<td>Under development: Not open for comment. Do not cite</td>
<td></td>
<td></td>
<td></td>
<td>Not under active development</td>
<td>Under Development</td>
<td>1.43</td>
<td>Included in OECD Work Plan</td>
</tr>
</tbody>
</table>
</div>
</div>
<!-- Abstract Section, text as generated by author -->
<div id="abstract">
<h2>Abstract</h2>
<p>It is well established that bioactivation of xenobiotics to reactive intermediates that covalently bind to proteins presents a major mechanism by which xenobiotics may cause proximal tubule injury. Examples for compounds that form covalent protein adducts in proximal tubule cells include haloalkenes (e.g. trichloroethylene, tetrachloroethylene, hexachloro-1,3-butadiene, chloroform), quinones (derived from e.g. hydroquinone, bromobenzene, 4-aminophenol), cephalosporins, and N-(3,5-dichlorophenyl)succinimide [1-6]. Covalent interaction of a chemical or a metabolite with cellular proteins represents the molecular initiating event (MIE) that triggers perturbation of cellular functions, of which mitochondrial dysfunction (KE1) leading to ATP depletion (KE2) appears to be most critical for proximal tubule cell death (KE3) by apoptosis and/or necrosis [5, 7-10]. Tubular obstruction and inflammatory responses to proximal tubule injury including activation of complement may cause secondary toxicity and thus amplify kidney injury, resulting in a progressive decline in kidney function (evidenced by e.g. rise in serum creatinine and blood urea nitrogen) (AO).</p>
<h2>Abstract</h2>
<hr>
<p>It is well established that bioactivation of xenobiotics to reactive intermediates that covalently bind to proteins presents a major mechanism by which xenobiotics may cause proximal tubule injury. Examples for compounds that form covalent protein adducts in proximal tubule cells include haloalkenes (e.g. trichloroethylene, tetrachloroethylene, hexachloro-1,3-butadiene, chloroform), quinones (derived from e.g. hydroquinone, bromobenzene, 4-aminophenol), cephalosporins, and N-(3,5-dichlorophenyl)succinimide [1-6]. Covalent interaction of a chemical or a metabolite with cellular proteins represents the molecular initiating event (MIE) that triggers perturbation of cellular functions, of which mitochondrial dysfunction (KE1) leading to ATP depletion (KE2) appears to be most critical for proximal tubule cell death (KE3) by apoptosis and/or necrosis [5, 7-10]. Tubular obstruction and inflammatory responses to proximal tubule injury including activation of complement may cause secondary toxicity and thus amplify kidney injury, resulting in a progressive decline in kidney function (evidenced by e.g. rise in serum creatinine and blood urea nitrogen) (AO).</p>
<br>
</div>
<!-- Background Section, text as generated by author -->
<div id="background">
<br>
</div>
<!-- AOP summary, includes summary of each of the events associated with this aop -->
</p><p>Covalent protein alkylation is a feature of many cytotoxic drugs but the overall extent of binding does not adequately distinguish toxic from non-toxic binding. <sup id="cite_ref-27" class="reference"><a href="#cite_note-27">[27]</a></sup> Interestingly, some chemicals significantly alkylate proteins without causing toxicity, which suggests that only alkylation of a specific protein subset critical subset contributes to injury. Indeed, Codreanu presented an inventory of proteins affected by electrophile-mediated alkylation in intact cells and suggested that non-toxic covalent binding largely affects cytoskeletal protein components, whereas toxic covalent binding induces lethal injury by targeting factors involved in protein synthesis and catabolism and possibly mitochondrial electron transport. <sup id="cite_ref-Codreanu_2014_3-2" class="reference"><a href="#cite_note-Codreanu_2014-3">[3]</a></sup>
In vitro covalent binding studies to macromolecules have been used to elucidate the biochemical mechanisms of chemical-induced toxicity. Experimental work with kidney epithelial cells by Chen et al suggested that following alkylation of cellular macromolecules as initial cytotoxic event both sulfhydryl depletion and lipid peroxidation are components of the cytotoxic mechanism <sup id="cite_ref-28" class="reference"><a href="#cite_note-28">[28]</a></sup> Dennehy et al have analyzed the protein targets in nuclear and cytoplasmic proteomes from human embryonic kidney cells (HEK293) treated in vitro with two biotin-tagged, thiol-reactive electrophiles and mapped the adducts. Certain protein families appeared particularly susceptible to alkylation. <sup id="cite_ref-29" class="reference"><a href="#cite_note-29">[29]</a></sup> Shin et al have identified protein targets of two biotin-tagged model electrophiles in human liver microsomes through LC-MS-MS and showed that different target selectivities of the two electrophile probes correlated with different biological outcomes and that alkylation reactions of specific targets could be quantified. <sup id="cite_ref-30" class="reference"><a href="#cite_note-30">[30]</a></sup>
</p>
<br>
<br>
<!-- end Evidence for Perturbation of This Event by Stressors -->
</div>
<h4>Evidence Supporting Applicability of this Event</h4>
<br>
<divid="overall_assessment">
<h2>Overall Assessment of the AOP</h2>
<!-- loop to find taxonomic applicability under event -->
<p><strong>Concordance of dose-response relationships</strong></p>
<p>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.</p>
<p> </p>
<p><strong>Temporal concordance among the key events and adverse outcome</strong></p>
<p>The individual KEs are shown to occur prior to or concomitant with the onset of nephrotoxicity.</p>
<p> </p>
<p><strong>Strength, consistency, and specificity of association of adverse outcome and initiating event</strong></p>
<p>The scientific evidence on the association between protein alkylation by reactive intermediates and kidney toxicity (AO) is strong and consistent. The MIE is not specific for kidney toxicity and is well established to lead to damage to other organs, whereby the site of toxicity is largely determined by the toxicokinetics of the parent compound or active metabolite.</p>
<p> </p>
<p><strong>Biological plausibility, coherence, and consistency of the experimental evidence</strong></p>
<p>The described AOP is biologically plausible, coherent and well supported by experimental data.</p>
<p> </p>
<p><strong>Alternative mechanism(s) that logically present themselves and the extent to which they may distract from the postulated AOP</strong></p>
<p>There are no alternative mechanism(s) that logically present themselves, although a contribution of other mechanisms such as generation of oxidative stress to the overall AO is possible.</p>
<p> </p>
<p><strong>Uncertainties, inconsistencies and data gaps</strong></p>
<p>This AOP is plausible and consistent with general biological knowledge. However, there is currently little understanding as to which target proteins are critical to toxicity mediated by alkalation damage. Quantitative information on dose response-relationships as well as response-response relationships for upstream and downstream KEs is needed to support its applicability for the development of alternative in vitro tests for nephrotoxicity testing.</p>
<!-- life stages -->
<div>
</div>
<!-- end life stages -->
<!-- sex terms -->
<div>
</div>
<!-- end sex terms -->
<h3>Quantitative Consideration</h3>
<p>Quantitative data on KERs between upstream and downstream KE are still lacking.</p>
<div>
<p>Human, rat and mouse <sup id="cite_ref-EPA_2010_11-0" class="reference"><a href="#cite_note-EPA_2010-11">[11]</a></sup>
<h2>Considerations for Potential Applications of the AOP (optional)</h2>
<p>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).</p>
</div>
<div id="references">
<h2>References</h2>
<p>1. Birner, G., et al., <em>Metabolism of tetrachloroethene in rats: identification of N epsilon-(dichloroacetyl)-L-lysine and N epsilon-(trichloroacetyl)-L-lysine as protein adducts.</em> Chem Res Toxicol, 1994. <strong>7</strong>(6): p. 724-32.</p>
<p>2. Pahler, A., et al., <em>Generation of antibodies to Di- and trichloroacetylated proteins and immunochemical detection of protein adducts in rats treated with perchloroethene.</em> Chem Res Toxicol, 1998. <strong>11</strong>(9): p. 995-1004.</p>
<p>3. Kleiner, H.E., et al., <em>Immunochemical detection of quinol--thioether-derived protein adducts.</em> Chem Res Toxicol, 1998. <strong>11</strong>(11): p. 1283-90.</p>
<p>4. Lau, S.S., <em>Quinone-thioether-mediated nephrotoxicity.</em> Drug Metab Rev, 1995. <strong>27</strong>(1-2): p. 125-41.</p>
<p>5. Tune, B.M., <em>Nephrotoxicity of beta-lactam antibiotics: mechanisms and strategies for prevention.</em> Pediatr Nephrol, 1997. <strong>11</strong>(6): p. 768-72.</p>
<p>6. Griffin, R.J. and P.J. Harvison, <em>In vivo metabolism and disposition of the nephrotoxicant N-(3, 5-dichlorophenyl)succinimide in Fischer 344 rats.</em> Drug Metab Dispos, 1998. <strong>26</strong>(9): p. 907-13.</p>
<p>7. Groves, C.E., et al., <em>Pentachlorobutadienyl-L-cysteine (PCBC) toxicity: the importance of mitochondrial dysfunction.</em> J Biochem Toxicol, 1991. <strong>6</strong>(4): p. 253-60.</p>
<p>8. Chen, Y., et al., <em>Role of mitochondrial dysfunction in S-(1,2-dichlorovinyl)-l-cysteine-induced apoptosis.</em> Toxicol Appl Pharmacol, 2001. <strong>170</strong>(3): p. 172-80.</p>
<p>9. Hill, B.A., T.J. Monks, and S.S. Lau, <em>The effects of 2,3,5-(triglutathion-S-yl)hydroquinone on renal mitochondrial respiratory function in vivo and in vitro: possible role in cytotoxicity.</em> Toxicol Appl Pharmacol, 1992. <strong>117</strong>(2): p. 165-71.</p>
<p>10. Aleo, M.D., et al., <em>Toxicity of N-(3,5-dichlorophenyl)succinimide and metabolites to rat renal proximal tubules and mitochondria.</em> Chem Biol Interact, 1991. <strong>78</strong>(1): p. 109-21.</p>
<td><a href="/aops/38">Aop:38 - Protein Alkylation leading to Liver Fibrosis</a></td>
<td>MolecularInitiatingEvent</td>
</tr>
<tr>
<td><a href="/aops/258">Aop:258 - Renal protein alkylation leading to kidney toxicity</a></td>
<td>MolecularInitiatingEvent</td>
</tr>
</tbody>
</table>
</div>
<!-- event text -->
<h4>How this Key Event Works</h4>
<p>Alkylation is the transfer of an alkyl group from one molecule to another. The alkyl group may be transferred as an alkyl carbocation, a free radical, a carbanion or a carbene (or their equivalents). Protein alkylation is the addition of an alkyl group to a protein amino acid. An alkyl group is any group derived from an alkane by removal of one hydrogen atom.
<p>Human, rat and mouse <sup id="cite_ref-EPA_2010_11-0" class="reference"><a href="#cite_note-EPA_2010-11">[11]</a></sup>
</p>
<h4>Key Event Description</h4>
<p>Alkylation is the transfer of an alkyl group from one molecule to another. The alkyl group may be transferred as an alkyl carbocation, a free radical, a carbanion or a carbene (or their equivalents). Protein alkylation is the addition of an alkyl group to a protein amino acid. An alkyl group is any group derived from an alkane by removal of one hydrogen atom.
Alkylating agents are highly reactive chemicals that introduce alkyl groups into biologically active molecules and thereby prevent their proper functioning. Alkylating agents are classified according to their nucleophilic or electrophilic character. Nucleophilic alkylating agents deliver the equivalent of an alkyl anion (carbanion). These compounds typically can add to an electron-deficient carbon atom such as at a carbonyl group. Electrophilic alkylating agents deliver the equivalent of an alkyl cation. Alkyl halides can also react directly with amines to form C-N bonds; the same holds true for other nucleophiles such as alcohols, carboxylic acids, thiols, etc. Alkylation with only one carbon is termed methylation. <sup id="cite_ref-1" class="reference"><a href="#cite_note-1">[1]</a></sup> <sup id="cite_ref-2" class="reference"><a href="#cite_note-2">[2]</a></sup>
</p><p>Covalent protein alkylation by reactive electrophiles was identified as a key triggering event in chemical toxicity over 40 years ago and these reactions remain a major cause of chemical-induced toxicity. Interestingly, some chemical molecules produce significant protein covalent binding without causing toxicity, which suggests that only a critical subset of protein alkylation events contributes to injury.
The study by Codreanu et al. (2014) describes an inventory of electrophile- mediated protein damage in intact cells and suggests that non-toxic covalent binding may largely be survivable damage to cytoskeletal components, whereas toxic covalent binding produces lethal injury by targeting protein synthesis and catabolism and possibly mitochondrial electron transport. <sup id="cite_ref-Codreanu_2014_3-0" class="reference"><a href="#cite_note-Codreanu_2014-3">[3]</a></sup>
</p><p>High Performance Liquid Chromatography – electrospray tandem mass spectrometry (HPLC-ESI-MS/MS) is the most popular MS technique. It combines the separation ability of HPLC along with the sensitivity and specificity of detection from MS. One of the advantages of HPLC-MS is that it allows samples to be rapidly desalted online, so no sample preparation is required unlike samples for GC-MS. Electrospray ionisation can produce singly or multiply charged ions. Typically high molecular weight compounds have multiple charges i.e. peptides and proteins. This technique is particularly suited to analysing polar molecules of mass <2000 Dalton and requires no prior derivatisation in most applications.
</p><p>MALDI-TOF/MS (Matrix Assisted Laser Desorption/Ionization Time of Flight Mass Spectrometry)
</p><p>Matrix-assisted laser desorption/ionization (MALDI) is a soft ionization technique used in mass spectrometry, allowing the analysis of biomolecules (biopolymers such as DNA, proteins, peptides and sugars) and large organic molecules (such as polymers, dendrimers and other macromolecules), which tend to be fragile and fragment when ionized by more conventional ionization methods. MALDI methodology is a three-step process. First, the sample is mixed with a suitable matrix material and applied to a metal plate. Second, a pulsed laser irradiates the sample, triggering ablation and desorption of the sample and matrix material. Finally, the analyte molecules are ionized by being protonated or deprotonated in the hot plume of ablated gases, and can then be accelerated into whichever mass spectrometer is used to analyse them. <sup id="cite_ref-10" class="reference"><a href="#cite_note-10">[10]</a></sup>
</p><p><br />
</p><p></em>
</p>
<br>
<h4>References</h4>
<ol class="references">
<h4>References</h4>
<ol class="references">
<li id="cite_note-1"><span class="mw-cite-backlink"><a href="#cite_ref-1">↑</a></span> <span class="reference-text">The European Bioinformatics Institute <a rel="nofollow" target="_blank" class="external free" href="http://www.ebi.ac.uk/QuickGO/GTerm?id=GO:0008213">http://www.ebi.ac.uk/QuickGO/GTerm?id=GO:0008213</a> (accessed on 20 January 2016).</span>
</li>
<li id="cite_note-2"><span class="mw-cite-backlink"><a href="#cite_ref-2">↑</a></span> <span class="reference-text">NLM, Medical Subject Headings, National Library of Medicine, <a rel="nofollow" target="_blank" class="external free" href="http://www.nlm.nih.gov/cgi/mesh/2011/MB_cgi?mode=&term=Alkylating+agents">http://www.nlm.nih.gov/cgi/mesh/2011/MB_cgi?mode=&term=Alkylating+agents</a> (accessed on 20 January 2016).</span>
</li>
<li id="cite_note-Codreanu_2014-3"><span class="mw-cite-backlink">↑ <sup><a href="#cite_ref-Codreanu_2014_3-0">3.0</a></sup> <sup><a href="#cite_ref-Codreanu_2014_3-1">3.1</a></sup> <sup><a href="#cite_ref-Codreanu_2014_3-2">3.2</a></sup></span> <span class="reference-text">Codreanu, S.G. et al. (2014), Alkylation damage by lipid electrophiles targets functional protein systems, Molecular & Cellular Proteomics, vol. 13, no. 3, pp.849–859.</span>
</li>
<li id="cite_note-4"><span class="mw-cite-backlink"><a href="#cite_ref-4">↑</a></span> <span class="reference-text">Grattagliano, I. et al. (2009), Biochemical mechanisms in drug-induced liver injury: certainties and doubts, World J Gastroenterol, vol. 15, no. 39, pp. 4865-4876.</span>
<li id="cite_note-Kehrer2000-6"><span class="mw-cite-backlink">↑ <sup><a href="#cite_ref-Kehrer2000_6-0">6.0</a></sup> <sup><a href="#cite_ref-Kehrer2000_6-1">6.1</a></sup></span> <span class="reference-text">Kehrer, J.P. and S. Biswal (2000), The Molecular Effects of Acrolein, Toxicol. Sciences,vol.57,pp.6-15.</span>
</li>
<li id="cite_note-7"><span class="mw-cite-backlink"><a href="#cite_ref-7">↑</a></span> <span class="reference-text">Schopfer, F.J., C. Cipollina and B.A. Freeman (2011), Formation and Signaling Actions of Electrophilic Lipids, Chem Rev, vol. 111, no. 10,pp.5997–6021.</span>
</li>
<li id="cite_note-8"><span class="mw-cite-backlink"><a href="#cite_ref-8">↑</a></span> <span class="reference-text">Zhang F et al. (2005), Differential adduction of proteins vs. deoxynucleosides by methyl methanesulfonate and 1-methyl-1-nitrosourea in vitro, Mass Spectrom, vol 19, no. 4, pp. 438–448.</span>
</li>
<li id="cite_note-9"><span class="mw-cite-backlink"><a href="#cite_ref-9">↑</a></span> <span class="reference-text">Gundry, R.L. et al. (2009), Preparation of proteins and peptides for mass spectrometry analysis in a bottom-up proteomics workflow, Curr Protoc Mol Biol, chapter 10, section VI, unit 10.25, pp. 1-23.</span>
</li>
<li id="cite_note-10"><span class="mw-cite-backlink"><a href="#cite_ref-10">↑</a></span> <span class="reference-text">Kislinger, T. et al. (2005), Analysis of protein glycation products by MALDI-TOF/MS, Ann N Y Acad Sci, vol. 1043, pp. 249-259.</span>
</li>
<li id="cite_note-EPA_2010-11"><span class="mw-cite-backlink">↑ <sup><a href="#cite_ref-EPA_2010_11-0">11.0</a></sup> <sup><a href="#cite_ref-EPA_2010_11-1">11.1</a></sup></span> <span class="reference-text">EPA Toxicological review of Carbon Tetrachloride (CAS No. 56-23-5). March 2010 EPA/635/R-08/005F available at: <a rel="nofollow" target="_blank" class="external free" href="http://cfpub.epa.gov/ncea/iris/iris_documents/documents/toxreviews/0020tr.pdf">http://cfpub.epa.gov/ncea/iris/iris_documents/documents/toxreviews/0020tr.pdf</a> (accessed 24.10.2015)</span>
</li>
<li id="cite_note-12"><span class="mw-cite-backlink"><a href="#cite_ref-12">↑</a></span> <span class="reference-text">Auerbach, S.S. et al. (2008), A comparative 90 day toxicity study of allyl acetate, allyl alcohol and acrolein, Toxicology, Vol. 253, No.1-3, pp.79–88.</span>
</li>
<li id="cite_note-13"><span class="mw-cite-backlink"><a href="#cite_ref-13">↑</a></span> <span class="reference-text">Huang, L. et al. (2008), Genes related to apoptosis predict necrosis of the liver as a phenotype observed in rats exposed to a compendium of hepatotoxicants, BMC Genomics, vol. 9: 288.</span>
</li>
<li id="cite_note-14"><span class="mw-cite-backlink"><a href="#cite_ref-14">↑</a></span> <span class="reference-text">Mohammad, M.K. et al. (2012), Acrolein cytotoxicity in hepatocytes involves endoplasmic reticulum stress, mitochondrial dysfunction and oxidative stress, Toxicol Appl Pharmacol, vol. 265, no. 1, pp. 73-82.</span>
</li>
<li id="cite_note-15"><span class="mw-cite-backlink"><a href="#cite_ref-15">↑</a></span> <span class="reference-text">Yamada T et al., (2013), A category approach to predicting the repeated-dose hepatotoxicity of allyl esters, Regulatory Toxicology and Pharmacology, vol. 65, no. 2, pp. 189–195.</span>
</li>
<li id="cite_note-16"><span class="mw-cite-backlink"><a href="#cite_ref-16">↑</a></span> <span class="reference-text">Basu, S. (2003), Carbon tetrachloride-induced lipid peroxidation: eicosanoid formation and their regulation by antioxidant nutrients, Toxicology,vol.189, no.1-2, pp. 113-127.</span>
</li>
<li id="cite_note-17"><span class="mw-cite-backlink"><a href="#cite_ref-17">↑</a></span> <span class="reference-text">Calabrese, E.J., L.A. Baldwin and H.M. Mehendale (1993), G2 subpopulation in rat liver induced into mitosis by low-level exposure to carbon tetrachloride: an adaptive response, Toxicol Appl Pharmacol, vol. 121. no. 1, pp. 1-7.</span>
<li id="cite_note-19"><span class="mw-cite-backlink"><a href="#cite_ref-19">↑</a></span> <span class="reference-text">Knockaert, L. et al. (2012), Carbon tetrachloride-mediated lipid peroxidation induces early mitochondrial alterations in mouse liver, Lab Invest, vol. 92, no. 3, pp. 396-410.</span>
</li>
<li id="cite_note-20"><span class="mw-cite-backlink"><a href="#cite_ref-20">↑</a></span> <span class="reference-text">Lee Kwang-Jong et al. (2004), Induction of molecular chaperones in carbon tetrachloride-treated rat liver: implications in protection against liver damage, Cell Stress Chaperones, vol. 9, no. 1, pp. 58-68.</span>
</li>
<li id="cite_note-21"><span class="mw-cite-backlink"><a href="#cite_ref-21">↑</a></span> <span class="reference-text">Li, Xiaowei et al. (2014), NMR-based metabonomic and quantitative real-time PCR in the profiling of metabolic changes in carbon tetrachloride-induced rat liver injury, J Pharm Biomed Anal; vol. 89, pp.42-49.</span>
</li>
<li id="cite_note-22"><span class="mw-cite-backlink"><a href="#cite_ref-22">↑</a></span> <span class="reference-text">Manibusan, M.K., M. Odin and D.A. Eastmond (2007), Postulated carbon tetrachloride mode of action: a review, J Environ Sci Health C Environ Carcinog Ecotoxicol Rev, vol. 25, no. 3,pp. 185-209.</span>
</li>
<li id="cite_note-23"><span class="mw-cite-backlink"><a href="#cite_ref-23">↑</a></span> <span class="reference-text">Masuda, Y. (2006), [Learning toxicology from carbon tetrachloride-induced hepatotoxicity],Yakugaku Zasshi, vol. 126, no. 10, pp. 885-899.</span>
</li>
<li id="cite_note-24"><span class="mw-cite-backlink"><a href="#cite_ref-24">↑</a></span> <span class="reference-text">Nagano, K. et al. (2007), Inhalation carcinogenicity and chronic toxicity of carbon tetrachloride in rats and mice, Inhal Toxicol, vol 19, no. 13, pp. 1089-1103.</span>
<li id="cite_note-26"><span class="mw-cite-backlink"><a href="#cite_ref-26">↑</a></span> <span class="reference-text">Weber, L.W., M. Boll and A. Stampfl (2003), Hepatotoxicity and mechanism of action of haloalkanes: carbon tetrachloride as a toxicological model, Crit Rev Toxicol, vol. 33,
no. 2, pp. 105-136.</span>
</li>
<li id="cite_note-27"><span class="mw-cite-backlink"><a href="#cite_ref-27">↑</a></span> <span class="reference-text">Bauman, J.N. et al. (2009), Can in vitro metabolism-dependent covalent binding data distinguish hepatotoxic from nonhepatotoxic drugs? An analysis using human
hepatocytes and liver S-9 fraction, Chem Res Toxicol, vol. 22, no. 2, pp. 332-340.</span>
</li>
<li id="cite_note-28"><span class="mw-cite-backlink"><a href="#cite_ref-28">↑</a></span> <span class="reference-text">Chen, Q. et al. (1990), The mechanism of cysteine conjugate cytotoxicity in renal epithelial cells. Covalent binding leads to thiol depletion and lipid peroxidation, J Biol Chem, vol. 265, no. 35, pp. 21603-21611.</span>
</li>
<li id="cite_note-29"><span class="mw-cite-backlink"><a href="#cite_ref-29">↑</a></span> <span class="reference-text">Dennehy, M.K. et al. (2006), Cytosolic and nuclear protein targets of thiol-reactive electrophiles, Chem Res Toxicol, vol. 19, no. 1, pp. 20-29.</span>
</li>
<li id="cite_note-30"><span class="mw-cite-backlink"><a href="#cite_ref-30">↑</a></span> <span class="reference-text">Shin, N.Y. et al. (2007), Protein targets of reactive electrophiles in human liver microsomes, Chem Res Toxicol, vol. 20, no. 6, pp. 859-867.</span>
<td><a href="/aops/48">Aop:48 - Binding of agonists to ionotropic glutamate receptors in adult brain causes excitotoxicity that mediates neuronal cell death, contributing to learning and memory impairment.</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/77">Aop:77 - Nicotinic acetylcholine receptor activation contributes to abnormal foraging and leads to colony death/failure 1</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/78">Aop:78 - Nicotinic acetylcholine receptor activation contributes to abnormal role change within the worker bee caste leading to colony death failure 1</a></td>
<td>KeyEvent</td>
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<tr>
<td><a href="/aops/79">Aop:79 - Nicotinic acetylcholine receptor activation contributes to impaired hive thermoregulation and leads to colony loss/failure</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/80">Aop:80 - Nicotinic acetylcholine receptor activation contributes to accumulation of damaged mitochondrial DNA and leads to colony loss/failure</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/87">Aop:87 - Nicotinic acetylcholine receptor activation contributes to abnormal foraging and leads to colony loss/failure </a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/3">Aop:3 - Inhibition of the mitochondrial complex I of nigro-striatal neurons leads to parkinsonian motor deficits</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/144">Aop:144 - Endocytic lysosomal uptake leading to liver fibrosis</a></td>
<td>KeyEvent</td>
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<tr>
<td><a href="/aops/178">Aop:178 - Nicotinic acetylcholine receptor activation contributes to mitochondrial dysfunction and leads to colony loss/failure</a></td>
<td>KeyEvent</td>
</tr>
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<td><a href="/aops/200">Aop:200 - Estrogen receptor activation leading to breast cancer </a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/273">Aop:273 - Mitochondrial complex inhibition leading to liver injury</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/377">Aop:377 - Dysregulated prolonged Toll Like Receptor 9 (TLR9) activation leading to Multi Organ Failure involving Acute Respiratory Distress Syndrome (ARDS)</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/437">Aop:437 - Inhibition of mitochondrial electron transport chain (ETC) complexes leading to kidney toxicity</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/423">Aop:423 - Toxicological mechanisms of hepatocyte apoptosis through the PARP1 dependent cell death pathway </a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/479">Aop:479 - Mitochondrial complexes inhibition leading to left ventricular function decrease via increased myocardial oxidative stress</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/480">Aop:480 - Mitochondrial complexes inhibition leading to heart failure via decreased ATP production</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/481">Aop:481 - AOPs of amorphous silica nanoparticles: ROS-mediated oxidative stress increased respiratory dysfunction and diseases.</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/509">Aop:509 - Nrf2 inhibition leading to vascular disrupting effects through activating apoptosis signal pathway and mitochondrial dysfunction</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/511">Aop:511 - The AOP framework on ROS-mediated oxidative stress induced vascular disrupting effects </a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/256">Aop:256 - Inhibition of mitochondrial DNA polymerase gamma leading to kidney toxicity</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/258">Aop:258 - Renal protein alkylation leading to kidney toxicity</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/464">Aop:464 - Calcium overload in dopaminergic neurons of the substantia nigra leading to parkinsonian motor deficits</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/500">Aop:500 - Activation of MEK-ERK1/2 leads to deficits in learning and cognition via ROS and apoptosis</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/205">Aop:205 - AOP from chemical insult to cell death</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/335">Aop:335 - AOP for urothelial carcinogenesis due to chemical cytotoxicity by mitochondrial impairment </a></td>
<td><a href="/aops/130">Aop:130 - Phospholipase A2 (LPLA2) inhibitors leading to hepatotoxicity</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/497">Aop:497 - ERa inactivation alters mitochondrial functions and insulin signalling in skeletal muscle and leads to insulin resistance and metabolic syndrome</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/34">Aop:34 - LXR activation leading to hepatic steatosis</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/447">Aop:447 - Kidney failure induced by inhibition of mitochondrial electron transfer chain through apoptosis, inflammation and oxidative stress pathways</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/207">Aop:207 - NADPH oxidase and P38 MAPK activation leading to reproductive failure in Caenorhabditis elegans</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/476">Aop:476 - Adverse Outcome Pathways diagram related to PBDEs associated male reproductive toxicity</a></td>
<td><a href="/aops/622">Aop:622 - Calcineurin inhibitor induced nephrotoxicity leading to kidney failure</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/624">Aop:624 - Increased 11β-Hydroxysteroid dehydrogenase type 1 activity leading to MASLD progression via insulin resistance-associated mitochondrial dysfunction</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/627">Aop:627 - Increased 11β-Hydroxysteroid dehydrogenase type 1 activity leading to MASLD progression via lipogenesis-associated mitochondrial dysfunction</a></td>
<td>KeyEvent</td>
</tr>
</tbody>
</table>
</div>
<div>
<!-- loop to find all aops that use this event -->
<p>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).</p>
<p><strong>- Revision of AOP3 (Project: </strong><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><strong>)</strong>: 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. <strong>- Not endorsed</strong></p>
<h4>Key Event Description</h4>
<p>Mitochondrial dysfunction is a consequence of inhibition of the respiratory chain leading to oxidative stress.</p>
<p>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).</p>
<p>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.</p>
<p>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).</p>
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.</p>
<p>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).</p>
<p><strong>Summing up:</strong> 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.</p>
<h4>How it is Measured or Detected</h4>
<p>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.</p>
<p>I. Mitochondrial dysfunction assays assessing a loss-of function.</p>
<p>1. Cellular oxygen consumption.</p>
<p>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 O<sub>2</sub> 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).</p>
<p><strong>- Revision of AOP3 (Project: </strong><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><strong>):</strong> 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). <strong>- Not endorsed</strong> </p>
<p>3. Enzymatic activity of the electron transport system (ETS).</p>
<p>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).</p>
<p>4. ATP content.</p>
<p>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).</p>
<p><strong>Determination of mitochondrial ATP production based on extracellular flux analysis </strong></p>
<p>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. </p>
<p><strong>Calculation of mitochondrial ATP production </strong></p>
<p>Mito ATP production rate (pmol ATP/min) = OCRATP (pmol O2/min) × 2 × P/O </p>
<p>OCR_ATP: ATP-coupled portion of OCR. </p>
<p>Factor 2: each O₂ molecule contains two oxygen atoms. </p>
<p>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). </p>
<p><strong>Limitations</strong> </p>
<p>P/O ratio varies by substrate (glucose vs. fatty acids), cell type, and conditions. Fixed values are approximations. </p>
<p>Non-mitochondrial oxygen consumption (oxidases, peroxidases, etc.) can confound OCR, hence use of ETC inhibitors. </p>
<p>PER vs. ECAR: CO₂-driven acidification must be corrected to avoid overestimating glycolytic ATP. </p>
<p>Normalization: results are usually expressed per cell, protein content, DNA, or mitochondrial mass — interpretation depends on normalization method. </p>
<p><span style="font-size:12.0pt"><span style="font-family:Arial"><span style="color:#212529"><span style="background-color:white"><strong>- Not endorsed</strong></span></span></span></span></p>
</div>
<p><br />
II. Mitochondrial dysfunction assays assessing a gain-of function.</p>
<p>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).</p>
<p>2. mtDNA damage as a biomarker of mitochondrial dysfunction.</p>
<p>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).</p>
<p>3. Generation of ROS and resultant oxidative stress.</p>
<p>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.</p>
<p>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.</p>
<p>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.</p>
<p>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.</p>
<p>e. Detection of hydrogen peroxide (H<sub>2</sub>O<sub>2</sub>) 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 H<sub>2</sub>O<sub>2</sub> form increasing amounts of fluorescent product (Tarpley et al., 2004).</p>
<p>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></p>
<p><strong>Assay Type & Measured Content</strong></p>
</td>
<td><strong>Description</strong></td>
<td><strong>Dose Range Studied</strong></td>
<td>
<p><strong>Assay Characteristics</strong></p>
<p><strong>(Length/Ease of use/Accuracy)</strong></p>
</td>
</tr>
<tr>
<td>
<p><strong>Rhodamine 123 Assay</strong></p>
<p>Measuring Mitochondrial membrane potential (MMP) and its collapse </p>
<p>(Shaki et al., 2012)</p>
</td>
<td>
<p>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.</p>
<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>
<p>Measuring cellular glutathione (GSH) status; ratio of GSH/GSSG</p>
<p>(Owen & Butterfield, 2010; Shaki et al., 2013)</p>
</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>
<p>Short / easy</p>
<p>Low accurancy</p>
</td>
</tr>
<tr>
<td>
<p><strong>TBARS Assay</strong></p>
<p>Quantification of lipid peroxidation</p>
<p>(Yuan et al., 2016)</p>
</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>
<p>Increase in mitochondrial Ca<sup>2+</sup> influx</p>
<p>(Pozzan & Rudolf, 2009)</p>
</td>
<td>Together with GFP, the aequorin moiety acts as Ca<sup>2+</sup> sensor <em>in vivo</em>, 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>
<p>Short / easy</p>
<p>Low accurancy</p>
</td>
</tr>
<tr>
<td>
<p><strong>Western blot & immunostaining analyses</strong></p>
<p>Measuring cytochrome c release</p>
(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>
<p>Short / easy</p>
<p>Medium accurancy</p>
</td>
</tr>
<tr>
<td>
<p><strong>Quantikine Rat/Mouse Cytochrome c Immunoassay</strong></p>
<p>Measuring cytochrome c release</p>
<p>(Shaki et al., 2012)</p>
</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>
<p>Short / easy</p>
<p>Low accurancy</p>
</td>
</tr>
<tr>
<td>
<p><strong>Membrane potential and cell viability – Flow Cytometry</strong></p>
<p>Measuring cytochrome c release</p>
<p>(Kruidering et al., 1997)</p>
</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 <em>g</em>. 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<br />
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 <em>et al.</em>, 1993)”</td>
<td> </td>
<td>
<p>Short / easy</p>
<p>Medium accurancy</p>
</td>
</tr>
</tbody>
</table>
<p> </p>
<p> </p>
<!-- event text -->
<h4>References</h4>
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<h4><a href="/events/40">Event: 40: Mitochondrial ATP production, decreased</a></h4>
<h5>Short Name: Mitochondrial ATP production, decreased</h5>
<td><a href="/aops/26">Aop:26 - Calcium-mediated neuronal ROS production and energy imbalance</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/245">Aop:245 - Reduction in photophosphorylation leading to growth inhibition in aquatic plants</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/258">Aop:258 - Renal protein alkylation leading to kidney toxicity</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/387">Aop:387 - Deposition of ionising energy leading to population decline via mitochondrial dysfunction</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/480">Aop:480 - Mitochondrial complexes inhibition leading to heart failure via decreased ATP production</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/564">Aop:564 - DBDPE-induced inhibition of mitochondrial complex Ⅰ leading to population decline via neurotoxicity and metabotoxicity.</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/590">Aop:590 - Increase of intracellular copper leading to cuproptosis via interference with energy metabolism </a></td>
<td>KeyEvent</td>
</tr>
</tbody>
</table>
</div>
<div>
<div>
<h4><a href="/events/40">40: Decrease, Mitochondrial ATP production</a><br></h4>
<h5>Short Name: Decrease, Mitochondrial ATP production</h5>
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<h4>Domain of Applicability</h4>
<p>All animals with kidneys containing renal proximal tubules.</p>
<h4>Key Event Description</h4>
<p style="text-align:justify">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).</p>
<p><strong>Cell Death and Cytotoxicity</strong></p>
<p style="text-align:justify">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).</p>
<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>
<p>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).</p>
<p>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).</p>
<p>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.</p>
<p>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).</p>
<td><strong>Assay Type & Measured Content</strong></td>
<td><strong>Description</strong></td>
<td><strong>Dose Range Studied</strong></td>
<td>
<p><strong>Assay Characteristics</strong></p>
<strong>(Length/Ease of use/Accuracy)</strong></td>
</tr>
<tr>
<td>
<p><strong>Kidney function assay</strong></p>
<p>Measuring total urinary protein, albumin, transferrin, b2-microglobulin, retinolbinding protein, brush border tubular antigens, N-acetyl-b-Dglucosaminidase activity, serum and urinary creatine</p>
<p> </p>
(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>“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>
<p><strong>MTT Assay (cytotoxicity)</strong></p>
<p>Measuring Cell Viability</p>
(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>
<p>50, 100 and 500 μM of uranyl acetate;</p>
0-1000µM U</td>
<td>
<p>Long</p>
<p>Easy/Difficult</p>
<p>High accuracy (mathematical measurement)</p>
Medium Precision</td>
</tr>
<tr>
<td>
<p><strong>LDH Cytotoxicity Assay</strong></p>
<p>Measuring Necrosis via Lactate Dehydrogenase release</p>
(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>
<p><strong>Caspase-3 and -8 colorimetric assay, Caspase-9 fluoresceine assay</strong></p>
<p>Measuring apoptosis initiation and execution via caspases 3, 8, 9 activity</p>
(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>
<p>“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)</p>
<p>“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)</p>
<p>“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)</p>
<p>“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)</p>
<p>“In the kidney, Cd mainly affects PCT cells. This damage manifests clinically as low molecular weight proteinuria,<br />
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)</p>
<!-- end event text -->
</div>
<h4>References</h4>
<p style="margin-left:30px"> </p>
<p>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></p>
<p>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</p>
<p>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</p>
<p>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</p>
<p>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</p>
<p>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</p>
<p>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</p>
<p>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]</p>
<p>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</p>
<p>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</p>
<p>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</p>
<p>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.</p>
<p>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</p>
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<p>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</p>
<p>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</p>
<p>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</p>
<p>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</p>
<p>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</p>
<p>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</p>
<p>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></p>
<p>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</p>
<p>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</p>
<p>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]</p>
<p>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</p>
<p>Sabath, E., & Robles-Osorio, M. L. (2012). Renal health and the environment: Heavy metal nephrotoxicity. Revista Nefrologia, doi:10.3265/Nefrologia.pre2012.Jan.10928</p>
<p>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</p>
<p>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</p>
<p>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</p>
<p>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</p>
<p>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]</p>
<p>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]</p>
<p>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</p>
<p>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</p>
<p>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</p>
<td><a href="/aops/128">Aop:128 - Kidney dysfunction by decreased thyroid hormone</a></td>
<td>AdverseOutcome</td>
</tr>
<tr>
<td><a href="/aops/256">Aop:256 - Inhibition of mitochondrial DNA polymerase gamma leading to kidney toxicity</a></td>
<td>AdverseOutcome</td>
</tr>
<tr>
<td><a href="/aops/257">Aop:257 - Receptor mediated endocytosis and lysosomal overload leading to kidney toxicity</a></td>
<td>AdverseOutcome</td>
</tr>
<tr>
<td><a href="/aops/258">Aop:258 - Renal protein alkylation leading to kidney toxicity</a></td>
<td>AdverseOutcome</td>
</tr>
<tr>
<td><a href="/aops/377">Aop:377 - Dysregulated prolonged Toll Like Receptor 9 (TLR9) activation leading to Multi Organ Failure involving Acute Respiratory Distress Syndrome (ARDS)</a></td>
<td>KeyEvent</td>
</tr>
<tr>
<td><a href="/aops/437">Aop:437 - Inhibition of mitochondrial electron transport chain (ETC) complexes leading to kidney toxicity</a></td>
<td>AdverseOutcome</td>
</tr>
<tr>
<td><a href="/aops/447">Aop:447 - Kidney failure induced by inhibition of mitochondrial electron transfer chain through apoptosis, inflammation and oxidative stress pathways</a></td>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Higher order animals (mammals) with functional and complete kidneys </span></span></p>
<!-- event text -->
<h4>Key Event Description</h4>
<p style="text-align:justify"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">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. </span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">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 (Cd<sup>2+</sup>, Pb<sup>2+</sup>, Pt<sup>2+</sup>, 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<sup>+2</sup> binding to anionic sites of proximal tubular epithelial brush borders (Shaki et al., 2012). </span></span></p>
<p style="text-align:justify"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">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 <span style="font-size:11.0pt">µ</span>g/kg to 9 <span style="font-size:11.0pt">µ</span>g/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 <span style="color:black">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.</span></span></span></p>
<td><strong><span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif">Assay Type & Measured Content</span></span></strong></td>
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<p><span style="font-size:12.0pt"><span style="font-family:"Times New Roman""><strong>Description</strong></span></span></p>
</td>
<td><strong><span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif">Dose Range Studied</span></span></strong></td>
<td>
<p><span style="font-size:12.0pt"><span style="font-family:"Times New Roman""><strong>Assay Characteristics</strong></span></span></p>
<p><span style="font-size:12.0pt"><span style="font-family:"Times New Roman""><strong>(Length/Ease of use/Accuracy)</strong></span></span></p>
</td>
</tr>
<tr>
<td>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><strong>Kidney Function Assay</strong></span></span></p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Measuring total urinary protein, albumin, transferrin, b2-microglobulin, retinolbinding protein, brush border tubular antigens, N-acetyl-b-Dglucosaminidase activity, serum and urinary creatine</span></span></p>
<p><span style="font-family:"Times New Roman",serif; font-size:12pt">(de Burbure et al., 2003)</span></p>
</td>
<td><span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif">“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)</span></span></td>
<td><span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif">“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”</span></span></td>
<td> </td>
</tr>
<tr>
<td>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><strong>NAG Assay</strong></span></span></p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Measuring N-acetyl-b-D-Glucosaminidase urinary content</span></span></p>
<span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif">(Lim et al., 2016)</span></span></td>
<td><span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif">“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 (</span></span><span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif"><a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4780232/#b13-tr-32-057" style="color:#0563c1; text-decoration:underline">13</a></span></span><span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif">,<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 (</span></span><span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif"><a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4780232/#b13-tr-32-057" style="color:#0563c1; text-decoration:underline">13</a></span></span><span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif">,<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)</span></span></td>
<td>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><strong>Pb: 0.0221ppm</strong><br />
(converted from blood Pb <span style="font-size:11.0pt">µg/dL)</span></span></span></p>
<strong><span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif">Cd: 1.08ppm</span></span></strong><br />
<span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif">(converted from Urinary Cd μg/g creatinine)</span></span></td>
<td><span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif">Fast, easy, accurate</span></span></td>
</tr>
<tr>
<td>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><strong>Kidney Dysfunction Assay </strong></span></span></p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Measuring BUN and creatinine serum blood levels</span></span></p>
<span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif">(Shaki et al., 2012)</span></span></td>
<td><span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif">“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<br />
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)</span></span></td>
<td><span style="font-size:12.0pt"><span style="font-family:"Times New Roman",serif">Fast, easy, medium accuracy </span></span></td>
</tr>
</tbody>
</table>
<p> </p>
<h4>References</h4>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="color:black">Al Dera, H. S. (2016). Protective effect of resveratrol against aluminum chloride induced nephrotoxicity in rats.<em> Saudi Med J, 37</em>(4), 369-378. doi:10.15537/smj.2016.4.13611</span></span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="color:black">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.<em> International Journal of Environmental Research and Public Health, 16</em>, 247. doi:10.3390/ijerph16020274</span></span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="color:black">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</span></span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Ballatori, N. (2002). Transport of toxic metals by molecular mimicry.<em> Environmental Health Perspectives, 110</em>, 689-694. doi:10.1289/ehp.02110s5689</span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Barnes, P., Yeboah, J. K., Gbedema, W., Saahene, R. O., & Amoani, B. (2020). Ameliorative effect of <em>vernonia amygdalina</em> plant extract on heavy metal-induced LIver and kidney dysfunction in rats.<em> Advances in Pharmacological and Pharmaceutical Sciences, 2020</em>, 1-7. doi:10.1155/2020/2976905</span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">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</span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="background-color:white"><span style="color:#303030">Bonventre, J. V., Vaidya, V. S., Schmouder, R., Feig, P., & Dieterle, F. (2010). Next-generation biomarkers for detecting kidney toxicity. </span></span><em><span style="background-color:white"><span style="color:#303030">Nature biotechnology</span></span></em><span style="background-color:white"><span style="color:#303030">, <em>28</em>(5), 436–440. </span></span><a href="https://doi.org/10.1038/nbt0510-436" style="color:#0563c1; text-decoration:underline"><span style="background-color:white">https://doi.org/10.1038/nbt0510-436</span></a></span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="color:black">Brzoska, M. M., Kaminski, M., Supernak-Bobko, D., Zwierz, K., & Moniuszko-Jakoniuk, J. (2003). </span><span style="color:black">Changes in the strucutre and function of the kidney of rats chronically exposed to cadmium. I. biochemical and histopathological studies.<em> Arch.Toxicol., 77</em>, 344-352. doi:10.1007/s00204-003-0451-1</span></span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="color:black">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.<em> Cell Biology International, 41</em>, 1356-1366. doi:10.1002/cbin.10871</span></span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="color:black">Chtourou, Y., Garoui, E. m., Boudawara, T., & Zeghal, N. (2014). </span><span style="color:black">Protective role of silymarin against manganese-induced nephrotoxicity and oxidative stress in rat.<em> </em></span><em><span style="color:black">Environ Toxicol, 29</span></em><span style="color:black">, 1147-1154. doi:10.1002/tox.21845</span></span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="color:black">Durante, P., Romero, F., Perez, M., Chavez, M., & Parra, G. (2010). </span><span style="color:black">Effect of uric acid on nephrotoxicity induced by mercuric chloride in rats.<em> Toxicology and Industrial Health, 26</em>(3), 163-174. doi:10.1177/0748233710362377</span></span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="color:black">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.<em> Evidence-Based Complementary and Alternative Medicine, </em>(424692), 1-19. doi:10.1155/2013/424692</span></span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="color:black">Goldman, M., Yaari, A., Doshnitzki, Z., Cohen-Luria, R., & Moran, A. (2006). Nephrotoxicity of uranyl acetate: Effect on rat kidney brush border membrane vesicles.<em> Archives of Toxicology, 80</em>(7), 387-393. doi:10.1007/s00204-006-0064-6</span></span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="background-color:white"><span style="color:#212121">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.</span></span></span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="color:black">Keith, S., Faroon, O., N., R., Scinicariello, F., Wilbur, S., Ingerman, L., . . . Diamond, G. (2013). <em>Toxicological profile for uranium.</em> </span><span style="color:black">U.S. Department of Health and Human Services. Agency for Toxic Substances and Disease Registry.</span></span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="color:black">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.<em> </em></span><em><span style="color:black">Environ Sci Pollut Res, 21</span></em><span style="color:black">, 12040-12049. doi:10.1007/s11356-014-3142-y</span></span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">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</span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Sabath, E., & Robles-Osorio, M. L. (2012). Renal health and the environment: Heavy metal nephrotoxicity. Revista Nefrologia, doi:10.3265/Nefrologia.pre2012.Jan.10928</span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">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.<em> Archives of Toxicology, 81</em>(7), 495-504. doi:10.1007/s00204-006-0173-2</span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Shaki, F., Hosseini, M. J., Ghazi-Khansari, M., & Pourahmad, J. (2012). Toxicity of depleted uranium on isolated rat kidney mitochondria.<em> Biochimica Et Biophysica Acta - General Subjects, 1820</em>(12), 1940-1950. doi:10.1016/j.bbagen.2012.08.015</span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">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.<em> Toxicology and Industrial Health, 34</em>(7), 469-480. doi:10.1177/0748233718770293</span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">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.<em> Metallomics, 9</em>, 1786. doi:10.1039/c7mt00271h</span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">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.<em> Biological Trace Element Research, 189</em>, 95-108. doi:10.1007/s12011-018-1443-6</span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Weber, E. J., Himmelfarb, J., & Kelly, E. J. (2017). Concise review: Current emerging biomarkers of nephrotoxicity.<em> Curr Opin Toxicol., 4</em>, 16-21. doi:10.1016/j.cotox.2017.03.002</span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="color:black">Yeh, Y., Lee, Y., Hsieh, Y., & Hwang, D. (2011). Dietary taurine reduces zinc-induced toxicity in male wistar rats.<em> Journal of Food Science, 76</em>(4), 90-98. doi:10.1111/j.1750-3841.2011.02110.x</span></span></span></p>
<p style="margin-left:30px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="color:black">Zamora, L. M., Tracy, B. L., Zielinski, J. M., Meyerhof, D. P., & Moss, M. A. (1998). Chronic ingestion of uranium in drinking water: A study of kidney bioeffects in humans.<em> Toxicological Sciences, 43</em>(1), 68-77. doi:10.1006/toxs.1998.242</span></span></span></p>
<h4>Evidence Supporting Applicability of this Relationship</h4>
<div>
</div>
<div>
</div>
<div>
</div>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">The domain of applicability only includes vertebrates, as invertebrates and non-animals do not have kidneys (Mahasen, 2016).</span></span></p>
<h4>Key Event Relationship Description</h4>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">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).</span></span></p>
<h4>Evidence Supporting this KER</h4>
<strong>Biological Plausibility</strong>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">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 (<span style="background-color:white">Lentini et al., 2017</span>). 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).</span></span></p>
<h3>Weight of Evidence Summary</h3>
<p><strong>Concordance of dose-response relationships</strong></p>
<p>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.</p>
<p> </p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">...</span></span></p>
<p><strong>Temporal concordance among the key events and adverse outcome</strong></p>
<p>The scientific evidence on the association between protein alkylation by reactive intermediates and kidney toxicity (AO) is strong and consistent. The MIE is not specific for kidney toxicity and is well established to lead to damage to other organs, whereby the site of toxicity is largely determined by the toxicokinetics of the parent compound or active metabolite.</p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">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). </span></span></p>
<p> </p>
<p><strong>Biological plausibility, coherence, and consistency of the experimental evidence</strong></p>
<p>The described AOP is biologically plausible, coherent and well supported by experimental data.</p>
<p> </p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="background-color:white">“Chronic exposure: </span><span style="background-color:white">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.” (</span><span style="background-color:white">Lentini et al., 2017)</span></span></span></p>
<p><strong>Alternative mechanism(s) that logically present themselves and the extent to which they may distract from the postulated AOP</strong></p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"> “<span style="background-color:white">Acute exposure: </span><span style="background-color:white">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 (</span><a href="https://www.spandidos-publications.com/10.3892/mmr.2017.6389#b17-mmr-15-05-3413" style="color:blue; text-decoration:underline"><span style="background-color:white">17</span></a><span style="background-color:white">).</span></span></span></p>
<p>There are no alternative mechanism(s) that logically present themselves, although a contribution of other mechanisms such as generation of oxidative stress to the overall AO is possible.</p>
<p> </p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="background-color:white"> “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 (</span><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><span style="background-color:white">).” (Lentini et al., 2017)</span></span></span></p>
<p><strong>Uncertainties, inconsistencies and data gaps</strong></p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="background-color:white"><span style="color:black">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 [</span></span><a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5454951/#B11-ijms-18-01039" style="color:blue; text-decoration:underline"><span style="background-color:white"><span style="color:#2f4a8b">11</span></span></a><span style="background-color:white"><span style="color:black">,</span></span><a href="https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5454951/#B12-ijms-18-01039" style="color:blue; text-decoration:underline"><span style="background-color:white"><span style="color:#2f4a8b">12</span></span></a><span style="background-color:white"><span style="color:black">]. 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)</span></span></span></span></p>
<p>This AOP is plausible and consistent with general biological knowledge. However, there is currently little understanding as to which target proteins are critical to toxicity mediated by alkalation damage. Quantitative information on dose response-relationships as well as response-response relationships for upstream and downstream KEs is needed to support its applicability for the development of alternative in vitro tests for nephrotoxicity testing.</p>
<h3>Quantitative Consideration</h3>
<p>Quantitative data on KERs between upstream and downstream KE are still lacking.</p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif"><span style="background-color:white">“In the rat, an acute perfusion of Cd2+ caused hypercalciuria, hyperphosphaturia and hypokaliuria without modification of glomerular filtration rate (GFR) [</span><a href="https://www.karger.com/Article/FullText/83981#ref1" style="color:blue; text-decoration:underline">1</a><span style="background-color:white">]. 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 [</span><a href="https://www.karger.com/Article/FullText/83981#ref13" style="color:blue; text-decoration:underline">13</a><span style="background-color:white">], 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)</span></span></span></p>
<strong>Uncertainties and Inconsistencies</strong>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">There are no currently known inconsistencies or uncertainties for this relationship.</span></span></p>
</div>
<h4>Quantitative Understanding of the Linkage</h4>
<strong>Response-response relationship</strong>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">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).</span></span></p>
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<h2>Considerations for Potential Applications of the AOP (optional)</h2>
<hr>
<p>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).</p>
<strong>Known modulating factors</strong>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">There are several known modulating factors of the relationship between renal tubular cytotoxicity and kidney failure. One modulator of this relationship is age. </span></span></p>
</div>
<strong>Known Feedforward/Feedback loops influencing this KER</strong>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">There are no known feedforward/feedback loops that influence this relationship.</span></span></p>
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<div id="references">
<h2>References</h2>
<hr>
<p>1. Birner, G., et al., <em>Metabolism of tetrachloroethene in rats: identification of N epsilon-(dichloroacetyl)-L-lysine and N epsilon-(trichloroacetyl)-L-lysine as protein adducts.</em> Chem Res Toxicol, 1994. <strong>7</strong>(6): p. 724-32.</p>
<h4>References</h4>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Lentini, P., Zanoli, L., Granata, A., Signorelli, S. S., Castellino, P., & Dell'aquila, R. (2017). </span></span></p>
<p>2. Pahler, A., et al., <em>Generation of antibodies to Di- and trichloroacetylated proteins and immunochemical detection of protein adducts in rats treated with perchloroethene.</em> Chem Res Toxicol, 1998. <strong>11</strong>(9): p. 995-1004.</p>
<p style="margin-left:48px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Kidney and heavy metals - the role of environmental exposure (review).<em> Molecular Medicine Reports, 15</em>(3413), 3419. doi:10.3892/mmr.2017.6389</span></span></p>
<p>3. Kleiner, H.E., et al., <em>Immunochemical detection of quinol--thioether-derived protein adducts.</em> Chem Res Toxicol, 1998. <strong>11</strong>(11): p. 1283-90.</p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Mahasen, L. M. A. (2016). Evolution of the kidney.<em> Anatomy Physiol. Biochem. Int. J., 1</em>(1), </span></span></p>
<p>4. Lau, S.S., <em>Quinone-thioether-mediated nephrotoxicity.</em> Drug Metab Rev, 1995. <strong>27</strong>(1-2): p. 125-41.</p>
<p style="margin-left:48px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">555554. doi:10.19080/APBIJ.2016.01.555554</span></span></p>
<p>5. Tune, B.M., <em>Nephrotoxicity of beta-lactam antibiotics: mechanisms and strategies for prevention.</em> Pediatr Nephrol, 1997. <strong>11</strong>(6): p. 768-72.</p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Orr, S. E., & Bridges, C. C. (2017). Chronic kidney disease and exposure to nephrotoxic </span></span></p>
<p>6. Griffin, R.J. and P.J. Harvison, <em>In vivo metabolism and disposition of the nephrotoxicant N-(3, 5-dichlorophenyl)succinimide in Fischer 344 rats.</em> Drug Metab Dispos, 1998. <strong>26</strong>(9): p. 907-13.</p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">metals.<em> International Journal of Molecular Sciences, 18</em> doi:10.3390/ijms18051039</span></span></p>
<p>7. Groves, C.E., et al., <em>Pentachlorobutadienyl-L-cysteine (PCBC) toxicity: the importance of mitochondrial dysfunction.</em> J Biochem Toxicol, 1991. <strong>6</strong>(4): p. 253-60.</p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Priante, G., Gianesello, L., Ceol, M., Del Prete, D., & Anglani, F. (2019). Cell death in the </span></span></p>
<p>8. Chen, Y., et al., <em>Role of mitochondrial dysfunction in S-(1,2-dichlorovinyl)-l-cysteine-induced apoptosis.</em> Toxicol Appl Pharmacol, 2001. <strong>170</strong>(3): p. 172-80.</p>
<p style="margin-left:48px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">kidney.<em> International Journal of Molecular Sciences, 20</em>(14), 3598. doi: 10.3390/ijms20143598. doi:10.3390/ijms20143598</span></span></p>
<p>9. Hill, B.A., T.J. Monks, and S.S. Lau, <em>The effects of 2,3,5-(triglutathion-S-yl)hydroquinone on renal mitochondrial respiratory function in vivo and in vitro: possible role in cytotoxicity.</em> Toxicol Appl Pharmacol, 1992. <strong>117</strong>(2): p. 165-71.</p>
<p><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Sano, K., Fujigaki, Y., Miyaji, T., Ikegaya, N., Ohishi, K., Yonemura, K., & Hishida, A. (2000). </span></span></p>
<p>10. Aleo, M.D., et al., <em>Toxicity of N-(3,5-dichlorophenyl)succinimide and metabolites to rat renal proximal tubules and mitochondria.</em> Chem Biol Interact, 1991. <strong>78</strong>(1): p. 109-21.</p>
<p style="margin-left:48px"><span style="font-size:12pt"><span style="font-family:"Times New Roman",serif">Role of apoptosis in uranyl acetate-induced acute renal failure and acquired resistance to uranyl acetate.<em> Kidney International, 57</em>(4), 1560-1570. doi:10.1046/j.1523-1755.2000.00777.x</span></span></p>