs and other toxins) to CI has been determined quantitatively and I50, and KI values are available. Potency relative to that of rotenone has been determined under the same conditions in beef mitochondria or submitochondrial particles using the ratio of the KI values, when they were available (Degli Esposti, 1998; Okun et al., 1999). Rotenone I50 value is defined as 20 nM (Okun et al., 1999). Example of a quantitative evaluation of concentration-dependent CI inhibition by rotenone (Fig. 1 from Barrientos and Moraes, 1999). <a class="image" href="/wiki/index.php/File:KER_1Fig._1.jpg"><img alt="KER 1Fig. 1.jpg" src="/wiki/images/d/da/KER_1Fig._1.jpg" style="height:280px; width:329px" /></a> Fig. 1. Fig.1. Effect of CI (NADH decylubiquinone reductase) inhibition on endogenous cell respiration. Cells were treated with different concentrations of rotenone for 4 h before measuring cell respiration in whole cells and CI activity in isolated mitochondria. Complete CI inhibition was achieved with 100 nM rotenone. The cell respiration was inhibited also in a dose-dependent manner but showed different inhibition kinetics and a saturation threshold. For comparison, the genetically-altered cell line HXC had an approximately 40% CI reduced activity and an approximately 80% residual cell respiration. HXC, human xenomitochondrial cybrids.   Time- and concentration-relationship of NADH oxidase inhibition by rotenone (Fig. 2. from Grivennikova et al., 1997). <a class="image" href="/wiki/index.php/File:KER1_Fig._2.jpg"><img alt="KER1 Fig. 2.jpg" src="/wiki/images/1/15/KER1_Fig._2.jpg" style="height:376px; width:461px" /></a> Fig. 2. Panel A and B: Time- and concentration-relationship of NADH oxidase inhibition by rotenone. The numbers on the curves indicate the final concentrations of rotenone (0, 20, 30, 40, 1000 nM). In Panel B: vo, zero-order rate of NADH oxidation in the absence of rotenone; vt, the `instant' values of the rates approximated within 10 s time intervals. Panel C: The dependence of first-order inhibition rate constant on the concentration of rotenone (for further description see Fig. 1 in Grivennikova et al., 1997).   Quantitative evaluation of the 1st KER: Binding of inhibitor to NADH-ubiquinone oxidoreductase (MIE; KE upstream) leads to its inhibition (KE downstream).   <table border="1"> <tbody> <tr> <td> MIE (KE upstream) Binding of inhibitor to NADH-ubiquinone oxidoreductase (nM) </td> <td> KE (downstream) Inhibition of CI (%, approximately) </td> <td> Comments (in vivo, in vitro or human studies) </td> <td> References </td> </tr> <tr> <td> Administration of rotenone at 2 mg/kg per day for 2 days resulted in free rotenone concentration of 20–30 nM in the brain. </td> <td> 75%   </td> <td> DA neuronal cell death determined after rotenone administration at 1 to 12 mg/kg per day, Sprague Dawley and Lewis rats infused continuously by jugular vein, 7days up to 5 weeks </td> <td> Betarbet et al., 2000   </td> </tr> <tr> <td> 20 nM rotenone Direct binding studies using bovine and Musca domestica sub-mitochondrial particles   </td> <td> 50% </td> <td> Binding studies that defined the I50 and Kd values for three classes of CI inhibitors (12 chemicals) including rotenone. </td> <td> Okun et al., 1999 </td> </tr> <tr> <td> Human skin fibroblasts exposed to 100 nM Rotenone for 72 hr </td> <td> 20% </td> <td> In the same experiment mitochondria morphology, motility was also evaluated. </td> <td> Koopman et al., 2007 </td> </tr> <tr> <td> 0-2.5 nM Rotenone 5/10 nM Rotenone Mesencephalic neurons were cultured from E14 C57/BL6 mouse embryos for 6 days and then treated with rotenone for 24 hr </td> <td> No effect 11% and33%, respectively </td> <td> Treatments with 5 or 10 nM rotenone killed 50% or 75% DA neurons respectively. </td> <td> Choi et al., 2008 </td> </tr> <tr> <td> 1-2.5-5-7.5-10-20 nM   1-10-20-80 nM     </td> <td> 10-20-35-50- 65-80 % 5- 75 %       </td> <td> In this study time course of the active and deactivated enzymes inhibition by rotenone and Piericidin A is study in a dose-dependent manner. Binding studies in sub-mitochondrial particles prepared from bovine heart after 20 min of exposure to rotenone. </td> <td> Grivennikova et al., 1997 </td> </tr> <tr> <td> 5-10 nM 20 nM 40 nM 100 nM 143B Cells (human osteo-sarcoma), exposed for 4 hrs to rotenone </td> <td> 55-78 % 80% 87% 100% </td> <td> In the same study similar experiments were performed using HXC cell line (see Fig. 1 above). </td> <td> Barrientos and Moraes 1999 </td> </tr> </tbody> </table>

The CI is well-conserved across species from lower organism to mammals. The central subunits of CI harboring the bioenergetic core functions are conserved from bacteria to humans. CI from bacteria and from mitochondria of Yarrowia lipolytica, a yeast genetic model for the study of eukaryotic CI (Kerscher et al., 2002) was analyzed by x-ray crystallography (Zickermann et al., 2015). However, the affinity of various chemicals to cause partial or total inhibition of CI activity across species is not well studied (except rotenone).

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430aef11e0> AOPWiki 2016-11-29T18:41:35

2017-08-25T09:35:19

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#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430af05af0> AOPWiki 2019-06-07T09:12:43

2019-06-07T09:12:43

<upstream-id>4f36d988-7052-420e-900c-79e907cdcfef</upstream-id> <downstream-id>d3784744-614a-4f81-a1a1-2ab9224b7522</downstream-id>

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430af265e8> AOPWiki 2019-06-07T09:28:14

2019-06-07T09:28:14

<upstream-id>d3784744-614a-4f81-a1a1-2ab9224b7522</upstream-id> <downstream-id>990f1b78-619a-4c89-800b-e785f12290b3</downstream-id> Cell death can help the maintenance of the organ, for example by avoiding cancer. However, in the KER, cell death will lead to necrotic tissue. If too many cells die, the tissue will fall apart. There are several types of necrotic tissue caused by cell death: Apoptosis (individual cell necrosis), spotty and focal necrosis, zonal necrosis confluent necrosis. Every type can have a different underlying disease. All types can occur by drug induced liver injury.(Krishna 2017).

It has been well established that cell death can lead to necrotic tissue. Reviews: (Guicciardi et al. 2013): Apoptosis and necrosis in the liver. (Luedde et al. 2014): Cell death in liver disease.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430af3aa20> AOPWiki 2019-01-07T09:12:27

2019-03-07T06:05:19

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#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430af51158> AOPWiki 2019-06-07T09:31:51

2019-06-07T09:31:51

<upstream-id>67f69bb7-f7be-42b5-a405-f7914052f76d</upstream-id> <downstream-id>a8e1ed8c-fea7-4f3f-ba6c-c0013875bbb6</downstream-id>

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430af72178> AOPWiki 2019-06-07T09:32:09

2019-06-07T09:32:09

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#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430af8e080> AOPWiki 2019-06-07T09:32:21

2019-06-07T09:32:21

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#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430afad688> AOPWiki 2019-06-15T11:06:56

2019-06-15T11:06:56

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#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430afd7280> AOPWiki 2019-06-15T11:07:58

2019-06-15T11:07:58

<upstream-id>ce0914e2-9c91-42dd-9830-32c380a8c53d</upstream-id> <downstream-id>e6f63097-04ff-4475-a589-8c9b046925e9</downstream-id>

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430aff11d0> AOPWiki 2019-06-15T11:08:33

2019-06-15T11:08:33

<upstream-id>ce0914e2-9c91-42dd-9830-32c380a8c53d</upstream-id> <downstream-id>4f36d988-7052-420e-900c-79e907cdcfef</downstream-id> In any cell type, including neurons, the protein homeostasis (proteostasis) plays a key role in cellular functions. There are two major systems involved in the removal of damaged cellular structures (e.g. defective mitochondria) and misfolded or damaged proteins, the ubiquitin-proteasome system (UPS) and the autophagy–lysosome pathway (ALP). These processes are highly energy demanding and highly susceptible to oxidative stress. Upon mitochondrial dysfunction UPS and ALP functions are compromised resulting in increased protein aggregation and impaired intracellular protein/organelles transport (e.g. Zaltieri et al., 2015; Song and Cortopassi, 2015; Fujita et al., 2014; Esteves et al., 2011; Sherer et al., 2002).

The weight of evidence supporting the relationship between mitochondrial dysfunction and impaired proteostasis, including the impaired function of UPS and ALP that results in decreased protein degradation and increase protein aggregation is strong.

The biological relationship between Mitochondrial dysfunction and Impaired proteostasis (unbalanced protein homeostasis) that involves dysregulation of proteins degradation (misfolded or damaged) as well as removal of cell organelles is partly understood. Under physiological conditions, mechanisms by which proteostasis is ensured include regulated protein translation, chaperone assisted protein folding and functional protein degradation pathways. Under oxidative stress, the proteostasis function becomes burdened with proteins modified by ROS (Powers et al., 2009; Zaltieri et al., 2015). These changed proteins can lead to further misfolding and aggregation of proteins (especially in non-dividing cells, like neurons). Particularly in DA cells, oxidative stress from dopamine metabolism and dopamine auto-oxidation may selectively increase their vulnerability to CI inhibitors (such as rotenone) and cause additional deregulation of protein degradation (Lotharius and Brundin, 2002; Esteves et al., 2011). As most oxidized proteins get degraded by UPS and ALP (McNaught and Jenner, 2001), mitochondrial dysfunction and subsequent deregulation of proteostasis play a pivotal role in the pathogenesis of PD (Dagda et al., 2013; Pan et al., 2008; Fornai et al., 2005; Sherer et al., 2002). It is also well documented that increased oxidative stress changes the protein degradation machinery and leads to a reduction of proteasome activity (Lin and Beal, 2006; Schapira, 2006).

Based on the existing in vitro and in vivo data it is suggested that mitochondrial dysfunction impairs protein homeostasis through oxidative and nitrosative stress resulting in protein aggregation, disruption of microtubule assembly and damaged intracellular transport of proteins and cell organelles. Mitochondrial dysfunction by rotenone or MPP+ reduces UPS activity: <ul> <li>Mitochondrial dysfunction induced by systemic and chronic CI inhibition by rotenone, results in a selective inhibition of proteasomal function in the midbrain (not in cortical or striatal homogenates) of rats that had lost the TH-positive terminals in the striatum. Initially, proteasomal activity showed an acute increase prior to a decrease by 16-31 %, during chronic rotenone exposure (3.0 mg/kg/day, through subcutaneous osmotic pump during 5 weeks). In the same animals a significant and selective increase in ubiquitinated proteins was observed (~ 25%) in the ventral midbrain of lesioned rats, indicating an increase in the proteins levels that have been marked for degradation by UPS. These results were confirmed immunocyto-chemically, pointing out that ubiquitin levels were elevated selectively in DA neurons present in SNpc (Betarbet et al., 2006).</li> </ul> <ul> <li>Nigral neurons in chronically rotenone-treated rats (up to 5 weeks, continuous intrajugular infusion of rotenone at 2.5 mg/kg/day) accumulate fibrillar cytoplasmic inclusions that contain ubiquitin and α-synuclein (the main protein of Lewy bodies observed in PD) (qualitative data, obtained by immuno-electron microscopy) (Betarbet et al., 2000).</li> <li>Inhibition of proteasomal function was also observed in in vitro systems using SK-N-MC human neuroblastoma. Exposure to 5 nM rotenone, for up to 4 weeks caused 60% increase in the levels of ubiquinated proteins, suggesting that chronic exposure to rotenone increased the level of misfolded or oxidized proteins targeted for degradation by UPS (Betarbet et al., 2006).</li> </ul> <ul> <li>To determine whether rotenone-induced proteasomal inhibition was due to CI inhibition or direct effects of rotenone on the UPS, proteasomal activity was determined in SKN-MC cells expressing the rotenone-insensitive single-subunit NADH dehydrogenase of Saccharomyces cerevisiae (NDI1), which acts as a "replacement" for the entire CI in mammalian cells (Bai et al., 2001; Seo et al., 2000, 2002). The obtained results confirmed that rotenone-induced proteasomal dysfunction is due to CI inhibition and not to direct effects of rotenone on proteasomal function (Betarbet et al., 2006). In the same study the decreased proteasomal activity and an accumulation of ubiquitinated proteins was completely prevented by continuous treatment with α-tocopherol (62.5 μM added 1 week prior to and continuously thereafter along with 5 nM rotenone) (qualitative data), confirming that oxidative damage played a major role in rotenone-induced proteasomal dysfunction rather than bioenergetic defects. Indeed, chronic, low levels of rotenone exposure did not changed significantly ATP levels (111.5 ± 1.5% of control), but produced ROS (not shown in this study). Similar results were published by Shamoto-Nagai's group (Shamoto-Nagai et al. 2003).</li> </ul> <ul> <li>Rotenone significantly lowered UPS activity in a concentration dependent manner in HEK (human embryonic kidney cells) and SK-N-MC human neuroblatoma cells even after 24 h exposure to doses as low as 10 nM. It caused a reduction in the 20S proteasome activity (by 5-25%) and of the 20S proteasome subunit (by 20-60%) (as shown by increase of GFP-U fluorescence) (Chou et al., 2010). Similar results were obtained using other pesticides that inhibit CI, including pyridaben and fenazaquin (Wang et al., 2006). This effect was mediated by oxidative stress as anti-oxidants, such as butylated-hydroxy toluene (BHT), and catalase attenuated rotenone-induced UPS inhibition. Additionally, nitric oxide (NO) and peroxinitrite contributed to this effect as well, since neuronal nitric oxide synthase (nNOS) inhibitor (LNMMA) attenuated rotenone-induced proteasome inhibition by 20% (Chou et al., 2010) indicating that both oxidative and nitrative stress can directly inhibit the proteasome activity through increased degradation of proteasome subunits. The same mechanisms of proteasome inhibition were suggested by many other studies (e.g. Szweda et al, 2002; Osna et al., 2004; Shamoto-Nagai et al., 2003).</li> </ul> <ul> <li>CI inhibition-induced proteasomal dysfunction has been reported in ventral mesencephalic cultures following acute rotenone or MPP+ exposure (Hoglinger et al., 2003). In DA neurones derived from rat (embryonic day 15.5) ventral mesencephalon, it has been showen that proteasome inhibition (by 100 nm epoxomicin) exacerbated the neurotoxicity of CI inhibitors (by mean of rotenone 30 nM, or MPP+ 3 µM, for 24 hr). All three proteasomal peptidase activities (i.e., chymotrypsin (CT)-like, trypsin (T)-like, and peptidylglutamyl-peptide hydrolase (PGPH) activity) significantly decreased in cultures upon 6 hr treatment with 30 nM rotenone (by 50+-60%) or 30 µM MPP+ (by 25-30%) (Hoglinger et al., 2003).</li> </ul> <ul> <li>CI inhibition-induced proteasomal dysfunction has been reported in human SH-SY5Y neuroblastoma cells following acute rotenone exposure (Shamoto-Nagai et al., 2003). After 96 hr of incubation with 25 or 50 nM rotenone, the activity was reduced respectively to 28.7% and 21.9% of control, and adding ATP did not increase the activity. After 120 hr, the activity was virtually undetectable (with or without added ATP). On the contrary, the levels of the proteins composing proteasome did not change with rotenone treatment (Shamoto-Nagai et al., 2003).</li> </ul> <ul> <li>The ability of rotenone to cause proteasome inhibition via disruption of microtubules (MT) assembly has been also documented. In human embryonic kidney (HEK) and neuroblastoma SK-N-MC cells rotenone (10-100-100 nM, 24 hr) was found to inhibit 26S UPS activity (by 25%, at 10 nM) (Chou et al., 2010). Rotenone was found to interfere with MT assembly at concentrations as low as 10 nM, providing evidence that there could be additional mechanisms implicated in the rotenone induced UPS inhibition, possibly mediated by nitric oxide (NO). In the same study, nocodazole, a MT disrupter (positive control), strongly inhibited the UPS activity (e.g., 10 µM nocodazole caused ~80% decrease of 26S UPS activity) (Chou et al., 2010).</li> </ul> • Oxidative stress triggered by the MPP+ inhibited CI (1 mM, for 2-6-24 hr) led to a decrease in proteolytic activity, as shown in NT2 human teratocarcinoma cells containing mitochondrial DNA (ρ+) and NT2 cells depleted of mtDNA (ρ0) (Domingues et al., 2008). In particular, MPP+ (1 mM, 2 hr) elevated ubiquitinylated protein content (by ~3 fold compared to untreated Ctr), and after 24 hr induced a significant decrease of chymotrypsin-like activity (by ~30%) and peptidyl-glutamyl peptide hydrolytic-like activity (by ~75%) compared to untreated cells (Domingues et al., 2008). <ul> <li>Mice following continuous MPTP infusion (1-5-30 mg/kg daily) exhibited inhibition of the UPS (respectively by 40-50-60%) and increased inclusions of ubiquitin and α-synuclein in the neurons in the substantia nigra (Fornai et al., 2005).</li> </ul> <ul> <li>A mouse model of mitochondrial CI deficiency (Ndufs4-/- mice) showed an impaired 20S proteasomal activity (by ~50%), leading to increased ubiquitin protein levels (by ~40%) in the substantia nigra (not in cortex and hippocampus), increased of ubiquitin+/TH+ neurons (by ~2 fold, compared to WT mice), and increased ubiquitinated neurofilaments in the midbrain (values of 1.2 - 2.8 vs 1.0 in WT) (Song and Cortopassi, 2015).</li> </ul> Human studies. <ul> <li>PD patients appear to have an impaired UPS. The presence of aggregated, poly-ubiquitinated proteins in Lewy Bodies indicates that proteolytic dysfunction and proteo-toxicity are critical steps in the pathogenic cascade of PD (Betarbet et al., 2005). In this regard, impairment of proteasomal activity and reduced expression of proteasomal subunits have been reported selectively in subtantia nigra of sporadic PD post-mortem brains (McNaught et al., 2003; McNaught and Jenner, 2001). In particular, in PD, there was a 40.2% reduction in the amount of α-subunits in the SNc. On the opposite α-subunits levels were increased by 9.2% in the cerebral cortex and by 29.1% in the striatum in PD compared to Ctr (McNaught et al., 2003). Chymotrypsin-like, trypsin-like, and peptidyl glutamyl-peptide hydrolytic (PGPH) 20/26S proteasomal activities were significantly decreased in the substantia nigra (by 43.9%, 45.9%, and 44.6% respectively) (not in the cortex or striatum) in PD patients. At the same time, in PD there was a marked increase in the levels of PA700 subunits (the 19S regulatory complex of the 26S proteasome) in the frontal cortex and/or the striatum compared to controls, while in the SNpc PA700 subunits resulted decreased up 33%, whereas levels of nigral PA28 were almost undetectable in both normal and PD subjects (McNaught et al., 2003).</li> </ul> <ul> <li>Steady-state levels of soluble AF-6 (modulates parkin ubiquitin-ligase activity) have been found significantly lower in the caudate/putamen (~66% lower) as well as in the SN of PD patients (~66% lower). AF-6 was also detected in &sim;25% of mature Lewy bodies and in occasional Lewy neurites in the substantia nigra of the four PD brains analysed, and may contribute to the disruption of mitochondrial homeostasis (Haskin et al. 2013).</li> </ul> <ul> <li>HDAC6 has recently been identified by immunocytochemistry as a constituent in Lewy bodies of PD and dementia with LBs (DLB), as well as in glial cytoplasmic inclusions in multiple system atrophy (MSA) (Kawaguchi et al. 2003; Miki et al. 2011; Chiba et al. 2012). HDAC6 is considered a sensor of proteasomal inhibition and a cellular stress surveillance factor. Upon proteasomal inhibition, HDAC6 is relocated and recruited to polyubiquitin-positive aggresomes. HDAC6 inhibition elicits tubulin acetylation and restores microtubule (MT)-dependent transport mechanisms in neurons (Richter-Landsberg and Leyk, 2013).</li> </ul> <ul> <li>Basal activity of 20S proteasome was significantly reduced (by ~33%) in PD as compared to control fibroblasts. Higher accumulation of ubiquitinated proteins (by ~2 fold), representative of impaired 26S proteasome function, were found in PD as compared to Ctr cells at baseline. In the presence of rotenone (20 and 500 μM, 6 hr) PD-derived fibroblasts showed a higher induction of 20S proteasome activity (~15% higher) as compared to Ctr fibroblasts, with no significant changes in autophagy (except from increased LC3-II accumulation in both groups after exposure to 500 μM rotenone) (Ambrosi et al., 2014).</li> </ul>   Mitochondrial dysfunction by rotenone or MPP+ deregulates ALP activity <ul> <li>Exposure to rotenone (10 μM, 24 hr) induced neurotoxicity in human neuronal SH-SY5Y cells (number of dead cells was 8 folds higher than Ctr group) and pre-treatment with rapamycin (3 μM, 48 hrs) (strong inducers of autophagy) robustly protected against rotenone-mediated toxicity (number of dead cells was 3 folds higher than Ctr group) and this was due to the induction of autophagy. Indeed, suppression of autophagy (by silencing of Atg5) blocked the neuroprotection of rapamycin (Pan et al., 2009).</li> </ul> <ul> <li>Similar results were produced using kaempferol (6 μM, 1 hr prior addition of rotenone) and rotenone (50 nM, max up to 24 hr) on SH-SY5Y cells. Kaempferol was found to counteract rotenone-induced effects (see KER2) and these protective effects were related to induction of autophagy (6 hr kaempferol induced LC3-II formation, as shown by Western blot) (Filomeni et al., 2012).</li> </ul> <ul> <li>Treatment of SH-SY5Y cells with high doses of rotenone (500 nM, 48 hr) induced Atg5–Atg12 dependent autophagy, which leads to lysosomal dysfunction, increased p62 levels, and an aberrant accumulation of α-synuclein (Pan et al., 2009; Dadakhujaev et al., 2010). In particular, in α-synuclein expressing SH-SY5Y cells Atg5–Atg12 were increased by addition of rotenone and rapamycin (100 nM, 48 hr). Co-treatment with rotenone and autophagy inhibitors (e.g., 3-MA, bafilomycin or wortmannin) similarly diminished the level of Atg5–Atg12 in α-synuclein expressing cells (western blot analyses) (Dadakhujaev et al., 2010).</li> </ul> <ul> <li>A few studies have suggested that rotenone can act as an inducer of autophagic flux. For instance, treating human embryonic kidney cells (HEK 293) and U87 glioma cells with rotenone (50 μM, for 0-72 hr) caused cell death (in HEK 293 cells, rotenone induced 30% cell death, after 72 hr; in U87 cells, 40%) by upregulating autophagy and mitophagy (as shown by increase of cells with AVOs (indicative of autophagosomes and autolysosomes, analysed by flow cytometry): by ~14% in HEK 293 cells, and by ~20% in U87 cells, as compared to untreated cells, 0%), a process that is supposed to be triggered by mitochondrial superoxide (Chen et al., 2007).</li> </ul> <ul> <li>Increased autophagic flux has been observed in SH-SY5Y cells and primary cortical neurons treated respectively with 1 μM and 250 nM of rotenone. Rotenone elicited increases in autophagy (~ 2 folds vs Ctr) and mitophagy (i.e., as shown by the percentage of GFP-LC3 puncta colocalizing with mitochondria (~ 4 folds vs Ctr), indicating a preferential increase in “mitophagosomes” relative to total autophagosomes. Additionally, rotenone induced a decrease in p62 (SQSMT1), levels (~40% decrease with 250 nM), consistent with increased autophagic flux. This effect was reversed by co-treating cells with bafilomycin A2, a specific inhibitor of vacuolar-type H(+)-ATPase, or by RNAi (knockdown of ATG7 and ATG8/LC3). The mechanism by which LC3 recognizes damaged mitochondria in rotenone-treated neurons involves, among others, the externalization of cardiolipin and recruitment of LC3 at the mitochondria initiating rotenone induced-mitophagy and lysosomal-mediated degradation of mitochondria (Chu et al., 2013).</li> </ul> <ul> <li>In the study by Wu et al., (2015) chronically rotenone-treated rats (male Lewis rats received rotenone 1mg/kg subcutaneously twice a day for 8 weeks) had a robust loss of TH+ neurons in striatum (~50%) and in SNpc (~30%). However, in the remaining DA neurons of SNpc, cytoplasmic inclusions containing α-synuclein were observed (~7% of α-synuclein+/TH+ cells vs ~2% in Ctr), probably due to rotenone-induced decreased degradation of the autophagosomes (upregulation of LC3-II by ~30%, Beclin 1 by ~10%, and p62 by ~150%, after 24 hr rotenone) indicating decreased ALP function. Compared with the control group, the nigral DA neurons of the rotenone-treated group exhibited an increased diffuse distribution of LAMP2 (~15% vs ~25% Ctr) and cathepsin D (~22% vs ~60% Ctr) instead of punctuate pattern, indicating impaired lysosome integrity and a redistribution of cathepsin D from lysosomes to the cytosol. In parallel in vitro studies by the same group showed that PC12 cells exposed to rotenone (500 nM for 24 hr) underwent increased protein levels (but not mRNA levels) of α-synuclein (~4.5 folds vs Ctr), indicating an impairment of protein degradation. In TEM pictures, the majority of neurons displayed mitochondrial swelling, crista fragmentation, and accumulation of double membrane structures containing damaged mitochondria, which were stalled autophagosomes (Wu et al., 2015).</li> </ul> Similar results, showing impaired autophagic flux resulting in α-synuclein accumulation and the rupture of lysosomes in neuronal cell lines exposed to rotenone have been described in many other studies (e.g. Mader et al., 2012; Sarkar et al., 2014). <ul> <li>Rotenone produced bidirectional effects on macroautophagy (decrease or increase). This may be attributed to differences in the dosage, the duration, and cell type which can produce variable levels of ROS and mitochondrial damage (Pan et al., 2009; Dadakhujaev et al., 2010; Chen et al., 2007; Filomeni et al., 2012; Mader et al., 2012).</li> </ul> <ul> <li>MPP+ (2.5 mM, 24 - 48 hr) increased autophagy (~14 folds increase vs Ctr, of LC3-II) and mitochondrial loss in SH-SY5Y cells (a DA neuronal cell line widely used as a cell culture model of PD) by increased MAP kinase signalling (MEK inhibition by UO126 reversed by both autophagy and mitochondrial loss elicited by MPP+) (Zhu et al., 2007).</li> </ul> <ul> <li>Another study from the same group showed that longer MPP+ treatment (250 μM, 2 weeks) induced formation of enlarged, coarse GFP-LC3 puncta, in a time- and dose-dependent manner (~1.8% of cells presenting coarse GFP-LC3 puncta, vs ~0.2% in Ctr, at 14 days with 250 μM rotenone) (Zhu et al., 2012).</li> </ul> <ul> <li>An in vitro study on MN9D cells (a fusion of embryonic ventral mesencephalic and neuroblastoma cells, used as a model of DA neurons) showed that MPP+ (50 μM, for 24 hr) blocked autophagic flux, as evidenced by increased steady-state levels of p62 (qualitative data, Western blot), increased of authophagic vacuoles numbers (~3 folds vs Ctr) along with lysosomal depletion and dysfunction presumably due to leakage of lysosomes, impaired lysosomal biogenesis, and increased proteasomal-mediated degradation of proteins (as shown by time-dependent increase of ubiquitinated proteins, by IC) (Lim et al., 2011).</li> </ul> <ul> <li>In another study human neuroblastoma BE-M17 cells were treated with MPP+ (0.25-2.5 mM, 24 hr); Lamp1 protein levels were decreased in a dose-dependent manner in MPP+-treated cells (by ~40% at 2.5 mM), without concomitant decreases in mRNA expression levels. Also, LC3-II increased in a dose-dependent manner with MPP+ treatment (~3000% increase at 2.5 mM vs Ctr), indicating lysosome depletion and autophagosome accumulation upon MPP+ treatment. These data were confirmed in vivo: lysosomal depletion and accumulation of autophagosomes (as shown by ~600% increase of LC3-II, and ~40% decrease of Lamp1, after 1 day of MPTP injection compared to saline) occurred also in MPTP-intoxicated mice (30 mg/kg/day, for 5 consecutive days) (Dehay et al., 2010).</li> </ul> <ul> <li>Other in vivo data support a negative role of MPTP on autophagic flux. Mice were i.p. injected with 2 mg/ml MPTP (30 mg/kg) for 7 days. Suppression of autophagic flux induced by MPTP (~20% reduction vs Ctr) was detrimental to neuronal survival (as shown by ~60% decrease of TH+ neurons). Treating mice with the autophagy inducer rapamycin after seven days of MPTP treatment (daily i.p. injections of 2 mg/ml MPTP (30 mg/kg) for 7 days, followed by 0.1 ml of 20 µg/ml rapamycin by i.v. for an additional 7 days), significantly increased the number of surviving dopamine neurons (~60% TH+ neurons vs ~30% with MPTP alone, as compared to Ctr 100%) and the levels of TH protein (~75% vs ~60% with MPTP alone, as compared to Ctr 100%) and decreased the levels of α-synuclein aggregates (~210% of α-synuclein protein level, vs ~300% with MPTP alone, as compared to Ctr 100%) (Liu et al., 2013).</li> </ul>   <ul> <li>Treating mice with the autophagy inducer rapamycin after seven days of MPTP treatment (daily i.p. injections of 2 mg/ml MPTP (30 mg/kg) for 7 days, followed by 0.1 ml of 20 µg/ml rapamycin by i.v. for an additional 7 days), significantly increased the number of surviving dopamine neurons (~75% of TH protein level vs ~60% with MPTP alone) and decreases the levels of α-synuclein aggregates (~210% of α-synuclein protein level, vs ~300% with MPTP alone) (Liu et al., 2013).</li> </ul> MPP+ induced dysregulation of macroautophagy in neurons is discussed in recently published reviews (e.g. Cherra et al., 2010; Jiang et al., 2010). The potential other mechanisms by which rotenone or MPTP induce mitochondrial dysfunction are further discussed in recent publications (e.g. Dagda et al., 2013; Esteves et al., 2011). Impaired UPS and ALP function leads to α-synuclein aggregation: α-synuclein is one of the most abundant neuronal proteins (Vekrellis et al., 2011). Several PD-related mutations and environmental toxicants cause autophagy dysfunction and lead to the accumulation of misfolded proteins in DA neurons, including α-synuclein. Both monomeric and aggregated forms of α-synuclein can be degraded by macroautophagy, whereas only wild-type α-synuclein (not Ala30Pro, Ala53Thr and Glu46Lys mutant forms) is degraded by the process of chaperone-mediated autophagy (CMA) (Vekrellis et al., 2011). <ul> <li>Rotenone-induced α-synuclein aggregation has the ability to inhibit proteasome activity due to its propensity to assemble into filaments (as reviewed in Zaltieri et al., 2015). In particular, expression of α-synuclein was found to inhibit proteasome activity in SH-SY5Y cells. Increased levels of GFP-CL1 band were observed in cells coexpressing GFP-CL1 and α-synuclein (~9000 arbitrary units (au) vs ~500 au in DMSO-Ctr), indicating that proteasome activity is inhibited effectively by expression of α-synuclein (Nonaka and Hasegawa, 2009).</li> </ul> <ul> <li>By using stable PC12 cell lines expressing wild-type (WT) or A53T mutant human α-synuclein it has been shown that cells expressing mutant α-synuclein showed: (1) disruption of the ubiquitin-dependent proteolytic system, manifested by small cytoplasmic ubiquitinated aggregates and by an increase in polyubiquitinated proteins (qualitative data); (2) marked accumulation of autophagic-vesicular structures (qualitative data); (3) reduction of lysosomal hydrolysis and chymotrypsin-like proteasomal function (by ~ 30%, compared to WT) (Stefanis et al., 2001).</li> </ul> <ul> <li>Rotenone- (or MPP+)-induced inhibition of CI results in calcium (Ca2+) release from mitochondria. Calcium rise and oxidative stress cooperatively can promote α-synuclein aggregation (Follett et al., 2013; Goodwin et al., 2013; Nath et al., 2011).</li> <li>For instance, to investigate the influence of raised Ca2+ in response to plasma membrane depolarization on the aggregation of a-synuclein, HEK293T and SH-SY5Y neuroblastoma cells have been used and depolarized by addition of KCl to the cell culture medium. After KCl treatment (50 mM) increase of cellular Ca2+ was observed (~90% increase 20 min after KCl treatment), leading to the formation of frequent perinuclear α-synuclein focal aggregates at 26–74 hr post-treatment (qualitative IC images). By adding TMO (a selective T-type Ca2+ channel blocker) no a-synuclein aggregates were detected (Follett et al., 2013).</li> </ul> <ul> <li>Similarly, increased intracellular free Ca2+ (obtained by treating cells with either calcium ionophore or thapsigargin) induced the formation of α-synuclein aggregates in α-synuclein-GFP-transfected 1321N1 glioma cells (~65% increase compared to Ctr-untreated cells) (Nath et al., 2011).</li> </ul> <ul> <li>On the other hand, α-synuclein can control mitochondrial calcium homeostasis by enhancing endoplasmic reticulum-mitochondria interactions. Silencing of endogenous α-synuclein (siRNA-α-syn) in HeLa cells was found to impair mitochondrial Ca2+ transients (~35% decrease compared to Ctr-scrambled siRNA) and morphology (Calì et al., 2012). Also, α-synuclein oligomerization exacerbates calcium dysregulation by increasing mitochondria permeability transition (Danzer et al., 2007). Therefore, it is possible that mitochondrial dysfunction-induced calcium rise precede the onset of α-synuclein accumulation leading to UPS inhibition (Chou et al., 2010).</li> </ul> <ul> <li>It has been demonstrated that rotenone increased the intracellular calcium levels, triggering aggregation and phosphorylation of α-synuclein in a calcium-dependent manner. The aggregation of α-synuclein in PC12 cells following rotenone exposure was observed in a dose and time-dependent manner (1, 10 and 100 nM for 48 hrs, 3 days, 1 and 3 weeks) (~4 fold increase of α-syn with 100 nM rotenone for 48 hr, vs Ctr; and also, ~2.5 fold increase of α-syn with 1 nM rotenone for 1 week, vs Ctr) as evaluated via a variety of methods, including western blotting, immunofluorescence and electron microscopy. The observed attenuation of autophagy and α-synuclein aggregation was reversed by scavenging calcium (by using the calcium chelator BAPTA at 10 μM). Aggregated α-synuclein is typically degraded by autophagy, but rotenone impaired this process (Yuan et al., 2015).</li> </ul> <ul> <li>Under physiological conditions, α-synuclein is degraded by both the proteasome and autophagy. Mutant α-synuclein inhibits ALP functioning by tightly binding to the receptor on the lysosomal membrane for autophagy pathway control (e.g. Pan et al., 2009; Betarbet et al., 2000).</li> </ul> <ul> <li>The strongest evidence supporting that mitochondrial dysfunction precedes the onset of α-synuclein pathology derives from studies on rotenone and MPTP in which repetitive exposure of rodents and monkeys to these chemicals via oral, intraperitoneal, intragastric, or nasal administration resulted in the pathological accumulation of α-synuclein in central as well as peripheral neurons (Cannon et al., 2009; Drolet et al., 2009; Mandel et al., 2004; Pan-Montojo et al., 2012 and 2010; Tristão et al., 2014). For example, male Lewis rats were injected with rotenone (2.0 mg/kg, i.p.) and sacrificed at 0, 4, 8, 16, or 32 h after injection and showed α-synuclein and poly-ubiquitin accumulation and aggregation (as shown by IHC data) (Cannon et al., 2009).</li> </ul> Drolet and colleagues injected rats with rotenone (2.0 mg/kg, 1.0 ml/kg, i.p. 5 injections/week for 6 weeks) and found formic acid-resistant α-synuclein aggregates in the small intestine myenteric plexus, particularly 6-months after the last rotenone injection (3.5 median, vs 2.0 in Ctr) (Drolet et al., 2009). Mandel et al. injected male C57-BL mice with MPTP (24 mg/kg/day, ip for 5 days) and found α-synuclein aggregates (IHC data), which were decreased by using the radical scavengers apomorphine (injected s.c. at 10 mg/kg/day) or epigallocatechin-3-gallate (EGCG, given alone orally, 2 mg/kg/d) for 10 days) or a combination of both (Mandel et al., 2004). Inhibition of the mitochondria respiratory chain induces oxidative stress that in turn leads to lipid peroxidation of cellular and vesicular membranes at synaptic sites, resulting in dysfunction of neurotransmitter release. These effects facilitate α-synuclein conformational changes, such as accumulation, and aggregation. It has been demonstrated that synaptic dysfunction (caused by mitochondrial dysfunction) triggered the accumulation of α-synuclein (Nakata et al., 2012). Also, alterations of mitochondrial fission or dynamics can reduce synaptic mitochondrial load and impair neuronal function by hindering the proper energy demand to ensure synaptic function. Mitochondrial behaviours, especially those regulated by neuronal activity and synapse location, determine their distribution in the axon (Obashi and Okabe, 2013). These observations support the idea that mitochondrial dysfunction can affect synaptic environment and consequently result in α-synuclein accumulation at synapses (Zaltieri et al., 2015). <ul> <li>It was found that continuous administration of MPTP produced formation of nigral inclusions immunoreactive for ubiquitin and α-synuclein (Fornai et al., 2005). Mice were implanted with osmotic pump to deliver MPTP-HCl. Delayed and prolonged inhibition of striatal proteasome activity (i.e., 40-50-60% inhibition of UPS) occurred after continuous MPTP administration (respectively, 1-5-30 mg/kg MPTP daily) for the indicated time periods (Fig. 1) (Fornai et al, 2005). Continuous MPTP infusions caused also a long-lasting activation of glucose uptake. Additionally, in mice lacking α-synuclein, the MPTP-induced inhibition of the UPS system and the production of inclusion bodies were reduced (e.g., Ctr mice showed ~40% inhibition of postglutamyl peptidase (PGPH) activity, vs ~13% inhibition observed in α-synuclein KO mice) (Fig. 2), suggesting that α-synuclein could play an important role in UPS inhibition induced by MPP+ (Fornai et al., 2005). These data suggest that continuous, low-level exposure of mice to MPTP causes a Parkinson-like syndrome in a α-synuclein-dependent manner (Fornai et al., 2005).</li> </ul> These results are supported by other studies showing that α-synuclein−/− mice are resistant to MPTP toxicity (Dauer et al., 2002; Drolet et al., 2004). MPTP exposure (0.5, 5, 50 µM, 48 hr) increases in a dose-dependent manner the α-synuclein protein level in mesencephalic neurons in culture (e.g., ~70% increase at 5 µM vs Ctr) (Duka et al., 2006). Increased expression of α-synuclein predisposes DA neuronal cells to proteasomal dysfunction (~50% decrease compared to Ctr-vector cells) (Sun et al., 2005). <ul> <li>Accumulation/overexpression of α-synuclein, both wild type and mutant, potentiates inhibition of proteasomal activity. Cells expressing mutant α-synuclein showed a reduction of lysosomal hydrolysis and chymotrypsin-like UPS function (by ~30%, compared to WT) (Stefanis et al., 2001).</li> </ul> <ul> <li>Proteasomal inhibition (by mean of lactacystin, a proteasome inhibitor, used at different concentrations for 24 hr) contributes to the accumulation of α-synuclein as it has been described by immunostaining in PC12 cells (Rideout et al., 2001) and in primary mesencephalic neurons (McNaught et al., 2002).</li> </ul> <ul> <li>α-Synuclein levels were selectively increased in the ventral midbrain (VMB) region of rotenone-infused rats with or without lesion (~ 110% increase vs Ctr) (Fig. 3) (Betarbet et al., 2006). Rotenone was administered up to 5 weeks, at 2.5 mg/kg/day. Additionally, 4 weeks of in vitro rotenone exposure (5 nM, on SK-N-MC human neuroblastoma cells) increased α-Synuclein levels by 24%, while lactacystin (9 μM, overnight) did not induce any detectable changes in α-synuclein levels. α-Tocopherol attenuated the rotenone-induced increase in α-synuclein (comparable to Ctr) (Fig. 4). Furthermore, levels of ubiquitinated proteins detected in solubilized protein fractions from SK-N-MC cells resulted increased (by 60%) with rotenone treatment (5 nM), and even more (by 484%) with rotenone combined with lactacystin (Fig. 5) (Betarbet et al., 2006).</li> </ul> <ul> <li>CI inhibition-induced proteasomal dysfunction has been reported in human SH-SY5Y neuroblastoma cells following acute rotenone exposure (Shamoto-Nagai et al., 2003). The proteasome activity decreased in the cells treated with rotenone (25 or 50 nM) in a time- and dose-dependent way. ATP addition restored the reduction of proteasome activity in the cells treated with 25 nM rotenone for 72 hr. However, after 96 hr of incubation with 25 or 50 nM rotenone, the activity was reduced respectively to 28.7% and 21.9% of control, and adding ATP did not increase the activity. After 120 hr, the activity was virtually undetectable (with or without added ATP) (Fig. 6). On the contrary, the levels of the proteins composing proteasome did not change with rotenone treatment (Shamoto-Nagai et al., 2003).</li> </ul>   Cytoskeletal damage further enhances disturbed proteostasis: <ul> <li>α-synuclein can trigger hyperphosphorylation of Tau. Treatment of primary mesencephalic neurons acutely (48 h) or subchronic treatment of wild-type (WT) mice with MPP+/MPTP results in selective dose-dependent hyperphosphorylation of Tau at Ser396/404 (p-Tau). The presence of α-synuclein was absolutely mandatory to observe MPP+/MPTP-induced increases in p-Tau levels, since no alterations in p-Tau were seen in transfected cells not expressing α-synuclein or in α-synuclein-/- mice. MPP+/MPTP also induced a significant accumulation of α-synuclein in both mesencephalic neurons and in WT mice striatum. Sub-chronic MPTP exposure increased phosphorylated-Tau in striatum of WT (but not α-Syn-/- mice) causing microtubule (MT) cytoskeleton instability that affects cellular microtubule transport (including axonal transport) (Qureshi et al., 2009; Duka et al., 2006). For instance, MPTP was found to elicit an increase of phosphorylated Tau at Ser262 by 2.8-, 4.5-, 4.6-, and 4.0-fold higher in 1, 5, 25, and 50 μM MPTP-treated cells than the basal level observed in Ctr/vehicle-treated cells, respectively. Additionally, MPTP caused a dose-dependent increase in the intracellular α-synuclein level in M17 human neuroblastoma cells (~3.5 fold increase in cells treated with 25 μM MPTP vs Ctr) (Qureshi and Paudel, 2009). These results were confirmed by other studies (e.g. Dauer et al., 2002; Drolet et al., 2004 etc.).</li> </ul> <ul> <li>α-synuclein accumulation followed by MT depolymerisation induces disruption in axonal transport, which leads to an accumulation of damaged organelles, aggregated/misfolded proteins and impaired vesicular release. Dopamine is leaking from the vesicles to the cytosol promoting an increase in oxidative stress, potentiated by dopamine oxidation (Feng, 2006; Kim et al., 2007). When microtubule network is disrupted, the amount of free tubulin increases, triggering α-synuclein fibrillization (Payton et al., 2001).</li> </ul> <ul> <li>Axonal transport might be impaired by misfolded α-synuclein through perturbation of microtubule assembly (Esposito et al., 2007; Lee et al., 2002; Chen et al., 2007 ), especially together with MAPT protein (Qureshi and Paudel, 2011; Giasson et al., 2003). It induces not only microtubule disruption but also impairs microtubule-dependent trafficking (Lee at al., 2006). MT-dependent transport is important for maintaining the Golgi structure, and thus, depolymerization of the MT leads to a specific pattern of Golgi fragmentation (Cole et al., 1996). When the MT network was disrupted by nocodazole treatment (5 µg/mL) or α-synuclein was overexpressed, this normally compact organelle was fragmented and dispersed (IC images) as shown in COS-7 cells (Lee at al., 2006). Similarly, overexpression of α-synuclein in differentiated SH-SY5Y cells caused Golgi fragmentation (e.g., ~190% increased fragmented Golgi at 12 m.o.i. (multiplicity of infection) of α-synuclein vs Ctr) (Lee at al., 2006).</li> </ul> <ul> <li>It was found that α-synuclein mutants associated with PD exhibit reduced transport in neurons, as shown in rat primary neuronal cortical cultures transfected with wild-type (WT), A53T or A30P α-synuclein. For instance, the rate of transport (expressed in µm/hr) was reduced of ~55% and ~60% after 3-4 hr for A30P and A53T respectively (vs Ctr-WT) (Saha et al., 2004).</li> </ul> <ul> <li>Damaged cytoskeletal proteins disrupt also mitochondrial trafficking. Mitochondria use cytoskeletal proteins as tracks for their directional movement (Nogales, 2000). The cytoskeletal system regulates not only mitochondrial movement but also their morphology and function. Therefore, damage to microtubules perturbs transport of mitochondria through axons, increasing their retrograde movement. These changes in mitochondria dynamics lead to a decrease of mitochondria numbers in axons and mitochondria accumulation in cell bodies (De vos et al., 2007; Miller and Sheetz, 2004). Depletion of mitochondria quantity and function in axons occurs in neurodegenerative disorders (Brownlees et al., 2002; Stamer et al., 2002). Since mitochondria are ATP suppliers and microtubules need ATP to accomplish their function, mitochondrial dysfunction has a profound effect on axonal transport and function (De Vos et al., 2008).</li> </ul> <ul> <li>Mitochondrial dysfunction may damage mitochondrial trafficking through calcium dysregulation. Cytosolic Ca2+ is one of the best-studied regulators of mitochondrial movement. Elevation of cytosolic Ca2+ stops both the anterograde and retrograde trafficking of mitochondria in neurons and in many cell lines. (Chang et al. 2006; Szabadkai et al. 2006). In H9c2 cells simultaneous measurements of free Ca2+ levels and mitochondrial dynamics showed that 50% reductions in mitochondrial movement occurred at concentrations of approximately 400 nM Ca2+, and a complete arrest in the low micromolar range (Yi et al. 2004; Saotome et al., 2008). These are indirect proofs suggesting that inhibition of CI, followed by mitochondrial dysfunction, could damage mitochondrial trafficking. Also, chronic exposure to rotenone (50 nM at different times of exposure) was reported to reduce mitochondrial movement in differentiated SH-SY5Y cells (e.g., ~30% reduction of mitochondrial movement (µm/sec) after 8 days of rotenone treatment vs Ctr) (Borland et al., 2008).</li> </ul> Human studies <ul> <li>In PD patient postmortem cortical tissues, levels of oligomeric α-synuclein in SNpc (~1000% vs Ctr samples) and expression of LC3-II levels (~130% vs Ctr samples) were up-regulated (Yu et al., 2009) (for further info, see the review from Vekrellis et al., 2011).</li> </ul> <ul> <li>The pathological observations in PD autopsy brains showed that LC3-II levels were elevated in the SNpc and amygdala of PD brain samples, suggesting an increase in macroautophagy (but they did not reach statistical significance). LC3 colocalized with α-synuclein in most LBs and Lewy neurites in PD SNpc as well as in small punctate α-synuclein immunoreactive inclusions (IC images) (Alvarez-Erviti et al., 2010).</li> </ul> <ul> <li>Analogously, another study reported that brain homogenates derived from the temporal cortex of dementia with LB (DLB) patients vs non-demented controls were characterized by higher levels of both mTor (~130% vs Ctr) and p-mTor (~ 10 folds higher than Ctr), and levels of Atg7 (molecular initiator of autophagy) were moderately reduced in DLB cases compared to Ctr (~ 40% lower than Ctr). Consistent with the studies in human brains, levels of both mTor and p-mTor were increased in the membrane fractions from brains of α-synuclein tg mice compared to non tg controls (respectively, by ~250% and ~200% vs Ctr), and levels of Atg7 were reduced in α-synuclein tg brains compared to non tg controls (~75% less than Ctr) (Crews et al., 2010).</li> </ul> <ul> <li>Another study showed that post-mortem brain samples derived from PD patients, compared to age-matched controls, presented significant reductions of LAMP1 (2069.10 ± 329.52), CatD (1809.35 ± 533.47), HSP73 (2604.92 ± 494.56), and 20S proteasome (1660.84 ± 229.87) calculated by optic density (OD) measures (Chu et al., 2009). These data globally indicate that the functions of both the UPS and ALP systems is compromised in PD patients.</li> </ul>

<ul> <li>The exact molecular link from mitochondrial dysfunction to disturbed proteostasis is not known. It is not clear which is the oxidative modification that drives the process.</li> </ul> <ul> <li>The sequence of events taking place after inhibition of CI is not entirely clear (Zaltieri et al., 2015). Some studies suggest that induced oxidative stress leads to α-synuclein aggregation that triggers proteosomal dysfunction (Betarbet et al., 2006). Such order of events is suggested to take place in vivo (McNaught and Jenner, 2001). However, in other studies opposite sequence of events is proposed suggesting that first proteosomal dysfunction take place that leads to α-synuclein aggregation.</li> </ul> A vicious circle is observed here as α-synuclein aggregation potentiates proteosomal dysfunction and v/v. In this vicious cycle it is difficult to establish exact quantitative relationship of these two events. <ul> <li>Whether α-synuclein is a substrate for proteasome remains controversial since both positive and negative data have been reported (Paxinou et al., 2001). Furthermore, polyubiquitination of α-synuclein, a prerequisite for 26S proteasomal degradation has yet to be reported (Stefanis et al., 2001). It is also not clear whether polyubiquitination of α-synuclein is necessary for its degradation. However, α-synuclein gets targeted by the UPS in the SHSY5Y neuroblastoma cell line. Phosphorylated α-synuclein gets targeted to mono- or di-ubiquitination in synucleinopathy brains (Hasegawa et al., 2002), but it is not clear if this modification can play any role in proteasomal degradation since monoubiquitination of proteins serves mainly as a signal for endocytosis or membrane trafficking.</li> </ul> <ul> <li>On the contrary to the increased α-synuclein levels observed in the midbrain, decreased α-synuclein levels were found in the cerebellums of PD patients when compared to controls, suggesting an imbalance of α-synuclein levels in different parts of the brain (Westerlund et al., 2008).</li> </ul> <ul> <li>Although mitochondrial alterations have been reported in PD patients (Ikawa et al., 2011) and disease models, it is not clear whether they represent a primary pathogenic mechanism. In particular, the critical interplay between mitochondrial dysfunction and oxidative stress, which has been widely reported in PD (Dias et al., 2013) and could constitute either a cause or a consequence of mitochondrial damage, hampers an effective comprehension of the above mentioned studies. Oxidative stress can constitute a bridge connecting mitochondrial dysfunction to the induction of α-synuclein misfolding, aggregation, and accumulation, but otherwise it may be also triggered by these latter events that in turn could induce mitochondrial alterations (Zhu and Chu, 2010; Dias et al., 2013).</li> </ul> <ul> <li>It is still unclear whether the involvement of α-synuclein in chronic MPTP toxicity reflects a physiological function for α-synuclein that has been activated in the wrong context, or whether α-synuclein produces an accidental pathogenicity that contributes to MPTP toxicity but is unrelated to the normal function of α-synuclein (Fornai et al., 2005).</li> </ul> <ul> <li>The inconsistent effects of MPP+ on autophagy (up or down regulation) are reported. It may be attributed to differences observed between immortalized cell lines and primary neurons, different timing or dose. While dysregulation of autophagy is always described, the direction is not clear. Further studies are required to clarify this issue.</li> </ul> <ul> <li>MPTP administration does not induce Lewy body formation (in contrast to rotenone) characteristic of PD, even after repeated injections (Drolet et al., 2004; Dauer et al., 2002).</li> </ul> <ul> <li>There is also controversy over whether the increase in autophagic markers is protective or, on the contrary, causative of neuronal death.</li> </ul> <ul> <li>MPP+ may have effects apart from CI inhibition, e.g., on microtubules but it is still unclear whether this is a primary effect. Indeed, MPP+ binds to microtubules in PC12 cells and inhibits their polymerization and stability (Cappelletti et al., 1999; Cappelletti et al., 2001).</li> </ul> <ul> <li>It is not clear whether microtubules disruption may be associated with α-synuclein aggregation since tubulin was shown to co-localize with α-synuclein in Lewy bodies. Furthermore, tubulin folding is dependent on ATP and GTP hydrolysis, and mitochondrial dysfunction with subsequent energy failure could trigger microtubules disruption. Cytoskeletal microtubule (MT) injury is likely to be responsible for altered rearrangement and movement of cell organelles, being a common feature of several neurodegenerative diseases including PD (Wade, 2009; Mattson et al., 1999).</li> </ul> <ul> <li>It is not clear whether rotenone could cause microtubules depolymerization in vivo and in vitro (Brinkley et al., 1974) by binding to the colchicine site on tubulin heterodimers (Marshall et al., 1978). Ren and Feng (2007) found that microtubule depolymerization induced by rotenone caused vesicle accumulation in the soma and kills neurons.</li> </ul>

As described in the studies above (Empirical support for linkage) a quantitative or semi-quantitative relationship has been established between rotenone-induced mitochondrial dysfunction and the impairment of UPS/ALP function. Below some representative studies are reported as examples for how such quantitative evaluations can be performed. <ul> <li>Human neuroblastoma SK-N-MC or human embryonic kidney (HEK) cells were exposed to rotenone at 100 nM for 24 or 48 hrs (for further details see Chou et al., 2010).</li> </ul> <ul> <li>PD patient-derived fibroblasts (vs Ctr fibroblasts) treated with rotenone (20 and 500 μM for 6 h for the evaluation of protein quality control system or 100 nM, 1 μM and 10 μM for 1 h for redox experiments) showed reduction of UPS function (as shown by higher induction of 20S proteasome activity in PD fibroblasts vs Ctr after both 20 and 500 μM rotenone administration). An increase of LC3-II accumulation in both groups (PD and Ctr) after exposure to 500 μM rotenone was observed suggesting that (Ambrosi et al. 2014).</li> </ul> <ul> <li>Human neuroblastoma cells (SK-N-MC) after short treatment with rotenone (1 week) elevated soluble α-synuclein protein (41 ± 16% increase) levels without changing mRNA levels, suggesting impairment of α-synuclein degradation via UPS. Chronic rotenone exposure (4 weeks) increased levels of insoluble α-synuclein (29 ± 9% increase) and ubiquitin (87 ± 14% increase) (Sherer et al., 2012).</li> </ul> <ul> <li>SHSY-5Y cells treated with rotenone (500 nM, 24 h) showed a ~2 fold increase in DCF fluorescence compared to untreated cells (indicative of intracellular ROS). Additionally, rotenone elevated cytosolic calcium (about 35-40% increase vs Ctr), ER-stress (about 45% increase vs Ctr), impaired UPS function (~3 fold increase of insoluble protein aggregate vs Ctr). Inhibition of Rac1 (Rho-like GTPase) mitigated the oxidative/nitrosative stress, prevented calcium-dependent ER-stress, and partially rescued UPS function (Pal et al. 2014).</li> </ul> <ul> <li>Human neuronal SH-SY5Y cells treated with rotenone (10 μM, for 24 hr showed accumulation of high molecular weight ubiquitinated bands (by immunoblotting – qualitative - assay), and increase of both mitochondrial- (~5 fold increase vs Ctr) and cytosolic- cytochrome c fractions (~1.2 fold increase vs Ctr). Rapamycin pre-treatment (3 μM, for 48 hr prior addition of rotenone) diminished rotenone-induced effects, as shown by enhanced degradation of ubiquitinated proteins, and reduced levels of cytosolic cytochrome c. Also, rapamycin promoted mitophagy (as shown by lysosome and mitochondria co-localization within the cells) (Pan et al. 2009).</li> </ul> Examples of quantitative evaluation of this KER   <a class="image" href="https://aopwiki.org/wiki/index.php/File:KER_3_Fig.1._proteosome_activity.jpg"><img alt="KER 3 Fig.1. proteosome activity.jpg" src="https://aopwiki.org/wiki/images/7/71/KER_3_Fig.1._proteosome_activity.jpg" style="height:577px; width:499px" /></a>   Fig.1. Dose and time dependent striatal proteasome activity after MPTP continuously infused upto 28 days measured by relative chymotrypsin-like, trypsin-like, and peptidyl-glutamyl-peptide hydrolysing (PGPH) proteasome activities in mice. Delayed and prolonged inhibition of proteasome activity after continuous MPTP administration (1, 5, or 30 mg/kg MPTP daily) for the indicated time periods. Asterisks indicate statistically significant differences (P _<0.05) from baseline proteasome activity (single asterisk) or from both baseline proteasome activity and activity after lower MPTP doses (1 and 5 mg/kg, daily, double asterisk; n =5 mice) (Fornai et al., 2005, Fig. 2 B).   <a class="image" href="https://aopwiki.org/wiki/index.php/File:KER3_Fig._2._.jpg"><img alt="KER3 Fig. 2. .jpg" src="https://aopwiki.org/wiki/images/2/29/KER3_Fig._2._.jpg" style="height:371px; width:438px" /></a> Fig. 2. Effect of α-synuclein deletion on MPTP toxicity. Proteasome activity in control and alpha-synuclein KO mice continuously infused for 28 days with MPTP (30 mg/kg of body weight daily, striatum concentration approximately 13 uM). Proteasome activities in the substantia nigra are depicted as percent of control (means +/- SEMs) as a function of time after beginning of the infusions (five mice per group). Asterisks indicate statistically significantly different values (P < 0.05) from controls (Fornai et al., 2005).   <a class="image" href="https://aopwiki.org/wiki/index.php/File:KER3_Fig._3._.jpg"><img alt="KER3 Fig. 3. .jpg" src="https://aopwiki.org/wiki/images/c/c5/KER3_Fig._3._.jpg" style="height:345px; width:580px" /></a> Fig. 3. α-Synuclein levels were selectively increased in the ventral midbrain (VMB) region of rotenone-infused rats with or without lesion. α-Synuclein levels, as determined from Western blot analysis, from rotenone-treated rats were expressed as a percentage of values from control vehicle-infused rats. Results are mean ± SEM (n = 3 control, 6 rotenone with lesion, 3 rotenone with no lesion) *P < 0.05 vs. vehicle-infused rats (from Betarbet et al., 2006, Fig. 3A).   <a class="image" href="https://aopwiki.org/wiki/index.php/File:KER3_Fig._4.jpg"><img alt="KER3 Fig. 4.jpg" src="https://aopwiki.org/wiki/images/4/4c/KER3_Fig._4.jpg" style="height:407px; width:513px" /></a> Fig. 4. Bar graph showing the effects of rotenone and lactacystin on α-synuclein levels after 4 weeks of rotenone exposure (5 nM) in vitro, on SK-N-MC human neuroblastoma cells. Rotenone alone increased α-synuclein levels, but lactacystin alone did not. α-Tocopherol attenuated the rotenone-induced increase in α-synuclein. Results are mean ± SEM (n = 4). *P < 0.05 vs. solvent-treated cells. CC, control cells; RC, rotenone-treated cells; C-Lac or CL, lactacystin treated cells; R-lac or RL, rotenone and lactacystin treated cells; R-AT, rotenone and α-tocopherol treated cells (from Betarbet et al., 2006, Fig. 5B).   <a class="image" href="https://aopwiki.org/wiki/index.php/File:KER3_Fig._5.png"><img alt="KER3 Fig. 5.png" src="https://aopwiki.org/wiki/images/e/ef/KER3_Fig._5.png" style="height:480px; width:500px" /></a> Fig.5. Levels of ubiquitinated proteins were estimated in solubilized protein fractions from SK-N-MC cells collected at the end of each week of rotenone treatment (5 nM), using gel electrophoresis and immunoblotting. Quantitative analysis demonstrated significant increases in ubiquitinated protein levels 4 weeks after rotenone treatment and after proteasomal inhibition with lactacystin. Band intensities were expressed as % of control. Results represent mean ± SEM. *P < 0.05 compared to control (from Betarbet et al., 2006, Fig. 8C). <a class="image" href="https://aopwiki.org/wiki/index.php/File:Fig._6.jpg"><img alt="Fig. 6.jpg" src="https://aopwiki.org/wiki/images/3/3c/Fig._6.jpg" style="height:244px; width:429px" /></a> Fig. 6. Effects of rotenone on the activity of proteasome. Proteasome activity in the cytoplasmic fraction of cells treated with 25 nM (A) or 50 nM (B) rotenone was measured fluorometrically in the absence (open triangles and circles) or presence (solid triangles and circles) of exogenously added ATP (2 mM) (from Shamoto-Nagai et al., 2003, Fig. 6).   <table border="1"> <tbody> <tr> <td> KE (upstream) Mitochondrial dysfunction </td> <td colspan="2"> KE3 (downstream) Impaired proteostasis UPS inhibition (% approx.) measured by: </td> <td> Comments </td> <td> References </td> </tr> <tr> <td> Rotenone (nM) (in vitro) </td> <td> 26S UPS activity </td> <td> + catalase (anti-oxidant) </td> <td> HEK cells exposed for 2 4hr </td> <td> Chou et al., 2010 </td> </tr> <tr> <td> 10 </td> <td> 24 </td> <td> Not done </td> <td>   </td> <td>   </td> </tr> <tr> <td> 100 </td> <td> 48 </td> <td> Increased UPS activity by 40% </td> <td>   </td> <td>   </td> </tr> <tr> <td> 1000 </td> <td> 60 </td> <td> Not done </td> <td>   </td> <td>   </td> </tr> <tr> <td>   </td> <td colspan="2"> 20S proteasome activity </td> <td> SK-N-MC human neuronal cell line (exposed for 24 hr) </td> <td> Chou et al., 2010 </td> </tr> <tr> <td> 1 </td> <td colspan="2"> 8 </td> <td>   </td> <td>   </td> </tr> <tr> <td> 50 </td> <td colspan="2"> 4 </td> <td>   </td> <td>   </td> </tr> <tr> <td> 100 </td> <td colspan="2"> 18 </td> <td>   </td> <td>   </td> </tr> <tr> <td> 500 </td> <td colspan="2"> 22 </td> <td>   </td> <td>   </td> </tr> <tr> <td> 1000 </td> <td colspan="2"> 24 </td> <td>   </td> <td>   </td> </tr> <tr> <td>   </td> <td colspan="2"> 20S proteasome immune-reactivity decrease </td> <td>   </td> <td>   </td> </tr> <tr> <td> 10 </td> <td colspan="2"> 22 </td> <td>   </td> <td>   </td> </tr> <tr> <td> 100 </td> <td colspan="2"> 48 </td> <td>   </td> <td>   </td> </tr> <tr> <td> 100 </td> <td colspan="2"> 70 </td> <td>   </td> <td>   </td> </tr> <tr> <td> MPTP (in vivo) </td> <td colspan="2"> Chymotrypsin-like UPS activities (at day 2) </td> <td>   </td> <td>   </td> </tr> <tr> <td> 1 mg/kg daily </td> <td colspan="2"> 20 </td> <td> Mice continuously infused with MPTP for 28 days </td> <td> Fornai et al., 2005 </td> </tr> <tr> <td> 5 mg/kg daily </td> <td colspan="2"> 30 </td> <td>   </td> <td>   </td> </tr> <tr> <td> 30 mg/kg daily </td> <td colspan="2"> 40 </td> <td>   </td> <td>   </td> </tr> <tr> <td>   </td> <td colspan="2"> Trypsin-like UPS activities (at day 2) </td> <td>   </td> <td>   </td> </tr> <tr> <td> 1 mg/kg daily </td> <td colspan="2"> 30 </td> <td>   </td> <td>   </td> </tr> <tr> <td> 5 mg/kg daily </td> <td colspan="2"> 40 </td> <td>   </td> <td>   </td> </tr> <tr> <td> 30 mg/kg daily </td> <td colspan="2"> 60 </td> <td>   </td> <td>   </td> </tr> <tr> <td>   </td> <td colspan="2"> Peptidyl-glutamyl-peptide hydrolysing (PGPH) UPS activities (at day 2) </td> <td>   </td> <td>   </td> </tr> <tr> <td> 1 mg/kg daily </td> <td colspan="2"> 20 </td> <td>   </td> <td>   </td> </tr> <tr> <td> 5 mg/kg daily </td> <td colspan="2"> 20 </td> <td>   </td> <td>   </td> </tr> <tr> <td> 30 mg/kg daily </td> <td colspan="2"> 30 </td> <td>   </td> <td>   </td> </tr> </tbody> </table>   Table. 1. These studies showed that rotenone caused a reduction in UPS activity (measured by 26S and 20S proteasome activity) in a dose-dependent manner. Further studies showed that rotenone increases proteasome subunit degradation, but does not alter synthesis (Western blot and RT-PCR studies, reviewed in Chou et al., 2010). Dose- and time- dependent striatal proteasome activity is also shown after MPTP continuously infused up to 28 days measured by relative chymotrypsin-like, trypsin-like, and peptidyl-glutamyl-peptide hydrolysing (PGPH) proteasome activities in mice (Fornai et al. 2005).   <ul> <li>PD patient-derived fibroblasts (vs Ctr fibroblasts) showed reduction of UPS function (by ~33%) and higher accumulation of ubiquitinated proteins (by ~2 fold) in PD as compared to control fibroblasts at baseline. Treatment with rotenone (20, 500 μM, 6hr) caused a higher induction of 20S proteasome activity in PD fibroblasts vs Ctr. An increase of LC3-II accumulation (indicative of autophagic vesicle accumulation) in both groups (PD and Ctr) after exposure to 500 μM rotenone was observed (Ambrosi et al. 2014).</li> </ul> <ul> <li>Human neuroblastoma cells (SK-N-MC) after short treatment with rotenone (1 week) elevated soluble α-synuclein protein (41 ± 16% increase) levels without changing mRNA levels, suggesting impairment of α-synuclein degradation via UPS. Chronic rotenone exposure (4 weeks) increased levels of insoluble α-synuclein (29 ± 9% increase) and ubiquitin (87 ± 14% increase) (Sherer et al., 2012).</li> </ul> <ul> <li>SHSY-5Y cells treated with rotenone (500 nM, 24 h) showed a ~2 fold increase in DCF fluorescence compared to untreated cells (indicative of intracellular ROS). Additionally, rotenone elevated cytosolic calcium (about 35-40% increase vs Ctr), ER-stress (about 45% increase vs Ctr), impaired UPS function (~3 fold increase of insoluble protein aggregate vs Ctr). Inhibition of Rac1 (Rho-like GTPase) mitigated the oxidative/nitrosative stress, prevented calcium-dependent ER-stress, and partially rescued UPS function (Pal et al. 2014).</li> </ul> <ul> <li>Human neuronal SH-SY5Y cells treated with rotenone (10 μM, for 24 hr showed accumulation of high molecular weight ubiquitinated bands (by immunoblotting – qualitative - assay), and increase of both mitochondrial- (~5 fold increase vs Ctr) and cytosolic- cytochrome c fractions (~1.2 fold increase vs Ctr). Rapamycin pre-treatment (3 μM, for 48 hr prior addition of rotenone) diminished rotenone-induced effects, as shown by enhanced degradation of ubiquitinated proteins, and reduced levels of cytosolic cytochrome c. Also, rapamycin promoted mitophagy (as shown by lysosome and mitochondria co-localization within the cells) (Pan et al. 2009).</li> </ul>

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b430b0385d0> AOPWiki 2016-11-29T18:41:35

2017-08-25T08:35:05

Mitochondrial complex inhibition leading to liver injury

Arthur Author

van der Stel, W. 1 1Leiden Academic Center for Drug Research, Leiden Univeristy, Leiden, The Netherlands

BY-SA

2.0

This AOP focusses on the inhibition of any of the mitochondrial complexes (I,II,III,IV or V) which eventually leads to the initiation of liver injury. In short, inhibition (fully or partially) of mitochondrial complexes that form the OXPHOS will lead to perturbation of the OXPHOS itself. OXPHOS perturbation will manifest as lower or no production of ATP via the mitochondria. Besides OXPHOS perturbation, the inhibition of the complexes will also lead to accumulation of the unused electrons and therefore ROS formation. Both process combined will trigger mitochondrial injury. One the most important additions to the AOPwiki by us would be to improve the mitochondrial injury key event with more detail, more method descriptions and more evidence. Finally when a certain threshold is reached mitochondria cannot coop with the induced stress and will trigger the cell death pathways (apoptosis and necrosis depending on ATP availability). Evidently, the cell death will proceed to necrotic tissue and in the end to liver injury.

Numerous hydrophobic, amphipathic compounds are known to inhibit the proton pumping NADH:ubiquinone oxidoreductase, also known as the ubiquinone reductase reaction of respiratory chain complex I (Fendel et al., 2008). However, the most studied examples of chemicals that inhibit CI are: rotenone and 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP) (Desplats et al., 2012; Lin et al., 2012; Sava et al., 2007). Both, rotenone (pesticide) and MPP+ (the active metabolite of MPTP) are well known to reproduce the anatomical, neurochemical, behavioural and neuropathological features of PD-like syndrome (Betarbet et al., 200; Greenamyre et al., 2001). Indeed, an overwhelming evidence has accumulated in the existing literature suggesting such a link and therefore these two inhibitors of CI will be discussed in the context of all KE identified in this AOP. 1. Rotenone affinity to complex I binding sites. Rotenone (a flavonoid, extracted from the several plants e.g. Derris scandens)is one of the most powerful, an irreversible inhibitor of CI, binding with high affinity to CI and is typically used to define the specific activity of this complex. Rotenone is extremely lipophilic, it crosses biological membrane easily and it gets into brain very rapidly. Rotenone inhibits 20 kDa subunit of complex I (PSST) labeling without effect on 36 kDa subunit of complex I (ND1) (Schuler and Casida, 2001). The interaction of rotenone with active ('pulsed') and thermally de-activated ('resting') membrane-bound Complex I as revealed by inhibition of NADH-ubiquinone- and ubiquinol-NAD+ reductase activities was studied. Ki = 1 x 10(-9) M, k(on) = 5 x 10(7) M-1 min-1 and k(off) = 0.02 min-1 (inhibitory effect of rotenone on NADH oxidation) and Ki = 2 x 10(-8) M (inhibition of reverse electron transfer) were determined for pulsed enzyme. The equilibrium between de-activated and active enzyme is reached (K approximately 100) after the slow strongly temperature-dependent de-activation process has completed. Rotenone partially prevents and reverses the enzyme de-activation. About two order of magnitude difference in affinity of rotenone to the active and de-activated forms of the enzyme was demonstrated (Grivennikova et al., 1997). Dose-dependent relative affinities of rotenone to the inhibitory site of CI is shown in Fig. 3B (for more detail Grivennikova et al., 1997). Most of the studies suggest that hydrophobic inhibitors like rotenone or Piericidin A most likely disrupt the electron transfer between the terminal Fe-S cluster N2 and ubiquinone (Fig. 3A).   <a class="image" href="/wiki/index.php/File:MIE_Fig._3A.jpg"><img alt="MIE Fig. 3A.jpg" src="/wiki/images/0/03/MIE_Fig._3A.jpg" style="height:283px; width:560px" /></a>   Fig. 3A. Rotenone structure and a schematic representation of its binding site (and other Rotenone-like compounds) to CI. IMS: inter-membrane space (based on Lummen, 1998)   <a class="image" href="/wiki/index.php/File:MIE_Fig._3B.jpg"><img alt="MIE Fig. 3B.jpg" src="/wiki/images/b/bd/MIE_Fig._3B.jpg" style="height:277px; width:505px" /></a>   Fig. 3B. Fig. 2. Relative affinities of rotenone to the inhibitory site(s) of Complex I. Panel (A): activated submitochondrial particles (SMP) (2.8 mg/ml, approx. 0.4 microM Complex I) were incubated in the standard reaction mixture for 20 min at 25oC and residual initial rate of NADH oxidation was measured. 100% correspond to the specific activity of 1.0 micromol/min per mg of protein. Panel (B): curve 1 (o), SMP (48 microg/ml, approx. 8 nM Complex I) were activated in the assay cuvette and pre-incubated with rotenone in the presence of gramicidin and 10 mM malonate for 20 min at 25oC and the residual NADH oxidase activity was then measured; black circle: the same as (o), except that pre-incubation with rotenone was made in the presence of 10 mM succinate (no gramicidin and malonate), 10 mM malonate and gramicidin were added simultaneously with 100 microM NADH to measure the residual activity. Curve 2, presents the reverse electron transfer activity and curve 3, de-activated SMP were preincubated with rotenone as described for curve 1(o) (for further details see Grivennikova et al., 1997). Panel (C): The same as Panel B, curve 3, except for enzyme concentration was 0.5 mg/ml and rotenone concentration range which was increased to show interaction of the inhibitor with de-activated enzyme. The activity was measured after 200-fold dilution into the assay mixture. All the continues lines corresponds to the theoretical titration curves for the reversible single site inhibition with Ki values of 1 nM, 20 nM and 80 nM for the curves 1, 2 and 3, respectively (for further details see Grivennikova et al., 1997). 2. MPTP affinity to complex I binding sites. MPTP is not directly binding to CI and it is therefore non-toxic to DA neurons. MPTP exerts its toxicity after it is metabolized by mono-amino-oxidase, type B (MAO B), in astrocytes to 1-methyl-4-phenylpyridinium (MPP+). This metabolite binds to CI, and is toxic. MPP+ is a good substrate for dopamine transporters (DAT), expressed selectively by DA neurons (Greenamyre et al (2001). Due to both a positive charge and an amphoteric character, MPP+ specifically accumulates in mitochondria, where despite a lower affinity to the binding site of complex I than rotenone, it reaches high enough intra-mitochondrial concentrations to inhibit CI activity (Ramsay et al., 1991). The binding affinity of MPP+ is low (mM range), and it can be totally reversed by washing out.  Competitive binding experiments with rotenone and MPP+ suggest that the two compounds bind to the same site of the CI (Ramasay et al., 1991). Schuler and Casida (2001) reported that MPP+ inhibits PSST and elevates ND1 labelling subunits of the mitochondrial complex I. 3. General characteristics of other complex I inhibitors. There is a variety of CI inhibitors, both naturally occurring besides rotenone such as Piericidin A (from Streptomyces mobaraensis), acetogenins (from various Annonaceae species) as well as their derivatives, and synthetically manufactured compounds like pyridaben and various piperazin derivatives (Ichimaru et al. 2008). They have been used to probe the catalytic activity of complex I especially in order to clarify its ubiquinone binding site and indeed, most of these compounds inhibit the electron transfer step from the Fe-S clusters to ubiquinone (Friedrich et al. 1994). Therefore, classification of CI inhibitors is based on their types of action. Type A inhibitors, like piericidin A, 2-decyl-4-quinazolinyl amine (DQA), annonin VI and rolliniastatin-1 and -2, are considered to be antagonists of the ubiquinone substrate. For piericidin A, it has been shown that it inhibits NADH:Q2 activity in a partially competitive manner. Contrary to type A, type B inhibitors, like the commonly used rotenone, have hydrogen-bonding acceptors only in the cyclic head of the molecule and are non-competitive towards UQ (ubiquinone), but are believed to displace the semiquinone intermediate during the catalysis (Fig. 2). Finally, inhibitors classified as type C, like stigmatellin and capsaicin, form a third group of hydrophobic CI inhibitors that are believed to act as antagonists of reduced ubiquinone (Degli Esposti 1998, Friedrich et al. 1994, Haefeli 2012) (Fig. 2). Competition studies with representatives of all three different types of inhibitors revealed that type A and B and type B and C, but not type A and C, compete with each other for binding. This led to a suggestion that all CI inhibitors acting at the ubiquinone binding pocket share a common binding domain with partially overlapping sites (Okun et al. 1999). Some inhibitors bind to the outside of the ubiquinone reduction site and do not fit the preceding classification. Examples of such compounds are ADP-ribose, which competes for substrate binding at the NADH site (Zharova and Vinogradov, 1997), and diphenyleneiodonium (DPI) that covalently binds to reduced flavin mononucleotide (FMN) in the hydrophilic part of the enzyme blocking the electron transfer to the Fe-S clusters (Majander et al., 1994). There are also new, commercially available insecticides/acaricides with potential to inhibit mitochondrial respiration such as benzimidazole, bullatacin, 6-chlorobenzothiadiazole, cyhalothrin, Fenazaquin Fenpyroximate, Hoe 110779, Pyridaben, Pyrimidifen, Sandoz 547A, Tebufenpyrad and Thiangazole (Greenamyre et al., 2001). It is clear that they are capable of inhibiting the mammalian CI of mitochondrial respiratory chain, by binding to and blocking ubiquinone-dependent NADH oxidation with high efficacy (Lummen, 1998).

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