Inhibition of mitochondrial DNA polymerase gamma (Pol gamma)

147127-20-6

SGOIRFVFHAKUTI-ZCFIWIBFSA-N

Tenofovir

Phosphonic acid, [[(1R)-2-(6-amino-9H-purin-9-yl)-1-methylethoxy]methyl]-

DTXSID9040132

202138-50-9

VCMJCVGFSROFHV-PFEQFJNWSA-N

Tenofovir disoproxil fumarate

DTXSID5050426

106941-25-7

SUPKOOSCJHTBAH-UHFFFAOYSA-N

Adefovir

DTXSID6046214

142340-99-6

WOZSCQDILHKSGG-UHFFFAOYSA-N

Adefovir dipivoxil

Propanoic acid, 2,2-dimethyl-, [[[2-(6-amino-9H-purin-9-yl)ethoxy]methyl]phosphinylidene]bis(oxymethylene) ester

DTXSID5046487

113852-37-2

VWFCHDSQECPREK-LURJTMIESA-N

Cidofovir

Phosphonic acid, [[(1S)-2-(4-amino-2-oxo-1(2H)-pyrimidinyl)-1-(hydroxymethyl)ethoxy]methyl]- (S)-1-(3-hydroxy-2-phosphonomethoxypropyl)cytosine (S)-HPMPC 1-(S)-(3-Hydroxy-2-phosphonylmethoxypropyl)cytosine 1-[(S)-3-Hydroxy-2-(phosphonomethoxy)propyl]cytosine Cidovir Phosphonic acid, [[2-(4-amino-2-oxo-1(2H)-pyrimidinyl)-1-(hydroxymethyl)ethoxy]methyl]-, (S)- Phosphonic acid, P-[[(1S)-2-(4-amino-2-oxo-1(2H)-pyrimidinyl)-1-(hydroxymethyl)ethoxy]methyl]- Vistide

DTXSID3043734

7440-43-9

BDOSMKKIYDKNTQ-UHFFFAOYSA-N

Cadmium

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

DTXSID1023940

7439-97-6

QSHDDOUJBYECFT-UHFFFAOYSA-N

Mercury

Liquid silver Mercure MERCURIC METAL TRIPLE DISTILLED mercurio Mercury element Quecksilber Quicksilver UN 2024 UN 2809

DTXSID1024172

7440-61-1

JFALSRSLKYAFGM-UHFFFAOYSA-N

Uranium

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

DTXSID1042522

7440-22-4

BQCADISMDOOEFD-UHFFFAOYSA-N

Silver

Ag Nanopaste NPS-J 90 Ag Sphere 2 Ag-C-GS Algaedyn Arctic Silver 3 Argentum Astroflake 5 Carey Lea silver Colloidal silver Dotite XA 208 Du Pont 4943 ECM 100AF4810 Enlight 600 Enlight silver plate 600 Epinall Finesphere SVND 102 Fordel DC FP 5369-502 Jelcon SH 1 Jungindai Takasago 300 KS (metal) LCP 1-19SFS Metz 3000-1 Nanomelt AGC-A Nanomelt Ag-XA 301 Nanomelt Ag-XF 301 Nanomelt Ag-XF 301H Nanopaste NPS-J 90 Perfect Silver Puff Silver X 1200 RT 1710S-C1 SD (metal) Shell Silver Silbest E 20 Silbest F 20 Silbest J 18 Silbest TC 12 Silbest TC 20E Silbest TC 25A Silbest TCG 1 Silbest TCG 7 Silcoat AgC 103 Silcoat AgC 2011 Silcoat AgC 209 Silcoat AgC 2190 Silcoat AgC 222 Silcoat AgC 2411 Silcoat AgC 74T Silcoat AgC-A Silcoat AgC-AO Silcoat AgC-B Silcoat AgC-BO Silcoat AgC-D Silcoat AgC-G Silcoat AgC-GS Silcoat AgC-L Silcoat AgC-O Silcoat GS Silcoat RF 200 Silflake 135 Silsphere 514 Silver atom Silver element Silver Flake 1 Silver Flake 25 Silver Flake 52 Silver Flake 7A SILVER FLAKES Silver metal Silvest TCG 11N Technic 299 Technic 450 Techno Alpha 175

DTXSID4024305

7440-38-2

RQNWIZPPADIBDY-UHFFFAOYSA-N

Arsenic

As Arsenic black ARSENIC METAL arsenico Grey arsenic UN 1558

DTXSID4023886

7440-57-5

PCHJSUWPFVWCPO-UHFFFAOYSA-N

Gold

AGC Micro Britecote Burnish Gold C.I. Pigment Metal 3 Colloidal gold Finesphere Gold W 011 Furuuchi 8560 Gold black Gold element Gold Flake Gold Leaf Keradec Palegold 5550 Perfect Gold Shell Gold Technic 504

DTXSID3064697

15663-27-1

DQLATGHUWYMOKM-UHFFFAOYSA-L

Cisplatin

Cis Platinum, diamminedichloro-, (SP-4-2)- Abiplatin Biocisplatinum Briplatin cis-DDP cis-Diaminedichloroplatinum cis-Diaminedichloroplatinum(II) cis-Diaminodichloroplatinum(II) cis-Diamminedichloroplatinum cis-Diamminedichloroplatinum(II) cis-Dichlorodiamineplatinum(II) cis-Dichlorodiammineplatinum cis-Dichlorodiammineplatinum(II) Cismaplat cis-Platin cisplatine cis-Platine cisplatino cis-Platinous diaminodichloride Cisplatinum cis-Platinum cis-Platinum diaminodichloride cis-Platinum II cis-Platinum(II) diaminodichloride cis-Platinum(II) diamminedichloride cis-Platinumdiamine dichloride cis-Platinumdiammine dichloride Cisplatyl Citoplatino Lederplatin lipoplatin Neoplatin NSC 119875 Platamine Platiblastin Platidiam Platinex Platinol Platinol AQ Platinoxan Platinum, diamminedichloro-, cis- Platistin Platosin SPI 077B103 cis-Dichlorodiamine platinum cis-Dichloro diaminoplatinum II

DTXSID4024983

7439-96-5

PWHULOQIROXLJO-UHFFFAOYSA-N

Manganese

Colloidal manganese Cutaval Manganese element Manganese fulleride Manganese metal alloy Manganese-55 manganeso

DTXSID2024169

7440-47-3

VYZAMTAEIAYCRO-UHFFFAOYSA-N

Chromium

Alpaste RRA 030 Alpaste RRA 050 Chromium element Chromium metal

DTXSID3031022

7440-66-6

HCHKCACWOHOZIP-UHFFFAOYSA-N

Zinc

Zn Asarco L 15 C.I. Pigment Black 16 Merrillite NC-Zinc Rheinzink Stapa TE Zinc AT UF (metal) UN 1436 Zinc dust Zinc Dust 3 Zinc Dust 500 mesh Zinc Dust LS 2 Zinc Dust MCS Zinc Flakes GTT ZINC METAL ZINC MOSSY ZINC STRIP ZINC, MOSSY Zincsalt GTT

DTXSID7035012

7429-90-5

XAGFODPZIPBFFR-UHFFFAOYSA-N

AZDRQVAHHNSJOQ-UHFFFAOYSA-N

Aluminum

Aisin Metal Fiber Al 050P-H24 ALC Fine Alcan XI 1391 Almi-Paste SSP 303AR Aloxal 3010 Alpaste 00-0506 Alpaste 0100M Alpaste 0100MA Alpaste 0100M-C Alpaste 0200M Alpaste 0200T Alpaste 0230M Alpaste 0230T Alpaste 0241M Alpaste 0300M Alpaste 0500M Alpaste 0539X Alpaste 0620MS Alpaste 0625TS Alpaste 0638-70C Alpaste 0700M Alpaste 0780M Alpaste 0900M Alpaste 100M Alpaste 100MS Alpaste 100MSR Alpaste 1100M Alpaste 1100MA Alpaste 1100N Alpaste 1100NA Alpaste 1109MA Alpaste 1109MC Alpaste 1200M Alpaste 1200T Alpaste 1260MS Alpaste 1500MA Alpaste 1700NL Alpaste 1810YL Alpaste 1830YL Alpaste 1900M Alpaste 1900XS Alpaste 1950M Alpaste 1950N Alpaste 210N Alpaste 2172EA Alpaste 2173 Alpaste 240T Alpaste 241M Alpaste 417 Alpaste 46-046 Alpaste 4-621 Alpaste 4919 Alpaste 50-63 Alpaste 50-635 Alpaste 51-148B Alpaste 51-231 Alpaste 5205N Alpaste 5207N Alpaste 52-509 Alpaste 52-568 Alpaste 5301N Alpaste 5302N Alpaste 53-119 Alpaste 5422NS Alpaste 54-452 Alpaste 54-497 Alpaste 54-542 Alpaste 55-516 Alpaste 55-519 Alpaste 55-574 Alpaste 5620NS Alpaste 5630NS Alpaste 5640NS Alpaste 56-501 Alpaste 5650NS Alpaste 5653NS Alpaste 5654NS Alpaste 5680N Alpaste 5680NS Alpaste 60-600 Alpaste 60-760 Alpaste 60-768 Alpaste 62-356 Alpaste 6340NS Alpaste 6370NS Alpaste 6390NS Alpaste 640NS Alpaste 65-388 Alpaste 66NLB Alpaste 710N Alpaste 7130N Alpaste 7160N Alpaste 7160NS Alpaste 725N Alpaste 740NS Alpaste 7430NS Alpaste 7580NS Alpaste 7620NS Alpaste 7640NS Alpaste 7670M Alpaste 7670NS Alpaste 7675NS Alpaste 7679NS Alpaste 7680N Alpaste 7680NS Alpaste 76840NS Alpaste 7730N Alpaste 7770N Alpaste 7830N Alpaste 8004 Alpaste 8080N Alpaste 8260NAR Alpaste 891K Alpaste 91-0562 Alpaste 92-0592 Alpaste 93-0595 Alpaste 93-0647 Alpaste 94-2315 Alpaste 95-0570 Alpaste 96-0635 Alpaste 96-2104 Alpaste 97-0510 Alpaste 97-0534 Alpaste AW 520B Alpaste AW 612 Alpaste AW 9800 Alpaste F 795 Alpaste FM 7680K Alpaste FX 440 Alpaste FX 910 Alpaste FZ 0534 Alpaste FZU 40C Alpaste G Alpaste HR 8801 Alpaste HS 2 Alpaste J Alpaste K 9800 Alpaste MC 666 Alpaste MC 707 Alpaste MF 20 Alpaste MG 01 Alpaste MG 1000 Alpaste MG 1300 Alpaste MG 500 Alpaste MG 600 Alpaste MH 6601 Alpaste MH 8801 Alpaste MH 9901 Alpaste MR 7000 Alpaste MR 9000 Alpaste MS 630 Alpaste N 1700NL Alpaste NS 7670 Alpaste O 100N Alpaste O 2130 Alpaste O 300M Alpaste P 0100 Alpaste P 1950 Alpaste S Alpaste SAP 110 Alpaste SAP 414P Alpaste SAP 550N Alpaste SCR 5070 Alpaste TCR 2020 Alpaste TCR 2060 Alpaste TCR 2070 Alpaste TCR 3010 Alpaste TCR 3030 Alpaste TCR 3040 Alpaste TCR 3130 Alpaste TD 200T Alpaste UF 500 Alpaste WB 0230 Alpaste WD 500 Alpaste WJP-U 75C Alpaste WX 0630 Alpaste WX 7830 Alpaste WXA 7640 Alpaste WXM 0630 Alpaste WXM 0650 Alpaste WXM 0660 Alpaste WXM 1415 Alpaste WXM 1440 Alpaste WXM 5422 Alpaste WXM 760b Alpaste WXM 7640 Alpaste WXM 7675 Alpaste WXM-T 60B Alpaste WXM-U 75 Alpaste WXM-U 75C Altop X Aluchrome Ultrafin Super Alumat 1600 Alumet H 30 aluminio Aluminium Aluminium Flake Aluminum 27 Aluminum atom Aluminum element Aluminum Flake PCF 7620 Aluminum granules ALUMINUM METAL/GRANULE ALUMINUM PASTE ALUMINUM PIGMENT ALUMINUM TURNINGS Alumi-paste 640NS Alumipaste 91-0562 Alumipaste 98-1822T Alumipaste AW 620 Alumipaste CR 300 Alumipaste GX 180A Alumipaste GX 201A Alumipaste HR 7000 Alumipaste HR 850 Alumipaste MG 11 Alumipaste MH 8801 Aquamet NPW 2900 Aquapaste 205-5 Aquasilver LPW Astroflake 40 Astroflake Black N 020 Astroflake Black N 070 Astroflake LG 40 Astroflake LG 70 Astroflake Silver N 040 Astroshine NJ 1600 Astroshine T 8990 Atomizalumi VA 200 C.I. PIGMENT METAL 1 Chromal IV Chromal X Decomet 1001/10 Decomet 2018/10 Decomet High Gloss Al 1002/10 Ecka AS 081 Eckart 9155 Eterna Brite 301-1 Eterna Brite 601-1 Eterna Brite 651-1 Eterna Brite EBP 251PA Eterna Brite Primier 251PA Ferro FX 53-038 Friend Color F 500GR-W Friend Color F 500WT Friend Color F 700RE-W Friend Color F 701RE-W Hi Print 60T High Print 60T Hisparkle HS 2 Hydro Paste 8726 Hydrolac WHH 2153 Hydrolan 3560 Hydrolux Reflexal 100 Hydroshine WS 1001 JISA 51010P Kryal Z Lansford 243 LE Sheet 800 Leafing Alpaste LG-H Silver 25 Lunar Al-V 95 Metallux 161 Metallux 2154 Metallux 2192 Metalure Metalure 55350 Metalure L 55350 Metalure L 59510 Metalure W 2001 Metapor Metasheen 1800 Metasheen HR 0800 Metasheen KM 100 Metasheen KM 1000 Metasheen Slurry 1807 Metasheen Slurry 1811 Metasheen Slurry KM 100 Metax G Metax S Mirror Glow 1000 Mirror Glow 600 Mirrorsheen Noral Aluminium Noral Ink Grade Aluminium Obron 10890 Offset FM 4500 Puratronic Reflexal 145 Reynolds 400 Reynolds 4-301 Reynolds 4-591 Reynolds 667 SAP 260PW-HS SAP-FM 4010 SBC 516-20Z Scotchcal 7755SE Serumekku Setanium 50MIS-H8 Siberline ET 2025 Siberline ST 21030E1 Silvar A Silver VT 522 Silverline SSP 353 Silvex 793-20C Sparkle Silver 3141ST Sparkle Silver 3500 Sparkle Silver 3641 Sparkle Silver 5000AR Sparkle Silver 516AR Sparkle Silver 5242AR Sparkle Silver 5245AR Sparkle Silver 5271AR Sparkle Silver 5500 Sparkle Silver 5745 Sparkle Silver 7000AR Sparkle Silver 7005AR Sparkle Silver 7500 Sparkle Silver 960-25E1 Sparkle Silver E 1745AR Sparkle Silver L 1526AR Sparkle Silver Premier 751 Sparkle Silver SS 3130 Sparkle Silver SS 5242AR Sparkle Silver SS 5588 Sparkle Silver SSP 132AR Special PCR 507 Splendal 6001BG Spota Mobil 801 SSP 760-20C Stapa Aloxal PM 2010 Stapa Aloxal PM 3010 Stapa Aloxal PM 4010 Stapa Hydrolac BG 8n.1 Stapa Hydrolac BGH Chromal X Stapa Hydrolac PM Chromal VIII Stapa Hydrolac W 60NL Stapa Hydrolac WH 16 Stapa Hydrolac WH 66NL Stapa Hydrolux 2192 Stapa Hydrolux 8154 Stapa IL Hydrolan 2192-55900G Stapa Metallic R 607 Stapa Metallux 1050 Stapa Metallux 211 Stapa Metallux 212 Stapa Metallux 2196 Stapa Metallux 274 Stapa Mobilux 181 Stapa Offset 3000 Stapa PV 10 Stapa VP 46432G Starbrite 2100 Super Fine 18000 Super Fine 22000 Supramex 2022 Toyo Aluminum 02-0005 Toyo Aluminum 93-3040 Transmet K 102HE Tufflake 3645 Tufflake 5843 UN 1396 US Aluminum 809 Valimet H 2 Valimet H 3 White Silver 7080N White Silver 7130N

DTXSID3040273

CL:1000507

CL kidney tubule cell

UBERON:0002113

UBERON kidney

GO:0005739

GO mitochondrion

GO:0008219

GO cell death

Q000633

MESH toxicity

WIKI increased

WIKI occurrence

WIKI functional change

Tenofovir

2017-10-25T07:45:32

Tenofovir disoproxil fumarate

2017-10-25T07:46:05

Adefovir

2017-10-25T07:46:24

Adefovir dipivoxil

2017-10-25T07:46:40

Cidofovir

2017-10-25T07:47:22

Cadmium

2017-10-25T08:33:12

Mercury

2016-11-29T18:42:19

Uranium

2021-08-05T14:28:50

Silver

2022-02-03T11:20:11

Arsenic

2021-04-27T00:15:21

Gold

2022-02-07T15:25:56

Nanoparticles and Micrometer Particles

2022-02-04T13:43:43

Cisplatin

2022-02-03T11:34:57

Manganese

2022-02-04T14:47:23

Chromium

2022-02-03T11:22:01

Zinc

2022-02-04T15:05:00

Aluminum

2022-02-04T14:42:11

WikiUser_15

ApacheUser Sprague-Dawley

WCS_9606

common toxicological species human

10090

NCBI mouse

10116

NCBI rat

WCS_7227

common ecological species Drosophila melanogaster

6239

NCBI Caenorhabditis elegans

WikiUser_4

Wikiuser: Blandesmann Human, rat, mouse

Inhibition of mitochondrial DNA polymerase gamma (Pol gamma)

Inhibition, mitochondrial DNA polymerase gamma (Pol gamma)

Molecular

AOPWiki 2017-10-25T07:48:27

2017-10-25T07:48:27

Depletion, mtDNA

Cellular

AOPWiki 2017-10-25T07:49:17

2017-10-25T07:49:17

Increase, Cytotoxicity (renal tubular cell)

Cellular

The renal proximal tubule is a crucial section of the nephron, responsible for the bulk of its reabsorption capabilities. About 60-70% of glomerular filtrate such as water, small molecules, and important ions, as well as nearly all the filtered amino acids, small peptides, and glucose are reabsorbed in the proximal tubule (Carson, 2019). The process of solute reabsorption is highly energetically expensive, making the proximal tubules the renal region of highest oxygen consumption. The microvilli, densely packed to form the brush border apical surface of the tubules, have abundant elongated mitochondria to sustain the energetic demand of their function (Carlson, 2019). The introduction of heavy metals into the kidneys causes aggregation in the proximal tubules due to their high mitochondrial content, leading to inhibition of the electron transport chain and reactive oxygen species (ROS) production. This area is particularly susceptible to heavy metal toxicity due to the abundance of mitochondria, as well as the fact that, regardless of toxicity, approximately 70% of cation absorption and transport passes through the proximal tubules (Barbier et al., 2005). Some heavy metal transport into the proximal tubules is conducted by MRP-1 and MRP-2 (ATP binding cassette-multidrug resistance proteins), and characterize toxicity by GSH depletion as some metals such as arsenic bind GSH and increased oxidative stress induced by free radicals (Sabath & Robles-Osorio, 2012). This oxidative stress causes disruption to mitochondrial homeostasis and mitophagy in proximal tubular epithelial cells by altering PPAR (peroxisome proliferator-activated receptor) (Small et al., 2018). At high enough concentrations of toxic heavy metals they can lead to cytotoxicity and cell death. An issue with assessment of kidney function is that the kidneys notoriously compensate for loss of function, leading to the appearance of adverse affects only at a late onset when there is very severe levels of damage (de Burbure et al., 2003). Cell Death and Cytotoxicity Cell death is a variety of processes defined by a cell ceasing to perform its function. This could happen by a variety of mechanisms. Apoptosis is a programmed physiological sequence leading to controlled cell death deemed necessary for the fitness and survival of the organism (cell is redundant, dysfunctional, cancerous, etc.) (Choi et al., 2019). Apoptosis, in the case of DNA damage, can be induced by free radicals produced as a result of heavy metal exposure, as shown in ex-vivo studies (Miller et al., 2002). Another cause by heavy metal exposure is physical and structural damage to mitochondria, damaging cellular metabolism and ATP production. There are many possible stressors that may lead to cell death, the effects exhibited depend on the cell type and the severity of the stress (Liu et al., 2018). Some modes of cell death include: apoptosis (programmed cell death), necrosis (uncontrolled cell death), and aging-caused cell death, known as senescent death  (Liu et al., 2018). <div>Apoptosis, also referred to as programmed cell death, is the predetermined procedure by which an organism disposes of cells that are no longer productive (Liu et al., 2018; Elmore, 2007). Apoptosis biochemically  manifests as cytoplasmic shrinkage, cytoskeleton collapse, chromatin condensation (pyknosis), nuclear fragmentation (karyorrhexis), mitochondrial dysfunction, cytochrome c release, altered Bcl-2 family protein expression or activation, plasma membrane blebbing, and in larger cells, the formation of apoptotic bodies. The surface of cells undergoing apoptosis is chemically altered to signal nearby cells and macrophages that then rapidly engulf them before they spill their contents (Alberts et al., 2014; Choi et al., 2019). Apoptosis occurs in three general phases: initiation, effector, and final. Variation can be seen as the initiation phase is dependant on stimuli, and there are two effector phase modes; an extrinsic and intrinsic pathways. Regardless of the pathway of the first 2 phases, the final stage of apoptosis is caspase-3 activation (Priant et al., 2019). The initiation and execution of apoptosis and other cell death processes is induced by the proteolytic activity of caspase as it cleaves the aspartic acid residues of proteins. The caspases can be broadly divided into two groups: those that are mainly involved in apoptosis (caspase-2, -3, -6, -7, -8, -9, and -10) and those related to caspase-1, whose primary role appears to be cytokine processing and pro-inflammatory cell death (caspase-1, -4, -5, -11, -12, -13, and -14). The apoptotic caspases can further be divided into initiator caspases (caspase-2, -8, -9, and -10) and executioner caspases (caspase-3, -6, and-7) (Fink & Cookson, 2005). Once the initial caspase activation occurs the resultant caspase cascade is irreversible (Alberts et al., 2014).</div> <div> </div> The extrinsic pathway, also known as the death receptor-mediated pathway, involves the ligation of death receptors determining the activation of caspase-8. Caspase-8 further activates downstream caspases leading to apoptosis (Priante et al., 2019). This pathway is triggered by extracellular signalling proteins binding to cell-surface death receptors. A well understood example of this process is the activation of the Fas receptor on the surface of a target cell by Fas ligand (FasL) on the surface of a cytotoxic lymphocyte (Alberts et al., 2014). In this process, the cytosolic Fas death receptor binds intracellular adaptor proteins. This complex then binds initiator, caspases, primarily caspase-8, forming a death-inducing signalling complex (DISC). The initiator caspases, once dimerized and activated in the DISC, activate downstream executioner caspases to induce apoptosis (Nair et al., 2014). In some cells, the extrinsic pathway recruits the intrinsic apoptotic pathway to amplify the caspase cascade. These pathways are linked by caspase-8, that triggers the caspase cascade and the protein, Bid (Priante et al., 2019; Alberts et al., 2014). Type I cells act independent of mitochondria for the induction of Fas death receptor-mediated apoptosis, and have therefore optimized the extrinsic pathway. Thymocytes or cells responsible for the immune system in general, for example, are expected to signal each other or target cells through membrane bound ligands, like FasL and TRAIL (Ozoren and El-Deiry, 2002). The intrinsic pathway is often referred to as the mitochondrial pathway of apoptosis. Pro-apoptotic Bcl-2 family proteins, Bax and Bak, create pores on the outer mitochondrial membrane, determining the release of apoptogenic factors, such as cytochrome c. In the cytosol, cytochrome c binds to, and stimulates, conformational modifications in the adaptor protein, Apaf-1, thus leading to the enrolment and activation of caspase-9. Caspase-9 further activates executioner caspases to elicit apoptosis (Priante et al., 2019). Type II cells are mitochondria-dependent, where the mitochondria are crucial to ensure successful apoptosis. For example, liver and kidney cells are responsible for the detoxification of the blood from chemicals toxicants, many of which are cytotoxic and genotoxic agents known to predominantly activate the intrinsic pathway (Ozoren and El-Deiry, 2002). In a study conducted by Eichler et al. (2006), cultured murine podocytes were incubated for three days with arsenite, cadmiuim, or mercury, as well as an equimolar combination of the three to test the modes and extent of apoptosis induced by the exposure. It was seen that the mix of metal exposure showed significantly fewer apoptotic affects, indicating an antagonistic affect of the metals over an additive or synergistic toxicity. It was also seen that the apoptosis observed in the separate metal tests showed a ~400% increase of caspase 8 activity as well as ~500% upregulation of Fas, factors of the extrinsic pathway. No significant change was seen to the intrinsic pathway factors. The results of this experiment indicate that heavy metals favour extrinsic apoptosis as their method of cytotoxicity. Necrosis is characterized as passive, accidental cell death resulting from environmental perturbation with uncontrolled release of inflammatory cellular contents (Fink & Cookson, 2005). Contrastingly, apoptosis is an active, intentional, programmed process of autonomous cellular dismantling that avoids eliciting inflammation. These modes would then be categorized into Accidental Cell Death (ACD) and Regulated Cell Death (RCD), respectively fitting necrosis and apoptosis (Choi et al., 2019). Necrosis biochemically manifests through plasma membrane rupture, cell swelling and lysis, energy decline, DAMP release, and emptying of cell contents (Choi et al., 2019; Thiebault et al., 2007). The caspases governing inflammatory cell death, such as necrosis, are caspases-1, -4, -5, -11, -12, -13, and -14 (Fink and Cookson, 2005). Cell fate could be decided by a number of factors. For instance, ATP is required for the execution of apoptosis, so, when lacking, apoptosis is disabled, making the mode of cell death ATP dependent (Shaki et al., 2012). Between apoptosis and necroptosis, cell fate is influenced primarily by the availability of caspase-8 and the cellular or X-linked inhibitors of apoptosis proteins (cIAP1, cIAP2, XIAP). Thiebault et al. (2007) studied the mechanism of cell mortality induced by uranium in NRK-52E cells and found that after low exposure to uranium (below the CI50 concentration, 500µL), apoptotic cell death was observed, whereas higher exposure to uranium resulted in necrotic cell death. Multiple types of death can be observed simultaneously in tissues exposed to the same stimulus, and the local intensity of a particular stimulus may influence the cell death mechanism (Fink and Cookson, 2005).

<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> Kidney function assay Measuring total urinary protein, albumin, transferrin, b2-microglobulin, retinolbinding protein, brush border tubular antigens, N-acetyl-b-Dglucosaminidase activity, serum and urinary creatine   (de Burbure et al., 2003)</td> <td>“All analyses of a given parameter were performed under similar experimental conditions in the same laboratories within 6mo of collection. Total urinary protein (Prot-T-U) was determined by the Coomassie blue G250 binding method. Albumin (Alb-U), transferrin (Transf-U), β2-microglobulin (β2m-U), and retinolbinding protein (RBP-U) in urine were quantified by latex immunoassay (Bernard & Lauwerys, 1983). Acceptable limits for precision and accuracy of measurements and external quality controls were the same as those described in the Cadmibel study (Lauwerys et al., 1990). The brush border tubular antigens (BBA-U) were analyzed by a sandwich enzyme-linked immunoassay using monoclonal antibodies (Mutti et al., 1985). The total activity of N-acetyl-β-Dglucosaminidase (NAG-T-U) in urine was determined colorimetrically using a kit (PPR Diagnostics Ltd.) as described elsewhere (Price et al., 1996). Only total NAG (NAG-T) was used for the purpose of this study. Serum and urinary creatinine (Creat-U) were measured by the methods of Heinegard and Tiderström (1973), and Jaffé, respectively (Henry, 1965).” (de Burbure et al., 2003)</td> <td>“The soil contamination in the area varied from 100 to 1700ppm lead (with values higher than 1000ppm in the immediate vicinity of the factories), 0.7 to 233ppm cadmium, and 101 to 22,257ppm zinc, with the highest concentrations being recorded within 500 m of the 2 factories”</td> <td> </td> </tr> <tr> <td> N-ACETYL-b-D-GLUCOSAMINIDASE (NAG) ASSAY Measuring NAG urinary content (Lim et al., 2016)</td> <td>“Urinary NAG activity was measured by using NAG Quantitative Kit (Shionogi, Osaka, Japan). After storing a synthetic substrate solution (1 mL) at 37°C for five minutes, the solution was mixed with the supernatant of the urine samples (50 mL) received after centrifugation. After storing it at 37°C for 15 min, stopping solution (2 mL) was added to and mixed with it. By using a spectrophotometer, its fluorescence intensities were measured with a wavelength of 580 nm (<a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4780232/#b13-tr-32-057">13</a>,<a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4780232/#b14-tr-32-057">14</a>). Urinary β2-MG was measured by using Enzygnost β2-MG Micro Kit (Behring Institute, Mannheim, Germany). Its method used the principle of solid phase enzyme-linked immunosorbent assay (ELISA). Monoclonal anti-β2-MG antibody and anti-2-MG-horseradish peroxidase conjugate solution were used. After that, color intensities were measured with a wavelength of 450 nm by using a spectrophotometer (<a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4780232/#b13-tr-32-057">13</a>,<a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4780232/#b14-tr-32-057">14</a>).” (Lim et al., 2016)</td> <td>Cd & Pb</td> <td>Fast, easy, accurate</td> </tr> <tr> <td> MTT Assay (cytotoxicity) Measuring Cell Viability (Thiebault et al., 2007; Shaki et al., 2012)</td> <td>This assay is a quantitative and sensitive method of detection of cell proliferation, measuring the growth rate of cells via activity and absorbance. It relies on the reduction of MTT (yellow, water-soluble tetrazolium dye) by mitochondrial dehydrogenases, to purple colored formazan crystals. The samples are then analyzed via spectrophotometry (550 nm). This assay can also be used to asses electron transport function.</td> <td> 50, 100 and 500 μM of uranyl acetate; 0-1000µM U</td> <td> Long Easy/Difficult High accuracy (mathematical measurement) Medium Precision</td> </tr> <tr> <td> LDH Cytotoxicity Assay Measuring Necrosis via Lactate Dehydrogenase release (Thiebault et al., 2007)</td> <td>LDH is released into extracellular space when the plasma membrane is damaged. To detect the leakage of LDH into cell culture medium as a measurement of membrane integrity, a tetrazolium salt is used in this assay. LDH oxidizes lactate to generate NADH, which then reacts with WST to generate a yellow colour. LDH activity can then be quantified by spectrophotometer or plate reader. </td> <td>15, 30 µM Cd</td> <td>Fast, easy, high accuracy</td> </tr> <tr> <td> Caspase-3 and -8 colorimetric assay, Caspase-9 fluoresceine assay Measuring apoptosis initiation and execution via caspases 3, 8, 9 activity (Thiebault et al., 2007)</td> <td>After cell lysate centrifugation, 10 µL of the supernatant was incubated with 80 µL of the caspase assay buffer and 10 µL of the colorimetric caspase-3 (Acetyl-asp-glu-val-asp-p-nitroanilide) or caspase-8 (Acetyl-ile-glu-thr-asp-p-nitroaniline) substrate. Plates were incubated for 90 min at 37° C and absorbance was read at 405 nm with a Statfax-2100 microplate reader. Fluorescence intensity of cell suspensions measuring caspase-9 activity was measured at an excitation wavelength of 490 nm and an emission wavelength of 530 nm with fluorescence spectrophotometer.</td> <td>0-800µM U</td> <td>Long, difficult, high accuracy</td> </tr> </tbody> </table> “Techniques such as micropuncture, microinjection [1, 6, 18] and microperfusion of isolated tubules [14] have made it possible to map the reabsorption of the heavy metals along the different segments of the nephron.” (Barbier et al., 2005) “Pb2+ , Hg2+ induced glomerular and tubular damage characterized by a reduced GFR, glycosuria, proteinuria and a rapid obstruction of the tubular system [13]” (Barbier et al., 2005) “Concerning chronic intoxication, most heavy metals (Cd2+ , Hg2+ , Pb2+ ) induced a Fanconi syndrome characterized by a decrease of the GFR, an increase in urinary flow rate, proteinuria, glycosuria, aminoaciduria and excessive loss of major ions.” (Barbier et al., 2005) “In the proximal tubule, Cd2+ has been shown to decrease phosphate and glucose transport by inhibiting the NaPi and the Na/glucose cotransporters respectively.” (Barbier et al., 2005) “In the kidney, Cd mainly affects PCT cells. This damage manifests clinically as low molecular weight proteinuria, aminoaciduria, bicarbonaturia, glycosuria and phosphaturia. Tubular damage markers such as alpha-1-microglobulin, beta-2-microglobulin, NAG and KIM-1 (kidney injury molecule-1) are useful in detecting early tubular damage.” (Sabath & Robles-Osorio, 2012)

All animals with kidneys containing renal proximal tubules.

CL:1000507

CL kidney tubule cell

Alberts, B., Johnson, A., Lewis, J., Raff, M., Roberts, K., & Walter, P. (2014). Molecular biology of the cell. New York: Garland Science. Retrieved from <a href="https://www.ncbi.nlm.nih.gov/books/NBK21054/" target="_blank">https://www.ncbi.nlm.nih.gov/books/NBK21054/</a> Barbier, O., Jcquillet, G., Tauc, M., Cougnon, M., & Poujeol, P. (2005). Effect of heavy metals on, and handling by, the  kidney. Nephron Physiology, 99, 105-110. doi:10.1159/000083981 Belyaeva, E. A., Sokolova, T. V., Emelyanova, L. V., & Zakharova, I. O. (2012). Mitochondrial electron transport chain in heavy metal-induced neurotoxicity : Effects of cadmium , mercury , and copper. The scientific world, 2012, 1-14. doi:10.1100/2012/136063 Carlson, B. M. (2019). The urinary system. The Human Body Academic Press, , 357-372. doi:https://doi.org/10.1016/B978-0-12-804254-0.00013-2 Choi, M. E., Price, D. R., Ryter, S. W., & Choi, A. M. K. (2019). Necroptosis: A crucial pathogenic mediator of human disease. JCI Insight, 4(15), 1-16. doi:10.1172/jci.insight.128834 Chomchan, R., Siripongvutikorn, S., Malyam, P., Saibandith, B., & Puttarak, P. (2018). Protective effect of selenium-enriched ricegrass juice against cadmium-induced toxicity and DNA damage in HEK293 kidney cells. Foods, 7, 81. doi:10.3390/foods7060081 De Burbure , C., Buchet , J., Bernard , A., Leroyer , A., Nisse , C., Haguenoer , J., Bergamaschi E., & Mutti, A. (2003). Biomarkers of Renal Effects in Children and Adults with Low Environmental Exposure to Heavy Metals. Journal of Toxicology and Environmental Health Part A, 66:9, 783-798, DOI: 10.1080/15287390306384 Fink, S. L., & Cookson, B. T. (2005). Apoptosis, pyroptosis, and necrosis: Mechanistic description of dead and dying eukaryotic cells. Infection and Immunity, 73(4), 1907-1916. doi:73/4/1907 [pii] Guéguen, Y., Suhard, D., Poisson, C., Manens, L., Elie, C., Landon, G., . . . Tessier, C. (2015). Low-concentration uranium enters the HepG2 cell nucleus rapidly and induces cell stress response. Toxicology in Vitro, 30, 552-560. doi:10.1016/j.tiv.2015.09.004 Hao, Y., Huang, J., Liu, C., Li, H., Liu, J., Zeng, Y., . . . Li, R. (2016). Differential protein expression in metallothionein protection from depleted uranium-induced nephrotoxicity. Scientific Reports, doi:10.1038/srep38942 Hao, Y., Ren, J., Liu, C., Li, H., Liu, J., Yang, Z., . . . Su, Y. (2014). Zinc protects human kidney cells from depleted uranium induced apoptosis. Basic & Clinical Pharmacology & Toxicology, 114, 271-280. doi:10.1111/bcpt.12167 Hinkle, P. M., Kinsella, P. A., & Osterhoudt, K. C. (1987). Cadmium uptake and toxicity via voltage-sensitive calcium channels. Journal of Biological Chemistry, 262(34), 16333-16337. 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:10.1016/j.toxlet.2009.03.014 Lim, H., Lim, J. A., Choi, J. H., Kwon, H. J., Ha, M., Kim, H., & Park, J. D. (2016). Associations of Low Environmental Exposure to Multiple Metals with Renal Tubular Impairment in Korean Adults. Toxicological research, 32(1), 57–64. doi:10.5487/TR.2016.32.1.057 Liu, S., Xu, L., Zhang, T., Ren, G., & Yang, Z. (2010). Oxidative stress and apoptosis induced by nanosized titanium dioxide in PC12 cells. Toxicology, 267, 172-177. doi:10.1016/j.tox.2009.11.012 Liu, X., Yang, W., Guan, Z., Yu, W., Fan, B., Xu, N., & Liao, D. J. (2018). There are only four basic modes of cell death, although there are many ad-hoc variants adapted to different situations. Cell & Bioscience, 8(1), 6. doi:10.1186/s13578-018-0206-6 Miller, A. C., Stewart, M., Brooks, K., Shi, L., & Page, N. (2002). Depleted uranium-catalyzed oxidative DNA damage: Absence of significant alpha particle decay. Journal of Inorganic Biochemistry, 91(1), 246-252. doi:10.1016/S0162-0134(02)00391-4 Miyayama, T., Arai, Y., Suzuki, N., & Hirano, S. (2013). Mitochondrial electron transport is inhibited by disappearance of metallothionein in human bronchial epithelial cells follwoing exposure to silver nitrate. Toxicology, 305, 20-29. doi:10.1016/j.tox.2013.01.004 Muller, D., Houpert, P., Cambar, J., & Henge-Napoli, M. (2006). Role of the sodium-dependent phosphate co-transporters and of the phosphate complexes of uranyl in the cytotoxicity of uranium in LLC-PK1 cells. Toxicology and Applied Pharmacology, 214, 166-177. doi:10.1016/j.taap.2005.12.016 Mezynska, M., Brzoska, M. M., Rogalska, J., & Galicka, A. (2019). Extract from aronia melanocarpa L. berries protects against cadmium-induced lipid peroxidation and oxidative damage to proteins and DNA in the liver: A study using a rat model of environmental human exposure to this xenobiotic. Nutrients, 11, 758. doi:10.3390/nu11040758 Nair, P., Lu, M., Petersen, S., & Ashkenazi, A. (2014). Chapter five - apoptosis initiation through the cell-extrinsic pathway. Methods in Enzymology, 544, 99-128. doi:<a href="https://doi.org/10.1016/B978-0-12-417158-9.00005-4">https://doi.org/10.1016/B978-0-12-417158-9.00005-4</a> Ozoren, N., & El-Deiry, W. S. (2002). WS. Defining characteristics of types I and II apoptotic cells in response to TRAIL.4(6), 551-557. doi:10.1038/sj.neo.7900270 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 Priante, G., Gianesello, L., Ceol, M., Del Prete, D., & Anglani, F. (2019). Cell death in the kidney. International Journal of Molecular Sciences, 20(14), 3598. doi: 10.3390/ijms20143598. doi:10.3390/ijms20143598 [doi] Rouas, C., Bensoussan, H., Suhard, D., Tessier, C., Grandcolas, L., Rebiere, F., . . . Gueguen, Y. (2010). Distribution of soluble uranium in the nuclear cell compartment at subtoxic concentrations. Chemical Research in Toxicology, 23(12), 1883-1889. doi:10.1021/tx100168c Sabath, E., & Robles-Osorio, M. L. (2012). Renal health and the environment: Heavy metal nephrotoxicity. Revista Nefrologia, doi:10.3265/Nefrologia.pre2012.Jan.10928 Santos, N. A. G., Catão, C. S., Martins, N. M., Curti, C., Bianchi, M. L. P., & Santos, A. C. (2007). Cisplatin-induced nephrotoxicity is associated with oxidative stress, redox state unbalance, impairment of energetic metabolism and apoptosis in rat kidney mitochondria. Archives of Toxicology, 81(7), 495-504. doi:10.1007/s00204-006-0173-2 Shaki, F., Hosseini, M. J., Ghazi-Khansari, M., & Pourahmad, J. (2012). Toxicity of depleted uranium on isolated rat kidney mitochondria. Biochimica Et Biophysica Acta - General Subjects, 1820(12), 1940-1950. doi:10.1016/j.bbagen.2012.08.015 Small, D. M., Sanchez, W. Y., Roy, S. F., Morais, C., Brooks, H. L., Coombes, J. S., . . . Gobe, G. (2018). N-acetyl-cysteine increases cellular dysfunction in progressive chronic kidney damage after acute kidney injury by dampening endogenousantioxidant responses. American Physiological Society - Renal Physiology, 314, F956-F968. doi:10.1152/ajprenal.00057.2017 Spreckelmeyer, S., Estrada-Ortiz, N., Prins, G. G. H., van der Zee, M., Gammelgaard, B., Sturup, S., . . . Casini, A. (2017). On the toxicity and transportation mechanisms of cisplatin in kidney tissues in comparison to a gold-based cytotoxic agent. Metallomics, 9, 1786. doi:10.1039/c7mt00271h Tad Eichler, Qing Ma, Caitlin Kelly, Jaya Mishra, Samir Parikh, Richard F. Ransom, Prasad Devarajan, William E. Smoyer, Single and Combination Toxic Metal Exposures Induce Apoptosis in Cultured Murine Podocytes Exclusively via the Extrinsic Caspase 8 Pathway, Toxicological Sciences, Volume 90, Issue 2, April 2006, Pages 392–399, <a href="https://doi.org/10.1093/toxsci/kfj106">https://doi.org/10.1093/toxsci/kfj106</a>Elmore, S. (2007). Apoptosis: A review of programmed cell death. Toxicologic Pathology, 35(4), 495-516. doi:779478428 [pii] Thiébault, C., Carrière, M., Milgram, S., Simon, A., Avoscan, L., & Gouget, B. (2007). Uranium induces apoptosis and is genotoxic to normal rat kidney (NRK-52E) proximal cells. Toxicological Sciences : An Official Journal of the Society of Toxicology, 98(2), 479-487. doi:kfm130 [pii] Turk, E., Kandemir, F. M., Yildirim, S., Caglayan, C., Kucukler, S., & Kuzu, M. (2019). Protective effect of hesperidin on sodium arsenite-induced nephrotoxicity and hepatotoxicity in rats. Biological Trace Element Research, 189, 95-108. doi:10.1007/s12011-018-1443-6 Yu, L., Li, W., Chu, J., Chen, C., Li, X., Tang, W., . . . Xiong, Z. (2021). Uranium inhibits mammalian mitochondrial cytochrome c oxidase and ATP synthase. Environmental Pollution, 271, 116377. doi:10.1016/j.envpol.2020.116377 Zhang, H., Chang, Z., Mehmood, K., Abbas, R. Z., Nabi, F., Rehman, M. U., . . . Zhou, D. (2018). 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   AOPWiki 2016-11-29T18:41:26

2022-03-03T15:14:42

Occurrence, Kidney toxicity

Organ

The kidneys are a crucial site of regulation of divalent cation levels in the plasma through filtration, reabsorption, and concentration (cite). On top of their excretion capabilities, the kidneys are also responsible for the production of hormones crucial for hematologic, cardiovascular, and skeletal muscle homeostasis (Bonventre et al., 2010). Nephrons are the functional units of the kidney and each kidney is made up of approximately 1 million nephrons (Bonventre et al., 2010). The nephrons are vital in reabsorption of these cations where 70% of transport has been shown to occur in the proximal tubule (Barbier et al., 2005). The kidneys are thought to be very susceptible to toxicity due to the increased concentration through their filtering structures with the tubular uptake mechanisms, specifically those of the proximal tubule, magnifying intracellular concentrations (Bonventre et al., 2010; Weber et al., 2017). Commonly, biomarkers like serum creatinine (sCr) and blood urea nitrogen (BUN) are utilized to identify kidney toxicity; however, these markers have been identified as nonspecific to the area of the kidney and slow in identification. Bonventre et al. (2010) has explored other biomarkers that may be used to identify segment specific injury. Proximal tubule injury can be identified using: albumin, RPB, NAG, clusterin, osteopontin, a1-microglobulin, and many others. Glomerulus damage can be identified through urinary Cystatin C, b2-microglobulin, a1-microglobulin, albumin, and more (Bonventre et al., 2010). These biomarkers do show some overlap between regions and can indicate damage to various areas of the nephron, though it is important to note the development of these specific techniques and therefore, the ability to develop more tailored and earlier identifying testing procedures. Since there are many essential metals for cellular function, there are also many transporters responsible for facilitating ionic entry into the cell and the designated cellular compartment (cite). Some of these transporters are very specific to a given metal and some are more diverse in the metals they handle, therefore, these transporters can facilitate the transport of toxic metals into the cell, often through mimickery exhibited by those metals (Ballatori, 2002). DMT1 (divalent metal transporter 1) is a strong example of such transporters. The introduction of toxic divalent cations (Cd2+, Pb2+, Pt2+, etc.) is highly problematic in the kidneys due to increased toxicity and occupancy of DMT1 limiting the transport of essential trace elements. DMT1 is an essential transport molecule that is highly expressed in the kidneys, and is responsible for transport of essential trace divalent cations, as well as highly toxic ones; this competition increases strain on the kidneys exposed to toxic heavy metals (Barbier et al., 2005; Ballatori, 2002). DMT1 has been shown to transport Fe, Zn, Mn, Co, Cd, Cu, Ni, and Pb via a proton-coupled, membrane potential dependant mechanism (Ballatori, 2002). Some toxic metals can also enter a cell by forming complexes that mimic endogenous molecules in their structure. Arsenate and vanadate, for example, act as phosphate mimics both for transport and metabolism, assaulting cellular function by the same mechanism as their initial entry; cromate, selenite and molybdate mimic sulfate in a similar way (Ballatori, 2002). Many of the identified transporters fooled by this mimicry have been localized to the brush border membrane of the renal proximal tubule and epithelial cells. Some divalent metals such as Cd, Ba, and Sr have been shown to enter cells through voltage gated calcium channels. Another important example focused on by Ballatori (2002) is the action of inorganic mercury and methyl mercury (MeHg) that were shown to have high affinity for reduced sulfhydryl groups. These groups are seen on the amino acid cysteine, and importantly on glutathione (GSH), a vital enzymatic antioxidant. MeHg mimics methionine to enter the cell, after which it binds to GSH, and interferes with ATP production (Ballatori, 2002). Uranium has been shown to enter the blood rapidly and then either form stable complexes with plasma proteins, due to its high affinity for phosphate, carboxyl and hydroxyl groups, or binds to bicarbonate in the blood (Keith et al., 2013). In the kidneys, uranium can be released from bicarbonate to combine with other small proteins in the kidney tubular walls, disrupting cellular function (Keith et al., 2013). Uranium has been seen to enter the glomerulus, where it is filtered, via endocytosis as UO+2 binding to anionic sites of proximal tubular epithelial brush borders (Shaki et al., 2012). To further understand the mode of action of heavy metals within the kidneys, many studies have been conducted to determine the specific region primarily damaged. It is also important to note that variation of results may be found in some studies as experimental conditions as well as other factors may influence the mode of action of some metals. Zamora et al. (1998) found that kidney function decrease and cytotoxicity increase were correlated with uranium ingestion. However, no glomerular injury was detected, indicating that chronic uranium ingestion in rats (0.004 µg/kg to 9 µg/kg body weight) damages the proximal tubule and not the glomerulus (Zamora et al., 1998). Homma-Takeda et al. (2013) identifies the kidneys as the major site of depleted uranium toxicity. Studying the kidneys of rats of varying ages, exposed to 0.1-2mg/kg uranyl acetate, they found that the younger kidneys did not flush the uranium out as well. Accumulation of uranium and its damages was seen in the S3 segment of the proximal tubules (Homma-Takeda et al., 2013). Shaki et al. (2012), assessed the mechanism of depleted uranium-induced nephrotoxicity that revealed damage to the mitochondria isolated from uranyl acetate treated rat kidney cells. The damage included oxidative stress, mitochondrial swelling, mitochondrial membrane potential collapse, cytochrome C release, impaired ATP production, and damage to the electron transport chain complexes. Utilizing rat renal brush border vesicles, Goldman et al. (2006) found that exposure to uranyl acetate induced decreased rates of glucose transport, in part due to a decreased number of sodium-coupled glucose transporters; this decreased the ability of the kidneys to reabsorb glucose properly. Berradi et al. (2008) assessed the red blood cell (RBC) count of rats drinking water containing 40mg DU/L and found that chronic exposure to DU causes RBC reduction, pointing to nephrotoxicity as the kidneys play a major role in RBC synthesis. Heavy metals consistently aggregate in the kidneys, and more specifically in the S3 segment of the proximal tubules. Evidence also suggests that uranium and other heavy metals induce nephrotoxicity after endocytosis into cells by disrupting the electron transport chain, inducing oxidative stress. The oxidative stress leads to mitochondrial dysfunction followed by, apoptosis at low doses of uranium and necrosis at  high doses of uranium. Finally, this induces renal injury and tissue damage to the proximal tubules, or nephrotoxicity.

<table border="1" cellpadding="1" cellspacing="1" style="width:500px"> <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> Kidney Function Assay Measuring total urinary protein, albumin, transferrin, b2-microglobulin, retinolbinding protein, brush border tubular antigens, N-acetyl-b-Dglucosaminidase activity, serum and urinary creatine (de Burbure et al., 2003) </td> <td>“All analyses of a given parameter were performed under similar experimental conditions in the same laboratories within 6mo of collection. Total urinary protein (Prot-T-U) was determined by the Coomassie blue G250 binding method. Albumin (Alb-U), transferrin (Transf-U), β2-microglobulin (β2m-U), and retinolbinding protein (RBP-U) in urine were quantified by latex immunoassay (Bernard & Lauwerys, 1983). Acceptable limits for precision and accuracy of measurements and external quality controls were the same as those described in the Cadmibel study (Lauwerys et al., 1990). The brush border tubular antigens (BBA-U) were analyzed by a sandwich enzyme-linked immunoassay using monoclonal antibodies (Mutti et al., 1985). The total activity of N-acetyl-β-Dglucosaminidase (NAG-T-U) in urine was determined colorimetrically using a kit (PPR Diagnostics Ltd.) as described elsewhere (Price et al., 1996). Only total NAG (NAG-T) was used for the purpose of this study. Serum and urinary creatinine (Creat-U) were measured by the methods of Heinegard and Tiderström (1973), and Jaffé, respectively (Henry, 1965).” (de Burbure et al., 2003)</td> <td>“The soil contamination in the area varied from 100 to 1700ppm lead (with values higher than 1000ppm in the immediate vicinity of the factories), 0.7 to 233ppm cadmium, and 101 to 22,257ppm zinc, with the highest concentrations being recorded within 500 m of the 2 factories”</td> <td> </td> </tr> <tr> <td> NAG Assay Measuring N-acetyl-b-D-Glucosaminidase urinary content (Lim et al., 2016)</td> <td>“Urinary NAG activity was measured by using NAG Quantitative Kit (Shionogi, Osaka, Japan). After storing a synthetic substrate solution (1 mL) at 37°C for five minutes, the solution was mixed with the supernatant of the urine samples (50 mL) received after centrifugation. After storing it at 37°C for 15 min, stopping solution (2 mL) was added to and mixed with it. By using a spectrophotometer, its fluorescence intensities were measured with a wavelength of 580 nm (<a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4780232/#b13-tr-32-057" style="color:#0563c1; text-decoration:underline">13</a>,<a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4780232/#b14-tr-32-057" style="color:#0563c1; text-decoration:underline">14</a>). Urinary β2-MG was measured by using Enzygnost β2-MG Micro Kit (Behring Institute, Mannheim, Germany). Its method used the principle of solid phase enzyme-linked immunosorbent assay (ELISA). Monoclonal anti-β2-MG antibody and anti-2-MG-horseradish peroxidase conjugate solution were used. After that, color intensities were measured with a wavelength of 450 nm by using a spectrophotometer (<a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4780232/#b13-tr-32-057" style="color:#0563c1; text-decoration:underline">13</a>,<a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4780232/#b14-tr-32-057" style="color:#0563c1; text-decoration:underline">14</a>).” (Lim et al., 2016)</td> <td> Pb: 0.0221ppm (converted from blood Pb µg/dL) Cd: 1.08ppm (converted from Urinary Cd μg/g creatinine)</td> <td>Fast, easy, accurate</td> </tr> <tr> <td> Kidney Dysfunction Assay Measuring BUN and creatinine serum blood levels (Shaki et al., 2012)</td> <td>“For studies in vivo rats were fasted overnight, then animals were divided into two groups, with six rats in each group. The control group (vehicle) received a single intraperitoneal (i.p.) injection of saline solution (1 ml per 100 g body weight). Uranyl acetate was dissolved in normal saline. Rats were treated with single intraperitoneal (i.p.) injections of UA in doses 0.5, 1 and 2 mg/kg body weight. These dosages was selected based on previous studies [28], which is sufficient to induce oxidative stress in kidney without causing death and none died within the duration of experiments. Blood urea nitrogen (BUN) and creatinine, marker of kidney dysfunction, were determined by commercial reagents (obtained from Parsazmoon Co., Iran). The rats were killed by decapitation 24 h after injection. The kidney were immediately removed and placed in ice-cold mitochondria isolation medium (0.225 M D-mannitol, 75 mM sucrose, and 0.2 mM EDTA, pH=7.4)” (Shaki et al., 2012)</td> <td>Control, 0.5, 1, 2 mg/kg Uranyl Acetate (UA) </td> <td>Fast, easy, medium accuracy </td> </tr> </tbody> </table>

Higher order animals (mammals) with functional and complete kidneys

UBERON:0002113

UBERON kidney

Not Specified Not Specified

Al Dera, H. S. (2016). Protective effect of resveratrol against aluminum chloride induced nephrotoxicity in rats. Saudi Med J, 37(4), 369-378. doi:10.15537/smj.2016.4.13611 Andjelkovic, M., Djordjevic, A. B., Antonijevic, E., Antonijevic, B., Stanic, M., Kotur-Stevuljevic, J., . . . Bulat, Z. (2019). Toxic effect of acute cadmium and lead exposure in rat blood, liver, and kidney. International Journal of Environmental Research and Public Health, 16, 247. doi:10.3390/ijerph16020274 Arzuaga , X., Rieth, S. H., Bathija, A. & Cooper, G. S. (2010) Renal Effects of Exposure to Natural and Depleted Uranium: A Review of the Epidemiologic and Experimental Data, Journal of Toxicology and Environmental Health, Part B, 13:7-8, 527-545, DOI:10.1080/10937404.2010.509015 Ballatori, N. (2002). Transport of toxic metals by molecular mimicry. Environmental Health Perspectives, 110, 689-694. doi:10.1289/ehp.02110s5689 Barnes, P., Yeboah, J. K., Gbedema, W., Saahene, R. O., & Amoani, B. (2020). Ameliorative effect of vernonia amygdalina plant extract on heavy metal-induced LIver and kidney dysfunction in rats. Advances in Pharmacological and Pharmaceutical Sciences, 2020, 1-7. doi:10.1155/2020/2976905 Barbier, O., Jcquillet, G., Tauc, M., Cougnon, M., & Poujeol, P. (2005). Effect of heavy metals on, and handling by, the  kidney. Nephron Physiology, 99, 105-110. doi:10.1159/000083981 Bonventre, J. V., Vaidya, V. S., Schmouder, R., Feig, P., & Dieterle, F. (2010). Next-generation biomarkers for detecting kidney toxicity. Nature biotechnology, 28(5), 436–440. <a href="https://doi.org/10.1038/nbt0510-436" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1038/nbt0510-436</a> Brzoska, M. M., Kaminski, M., Supernak-Bobko, D., Zwierz, K., & Moniuszko-Jakoniuk, J. (2003). Changes in the strucutre and function of the kidney of rats chronically exposed to cadmium. I. biochemical and histopathological studies. Arch.Toxicol., 77, 344-352. doi:10.1007/s00204-003-0451-1 Buelna-Chontal, M., Franco, M., Hernandez-Esquivel, L., Pavon, N., Rodriguez-Zalvala, J. S., Correa, F., . . . Chavez, E. (2017). CDP-choline circumvents mercury-induced mitochondrial damage and renal dysfunction. Cell Biology International, 41, 1356-1366. doi:10.1002/cbin.10871 Chtourou, Y., Garoui, E. m., Boudawara, T., & Zeghal, N. (2014). Protective role of silymarin against manganese-induced nephrotoxicity and oxidative stress in rat. Environ Toxicol, 29, 1147-1154. doi:10.1002/tox.21845 Durante, P., Romero, F., Perez, M., Chavez, M., & Parra, G. (2010). Effect of uric acid on nephrotoxicity induced by mercuric chloride in rats. Toxicology and Industrial Health, 26(3), 163-174. doi:10.1177/0748233710362377 García-Niño, W. R., Tapia, E., Zazueta, C., Zatarain-Barrón, Z. L., Hernández-Pando, R., Vega-García, C. C., & Pedraza-Chaverrí, J. (2013). Curcumin pretreatment prevents potassium dichromate-induced hepatotoxicity, oxidative stress, decreased respiratory complex I activity, and membrane permeability transition pore opening. Evidence-Based Complementary and Alternative Medicine, (424692), 1-19. doi:10.1155/2013/424692 Goldman, M., Yaari, A., Doshnitzki, Z., Cohen-Luria, R., & Moran, A. (2006). Nephrotoxicity of uranyl acetate: Effect on rat kidney brush border membrane vesicles. Archives of Toxicology, 80(7), 387-393. doi:10.1007/s00204-006-0064-6 Homma-Takeda S, Kokubo T, Terada Y, Suzuki K, Ueno S, Hayao T, Inoue T, Kitahara K, Blyth BJ, Nishimura M, Shimada Y. Uranium dynamics and developmental sensitivity in rat kidney. J Appl Toxicol. 2013 Jul;33(7):685-94. doi: 10.1002/jat.2870. Epub 2013 Apr 26. PMID: 23619997. Keith, S., Faroon, O., N., R., Scinicariello, F., Wilbur, S., Ingerman, L., . . . Diamond, G. (2013). Toxicological profile for uranium. U.S. Department of Health and Human Services. Agency for Toxic Substances and Disease Registry. Kharroubi, W., Dhibi, M., Mekni, M., Haouas, Z., Chreif, I., Neffati, F., . . . Sakly, R. (2014). Sodium arsenate induce changes in fatty acids profiles and oxidative damage in kidney of rats. Environ Sci Pollut Res, 21, 12040-12049. doi:10.1007/s11356-014-3142-y Lunyera, J., & Smith, S. R. (2017). Heavy metal nephropathy: Considerations for exposure analysis. Kidney International, 92, 548-550. doi:http://dx.doi.org/10.1016/j.kint.2017.04.043 Sabath, E., & Robles-Osorio, M. L. (2012). Renal health and the environment: Heavy metal nephrotoxicity. Revista Nefrologia, doi:10.3265/Nefrologia.pre2012.Jan.10928 Santos, N. A. G., Catão, C. S., Martins, N. M., Curti, C., Bianchi, M. L. P., & Santos, A. C. (2007). Cisplatin-induced nephrotoxicity is associated with oxidative stress, redox state unbalance, impairment of energetic metabolism and apoptosis in rat kidney mitochondria. Archives of Toxicology, 81(7), 495-504. doi:10.1007/s00204-006-0173-2 Shaki, F., Hosseini, M. J., Ghazi-Khansari, M., & Pourahmad, J. (2012). Toxicity of depleted uranium on isolated rat kidney mitochondria. Biochimica Et Biophysica Acta - General Subjects, 1820(12), 1940-1950. doi:10.1016/j.bbagen.2012.08.015 Soussi, A., Gargouri, M., & El Feki, A. (2018). Effects of co-exposure to lead and zinc on redox status, kidney variables and histopathology in adult albino rats. Toxicology and Industrial Health, 34(7), 469-480. doi:10.1177/0748233718770293 Spreckelmeyer, S., Estrada-Ortiz, N., Prins, G. G. H., van der Zee, M., Gammelgaard, B., Sturup, S., . . . Casini, A. (2017). On the toxicity and transportation mechanisms of cisplatin in kidney tissues in comparison to a gold-based cytotoxic agent. Metallomics, 9, 1786. doi:10.1039/c7mt00271h Turk, E., Kandemir, F. M., Yildirim, S., Caglayan, C., Kucukler, S., & Kuzu, M. (2019). Protective effect of hesperidin on sodium arsenite-induced nephrotoxicity and hepatotoxicity in rats. Biological Trace Element Research, 189, 95-108. doi:10.1007/s12011-018-1443-6 Weber, E. J., Himmelfarb, J., & Kelly, E. J. (2017). Concise review: Current emerging biomarkers of nephrotoxicity. Curr Opin Toxicol., 4, 16-21. doi:10.1016/j.cotox.2017.03.002 Yeh, Y., Lee, Y., Hsieh, Y., & Hwang, D. (2011). Dietary taurine reduces zinc-induced toxicity in male wistar rats. Journal of Food Science, 76(4), 90-98. doi:10.1111/j.1750-3841.2011.02110.x Zamora, L. M., Tracy, B. L., Zielinski, J. M., Meyerhof, D. P., & Moss, M. A. (1998). 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2022-03-04T10:58:19

Increase, Mitochondrial dysfunction

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). Metal-induced Mitochondrial Dysfunction Mitochondria are an important site of Ca2+ regulation and storage, taking up Ca2+ ions electrophoretically from the cytosol through a Ca2+ uniporter, which can then accumulate in the mitochondria (Roos et al., 2012; Orrenius et al., 2015). Similarities between calcium and metals, such as cadmium and lead, makes the entrance and accumulation of these metals into the mitochondria via calcium metals possible by mode of molecular mimicry (Mathews et al., 2013; Adiele et al., 2012). The outer mitochondrial membrane also contains the divalent metal transporter (DMT1), which allows for mitochondrial uptake of divalent metals such as Fe and Mn. When cells are under heavy metal-induced stress, DMT has been shown to be overexpressed in the mitochondrial membrane, making the mitochondria targets of metal toxicity and accumulation. Heavy metal exposure in aerobic organisms increases ROS formation through redox cycling, where metals with different valence states (Fe, Cu, Cr, etc.) directly produce ROS as they are reduced by cellular antioxidants and then react with oxygen (Shaki et al., 2012; Shaki et al., 2013; Pourahmad et al., 2006; Santos et al., 2007). The production of highly reactive hydroxyl radicals under mitochondrial oxidative stress and in the presence of transition metals occurs via the Fenton reaction or Haber-Weiss reaction (Hancock et al., 2001; Valko et al., 2005; Adam-Vizi et al., 2010). Metals and ROS are capable of damaging mitochondrial DNA as well as mechanisms of DNA repair and proliferation arrest (Valko et al., 2005). Metals and ROS have the potential to directly damage mitochondrial membranes and structure by binding to and oxidizing membrane lipids and proteins. This structural damage can collapse the MMP and lead to the opening of the MPTP (Orrenius et al., 2015; Roos et al., 2012; Pourahmad et al., 2006). Uranium and mercury, for example, have both been shown to directly inhibit the mitochondrial electron transport chain and interfere with ATP production (Shaki et al., 2012; Roos et al., 2012). Furthermore, as previously mentioned, metals have been shown to inhibit ROS-detoxifying enzymes. By binding to these enzymes, metals can inhibit their antioxidant functions, and cause an accumulation of ROS and increased synthesis of more antioxidant enzymes in order to combat the oxidative stress (Blajszczak and Bonini, 2017). 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 ). - Revision of AOP3 (Project: <a href="https://www.efsa.europa.eu/en/call/npefsaprev202402-development-aop-network-parkinsonian-motor-symptoms" rel="noreferrer noopener" target="_blank">NP/EFSA/PREV/2024/02</a>): 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. Quantitative assessment of ΔΨm in living cells is most commonly achieved through the use of cationic, lipophilic fluorescent probes that accumulate within the mitochondrial matrix in proportion to the electrochemical gradient (Leonard et al., 2014). Among these, tetramethylrhodamine derivatives such as TMRE (tetramethylrhodamine ethyl ester) and TMRM (tetramethylrhodamine methyl ester) are widely employed due to their reversible, potential-dependent distribution across the inner mitochondrial membrane (Scaduto and Grotyohann, 1999; Creed and McKenzie, 2019). When applied at non-quenching, nanomolar concentrations, these dyes allow linear and quantitative detection of ΔΨm, as fluorescence intensity directly correlates with mitochondrial polarization. Detection can be performed by flow cytometry for population-level quantification, by high-content microscopy for spatially resolved analysis, or by fluorescence plate readers for higher throughput (Wong and Cortopassi, 2002; Valdebenito and Dunchen, 2022). Quantitative interpretation requires the use of appropriate controls, typically involving treatment with protonophores such as FCCP or CCCP, which fully dissipate ΔΨm and thereby establish baseline fluorescence, and inhibitors such as oligomycin or antimycin A to reveal different components of mitochondrial respiration. In parallel, dyes such as JC-1 are also used, though their ratiometric readout is less sensitive at low potentials and more prone to artifacts compared with TMRE or TMRM (Leonard et al., 2022). For accurate normalization, measurements are often corrected for cell number, mitochondrial content, or total protein, and fluorescence changes are expressed relative to maximal depolarization. In addition to chemical probes, genetically encoded sensors, such as mitochondria-targeted fluorescent proteins fused to potential-sensitive domains, provide complementary tools for ΔΨm monitoring in live-cell and in vivo contexts (Leonard et al., 2022). - Not endorsed  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). <div> - Revision of AOP3 (Project: <a href="https://www.efsa.europa.eu/en/call/npefsaprev202402-development-aop-network-parkinsonian-motor-symptoms">NP/EFSA/PREV/2024/02</a>): Determination of mitochondrial ATP production based on extracellular flux analysis   The method is based on the detection of OCR (Oxygen Consumption Rate) that represents mitochondrial respiration as well as on the detection of ECAR (extracellular acidification rate) / proton efflux rate (PER): reflects extracellular acidification, a proxy for glycolysis (lactate release) plus contributions from CO₂/HCO₃⁻. PER is preferred over raw ECAR since it corrects for CO₂-derived acidification (Desousa et al., 2023; Espinosa et al., 2022). Application of inhibitors of individual complexes of the respiratory chain allows the detection of ATP-linked OCR: portion of oxygen consumption directly driving ATP synthesis (lost after ATP synthase inhibition) (Yoo et al., 2024). The proton leak & non-mitochondrial OCR represents remaining oxygen consumption after ATP synthase and electron transport chain inhibitor addition. The difference yields the ATP-coupled respiration component.   Calculation of mitochondrial ATP production  Mito ATP production rate (pmol ATP/min) = OCRATP (pmol O2/min) × 2 × P/O    OCR_ATP: ATP-coupled portion of OCR.   Factor 2: each O₂ molecule contains two oxygen atoms.   P/O ratio: number of ATP molecules synthesized per oxygen atom reduced. A mean P/O ≈ 2.75 is typically assumed (validated across many cell types but substrate- and condition-dependent) (Plitzko and Loesgen, 2018; Mookerjee et al., 2017; Motawe et al., 2024).    Limitations   P/O ratio varies by substrate (glucose vs. fatty acids), cell type, and conditions. Fixed values are approximations.   Non-mitochondrial oxygen consumption (oxidases, peroxidases, etc.) can confound OCR, hence use of ETC inhibitors.   PER vs. ECAR: CO₂-driven acidification must be corrected to avoid overestimating glycolytic ATP.   Normalization: results are usually expressed per cell, protein content, DNA, or mitochondrial mass — interpretation depends on normalization method.  - Not endorsed </div> 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). - Revision of AOP3 (Project: <a href="https://www.efsa.europa.eu/en/call/npefsaprev202402-development-aop-network-parkinsonian-motor-symptoms" rel="noreferrer noopener" target="_blank">NP/EFSA/PREV/2024/02</a>): Endogenous ROS formation by complex I: In mammals, complex I is a dominant site of mitochondrial ROS, especially via RET. In plants (Senkler et al. 2017; Maldonado), mitochondria contain alternative NAD(P)H dehydrogenases and an alternative oxidase (AOX) that bypass Complex I and III These pathways reduce ROS formation by preventing over-reduction of the ETC. Complex I still produces ROS, but generally less damaging due to AOX. Yeast: S. cerevisiae lacks a canonical Complex I entirely, relying instead on alternative NADH dehydrogenases. Consequently, mitochondrial ROS production from a Complex I-like source is absent. Other fungi with true Complex I (e.g., Neurospora crassa) do generate ROS similar to animals. - Not endorsed

UBERON:0000062

UBERON organ

CL:0000255

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2026-02-11T07:06:25

<upstream-id>7eb4ebd4-d969-4bbc-a93e-458d1e3358cd</upstream-id> <downstream-id>c2feac60-2270-423e-882a-8699dee6414f</downstream-id>

AOPWiki 2017-10-25T07:52:43

2017-10-25T07:52:43

<upstream-id>c2feac60-2270-423e-882a-8699dee6414f</upstream-id> <downstream-id>1f81f483-ff18-47c6-809b-7c4949d5fd5f</downstream-id>

AOPWiki 2024-02-28T14:53:17

2024-02-28T14:53:17

<upstream-id>1f81f483-ff18-47c6-809b-7c4949d5fd5f</upstream-id> <downstream-id>c225ef26-e858-4f65-ac79-71ffa68096de</downstream-id> <div> Mitochondria are an essential organelle in ROS production, ATP production, and have a crucial role in regulating both apoptotic and necrotic death (Bhatia, Capili, and Choi, 2020; Wang et al., 2016; Gao et al., 2019). This is particularly notable for renal tubular cells due to their high energy demands for filtration (Cohen, 1986). When mitochondria are damaged and their membrane potential is dissipated, the mitochondrial permeability transition pore (mPTP) is opened and they release cytochrome c (Cyt c), pro-apoptotic caspases, and apoptosis-inducing factor (AIF) (Mao et al., 2010; Galluzzi et al., 2018; Garrido et al., 2006). Cyt c is a protein that triggers apoptosis or necrosis by allosterically activating apoptosis-protease activating factor 1 (APAF 1) (Mao et al., 2010; Galluzzi et al., 2018; Garrido et al., 2006). APAF 1 is then able to cleave caspases 9 and 3, which lead to the induction of a caspase cascade that triggers apoptosis (Mao et al., 2010; Galluzzi et al., 2018; Garrido et al., 2006). Caspase-independent apoptosis is also triggered by the opening of the mPTP, causing the release of AIF which triggers DNA condensation into chromosomes and  degradation in a characteristic manner (Sevrioukova, 2011). Mitochondrial dysfunction therefore leads to not only further oxidative stress, but also to cell death (Bhatia, Capili, and Choi, 2020; Wang et al., 2016; Zhan et al., 2013). </div>

The evidence of biological plausibility between increased mitochondrial dysfunction and renal tubular cytotoxicity is strong, as it is detailed in numerous review articles (Bhatia, Capili, and Choi, 2020; Zhan et al., 2013; Gao et al., 2019; Garrido et al., 2006). Mitochondrial dysfunction includes the inhibition of calcium accumulation in the mitochondria, loss of mitochondrial membrane potential, the opening of the mitochondrial permeability transition pore, increased mitochondrial swelling, decreased ATP production, and mitochondrial depolarization (Bhatia, Capili, and Choi, 2020; Zhan et al., 2013; Gao et al., 2019; Garrido et al., 2006). The induction of the permeability transition of the mitochondrial inner membrane in particular leads to the opening of the mitochondrial permeability transition pore (Mao et al., 2010). This allows the release of several pro-apoptotic molecules to the cytosol, including Cyt c, apoptosis-inducing factor (AIF), and the pro-caspases-2, -3, and -9 (Garrido et al., 2006). Mitochondrial dysfunction can trigger apoptosis through the use of caspase-dependant and independent pathways (Bhatia, Capili, and Choi, 2020; Zhan et al., 2013; Gao et al., 2019; Garrido et al., 2006).  The caspase-dependant pathway of apoptosis induction requires Cyt c. Cyt c is an electron carrier in the mitochondrial electron transport chain (Mao et al., 2010). When released from the mitochondrial it is able to allosterically activate a protein called apoptosis protease activating factor-1 (APAF-1) (Mao et al., 2010). APAF-1 is required for the proteolysis of caspases-9 and -3, which are able of inducing a caspase cascade that results in apoptosis (Mao et al., 2010). The opening of the MPTP is also able to induce a cascade-independent method of cell death via the protein AIF (Mao et al., 2010; Sevrioukova, 2011). AIF is a protein that is normally located on the inner mitochondrial membrane of the mitochondria (Sevrioukova, 2011). When released to the cytosol, AIF undergoes proteolysis and is transported to the nucleus of the cell and induces the condensation of chromatin, as well as characteristic degradation of the cell’s DNA to induce apoptosis in a caspase-independent manner (Sevrioukova, 2011)

One article found that pig kidneys treated with varying concentrations of cisplatin for 10 minutes showed that mitochondrial membrane potential was significantly decreased at a concentration of only 10 μM and cell viability was not significantly decreased until 30 μM (Kruidering et al., 1997). Another article found that treatment with cadmium for 3 hours on human kidney cells caused significant inhibition of resting respiration rates at a concentration of 100 μM and significant inhibition of cell viability at a concentration of 500 μM (Belyaeva et al., 2012). An article examining rat brain mitochondria treated with uranium for 1 hour also showed that treatment with 50 μM uranium caused significant increase in mitochondrial membrane potential and treatment with 100 μM uranium induced a significant decrease in cell viability (Shaki et al., 2012). A study examining the effects of silver nitrate treatment on human bronchial epithelial cells, showed that treatment for 24 hours induced significant mitochondrial dysfunction at a dose of 1.0 μM and cell death was significantly increased at a dose of 5.0 μM (Miyayama et al., 2013). An article investigating the nephrotoxic effects of arsenic on isolated rat kidneys also found that mitochondrial membrane potential (MMP) collapse and Cyt c release were both significantly elevated when the rat kidney cells were treated with 50 and 100 μM for 1 hour, respectively (Hassani et al., 2015). <h3>Temporal concordance</h3> There are currently few articles available that show temporal concordance for mitochondrial dysfunction leading to renal tubular cytotoxicity. One article assessed the effect of uranyl nitrate treatment on normal rat kidney proximal cells and found that mitochondrial membrane potential was significantly decreased after 18 hours of treatment with 600 μM of uranyl nitrate while cell viability was not significant until 24 hours of treatment (Thiébault et al., 2007). <h3>Incidence concordance</h3> Another article found that treating human kidney cells with mercury for 3 hours caused significant inhibition of resting respiration rates and significant inhibition of cell viability both at a concentration of 50 μM (Belyaeva et al., 2012). Copper nanoparticles were also able to induce notable mitochondrial dysfunction and nephrotoxicity in pig kidneys treated for 12 hours with 20 and μg/mL (Zhang et al., 2018). Another article investigated the effects of treatment with varyiousheavy metal nanoparticles and micrometer particles on an alveolar type-II epithelial cell line (Karlsson et al., 2009). They found that iron(II) oxide nanoparticles had a 4.1-fold increase in  mitochondrial depolarization and a 3.5-fold increase in non-viable cells.

<div> <ol> <li>In an article studying the nephrotoxic effects of cisplatin treatment in rat kidneys, they found that treatment with 10 mg/kg bodyweight of cisplatin for 72 hours caused only a 0.3-fold decrease in the level of ATP content in the cell but induced a 1.8-fold increase in the level of executioner caspase-3 activity (Santos et al., 2007).</li> <li>In an article assessing the effects of depleted uranium on human kidney cells, mitochondrial membrane potential was increased significantly by only 3.9-fold when cells were treated with 500 μM for 24 hours (Hao et al., 2014). However, under the same conditions, cytotoxicity was elevated 6.5-fold. (Hao et al., 2014).</li> <li>A study assessing depleted uranium treatment on human embryonic kidney cells also found that only a 3.6-fold increase in membrane potential occurred for cells treated with depleted uranium, while a 5.6-fold increase was seen in cell death compared to the control (Hao et al., 2016).</li> <li>Another article investigated the effects of treatment with various heavy metal nanoparticles and micrometer particles on an alveolar type-II epithelial cell line. In addition, copper oxide nanoparticles and micrometer particles both showed 27.1-fold and 12.1-fold increases in mitochondrial depolarization while cytotoxicity was increased by 56.4-fold and 18.2-fold, respectively (Karlsson et al., 2009).</li> </ol> </div>

<div> There are several known modulating factors that affect the relationship between mitochondrial dysfunction and cell death. One known modulating factor of this variation is mitochondrial biogenesis (Jornayvaz and Shulman, 2010). Mitochondrial biogenesis is the ratio of growth, division, and recycling of existing mitochondria in the cell. Mitochondrial biogenesis itself is influenced by a variety of factors such as exercise levels, cell division, low temperature, caloric restriction, and as is discussed in this AOP, oxidative stress . As a result of changes in these factors, the mitochondrial content, as well as sizes and masses of each of the organelles, is altered . For example, it is generally accepted that increased exercise in an organism increases mitochondrial content and functioning, as there is an increased energy need in those organisms . Therefore, an organism with increased exercise habits would have a less steep slope in the relationship between mitochondrial dysfunction and cell death, as the mitochondrial dysfunction would not be able cause cell death as quickly, since mitochondrial content would be higher requiring a longer time period to accumulate sufficiently to induce apoptosis . This would allow the cell to undergo mitophagy of the injured mitochondria while still continuing to produce adequate energy to keep the cell alive . Similarly, if a cell had just divided, it would be less prepared and capable of dealing with mitochondrial dysfunction, which would allow it to accumulate much faster and would therefore increase the slope between mitochondrial dysfunction and cell death (Jornayvaz and Shulman, 2010). Some diseases are also known to modulate this relationship, such as early aging and diabetes (Pizzorno, 2014). These diseases increase the slope of the relationship between mitochondrial dysfunction leading to oxidative stress due to the fact that they induce faster accumulation of mitochondrial dysfunction via increased levels of oxidative stress and faster accumulation of mitochondrial DNA damage leading to earlier mitochondrial dysfunction (Nissanka and Moraes, 2018; Zelenka, Dvorak, and Alan, 2015; Wei et al., 2015; Kudryavtseva et al., 2016; Forbes and Thorburn, 2018; Schiffer and Friederich-Persson, 2017). </div>

There is a strong, positive correlation between the increase in mitochondrial dysfunction and the increase in cytotoxicity. This was demonstrated by an article which plotted MTT cell viability assay results and LDH concentration in the cell culture media which showed a negative correlation with a slope of -0.99 and a correlation coefficient of 0.97 (Wang et al., 2016).

There has not yet been an identified time-scale for this relationship. Further research will be required in order to determine the time-scale.

There are no feedforward or feedback loops that are known to influence this KER.

<div> The domain of applicability only includes vertebrates, as invertebrates and non-animals do not have kidneys (Mahasen, 2016). </div>

<div> Belyaeva, E. A., Sokolova, T. V., Emelyanova, L. V., & Zakharova, I. O. (2012). Mitochondrial electron transport chain in heavy metal-induced neurotoxicity : Effects of cadmium , mercury , and copper. Thescientificworld, 2012, 1-14. doi:10.1100/2012/136063 Bhatia, D., Capili, A., & Choi, M. E. (2020). Mitochondrial dysfunction in kidney injury, inflammation, and disease; potential therapeutic approaches. Kidney Res. Clin. Pract., 39(3), 244-258. doi:10.23876/j.krcp.20.082 Cohen, J. J. (1986). Relationship between energy requirements for na reabsorption and other renal functions. Kidney International, 29(1), 32-40. doi:10.1038/ki.1986.5 Forbes, J. M., & Thorburn, D. R. (2018). Mitochondrial dysfunction in diabetic kidney disease. Nature Review Nephrology, 14(5), 291. doi:10.1038/nrneph.2018.9 Gao, N., Huang, Z., Liu, H., Hou, J., & Liu, X. (2019). Advances on the toxicity of uranium to different organisms. Chemosphere, 237, 124548. doi:10.1016/j.chemosphere.2019.124548 Galluzzi, L., Vitale, I., Aaronson, S., & et al. (2018). Molecular mechanisms of cell death: Recommendations of the nomenclature committee on cell death 2018. Cell Death and Differentiation, 25, 486-541. doi:10.1038/s41418-017-0012-4 Garrido, C., Galluzzi, L., Brunet, M., Puig, P. E., Didelot, C., & Kroemer, G. (2006). Mecxhanisms of cytochrome c release from mitochondria. Cell Death and Differentiation, 13, 1423-1433. doi:10.1038/sj.cdd.4401950 Hao, Y., Huang, J., Liu, C., Li, H., Liu, J., Zeng, Y., . . . Li, R. (2016). Differential protein expression in metallothionein protection from depleted uranium-induced nephrotoxicity. Scientific Reports, doi:10.1038/srep38942 Hao, Y., Ren, J., Liu, C., Li, H., Liu, J., Yang, Z., . . . Su, Y. (2014). Zinc protects human kidney cells from depleted uranium induced apoptosis. Basic & Clinical Pharmacology & Toxicology, 114, 271-280. doi:10.1111/bcpt.12167 Hassani, S., Yaghoubi, H., Khosrokhavar, R., Jafarian, I., Mahayekhi, V., Housseini, M., & Shahraki, J. (2015). Mechanistic view for toxic effects of arsenic on isolated rat kidney and brain mitochondria. Biologia, 70(5), 683-689. doi:10.1515/biolog-2015-0081 Jornayvaz, F. R., & Shulman, G. I. (2010). Regulation of mitochondrial biogenesis. Essays Biochem., 47, 69-84. doi:10.1042/bse0470069 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:10.1016/j.toxlet.2009.03.014 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. Kudryavtseva, A. V., Krasnov, G. S., Dmitriev, A. A., Alekseev, B. Y., Kardymon, O. L., Sadritdinova, A. F., . . . Snezhkina, A. V. (2016). Mitochondrial dysfunction and oxidative stress in aging and cancer. Oncotarget, 7(29), 44879-44905. doi:10.18632/oncotarget.9821 Mahasen, L. M. A. (2016). Evolution of the kidney. Anatomy Physiol. Biochem. Int. J., 1(1), 555554. doi:10.19080/APBIJ.2016.01.555554 Mao, W., Zhang, N. N., Zhou, F. Y., Li, W. X., Liu, H. Y., Feng, J., . . . He, Z. J. (2010). Cadmium directly induced mitochondrial dysfunction of human embryonic kidney cells. Human and Experimental Toxicology, 30(8), 920-929. doi:10.1177/0960327110384286 Miyayama, T., Arai, Y., Suzuki, N., & Hirano, S. (2013). Mitochondrial electron transport is inhibited by disappearance of metallothionein in human bronchial epithelial cells follwoing exposure to silver nitrate. Toxicology, 305, 20-29. doi:10.1016/j.tox.2013.01.004 Nissanka, N., & Moraes, C. T. (2018). Mitochondrial DNA damage and reactive oxygen species in neurodegenerative disease. FEBS Lett., 592(5), 728-742. doi:10.1002/1873-3468.12956 Pizzorno, J. (2014). Mitochondria - fundamental to life and health. Integrative Medicine (Encinitas), 13(2), 8-15. Santos, N. A. G., Catão, C. S., Martins, N. M., Curti, C., Bianchi, M. L. P., & Santos, A. C. (2007). Cisplatin-induced nephrotoxicity is associated with oxidative stress, redox state unbalance, impairment of energetic metabolism and apoptosis in rat kidney mitochondria. Archives of Toxicology, 81(7), 495-504. doi:10.1007/s00204-006-0173-2 Schiffer, T. A., & Friederich-Persson, M. (2017). Mitochondrial reactive oxygen species and kidney hypoxia in the development of diabetic nephropathy. Front. Physiol., 8:211. doi:10.3389/fphys.2017.00211 Sevrioukova, I. F. (2011). Apoptosis-inducing factor: Structure, function, and redox regulation. Antioxid Redox Signal., 14(12), 2545-2579. doi:10.1089/ars.2010.3445 Shaki, F., Hosseini, M. J., Ghazi-Khansari, M., & Pourahmad, J. (2012). Toxicity of depleted uranium on isolated rat kidney mitochondria. Biochimica Et Biophysica Acta - General Subjects, 1820(12), 1940-1950. doi:10.1016/j.bbagen.2012.08.015 Thiébault, C., Carrière, M., Milgram, S., Simon, A., Avoscan, L., & Gouget, B. (2007). Uranium induces apoptosis and is genotoxic to normal rat kidney (NRK-52E) proximal cells. Toxicological Sciences : An Official Journal of the Society of Toxicology, 98(2), 479-487. doi:kfm130 [pii] Wang, Y., Wang, S., Jia, L., Zhang, L., Ba, J., Han, D., . . . Wu, Y. (2016). Nickel-refining fumes induced DNA damage and apoptosis of NIH/3T3 cells via oxidative stress. International Journal of Environmental Research and Public Health, 13(7), 629-644. doi:10.3390/ijerph13070629 Wei, Y., Zhang, Y., Cai, Y., & Xu, M. (2015). The role of mitochondria in mTOR-regulated longevity. Biol. Rev., 90, 167-181. doi:10.1111/brv.12103 Zelenka, J., Dvorak, A., & Alan, L. (2015). L-lactate protects skin fibroblasts against aging- asociated mitochondrial dysfunction via mitohormesis. Oxidative Medicine and Cellular Longevity, 2015 doi:10.1155/2015/351698 Zhan, M., Brooks, C., Liu, F., Sun, L., & Zheng, D. (2013). Mitochondrial dynamics: Regulatory mechanisms and emerging role in renal pathophysiology. Kidney International, 83, 568-581. doi:10.1038/ki.2012.441 Zhang, H., Chang, Z., Mehmood, K., Abbas, R. Z., Nabi, F., Rehman, M. U., . . . Zhou, D. (2018). 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 </div> AOPWiki 2024-02-28T14:53:52

2024-03-06T17:09:46

<upstream-id>c225ef26-e858-4f65-ac79-71ffa68096de</upstream-id> <downstream-id>0fe56e10-e666-4849-9560-c3b61bf7690c</downstream-id> Excessive renal tubular cytotoxicity, both apoptotic and necrotic, leads to the eventual failure of the kidneys (Priante et al., 2019). This is because the mass cytotoxicity of renal tubular cells leads to the inability of the nephrons to properly filter nutrients and waste from the blood (Pirante et al., 2019). The kidneys can make compensational adjustments to the nephrons to continue adequate filtration up to the loss of 75% of the nephrons, beyond this amount of nephron loss, the kidneys lose function (Orr and Bridges, 2017).

Renal tubular cells are very important functional units of the nephrons (Priante et al., 2019). The tubular cells are essential for the proper removal of waste material from the blood, as well as retaining essential nutrients, water, and salt levels for homeostatic blood content (Priante et al., 2019). The S3 segment of the proximal tubule in particular is highly susceptible to damage by environmental toxicants (Lentini et al., 2017). Apoptosis is the preferred method of cell death for renal tubule cells, as injured cells need to be removed without inducing an inflammatory response (Priante et al., 2019). By forming apoptotic bodies that can be recycled via phagocytes or epithelial cells, the kidney avoids the induction of an inflammatory response which causes the injury of surrounding, healthy cells. However, when apoptotic bodies are not phagocytosed quickly enough, their membranes can become damaged. This causes the apoptotic bodies to enter secondary necrosis, lysing and releasing their contents to the extracellular space. The immune cells will instigate an inflammatory response as a result, causing the injury to nearby tubular cells through the release of granule contents of by the immune cells (Priante et al., 2019). Remarkably, thanks to compensatory functional, molecular, and structural changes in the kidney, the remaining healthy nephrons are able to function adequately until more than 75% of them die (Orr and Bridges, 2017). After the loss of more than 75% of the nephrons however the remaining nephrons are no longer able to effectively remove environmental toxicants or waste from the filtrate, resulting in failed renal function (Orr and Bridges, 2017).

<h3>Dose Concordance</h3> ... <h3>Temporal concordance</h3> ...   <h3>Incidence concordance</h3> One article showed that rats treated once with 5 mg/kg uranyl acetate showed significantly increased proximal tubular cytotoxicity and significant increase in serum creatinine 3 days after the treatment (Sano et al., 2000).   <h3>Other Evidence</h3> “Chronic exposure: After ingestion or inhalation, cadmium is transported to the liver and to the kidney by metallothionein, which binds cadmium. Signs of cell apoptosis and cytokine pathway activation are common in this syndrome. A typical, chronic tubular-interstitial nephropathy is produced by the accumulation of this metal in the medulla and S1 segment of the proximal tubule.” (Lentini et al., 2017)  “Acute exposure: The ionized free form induces cellular toxicity reducing phosphate and glucose transport and inhibiting mitochondrial respiration, with membrane rupture of the proximal tubular cells of the nephron (<a href="https://www.spandidos-publications.com/10.3892/mmr.2017.6389#b17-mmr-15-05-3413" style="color:blue; text-decoration:underline">17</a>).  “Organic mercury gives skin manifestations and neurological disturbances such as hearing loss, paraesthesia and ataxia. Mercury-related kidney damage can due to tubular dysfunction with elevated urinary excretion of albumin, transferrin, retinol binding protein, and β-galactosidase and a nephrotic syndrome with membranous nephropathy pattern (<a href="https://www.spandidos-publications.com/10.3892/mmr.2017.6389#b21-mmr-15-05-3413" style="color:blue; text-decoration:underline">21</a>).” (Lentini et al., 2017) Orr and Bridges (2017) found that exposure to heavy metals **** “Indeed, it has also been suggested that exposure to heavy metals can negatively alter the function of the remaining functional nephrons [<a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5454951/#B11-ijms-18-01039" style="color:blue; text-decoration:underline">11</a>,<a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5454951/#B12-ijms-18-01039" style="color:blue; text-decoration:underline">12</a>]. These adverse effects could conceivably lead to additional and/or more rapid cell death and glomerulosclerosis, which would further reduce the functional renal mass of the patient.” (Orr & Bridges, 2017) “In the rat, an acute perfusion of Cd2+ caused hypercalciuria, hyperphosphaturia and hypokaliuria without modification of glomerular filtration rate (GFR) [<a href="https://www.karger.com/Article/FullText/83981#ref1" style="color:blue; text-decoration:underline">1</a>]. By contrast, a single, 20-fold lower dose of Pb2+, Hg2+ induced glomerular and tubular damage characterized by a reduced GFR, glycosuria, proteinuria and a rapid obstruction of the tubular system [<a href="https://www.karger.com/Article/FullText/83981#ref13" style="color:blue; text-decoration:underline">13</a>], illustrating that the pattern of nephrotoxicity differs between heavy metals. Therefore, Pb2+ and Hg2+ are more dangerous than Cd2+ because they induce an irreversible renal insufficiency even during acute intoxication.” (Barbier et al., 2005)

There are no currently known inconsistencies or uncertainties for this relationship.

There are several known modulating factors of the relationship between renal tubular cytotoxicity and kidney failure. One modulator of this relationship is age.

There is a defined response-response relationship for renal tubule cytotoxicity leading to kidney failure. The loss of 75% of the nephrons to damage is the threshold for kidney failure (Orr and Bridges, 2017). This is due to the ability of the kidneys to make changes in the structure and function of the remaining nephrons at a molecular level to compensate for the lost nephrons (Orr and Bridges, 2017). The kidneys are able to retain adequate functioning until only 25% of the original nephrons remain, at which point the compensatory changes cannot maintain kidney functioning and kidney failure is final (Orr and Bridges, 2017).

There are no known feedforward/feedback loops that influence this relationship.

The domain of applicability only includes vertebrates, as invertebrates and non-animals do not have kidneys (Mahasen, 2016).

Lentini, P., Zanoli, L., Granata, A., Signorelli, S. S., Castellino, P., & Dell'aquila, R. (2017). Kidney and heavy metals - the role of environmental exposure (review). Molecular Medicine Reports, 15(3413), 3419. doi:10.3892/mmr.2017.6389 Mahasen, L. M. A. (2016). Evolution of the kidney. Anatomy Physiol. Biochem. Int. J., 1(1), 555554. doi:10.19080/APBIJ.2016.01.555554 Orr, S. E., & Bridges, C. C. (2017). Chronic kidney disease and exposure to nephrotoxic metals. International Journal of Molecular Sciences, 18 doi:10.3390/ijms18051039 Priante, G., Gianesello, L., Ceol, M., Del Prete, D., & Anglani, F. (2019). Cell death in the kidney. International Journal of Molecular Sciences, 20(14), 3598. doi: 10.3390/ijms20143598. doi:10.3390/ijms20143598 Sano, K., Fujigaki, Y., Miyaji, T., Ikegaya, N., Ohishi, K., Yonemura, K., & Hishida, A. (2000). Role of apoptosis in uranyl acetate-induced acute renal failure and acquired resistance to uranyl acetate. Kidney International, 57(4), 1560-1570. doi:10.1046/j.1523-1755.2000.00777.x AOPWiki 2017-10-25T07:54:27

2022-03-08T11:46:18

Inhibition of mitochondrial DNA polymerase gamma leading to kidney toxicity

Angela Mally

Prof. Dr. Angela Mally Department of Toxicology University of Würzburg Versbacher Str. 9 97078 Würzburg Germany Phone/fax:          +49 931 31-81194 Email:                  <a href="mailto:mally@toxi.uni-wuerzburg.de">mally@toxi.uni-wuerzburg.de</a>

BY-SA

Under Development

1.43

1.0

This Adverse Outcome Pathway describes the sequential key events that link inhibition of mitochondrial DNA polymerase gamma (Pol gamma) to kidney toxicity. Nucleoside and nucleotide  (nucleos(t)ide) analogs are widely used as antiviral drugs for the effective treatment of viral infections including HIV and chronic Hepatitis B virus infections. As structural analogs of substrate nucleotides, these drugs act as chain terminators of viral DNA synthesis via competitive inhibition of reverse transcriptase or viral DNA polymerases, thereby blocking virus replication. Besides targeting viral enzymes, nucleos(t)ide antiviral agents are also substrates for human DNA polymerases, which may lead to moderate to life-threatening adverse drug reactions, including peripheral neuropathy, myopathy, lactic acidosis, and acute and chronic kidney injury [1-4]. Toxicity of antiviral nucleos(t)ides has been linked to mitochondrial dysfunction as a consequence of inhibition of mitochondrial DNA polymerase gamma (Pol gamma), a particular sensitive target, and associated inhibition of mtDNA replication [1, 3]. In the kidney, the proximal tubule is the main target of antiviral nucleos(t)ide drug toxicity due to active uptake via basolateral organic anion transporters (e.g. OAT1 and OAT3) expressed at this site [5, 6]. Based on the current mechanistic understanding, the subsequent sequence of key events (KE) leading to kidney injury as an adverse outcome can be described as inhibition of Pol gamma as the molecular initiating event (MIE), leading to mtDNA depletion (KE1), mitochondrial dysfuntion (KE2) and proximal tubule cell toxicity (KE3).

adjacent

Low

Moderate adjacent

Low

High adjacent

Low

High adjacent

Moderate

High

Not Specified Unspecific

Not Specified

All life stages

High

Mechanistic data on KEs and KERs in this AOP are derived from in vitro and in vivo studies in humans and rodents.  The described AOP presents a general mechanism leading to kidney toxicity in preclinical animal species and humans. The described AOP is not limited to a specific life stage or sex. The sequence of MIE and KEs in this AOP presents a universal mechanism by which nucleos(t)ide analogs are thought to cause toxicity not only in the kidney but also in other organs and tissues, including liver, heart, muscle and the nervous system [1, 3, 4, 7]. The tissue-specificity and severity of the response to a particular nucleos(t)ide analog is considered to be at least in part determined by toxicokinetic factors, most notably active uptake into and efflux from target cells, transport across the mitochondrial membrane and metabolic conversion into the active triphosphate form [5-8]. Nephrotoxicity presents a treatment-limiting toxicity for a number of nucleos(t)ide analogs (e.g. tenofovir, adefovir, cidofovir). Experimental evidence for inhibition of mitochondrial DNA polymerase gamma leading to kidney toxicity as an adverse outcome is comes from in vitro studies, studies in laboratory animals (rats and mice) as well as from reports of patients treated with these compounds. These studies show a strong association between mitochondrial toxicity and antiviral nucleos(t)ide induced nephrotoxicity [9-14], with some studies also demonstrating concomitant mtDNA depletion [9, 11, 12, 15].  The causal relationship between the MIE and the downstream KEs is further supported by studies investigating the mechanism of toxicity of nucleos(t)ide analogs in other cells and tissues. For instance, a significant reduction in mtDNA was observed in muscle biopsies of zidovudine-treated HIV positive patients with myopathy as compared non-HIV-patient controls [16].  Studies with isolated human DNA polymerases demonstrate increased sensitivity of Pol gamma to inhibition by antiretroviral nucleotides as compared to nuclear polymerases. Inhibition of mtDNA synthesis and loss of cell number was observed in a T-lymphoid leukemic cell line (Molt-4) treated with several anti-HIV and anti-HBV nucleoside analogs (d4T, 3'-deoxy-2',3'-didehydrothymidine; FLT, 3'-fluoro-3'-deoxythynidine; ddC, 2',3'-dideoxycytidine), which were also identified as potent inhibition of Pol gamma. However, a number of potent Pol gamma inhibitors did not cause significant effects on mtDNA synthesis and cell viability. Based on these findings, the authors concluded that there was no clear quantitative or qualitative correlation between the inhibition of isolated Pol gamma and inhibition of mitochondrial DNA synthesis in vitro, and moreover that these data are not predictive of in vivo toxicity.  It is however important to stress that toxicokinetics, most notably cellular uptake of the tested antivirals, were not considered in this assessment. Thus, it is likely that some of the most potent inhibitors of Pol gamma failed to induce mtDNA depletion and cytotoxicity in this cell model simply because of insufficient cellular uptake [17].  Mechanistic data on KEs and KERs in this AOP are derived from in vitro and in vivo studies in humans and rodents. The described AOP presents a general mechanism leading to kidney toxicity in preclinical animal species (rats, mice) and humans. The described AOP is not limited to a specific life stage or sex.

<table border="1" cellpadding="0" cellspacing="0"> <tbody> <tr> <td style="width:72px"> MIE / KE </td> <td style="width:132px"> Short name </td> <td style="width:227px"> Support </td> <td style="width:92px"> Essentiality </td> </tr> <tr> <td style="width:72px"> MIE </td> <td style="width:132px"> Inhibition, Pol gamma </td> <td style="width:227px"> Inhibition of mtDNA Pol gamma by antiviral nucleos(t)ides demonstrated using enzymatic assays  [2, 18-20]  </td> <td style="width:92px"> high </td> </tr> <tr> <td style="width:72px"> KE1 </td> <td style="width:132px"> Depletion, mtDNA </td> <td style="width:227px"> Loss of mtDNA observed in vitro, in laboratory animals and patients after treatment with antiviral nucleos(t)ides [9, 11, 12, 15, 21] </td> <td style="width:92px"> high </td> </tr> <tr> <td style="width:72px"> KE2 </td> <td style="width:132px"> Dysfunction, mitochondria </td> <td style="width:227px"> Changes in mitochondrial ultrastructure and/or function (e.g. mitochondrial enzyme activities) observed in vitro, in laboratory animals and kidney biopsies of patients after treatment with antiviral nucleos(t)ides [9] [10-14, 21, 22] </td> <td style="width:92px"> high </td> </tr> <tr> <td style="width:72px"> KE3 </td> <td style="width:132px"> Increase, Cytotoxicity </td> <td style="width:227px"> Cytotoxicity of antiviral nucleos(t)ides observed in a range of kidney cell models with the severity depending on cellular uptake [11-14, 21-24] </td> <td style="width:92px"> high </td> </tr> <tr> <td style="width:72px"> AO </td> <td style="width:132px"> Occurrence, Kidney Toxicity </td> <td style="width:227px"> Nephrotoxicity observed in laboratory animals and patients after treatment with antiviral nucleos(t)ides [9] [10-14, 25-28] [22] </td> <td style="width:92px">   </td> </tr> </tbody> </table>

Concordance of dose-response relationships This is still a qualitiative description of the pathway. There is at present no quantitative information on dose-response relationships. Experiments are underway to provide quantitative understanding of dose-response relationships and response-response relationships between upstream and downstream KEs. In establishing dose-response relationships, it needs to be considered that effective excision of nucleotides by proofreading exonuclease of DNA polymerase as a repair mechanism may affect downstream KEs [2].   Temporal concordance among the key events and adverse outcome The individual KEs are shown to occur prior to or concomitant with the onset of nephrotoxicity.   Strength, consistency, and specificity of association of adverse outcome and initiating event The scientific evidence on the association between inhibition of DNA Polymerase gamma (MIE) and kidney toxicity (AO) is strong and consistent. The MIE is not specific for kidney toxicity as is considered responsible for a range of adverse effects of antiviral nucleos(t)ide treatment, whereby the site of toxicity appears to be at least in part determined by the toxicokinetics of individual drugs.   Biological plausibility, coherence, and consistency of the experimental evidence Since antiviral nucleos(t)ide analogs are specifically designed to inhibit (viral) DNA polymerases or reverse transcriptase, off-target effects via interaction of human DNA polymerases are biologically plausible and consistent with the pharmacological MoA. The described AOP is biologically plausible, coherent and supported by experimental data.   Alternative mechanism(s) that logically present themselves and the extent to which they may distract from the postulated AOP There are no alternative mechanism(s) that logically present themselves, although a contribution of yet undefined off-target effects to the overall AO cannot be excluded.   Uncertainties, inconsistencies and data gaps This AOP is plausible and consistent with general biological knowledge. Quantitative information on dose response-relationships as well as repsonse-response relationships for upstream and downstream KEs is needed to support its applicability for the development of alternative in vitro tests for nephrotoxicity testing.

Quantitative data on KERs between upstream and downstream KE are still lacking.

The described AOP is intended to provide a mechanistic framework for the development of in vitro bioactivity assays capable of predicting quantitative points of departure for safety assessment with regard to nephrotoxicity. Such assays may form part of an integrated testing strategy to reduce the need for repeated dose toxicity studies (e.g.  OECD Guideline 407; OECD Guideline 407) and to aid in the design of new antiviral drugs.

High

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