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Key Event Title
Increase, Cytotoxicity (renal tubular cell)
|Level of Biological Organization|
|kidney tubule cell|
Key Event Components
|cell death||kidney tubule cell||increased|
Key Event Overview
AOPs Including This Key Event
|AOP Name||Role of event in AOP||Point of Contact||Author Status||OECD Status|
|α2u-globulin- renal adenomas/carcinomas||KeyEvent||Charles Wood (send email)||Under Development: Contributions and Comments Welcome|
|Inhibition of mitochondrial DNA polymerase gamma leading to kidney toxicity||KeyEvent||Angela Mally (send email)||Under development: Not open for comment. Do not cite||Under Development|
|Receptor mediated endocytosis and lysosomal overload leading to kidney toxicity||KeyEvent||Angela Mally (send email)||Under development: Not open for comment. Do not cite||Under Development|
|Renal protein alkylation leading to kidney toxicity||KeyEvent||Angela Mally (send email)||Not under active development||Under Development|
|Inhibition of Mt-ETC complexes leading to kidney toxicity||KeyEvent||Baki Sadi (send email)||Under development: Not open for comment. Do not cite|
Key Event Description
The renal proximal tubule is a crucial section of the nephron, responsible for the bulk of its reabsorption capabilities. About 60-70% of glomerular filtrate such as water, small molecules, and important ions, as well as nearly all the filtered amino acids, small peptides, and glucose are reabsorbed in the proximal tubule (Carson, 2019). The process of solute reabsorption is highly energetically expensive, making the proximal tubules the renal region of highest oxygen consumption. The microvilli, densely packed to form the brush border apical surface of the tubules, have abundant elongated mitochondria to sustain the energetic demand of their function (Carlson, 2019). The introduction of heavy metals into the kidneys causes aggregation in the proximal tubules due to their high mitochondrial content, leading to inhibition of the electron transport chain and reactive oxygen species (ROS) production. This area is particularly susceptible to heavy metal toxicity due to the abundance of mitochondria, as well as the fact that, regardless of toxicity, approximately 70% of cation absorption and transport passes through the proximal tubules (Barbier et al., 2005). Some heavy metal transport into the proximal tubules is conducted by MRP-1 and MRP-2 (ATP binding cassette-multidrug resistance proteins), and characterize toxicity by GSH depletion as some metals such as arsenic bind GSH and increased oxidative stress induced by free radicals (Sabath & Robles-Osorio, 2012). This oxidative stress causes disruption to mitochondrial homeostasis and mitophagy in proximal tubular epithelial cells by altering PPAR (peroxisome proliferator-activated receptor) (Small et al., 2018). At high enough concentrations of toxic heavy metals they can lead to cytotoxicity and cell death. An issue with assessment of kidney function is that the kidneys notoriously compensate for loss of function, leading to the appearance of adverse affects only at a late onset when there is very severe levels of damage (de Burbure et al., 2003).
Cell Death and Cytotoxicity
Cell death is a variety of processes defined by a cell ceasing to perform its function. This could happen by a variety of mechanisms. Apoptosis is a programmed physiological sequence leading to controlled cell death deemed necessary for the fitness and survival of the organism (cell is redundant, dysfunctional, cancerous, etc.) (Choi et al., 2019). Apoptosis, in the case of DNA damage, can be induced by free radicals produced as a result of heavy metal exposure, as shown in ex-vivo studies (Miller et al., 2002). Another cause by heavy metal exposure is physical and structural damage to mitochondria, damaging cellular metabolism and ATP production. There are many possible stressors that may lead to cell death, the effects exhibited depend on the cell type and the severity of the stress (Liu et al., 2018). Some modes of cell death include: apoptosis (programmed cell death), necrosis (uncontrolled cell death), and aging-caused cell death, known as senescent death (Liu et al., 2018).
The extrinsic pathway, also known as the death receptor-mediated pathway, involves the ligation of death receptors determining the activation of caspase-8. Caspase-8 further activates downstream caspases leading to apoptosis (Priante et al., 2019). This pathway is triggered by extracellular signalling proteins binding to cell-surface death receptors. A well understood example of this process is the activation of the Fas receptor on the surface of a target cell by Fas ligand (FasL) on the surface of a cytotoxic lymphocyte (Alberts et al., 2014). In this process, the cytosolic Fas death receptor binds intracellular adaptor proteins. This complex then binds initiator, caspases, primarily caspase-8, forming a death-inducing signalling complex (DISC). The initiator caspases, once dimerized and activated in the DISC, activate downstream executioner caspases to induce apoptosis (Nair et al., 2014). In some cells, the extrinsic pathway recruits the intrinsic apoptotic pathway to amplify the caspase cascade. These pathways are linked by caspase-8, that triggers the caspase cascade and the protein, Bid (Priante et al., 2019; Alberts et al., 2014). Type I cells act independent of mitochondria for the induction of Fas death receptor-mediated apoptosis, and have therefore optimized the extrinsic pathway. Thymocytes or cells responsible for the immune system in general, for example, are expected to signal each other or target cells through membrane bound ligands, like FasL and TRAIL (Ozoren and El-Deiry, 2002).
The intrinsic pathway is often referred to as the mitochondrial pathway of apoptosis. Pro-apoptotic Bcl-2 family proteins, Bax and Bak, create pores on the outer mitochondrial membrane, determining the release of apoptogenic factors, such as cytochrome c. In the cytosol, cytochrome c binds to, and stimulates, conformational modifications in the adaptor protein, Apaf-1, thus leading to the enrolment and activation of caspase-9. Caspase-9 further activates executioner caspases to elicit apoptosis (Priante et al., 2019). Type II cells are mitochondria-dependent, where the mitochondria are crucial to ensure successful apoptosis. For example, liver and kidney cells are responsible for the detoxification of the blood from chemicals toxicants, many of which are cytotoxic and genotoxic agents known to predominantly activate the intrinsic pathway (Ozoren and El-Deiry, 2002).
In a study conducted by Eichler et al. (2006), cultured murine podocytes were incubated for three days with arsenite, cadmiuim, or mercury, as well as an equimolar combination of the three to test the modes and extent of apoptosis induced by the exposure. It was seen that the mix of metal exposure showed significantly fewer apoptotic affects, indicating an antagonistic affect of the metals over an additive or synergistic toxicity. It was also seen that the apoptosis observed in the separate metal tests showed a ~400% increase of caspase 8 activity as well as ~500% upregulation of Fas, factors of the extrinsic pathway. No significant change was seen to the intrinsic pathway factors. The results of this experiment indicate that heavy metals favour extrinsic apoptosis as their method of cytotoxicity.
Necrosis is characterized as passive, accidental cell death resulting from environmental perturbation with uncontrolled release of inflammatory cellular contents (Fink & Cookson, 2005). Contrastingly, apoptosis is an active, intentional, programmed process of autonomous cellular dismantling that avoids eliciting inflammation. These modes would then be categorized into Accidental Cell Death (ACD) and Regulated Cell Death (RCD), respectively fitting necrosis and apoptosis (Choi et al., 2019). Necrosis biochemically manifests through plasma membrane rupture, cell swelling and lysis, energy decline, DAMP release, and emptying of cell contents (Choi et al., 2019; Thiebault et al., 2007). The caspases governing inflammatory cell death, such as necrosis, are caspases-1, -4, -5, -11, -12, -13, and -14 (Fink and Cookson, 2005). Cell fate could be decided by a number of factors. For instance, ATP is required for the execution of apoptosis, so, when lacking, apoptosis is disabled, making the mode of cell death ATP dependent (Shaki et al., 2012). Between apoptosis and necroptosis, cell fate is influenced primarily by the availability of caspase-8 and the cellular or X-linked inhibitors of apoptosis proteins (cIAP1, cIAP2, XIAP). Thiebault et al. (2007) studied the mechanism of cell mortality induced by uranium in NRK-52E cells and found that after low exposure to uranium (below the CI50 concentration, 500µL), apoptotic cell death was observed, whereas higher exposure to uranium resulted in necrotic cell death. Multiple types of death can be observed simultaneously in tissues exposed to the same stimulus, and the local intensity of a particular stimulus may influence the cell death mechanism (Fink and Cookson, 2005).
How It Is Measured or Detected
|Assay Type & Measured Content||Description||Dose Range Studied||
Assay Characteristics(Length/Ease of use/Accuracy)
Kidney function assay
Measuring total urinary protein, albumin, transferrin, b2-microglobulin, retinolbinding protein, brush border tubular antigens, N-acetyl-b-Dglucosaminidase activity, serum and urinary creatine(de Burbure et al., 2003)
|“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)||“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”|
N-ACETYL-b-D-GLUCOSAMINIDASE (NAG) ASSAY
Measuring NAG urinary content(Lim et al., 2016)
|“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 (13,14). 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 (13,14).” (Lim et al., 2016)||Cd & Pb||Fast, easy, accurate|
MTT Assay (cytotoxicity)
Measuring Cell Viability(Thiebault et al., 2007; Shaki et al., 2012)
|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.||
50, 100 and 500 μM of uranyl acetate;0-1000µM U
High accuracy (mathematical measurement)Medium Precision
LDH Cytotoxicity Assay
Measuring Necrosis via Lactate Dehydrogenase release(Thiebault et al., 2007)
|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.||15, 30 µM Cd||Fast, easy, high accuracy|
Caspase-3 and -8 colorimetric assay, Caspase-9 fluoresceine assay
Measuring apoptosis initiation and execution via caspases 3, 8, 9 activity(Thiebault et al., 2007)
|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.||0-800µM U||Long, difficult, high accuracy|
“Techniques such as micropuncture, microinjection [1, 6, 18] and microperfusion of isolated tubules  have made it possible to map the reabsorption of the heavy metals along the different segments of the nephron.” (Barbier et al., 2005)
“Pb2+ , Hg2+ induced glomerular and tubular damage characterized by a reduced GFR, glycosuria, proteinuria and a rapid obstruction of the tubular system ” (Barbier et al., 2005)
“Concerning chronic intoxication, most heavy metals (Cd2+ , Hg2+ , Pb2+ ) induced a Fanconi syndrome characterized by a decrease of the GFR, an increase in urinary flow rate, proteinuria, glycosuria, aminoaciduria and excessive loss of major ions.” (Barbier et al., 2005)
“In the proximal tubule, Cd2+ has been shown to decrease phosphate and glucose transport by inhibiting the NaPi and the Na/glucose cotransporters respectively.” (Barbier et al., 2005)
“In the kidney, Cd mainly affects PCT cells. This damage manifests clinically as low molecular weight proteinuria, aminoaciduria, bicarbonaturia, glycosuria and phosphaturia. Tubular damage markers such as alpha-1-microglobulin, beta-2-microglobulin, NAG and KIM-1 (kidney injury molecule-1) are useful in detecting early tubular damage.” (Sabath & Robles-Osorio, 2012)
Domain of Applicability
All animals with kidneys containing renal proximal tubules.
Evidence for Perturbation by Stressor
Belyaeva et al. (2012) investigated the effects of cadmium on cell viability of human kidney cells. When they observed the release of lactate dehydrogenase (LDH) after different incubation times they found that the kidney cells treated with 500 μM of cadmium released significant LDH (Belyaeva et al. 2012). Belyaeva et al. (2012) also looked at the LDH release in kidney cells treated with mercury and found that mercury treatment was more toxic than cadmium treatment, as it showed significant LDH release at a lower dosage of treatment.
Hinkle et al. (1987) treated rat pituitary gland neoplasm cells with cadmium to assess the cytotoxicity of the heavy metal treatment. Their results showed a dose-dependant decrease in association with cadmium treatment up to 15 μM (Hinkle et al., 1987).
In their study of the effects of cadmium treatment on human embryonic kidney cells, Chomchan et al. (2018) determined that cadmium treatment caused a dose-dependant decrease in cell viability when treating cells with 0 to 100 μM. The IC50 value determined in this study was 68.50 μg/mL (Chomchan et al., 2018).
Mezynska et al. (2019) treated rats with cadmium and observed the cytotoxicity of the liver. They found that the lipid peroxides (LPO) released by the treated cells was significantly increased as early as 3 months into 1 mg/kg treatment and was the highest at 10 months (Mezynska et al., 2019). When treated with 5 mg/kg of cadmium treatment was significant as early as 3 months and was the most affected at 10 months (Mezynska et al., 2019).
Belyaeva et al. (2012) conducted a study to determine the effect of mercury treatment on rat kidney cell (PC12 cells) viability and found that treatment with 50 μM of mercury for 24 hours resulted in significant lactate dehydrogenase (LDH) release.
Rouas et al. (2010) treated human embryonic kidney cells (HEK-293) with depleted uranium of varying concentrations for 24 or 48 hours to assess the effect on cell viability. They found that the cells showed a time- and dose-dependant increase in cytotoxicity, as the 24 hour treatment was not significant below 700 μM but increased up to 1000 μM. The 48 hour treatment was significant at as low as 100 μM and showed dose-dependant increase up to 1000 μM (Rouas et al., 2010).
Shaki et al. (2012) investigated the effects of uranium on rat kidney cell viability, finding that cytochrome c increases were significant in cells treated with 100 μM for 24 hours.
In their investigation of the effects of uranium treatment on human kidney cells, Hao et al. (2014) learned that cells treated with depleted uranium had significantly increased levels of caspase-3, caspase-8, and caspase-9. They also found that the treated cells released more mitochondrial cytochrome c and lactate dehydrogenase (LDH) and had increased levels of Bax, while Bcl-2 levels were decreased (Hao et al., 2014). These results indicate an increase in apoptosis in cells treated with depleted uranium (Hao et al., 2014). Hao et al. (2014) also directly observed cytotoxicity in the treated cells and found dose- and time-dependant decreases in cell viability when cells were treated with 0 to 700 μM of depleted uranium for 0 to 48 hours.
In a study conducted by Hao et al. (2016) they assessed the effects of depleted uranium on kidney mitochondria in human embryonic kidney cells. They found that the treated cells showed significant increases in mitochondrial damage and subsequent apoptosis (Hao et al., 2016).
Guéguen et al. (2015) treated human hepatocyte carcinoma cells with uranium to determine cytotoxicity. Their results showed that caspase 3/7 activity was significantly increased in a dose-dependant manner when cells were treated with 300 to 1000 μM for 4, 6, 12, to 24 hours (Guéguen et al., 2015).
Yu et al. (2021) investigated the effects of uranium on human kidney cells and found that the treated cells showed time- and dose-dependant decreases in cell viability when treated with concentrations from 1 to 10000 μM and when treated for 5 to 40 hours. The IC50 value of uranium was determined to be 520 μM for uranium in this study (Yu et al., 2021).
Muller et al. (2006) treated pig kidney cells with uranium and found both time- and dose-dependant increases in cytotoxicity when treated with 0 to 1.4 mM of uranium for 0 to 35 hours.
Miyayama et al. (2013) investigated the effects of silver treatment on human lung epithelial cells and found that silver showed a dose dependant decrease in viability which was significant between the doses 5 and 100 μM.
Turk et al. (2019) treated rats with arsenic and observed the kidneys for biochemical changes. Their results showed increased caspase-3 activity in the treated kidneys, indicating an increase in cellular death when the cells were treated with arsenic (Turk et al., 2019).
In their study of the effects of gold (III) treatment on rat kidney cells, Sprekelmeyer et al. (2017) found that the treated kidneys showed a dose-dependant decrease in viability when treated with 0 to 10 μM. The IC50 value determined for gold in this study was 4.3 μM (Spreckelmeyer et al., 2017).
Nanoparticles and Micrometer Particles
Zhang et al. (2018) conducted a study investigating the effects of copper nanoparticle treatment on pig kidney cells. Their results showed that pig kidney cells experience dose- and time-dependant decreases in cell viability when treated with 60 μg/mL of copper nanoparticles for 6 hours or more, or 20 μg/mL for 12 hours or more (Zhang et al., 2018).
Karlsson et al. (2009) investigated the effects of varying heavy metal nanoparticles and micrometer particles on human alveolar type-II epithelial cells and found that only copper nanoparticles, copper micrometer particles, and iron(II) nanoparticles caused a significant increase in cytotoxicity when used to treat cells. Copper nanoparticles were the most toxic treatment, causing complete cytotoxicity in the treated cells, while copper micrometer particles were only able to cause a 31% increase in cytotoxicity and iron(II) nanoparticles were only able to cause a 5% increase in non-viable cells (Karlsson et al., 2009).
Pan et al. (2009) investigated the effects of gold nanoparticles (Au1.4MS) on human cervix carcinoma cells and found that the treated cells experience a dose-dependant increase in cytotoxicity, which resulted in the determination of an IC50 of 48 μM. When they assayed the histological effect of the nanoparticles, Pan et al. (2009) also found that the treated cells showed increased cell death.
Liu et al. (2010) found that when rat kidney cells were treated with titanium oxide nanoparticles, they showed a time- and dose-dependant decrease in cell viability, with significance occurring at 6 hours for 100 μg/mL and 12 hours for 10 and 50 μg/mL.
Santos et al. (2007) investigated the effects of cisplatin treatment on rat kidneys and found that the treated rats had significantly elevated levels of caspase-3 activity, implying an increase in apoptosis in the treated cells.
Sprekelmeyer et al. (2017) also found that rat kidney cells treated with cisplatin showed dose dependant decreases in viability when treated with 0 to 100 μM. The IC50 value determined from this article was 17.0 μM (Spreckelmeyer et al., 2017).
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