Relationship: 2877: Conjugation, GSH leads to Depletion, GSH

AOPs Referencing Relationship

Evidence Supporting this KER

GSH is an antioxidant generated in various kinds of cells, however, in vertebrates, this takes place mainly in liver cells, from where it is exported to other cell types  (Lu 2013). Around 85% of free GSH is found in the cytoplasm, from where it is distributed to organelles such as mitochondria, which stores approximately 10% of the total GSH content, endoplasmic reticulum and extracellular space (Lu 2013; Aquilano, Baldelli, and Ciriolo 2014). Depletion of free GSH content happens because of the sulfhydryl group of the cysteine residue of this tripeptide reacts with xenobiotics during detoxification process, producing conjugates, which are secreted directly into the bile or converted to mercapturic acids and excreted into the urine, as well as due to the reaction with other reactive species as ROS. Nevertheless, unlike what happens to glycine and glutamate residues with oxidized glutathione (GSSG), which are recycled respectively from detoxification of xenobiotics and ROS-mediated oxidation, the cysteine molecule from GSH is excreted from the organism as a byproduct conjugated to the toxic molecule, causing, thereby, reduction of cellular levels of this limiting amino acid for the tripeptide production. In this way, restoration of regular intracellular GSH levels, via de novo synthesis and still from the reaction of reduction of oxidized glutathione (GSSG) ends up being hampered (X. Li 2009; Lushchak 2012; Gupta 2016; Aquilano, Baldelli, and Ciriolo 2014) and GSH levels are, consequently, depleted.

In vertebrate animals, chemicals such as ATZ (Egaas et al. 1993; Wiegand et al. 2001; Elia, Waller, and Norton 2002; Abel et al. 2004; McMullin et al. 2007; LeBlanc and Sleno 2011), DEM (Combes and Backof 1982; Kubal et al. 1995) and Hg (Stricks and Kolthoff 1953; Valko, Morris, and Cronin 2005) are metabolized by GST through reduced GSH-binding in phase II of biotransformation, generating GSH conjugates.

In vitro and in vivo data reveal that ATZ leads to GSH depletion through that pathway in various fish species. For instance, 25 μg/mL of this chemical uses up GSH neutrophil in common carp after 2 – 3 h of exposure (Wang et al. 2019). In young Prochilodus lineatus, after 24h of exposure at the concentrations of 2 and 10 μg/L, this herbicide did not display any effect on hepatic GSH levels, but, after 48 h, ATZ induced a significant decrease in the content of this biomarker (Santos and Martinez 2012). And in studies of acute toxicity, this chemical caused depletion of GSH level in both young catfish (Rhamdia quelen) (Mela et al. 2013) and in embryos of zebrafish (Danio rerio) (Adeyemi, da Cunha Martins-Junior, and Barbosa 2015) after 96 h of exposure, at the concentrations of 100 μg/L and 0.1 mM, respectively. Confirming these data, at longer exposures in adult female zebrafish, ATZ also underwent a decrease in ovary and liver GSH levels at concentrations above 1 and 10 μg/L, respectively, after 14 days of exposure (Jin et al. 2010). Similarly, common carp submitted to 1/5 of LC50 (96-h)  of ATZ for 40 days, showed GSH levels significantly (p < 0.05)  reduced in liver cells (Toughan et al. 2018). 

As in fishes, this drop in GSH levels is also observed in different organs and tissues of mammals. Albino male rats orally treated with ATZ (200 mg/Kg of body weight/day), for a period of 30 days, exhibited a decrease in brain, hippocampus and submandibular salivary gland GSH contents (Ahmed et al. 2022). Moreover, Sprague-Dawley male rats ATZ-exposed via gavage, for 30 days, showed reduction of total antioxidant capacity in a dose-dependent manner, as well as a significant decrease of free GSH level in testicles of these animals (Song et al. 2014). 

GSH-depleting agent DEM, likewise, is able to induce a drop in testicular GSH in BALB/c mice. 52 μM of this chemical intraperitoneally injected, during two weeks, leads to a significant reduction of free GSH levels in testicles of these animals (Kalia and Bansal 2008). (Kaur, Kalia, and Bansal 2006) had previously found evidences of relevant diminishment (p < 0.001) of GSH content and elevation of GSSG levels in testicles of this same animal strain daily submitted to DEM intraperitoneal injection, at 8.7 μM, for two weeks. 

Regarding Hg, several taxons also have their GSH levels affected in organs and varied tissues. Adult female zebrafish exposed to 15 and 30 μg/L for a period of 30 days, exhibited a reduction of GSH content in ovaries in a dose-dependent manner (Zhang et al. 2016). In male albino Wistar rats, a single dose (5 mg/Kg bw) of a mercury (II) chloride (HgCl2) solution subcutaneously administered, three times a week, for 60 days, was also able to negatively change GSH testicular content (El-Desoky et al. 2013). Nevertheless, the authors also noticed that animals treated with Spirulina platensis (300 mg/Kg bw), by gavage for 10 consecutive days, before mercury (II) chloride administration and continued up to 60 days along with HgCl2, did not suffer changes in GSH levels, emphasizing downstream KE essentiality, once this can be prevented. This GSH reduction is also seen in bird for Hg. Hy-Line Brown laying hens fed with four experimental diets containing gradual levels of mercury at 0.280, 3.325, 9.415 e 27.240 mg/Kg, respectively, for a period of 10 weeks, displayed GSH content considerably decreased in all Hg-treated groups (Ma et al. 2018).

Hence, it is noted that GSH depletion caused by chemicals happens in all stages of live in teleosts, as well as adult mammals and birds, showing that this KER is conserved among these taxa, which is expected, since this antioxidant participates in basic cellular processes in vertebrate organisms. However, the time necessary for this response varies depending on the different stages and among species, as well as it is dependent on the dose/concentration applied. Still, this is not surprising, because toxicokinetics for chemicals obviously differs among taxa and depends on some variables, such as uptake and solubility.

Quantitative Understanding of the Linkage

GSH depletion depends on the constant conjugation rate of GSH to a xenobiotic, from the initial GSH concentration and its synthesis and degradation rates. Several chemicals that undergo GSH conjugation at high concentrations cause depletion of GSH supplies in the liver and others tissues (D’Souza, Francis, and Andersen 1988; D’Souza and Andersen 1988; Csanády et al. 1996; Mulder and Ouwerkerk-Mahadevan 1997; Fennell and Brown 2001).

In this context, the global kinetic equation for GSH consumption through conjugation to xenobiotics, catalyzed by microsomal glutathione transferase 1 (mGST1), purified from rat liver can be defined by (Spahiu et al. 2017) (figure below.). In this equation, C is the electrophilic substrate, while E represents the enzyme and P serves as a GSH-conjugate. In relation to constants, k₂ is the rate for thiolate anion, k_-2 is the rate for the reverse process of thiolate anion, k₃ is the rate for the chemical step that is essentially irreversible, K_C is the dissociation constant for electrophilic substrate and K_G is the dissociation constant for GSH (Spahiu et al. 2017). 

Moreover, thiolate anion formation (k_obs) can be easily calculated through equations described by (Morgenstern et al. 2001). Kinetic parameters K_M e k_cat values for both electrophiles and GSH can also be determined according to the equations established by (Spahiu et al. 2017).Furthermore, nucleophilic reactivity (N) and electrophilicity (E) parameters of GSH have also been settled to a variety of Michael acceptors (Mayer and Ofial 2019). 

Velocity of conjugation, however, depends on the kind of GST involved and on the chemical, as well as the organism in which it takes place. For instance, the ATZ-GSH conjugate formed in GSTs from zebrafish embryos works in a time-dependent manner, although conjugation in the microsomal GST increased linearly by a factor of 23 up to 12 h of incubation time, whereas in the soluble GST the conversion rate increased more slowly and was higher by a factor of 5.8 after 24 h of incubation time than that at start (Wiegand et al. 2001). In rats, the estimated GSH conjugation rate constant with ATZ was 0.53 L/mmol/h, a value comparable to that for other chemicals that are largely conjugated by GSTs, even so less than known depleters such as ethylene dichloride (1.2 L/mmol/h) and allyl chloride (9.0 L/mmol/h). Although ATZ is mostly metabolized by GSH, the model estimated that 50% depletion of GSH is predicted to occur, but only after three daily doses of 500 mg ATZ/Kg  (McMullin et al. 2003).

In humans, intrahepatic glutathione concentration is predicted to be the lowest one, due to conjugation to the reactive intermediate NAPQI, at 6 h after 2 g of intravenous (IV) infusion administration of paracetamol and then to recover slowly. In addition, it responds in a time-dependent way. However, concentrations of glutathione were predicted to be markedly and progressively depleted when patients had an initial 2 g dose and then 1 g dose every 6 h (Geenen et al. 2013).

(Hughes, Miller, and Swamidass 2015), for example, constructed a model to predict the GSH reactivity to 1213 molecules and determined the percent depletion of GSH after 15 min incubation with each molecule. In this context, such a model can be easily used for investigation and initial selection of molecules that might impair fertility.

References

Lu, Shelly C. 2013. “Glutathione Synthesis.” Biochimica et Biophysica Acta 1830 (5): 3143–53.

Aquilano, Katia, Sara Baldelli, and Maria R. Ciriolo. 2014. “Glutathione: New Roles in Redox Signaling for an Old Antioxidant.” Frontiers in Pharmacology 5 (August): 196.

Li, Xianchun. 2009. “Glutathione and Glutathione-S-Transferase in Detoxification Mechanisms.” In General, Applied and Systems Toxicology. Chichester, UK: John Wiley & Sons, Ltd. https://doi.org/10.1002/9780470744307.gat166.

Lushchak, Volodymyr I. 2012. “Glutathione Homeostasis and Functions: Potential Targets for Medical Interventions.” Journal of Amino Acids 2012 (February): 736837.

Gupta, P. K. 2016. “Chapter 8 - Biotransformation.” In Fundamentals of Toxicology, edited by P. K. Gupta, 73–85. Academic Press.

Egaas, E., J. U. Skaare, N. O. Svendsen, M. Sandvik, J. G. Falls, W. C. Dauterman, T. K. Collier, and J. Netland. 1993. “A Comparative Study of Effects of Atrazine on Xenobiotic Metabolizing Enzymes in Fish and Insect, and of the Invitro Phase II Atrazine Metabolism in Some Fish, Insects, Mammals and One Plant Species.” Comparative Biochemistry and Physiology. Part C, Pharmacology, Toxicology & Endocrinology 106 (1): 141–49.

Wiegand, C., E. Krause, C. Steinberg, and S. Pflugmacher. 2001. “Toxicokinetics of Atrazine in Embryos of the Zebrafish (Danio Rerio).” Ecotoxicology and Environmental Safety 49 (3): 199–205.

Elia, A. C., W. T. Waller, and S. J. Norton. 2002. “Biochemical Responses of Bluegill Sunfish (Lepomis Macrochirus, Rafinesque) to Atrazine Induced Oxidative Stress.” Bulletin of Environmental Contamination and Toxicology 68 (6): 809–16.

Abel, Erika L., Shaun M. Opp, Christophe L. M. J. Verlinde, Theo K. Bammler, and David L. Eaton. 2004. “Characterization of Atrazine Biotransformation by Human and Murine Glutathione S-Transferases.” Toxicological Sciences: An Official Journal of the Society of Toxicology 80 (2): 230–38.

McMullin, Tami S., William H. Hanneman, Brian K. Cranmer, John D. Tessari, and Melvin E. Andersen. 2007. “Oral Absorption and Oxidative Metabolism of Atrazine in Rats Evaluated by Physiological Modeling Approaches.” Toxicology 240 (1-2): 1–14.

LeBlanc, André, and Lekha Sleno. 2011. “Atrazine Metabolite Screening in Human Microsomes: Detection of Novel Reactive Metabolites and Glutathione Adducts by LC-MS.” Chemical Research in Toxicology 24 (3): 329–39.

Combes, B., and B. Backof. 1982. “Effect of Diethyl Maleate on the Biliary Excretion Rate of Infused Sulfobromophthalein-Glutathione.” Biochemical Pharmacology 31 (16): 2669–74.

Kubal, G., D. J. Meyer, R. E. Norman, and P. J. Sadler. 1995. “Investigations of Glutathione Conjugation in Vitro by 1H NMR Spectroscopy. Uncatalyzed and Glutathione Transferase-Catalyzed Reactions.” Chemical Research in Toxicology 8 (5): 780–91.

Stricks, W., and I. M. Kolthoff. 1953. “Reactions between Mercuric Mercury and Cysteine and Glutathione. Apparent Dissociation Constants, Heats and Entropies of Formation of Various Forms of Mercuric Mercapto-Cysteine and -Glutathione.” Journal of the American Chemical Society 75 (22): 5673–81.

Valko, M., H. Morris, and M. T. D. Cronin. 2005. “Metals, Toxicity and Oxidative Stress.” Current Medicinal Chemistry 12 (10): 1161–1208.

Wang, Shengchen, Qiaojian Zhang, Shufang Zheng, Menghao Chen, Fuqing Zhao, and Shiwen Xu. 2019. “Atrazine Exposure Triggers Common Carp Neutrophil Apoptosis via the CYP450s/ROS Pathway.” Fish & Shellfish Immunology 84 (January): 551–57.

Santos, Thais G., and Cláudia B. R. Martinez. 2012. “Atrazine Promotes Biochemical Changes and DNA Damage in a Neotropical Fish Species.” Chemosphere 89 (9): 1118–25.

Mela, M., I. C. Guiloski, H. B. Doria, M. A. F. Randi, C. A. de Oliveira Ribeiro, L. Pereira, A. C. Maraschi, V. Prodocimo, C. A. Freire, and H. C. Silva de Assis. 2013. “Effects of the Herbicide Atrazine in Neotropical Catfish (Rhamdia Quelen).” Ecotoxicology and Environmental Safety 93 (July): 13–21.

Adeyemi, Joseph A., Airton da Cunha Martins-Junior, and Fernando Barbosa Jr. 2015. “Teratogenicity, Genotoxicity and Oxidative Stress in Zebrafish Embryos (Danio Rerio) Co-Exposed to Arsenic and Atrazine.” Comparative Biochemistry and Physiology. Toxicology & Pharmacology: CBP 172-173 (April): 7–12.

Jin, Yuanxiang, Xiangxiang Zhang, Linjun Shu, Lifang Chen, Liwei Sun, Haifeng Qian, Weiping Liu, and Zhengwei Fu. 2010a. “Oxidative Stress Response and Gene Expression with Atrazine Exposure in Adult Female Zebrafish (Danio Rerio).” Chemosphere 78 (7): 846–52.

Toughan, Hosam, Samah R. Khalil, Ashraf Ahmed El-Ghoneimy, Ashraf Awad, and A. Sh Seddek. 2018. “Effect of Dietary Supplementation with Spirulina Platensis on Atrazine-Induced Oxidative Stress- Mediated Hepatic Damage and Inflammation in the Common Carp (Cyprinus Carpio L.).” Ecotoxicology and Environmental Safety 149 (March): 135–42.

Ahmed, Yasmine H., Huda O. AbuBakr, Ismail M. Ahmad, and Zainab Sabry Othman Ahmed. 2022. “Histopathological, Immunohistochemical, And Molecular Alterations In Brain Tissue And Submandibular Salivary Gland Of Atrazine-Induced Toxicity In Male Rats.” Environmental Science and Pollution Research International 29 (20): 30697–711.

Song, Yang, Zhen Chao Jia, Jin Yao Chen, Jun Xiang Hu, and Li Shi Zhang. 2014. “Toxic Effects of Atrazine on Reproductive System of Male Rats.” Biomedical and Environmental Sciences: BES 27 (4): 281–88.

Kalia, Sumiti, and M. P. Bansal. 2008. “Diethyl Maleate-Induced Oxidative Stress Leads to Testicular Germ Cell Apoptosis Involving Bax and Bcl-2.” Journal of Biochemical and Molecular Toxicology 22 (6): 371–81.

Kaur, Parminder, Sumiti Kalia, and Mohinder P. Bansal. 2006. “Effect of Diethyl Maleate Induced Oxidative Stress on Male Reproductive Activity in Mice: Redox Active Enzymes and Transcription Factors Expression.” Molecular and Cellular Biochemistry 291 (1-2): 55–61.

Zhang, Qun-Fang, Ying-Wen Li, Zhi-Hao Liu, and Qi-Liang Chen. 2016. “Reproductive Toxicity of Inorganic Mercury Exposure in Adult Zebrafish: Histological Damage, Oxidative Stress, and Alterations of Sex Hormone and Gene Expression in the Hypothalamic-Pituitary-Gonadal Axis.” Aquatic Toxicology  177 (August): 417–24.

El-Desoky, Gaber E., Samir A. Bashandy, Ibrahim M. Alhazza, Zeid A. Al-Othman, Mourad A. M. Aboul-Soud, and Kareem Yusuf. 2013. “Improvement of Mercuric Chloride-Induced Testis Injuries and Sperm Quality Deteriorations by Spirulina Platensis in Rats.” PloS One 8 (3): e59177.

Ma, Yan, Mingkun Zhu, Liping Miao, Xiaoyun Zhang, Xinyang Dong, and Xiaoting Zou. 2018. “Mercuric Chloride Induced Ovarian Oxidative Stress by Suppressing Nrf2-Keap1 Signal Pathway and Its Downstream Genes in Laying Hens.” Biological Trace Element Research 185 (1): 185–96.

D’Souza, R. W., and M. E. Andersen. 1988. “Physiologically Based Pharmacokinetic Model for Vinylidene Chloride.” Toxicology and Applied Pharmacology 95 (2): 230–40.

D’Souza, R. W., W. R. Francis, and M. E. Andersen. 1988. “Physiological Model for Tissue Glutathione Depletion and Increased Resynthesis after Ethylene Dichloride Exposure.” The Journal of Pharmacology and Experimental Therapeutics 245 (2): 563–68.

Mulder, G. J., and S. Ouwerkerk-Mahadevan. 1997. “Modulation of Glutathione Conjugation in Vivo: How to Decrease Glutathione Conjugation in Vivo or in Intact Cellular Systems in Vitro.” Chemico-Biological Interactions 105 (1): 17–34.

Fennell, T. R., and C. D. Brown. 2001. “A Physiologically Based Pharmacokinetic Model for Ethylene Oxide in Mouse, Rat, and Human.” Toxicology and Applied Pharmacology 173 (3): 161–75.

Spahiu, Linda, Johan Ålander, Astrid Ottosson-Wadlund, Richard Svensson, Carina Lehmer, Richard N. Armstrong, and Ralf Morgenstern. 2017a. “Global Kinetic Mechanism of Microsomal Glutathione Transferase 1 and Insights into Dynamic Enzyme Activation.” Biochemistry 56 (24): 3089–98.

Csanády, G. A., P. E. Kreuzer, C. Baur, and J. G. Filser. 1996. “A Physiological Toxicokinetic Model for 1,3-Butadiene in Rodents and Man: Blood Concentrations of 1,3-Butadiene, Its Metabolically Formed Epoxides, and of Haemoglobin Adducts--Relevance of Glutathione Depletion.” Toxicology 113 (1-3): 300–305.

Morgenstern, R., R. Svensson, B. A. Bernat, and R. N. Armstrong. 2001. “Kinetic Analysis of the Slow Ionization of Glutathione by Microsomal Glutathione Transferase MGST1.” Biochemistry 40 (11): 3378–84.

Mayer, Robert J., and Armin R. Ofial. 2019. “Nucleophilicity of Glutathione: A Link to Michael Acceptor Reactivities.” Angewandte Chemie  58 (49): 17704–8.

McMullin, Tami, Jill Brzezicki, Brian Cranmer, John Tessari, and Melvin Andersen. 2003. “Pharmacokinetic Modeling of Disposition and Time-Course Studies With [ 14 C]Atrazine.” Journal of Toxicology and Environmental Health. Part A 66 (10): 941–64.

Geenen, Suzanne, James W. T. Yates, J. Gerry Kenna, Frederic Y. Bois, Ian D. Wilson, and Hans V. Westerhoff. 2013. “Multiscale Modelling Approach Combining a Kinetic Model of Glutathione Metabolism with PBPK Models of Paracetamol and the Potential Glutathione-Depletion Biomarkers Ophthalmic Acid and 5-Oxoproline in Humans and Rats.” Integrative Biology: Quantitative Biosciences from Nano to Macro 5 (6): 877–88.

Hughes, Tyler B., Grover P. Miller, and S. Joshua Swamidass. 2015. “Site of Reactivity Models Predict Molecular Reactivity of Diverse Chemicals with Glutathione.” Chemical Research in Toxicology 28 (4): 797–809.

Abarikwu, S. O., E. O. Farombi, and A. B. Pant. 2011. “Biflavanone-Kolaviron Protects Human Dopaminergic SH-SY5Y Cells against Atrazine Induced Toxic Insult.” Toxicology in Vitro: An International Journal Published in Association with BIBRA 25 (4): 848–58.

Singh, Mohan, Rajat Sandhir, and Ravi Kiran. 2010. “Oxidative Stress Induced by Atrazine in Rat Erythrocytes: Mitigating Effect of Vitamin E.” Toxicology Mechanisms and Methods 20 (3): 119–26.

Relationship: 2878: Depletion, GSH leads to Increased, Reactive oxygen species

AOPs Referencing Relationship

Evidence Supporting this KER

Biological plausibility for GSH depletion leading to ROS increase is rooted in the fact that this antioxidant is crucial to eliminate these reactive molecules from cells. When GSH is depleted from cytosol and mitochondria, there is an exaggerated accumulation of ROS, produced, mainly, by electron transport chain.

Empirical evidence shows that this KER is commonly registered in several animal models, including vertebrates, and this is because it is conserved among taxa. Additionally, as expected, in vitro and in vivo data gathered for the three chosen compounds highlight that.

(Tirmenstein et al. 2000), analyzing the relation between GSH depletion and ROS production in rat hepatocyte suspensions exposed to 5 mM DEM, for a period of 4 h, noted that GSH is used up, leading to overproduction and hyperaccumulation of ROS in mitochondria.  

Still in the same work, Tirmenstein et al. (2000) showed that at lower concentrations, DEM, an alkylating agent, does not interfere with ROS production, but it exhausts GSH at different levels, pointing up that decrease in GSH content is affected for that stressor at concentrations equal or lower to those that induce a rise in ROS levels. DEM at 0.1, 0.5, 1, 2.5 and 5 mM for five hours caused GSH depletion in hepatocytes at all concentrations in a dose-dependent manner. However, only 5 mM of the compound was able to consume GSH to the point that this antioxidant was kept below detection levels (4%) and led to overproduction of ROS.

In relation to ATZ, in PC12 cells (rat pheochromocytoma cell line), at 232 μM, the herbicide causes a decrease in GSH content followed by a rise in ROS levels after 24 h of exposure (Abarikwu et al. 2011). In in vivo models, ATZ-treated rat erythrocytes (300 mg/Kg body weight, daily) for 7, 14 and 21 days, displayed significant GSH consumption with concomitant increase in superoxide dismutase (SOD), catalase (CAT) and glutathione peroxidase (GPX) activities, which suggests a rise in ROS levels. And in BALB/c mice, ATZ doses (100, 200, or 400 mg/Kg body weight/daily) administered for 21 days led to a reduction in GSH content and increase of ROS levels in splenocytes in a dose-dependent manner (Gao et al. 2016).  

These data are in accordance with two other studies carried out with other experimental models. Human neuroblastoma SH-SY5Y cells exposed to ATZ (0.3 mM) for 24 h displayed both a drop in GSH content as well as ROS overproduction  (Abarikwu et al. 2011). Additionally, in zebrafish embryos exposed to atrazine at 0.1 mM for 96 h exhibited a decrease in GSH content followed by a rise in CAT enzyme activity, responsible for H2O2 scavenging, suggesting in this model that decrease in GSH content induces a rise in ROS levels (Adeyemi, da Cunha Martins-Junior, and Barbosa 2015). 

This same response pattern is observed using Hg as a stressor. Cultured human bronchial epithelial cells (BEAS-2B cell line) exposed to mercury (II) chloride (2, 4, 6, and 8 ppm) for 24 h, and monitored every 3 h in order to measure GSH levels, displayed a decrease in GSH content at all concentrations from the third hour in a dose-dependent manner. A 60% drop in GSH content was kept constant during all exposure time at 8 ppm, whereas all other concentrations induced a constant diminishment of GSH content from 12 to 24 h post-exposure, but not that high. Likewise, a dose-dependent pattern of ROS generation was observed in BEAS-2B cell line, but only after 24 h of exposure. The authors still exposed these cells to 8 ppm of mercury (for 3, 6, 12 and 24 h) and noted a quick increase in ROS levels from 12 h of exposure on, but only after 24 h it was observed a noticeable increase in ROS levels – 3 times greater than the control group (Park and Park 2007). 

The same response pattern in in vitro and in vivo models is found if GSH depletion  is specifically stimulated for the inhibition of its de novo synthesis, revealing the direct causality among KEs. In HT-22 mouse hippocampal cell line submitted to 50 µM buthionine sulfoximine (BSO), a traditional GSH synthesis inhibitor, leads to glutathione depletion so that  an initial increase in ROS levels takes place afterwards (Tan et al. 1998). (Armstrong et al. 2002) tracking GSH and ROS levels in human B lymphoma cell lines (PW) submitted to 1 mM BSO, for a long period of time (24, 48 and 72 h) concluded that GSH depletion is directly responsible for the increase of ROS levels and a drop in mitochondrial GSH content is a key factor for the exponential augmentation of these free radicals. 

Corroborating these data, adult rats treated with BSO 20 and 30 mM, for 10 days, diligently, showed a reduction of, respectively, 44.25 % and 60.14 % of liver GSH content, while H2O2 levels underwent an augmentation of 42 and 60%, in that order (Ford et al. 2006).

Thus, from this overview of experimental data, it is noted that GSH depletion needs to happen previously in the course of time so that ROS production is triggered in cells and tissues and, besides that, the greater the depletion, the more pronounced the increase in ROS levels. In addition, this assessment reveals that upstream KE is affected by stressor in doses equal or lower to those that unleash downstream KE, as well as the upstream KE is more frequent than the downstream one in equivalent stress degrees. In addition, this provides robustness to dose, time and incidence concordances for this KER. Just as important, the relation is also quite conserved through several taxa (Trachootham et al. 2008).

Quantitative Understanding of the Linkage

The close relation between GSH depletion and increase in ROS levels is a well-established biological process, which is a result of diverse experimental evidence.

Drop in GSH levels and increase in ROS generation changes cellular redox potential, which can be calculated by the Nernst equation (Han et al. 2006): 

where E_cell is cell electrochemical voltage, E^o is the electromotive force, R is molar gas constant, T is the temperature in Kelvin, F is the Faraday constant, n is the number of electrons transferred in the reaction, and Q is [GSH]2/[GSSG].

If GSH levels drop until a certain threshold (~30 - 40% of depletion) in mitochondria, there is an excessive H₂O₂ release in cells (Han et al. 2006) and, hence, ROS exacerbation.

For HT22 cells exposed to 50 µM BSO (for 10 h), ROS production occurs in two phases: an initial slow increase for the first 6 h, followed by a much higher rate. The latter high rate of increase in ROS only starts after the cellular GSH levels drop to nearly zero (Tan et al. 1998).

Moreover, isolated rat hepatocyte suspensions exposed to DEM (0.5, 1, 2.5 and 5 mM) for 5 h reach maximum levels of GSH depletion after 1 h of exposure (Tirmenstein et al. 2000), whereas the maximum increase in ROS levels is observed only after four hours at the two highest concentrations of each depleter.

GSH has its levels reduced by more than 95% in PW cells after around 8 h of exposure to BSO and reacher maximum depletion level at 48 h, when  mitochondrial GSH supplies become undetectable as well, whereas ROS levels undergo a slight increase only 24 h post-exposure and reaches maximum values after 60 h of treatment (Armstrong et al. 2002).

References

Tirmenstein, M. A., F. A. Nicholls-Grzemski, J. G. Zhang, and M. W. Fariss. 2000. “Glutathione Depletion and the Production of Reactive Oxygen Species in Isolated Hepatocyte Suspensions.” Chemico-Biological Interactions 127 (3): 201–17.

Abarikwu, Sunny O., Ebenezer O. Farombi, Mahendra P. Kashyap, and Aditya B. Pant. 2011. “Kolaviron Protects Apoptotic Cell Death in PC12 Cells Exposed to Atrazine.” Free Radical Research 45 (9): 1061–73.

Gao, Shuying, Zhichun Wang, Chonghua Zhang, Liming Jia, and Yang Zhang. 2016. “Oral Exposure to Atrazine Induces Oxidative Stress and Calcium Homeostasis Disruption in Spleen of Mice.” Oxidative Medicine and Cellular Longevity 2016 (November): 7978219.

Adeyemi, Joseph A., Airton da Cunha Martins-Junior, and Fernando Barbosa Jr. 2015. “Teratogenicity, Genotoxicity and Oxidative Stress in Zebrafish Embryos (Danio Rerio) Co-Exposed to Arsenic and Atrazine.” Comparative Biochemistry and Physiology. Toxicology & Pharmacology: CBP 172-173 (April): 7–12.

Park, Eun-Jung, and Kwangsik Park. 2007. “Induction of Reactive Oxygen Species and Apoptosis in BEAS-2B Cells by Mercuric Chloride.” Toxicology in Vitro: An International Journal Published in Association with BIBRA 21 (5): 789–94.

Tan, S., Y. Sagara, Y. Liu, P. Maher, and D. Schubert. 1998. “The Regulation of Reactive Oxygen Species Production during Programmed Cell Death.” The Journal of Cell Biology 141 (6): 1423–32.

Armstrong, J. S., K. K. Steinauer, B. Hornung, J. M. Irish, P. Lecane, G. W. Birrell, D. M. Peehl, and S. J. Knox. 2002. “Role of Glutathione Depletion and Reactive Oxygen Species Generation in Apoptotic Signaling in a Human B Lymphoma Cell Line.” Cell Death & Differentiation. https://doi.org/10.1038/sj.cdd.4400959.

Ford, Rebecca J., Drew A. Graham, Steven G. Denniss, Joe Quadrilatero, and James W. E. Rush. 2006. “Glutathione Depletion in Vivo Enhances Contraction and Attenuates Endothelium-Dependent Relaxation of Isolated Rat Aorta.” Free Radical Biology & Medicine 40 (4): 670–78.

Trachootham, Dunyaporn, Weiqin Lu, Marcia A. Ogasawara, Rivera-Del Valle Nilsa, and Peng Huang. 2008. “Redox Regulation of Cell Survival.” Antioxidants & Redox Signaling 10 (8): 1343–74.

Han, Derick, Naoko Hanawa, Behnam Saberi, and Neil Kaplowitz. 2006. “Mechanisms of Liver Injury. III. Role of Glutathione Redox Status in Liver Injury.” American Journal of Physiology. Gastrointestinal and Liver Physiology 291 (1): G1–7.

Singh, Mohan, Rajat Sandhir, and Ravi Kiran. 2010. “Oxidative Stress Induced by Atrazine in Rat Erythrocytes: Mitigating Effect of Vitamin E.” Toxicology Mechanisms and Methods 20 (3): 119–26.

Relationship: 2460: Increased, Reactive oxygen species leads to Increased, LPO

AOPs Referencing Relationship

Evidence Supporting this KER

Biological plausibility of this KER lies in the fact that reactive species, in excess, react and change macromolecules such as proteins, nucleic acids and lipids. Membrane lipids are particularly susceptible to damage by free radicals, as they are composed by unsaturated fatty acids (Su et al. 2019). Hence, increase in ROS production beyond antioxidant system defense capability of cells enables free circulation of molecules such as O2·−, HO·, H2O2, which removes electrons from membrane lipids and then triggers lipid peroxidation (Auten and Davis 2009; Su et al. 2019). 

Analyses performed to support this relation show that KER3 is unchained by the three previously selected xenobiotics, as well as it takes place in a conserved way among species. Connection among the KEs is observed in both in vitro experimental models and in vivo systems, including fishes, birds and mammals.

In cultures of rat hepatocytes, progressive ROS increase during 4 hours of treatment, triggered by DEM (5 mM), is followed by a continuous growth in levels of thiobarbituric acid reactive substances (TBARS), lipid peroxidation markers (Tirmenstein et al. 2000). This chemical depletes GSH content, leading to an augmentation of ROS levels and, consequently, to lipid peroxidation. In an in vivo model, 52 μM of DEM intraperitoneally injected in male Balb/c mice for two weeks caused a significant decrease in the GSH, increase in GSSG, ROS generation and increase in lipid peroxidation in testicles (Kalia and Bansal 2008).

ATZ (46.4 µM) causes an increase of 48.97% of ROS and of 12.5% in MDA content in cultures of Sertoli-Germ cells from Wistar rats (25–28 days old), after, respectively, 3 and 24 h post-exposure. At a higher concentration (232 µM), these cells reach a maximum peak of ROS production after 6h of exposure, while MDA generation gets to the peak only after 24 h of treatment (Abarikwu, Pant, and Farombi 2012). In in vivo model, ATZ (38.5, 77 e 154 mg/Kg bw/day) led to a decrease in total antioxidant capacity (TAC) in a dose-dependent manner in male Sprague-Dawley rats of Specific Pathogen Free (SPF) ATZ-treated for 30 days. Which indirectly suggests increase in ROS levels – and increased malondialdehyde (MDA) content in 154 mg/Kg (Song et al. 2014). 

In relation to Hg, it was found that male young Wistar rats exposed to an initial dose of 4.6 μg/Kg of this metal (with following doses of 0.07 μg/Kg/day) displayed an increase in ROS levels, followed by an elevation of MDA content in testicles and epididymis of these rats 60 days post-exposure (Rizzetti et al. 2017). Other assays still carried out with male rats showed that the heavy metal induces oxidative stress with a single subcutaneous dose of 5 mg/Kg, by a substantial diminishment of activity of the main testicle antioxidant enzymes: SOD, CAT and GPX. Consequently, blood hydroperoxide and testicle MDA levels rose in a relevant way (El-Desoky et al. 2013).

Furthermore, Hy-Line Brown laying hens fed with 4 experimental diets containing graded levels of Hg at 0.280, 3.325, 9.415, and 27.240 mg/Kg, respectively, for 10 weeks had GSH content significantly decreased in all Hg-treatment groups in ovaries, whilst SOD, CAT, GPX and glutathione reductase (GR) enzyme activities were significantly reduced, pointing to ROS accumulation. MDA content strongly increased in the 27.240-mg/Kg Hg group (Ma et al. 2018). 

Hence, it can be deduced that, as in other adjacent relations evaluated, there is also evidence here that upstream KE is initially required in order to downstream KE take place, which reaffirms time concordance. Besides this, data enhance dose and incidence concordances for this KER.

Quantitative Understanding of the Linkage

Mechanisms involving lipid peroxidation, such as that one caused by ROS accumulation in cells, have been investigated for decades (Tirmenstein et al. 2000; Yin, Xu, and Porter 2011; Su et al. 2019). For this reason, there is much experimental data about response-response relationships or a growth of upstream KE in relation to downstream KE.

This mechanism can be better understood through a process chain that consists of initiation, propagation and termination, as discussed by (Yin, Xu, and Porter 2011). In their review, these authors summarized a series of chemical reactions that develop during all this self-oxidation process and represent them in a schematic manner, as displayed in figure below.

Furthermore, although phospholipid oxidizability is lower, once their rate of diffusion in membranes is slower, the kinetics for this kind of reaction shown in figure follows the same law of velocity (steady-state rate) of homogeneous systems (equation below) (Yin, Xu, and Porter 2011). Oxygen consumption of the equation represents the rate of steady state, while rate of radical generation is defined by R_i, the constant of propagation rate is expressed as k_p and the termination rate constant for the reaction is called k_t.

-d[O] / dt = k_p / (2k_t)^1/2. [L-H] . R_i^1/2

For instance, empirical evidences show that rat hepatocytes begin ROS production after the first 30 minutes of DEM exposition (5 mM), growing linearly for all the remaining time, whereas the increase in products of lipid peroxidation (TBARS) starts only from the first hour of exposure (Tirmenstein et al. 2000).

References

Su, Lian-Jiu, Jia-Hao Zhang, Hernando Gomez, Raghavan Murugan, Xing Hong, Dongxue Xu, Fan Jiang, and Zhi-Yong Peng. 2019. “Reactive Oxygen Species-Induced Lipid Peroxidation in Apoptosis, Autophagy, and Ferroptosis.” Oxidative Medicine and Cellular Longevity 2019 (October): 5080843.

Auten, Richard L., and Jonathan M. Davis. 2009. “Oxygen Toxicity and Reactive Oxygen Species: The Devil Is in the Details.” Pediatric Research 66 (2): 121–27.

Tirmenstein, M. A., F. A. Nicholls-Grzemski, J. G. Zhang, and M. W. Fariss. 2000. “Glutathione Depletion and the Production of Reactive Oxygen Species in Isolated Hepatocyte Suspensions.” Chemico-Biological Interactions 127 (3): 201–17.

Kalia, Sumiti, and M. P. Bansal. 2008. “Diethyl Maleate-Induced Oxidative Stress Leads to Testicular Germ Cell Apoptosis Involving Bax and Bcl-2.” Journal of Biochemical and Molecular Toxicology 22 (6): 371–81.

Abarikwu, S. O., E. O. Farombi, and A. B. Pant. 2011. “Biflavanone-Kolaviron Protects Human Dopaminergic SH-SY5Y Cells against Atrazine Induced Toxic Insult.” Toxicology in Vitro: An International Journal Published in Association with BIBRA 25 (4): 848–58.

Rizzetti, Danize Aparecida, Caroline Silveira Martinez, Alyne Goulart Escobar, Taiz Martins da Silva, José Antonio Uranga-Ocio, Franck Maciel Peçanha, Dalton Valentim Vassallo, Marta Miguel Castro, and Giulia Alessandra Wiggers. 2017. “Egg White-Derived Peptides Prevent Male Reproductive Dysfunction Induced by Mercury in Rats.” Food and Chemical Toxicology: An International Journal Published for the British Industrial Biological Research Association 100 (February): 253–64.

El-Desoky, Gaber E., Samir A. Bashandy, Ibrahim M. Alhazza, Zeid A. Al-Othman, Mourad A. M. Aboul-Soud, and Kareem Yusuf. 2013. “Improvement of Mercuric Chloride-Induced Testis Injuries and Sperm Quality Deteriorations by Spirulina Platensis in Rats.” PloS One 8 (3): e59177.

Ma, Yan, Mingkun Zhu, Liping Miao, Xiaoyun Zhang, Xinyang Dong, and Xiaoting Zou. 2018. “Mercuric Chloride Induced Ovarian Oxidative Stress by Suppressing Nrf2-Keap1 Signal Pathway and Its Downstream Genes in Laying Hens.” Biological Trace Element Research 185 (1): 185–96.

Yin, Huiyong, Libin Xu, and Ned A. Porter. 2011. “Free Radical Lipid Peroxidation: Mechanisms and Analysis.” Chemical Reviews 111 (10): 5944–72.

Auten, Richard L., and Jonathan M. Davis. 2009. “Oxygen Toxicity and Reactive Oxygen Species: The Devil Is in the Details.” Pediatric Research 66 (2): 121–27.

Relationship: 2879: Increased, LPO leads to impaired, Fertility

AOPs Referencing Relationship

Evidence Supporting this KER

Biological plausibility of this KER lies in the fact that lipid peroxidation in gonad membranes induces morphological changes in seminiferous tubules, and degeneration of ovarian follicles and Sertoli and Leydig cells in testicles, damage to gametic cells, and, consequently, reduction of their viability. This directly affects animal reproductive capability, for it reduces quality and production of oocytes and spermatocytes, as well as decreases egg and sperm release (spawn), leading to a drop-in fertilization rate (Tillitt et al. 2010; Papoulias et al. 2014; Song et al. 2014; Dasmahapatra et al. 2020; Biswas et al. 2020; Mu et al. 2022). 

Just like the other KERs, the adjacent relation here assessed is also observed in different species and models used in toxicological studies. Nevertheless, evidence gathered here shows that occurrence of lipid peroxidation triggered by ATZ, DEM and Hg leading to fertility impairment is limited to fishes and mammals. In this case, mammals have their reproductive capability reduced mainly because of morphological changes and poor quality of gametes.

Sexually mature female Wistar rats treated by a daily gavage of 0, 5, 25 and 125 mg/Kg ATZ for 28 consecutive days exhibited lipid peroxidation and ovarian atresia significantly increased in a dose-dependent manner in  ATZ-treated animals (Zhao et al. 2014). In male Sprague-Dawley rats exposed to ATZ by gavage (0, 38.5, 77, and 154 mg/Kg bw/day) for 30 days had significant adverse effects on the reproductive system, with the animals showing a decreased level of total antioxidant capacity (TAC) in a dose-dependent manner, a depletion in GSH and an increased MDA content at the highest dose, followed by not only irregular and disordered arrangement of the seminiferous epithelium, but also a decreased number of spermatozoa and augmented spermatozoa abnormality rate in groups  treated with 77, and 154 mg/Kg of ATZ (Song et al. 2014).

Corroborating these data, Farombi et al. (2013) showed that male Wistar rats administered with ATZ at a dose equivalent to 120 mg/Kg body weight each day for 16 days displayed augmented MDA levels in testicles and epididymis, as well an increased number of sperm abnormalities and reduced sperm production, sperm motility and epididymal and testicular sperm numbers. Moreover, degeneration of seminiferous tubules in testicles with the presence of defoliation was noted as well (Farombi et al. 2013). In adult male Albino rats, ATZ (120 mg/Kg bw) causes significantly increased malondialdehyde (MDA) serum level and also diminished total antioxidant capacity (TAC), besides inducing a significant rise in sperm cell abnormalities (Abdel Aziz et al. 2018). In addition, pathological lesions such as disorganized seminiferous tubules with degenerated and irregularly arranged necrotized germinal cells were also reported (Abdel Aziz et al. 2018). This very ATZ dosage orally administered in rats caused an increase in MDA formation in the liver, testis and epididymis, along with an inhibition of GST activities, and decreased epididymal and testicular sperm number, sperm motility, daily sperm production and increased number of dead and abnormal sperm in animals (Adesiyan et al. 2011).

Kalia & Bansal (2008) found in their study that male Balb/c mice treated with DEM (52 μM) underwent a decrease in GSH content, and increased ROS generation and lipid peroxidation in testicles, followed by augmented apoptosis in germ cells, as well as a significant reduction in the number of these cells (Kalia and Bansal 2008). A lower dose of this compound (8.7 μM)  daily and intraperitoneally injected, for two weeks, resulted in depleted GSH and increase in testis GSSG levels. As a consequence sperm motility was decreased by  40%, and epididymal sperm count was significantly reduced in DEM-treated animals. Beyond that, fertility status was also affected by DEM exposure, with a 34% diminishment compared to the control group, and there was a reduction in litter size as well (Kaur, Kalia, and Bansal 2006).

Using Hg in order to induce oxidative stress, these kinds of results also occur in different taxa. Male Wistar rats continuously exposed to 0, 50 and 100 ppm Hg for 90 days through oral administration in the drinking water displayed a significant increase in testicular MDA, along with specific alterations in the histoarchitecture of testis, including disintegration of germinal epithelium of seminiferous tubules, detachment and degenerative changes of lining cells, increased space between the seminiferous tubules and their lumen enlarged in a dose-dependent manner (Boujbiha et al. 2009). Interestingly, Hg-treated males were mated with normal cyclic females and they showed a decline in reproductive performance. In another study, the heavy metal led to an increase in ROS and MDA levels in testes and epididymis 60 days post-exposure in Wistar rats submitted to a first dose of 4.6 μg/Kg and subsequent doses of 0.07 μg/Kg/day, as well as decreased sperm number, increased sperm transit time in epididymis and impaired sperm morphology (Rizzetti et al. 2017). In fishes, sublethal doses of mercury (II) chloride (0.04 and 0.12 ppm) for 30 days caused a significant increase in testicle lipid peroxidation, DNA fragmentation, and a decrease in sperm count, activity and motility in relation to the control group in African sharptooth catfish (Clarias gariepinus) (Ibrahim, Banaee, and Sureda 2019). In addition to that, histopathological alterations in testis sections including rupture of interlobular connective tissue, lessening of spermatogonia, derangement of spermatogenesis and low spermatozoa counting were also observed (Ibrahim, Banaee, and Sureda 2019). 
 

Quantitative Understanding of the Linkage

With regard to KER4, several studies have brought quantitative data concerning the negative correlation between lipid peroxidation and fertility disorders of vertebrate organisms (Gomez, Irvine, and Aitken 1998; Hsieh, Chang, and Lin 2006; Aitken et al. 2007; Abarikwu et al. 2010; Mihalas et al. 2017).

According to (Gomez, Irvine, and Aitken 1998), there is a negative relationship between malondialdehyde and 4-hydroxyalkenal production (MDA + 4-HA) and loss of motility in human spermatozoa. The higher the amount of these peroxidation products, the lower the cell motility. A negative correlation between sperm numbers and testicular and epididymal MDA levels (-0.85 and -0.68 correlation coefficient r, respectively) was also found by (Abarikwu et al. 2010) in rats exposed to ATZ for 7 and 16 days. Conversely, the authors observed a positive correlation between abnormal sperm rate and testicular and epididymal MDA levels (+0.78 and +0.89). 
Hsieh et al. (2006), assessing MDA levels and sperm quality of 51 subfertile men, were able to establish two formulas to associate lipid peroxidation with sperm concentration and motility, which are represented, respectively, by: 

MDA = - 0.0045 x sperm cell concentration + 2.23;

and

MDA = - 0.014 x sperm motility + 2.62. 

On the other hand, (Mihalas et al. 2017) brought important quantitative data about the direct relation between lipid peroxidation and reduction of quality in oocytes. Experimental evidences showed that the lipid peroxidation product 4-HNE, at 0, 5, 10, 20, 30 and 50 µM, induces a dose-dependent decrease in meiotic competence during in vitro oocyte maturation, as well as aneuploidies in germinal vesicle (GV) oocytes from 20 µM of 4-HNE. They still reported this happens because tubulins, component proteins of microtubules of the mitotic spindle, generate adducts with 4-HNE. 

References

Tillitt, Donald E., Diana M. Papoulias, Jeffrey J. Whyte, and Catherine A. Richter. 2010. “Atrazine Reduces Reproduction in Fathead Minnow (Pimephales Promelas).” Aquatic Toxicology  99 (2): 149–59.

Papoulias, Diana M., Donald E. Tillitt, Melaniya G. Talykina, Jeffrey J. Whyte, and Catherine A. Richter. 2014. “Atrazine Reduces Reproduction in Japanese Medaka (Oryzias Latipes).” Aquatic Toxicology  154 (September): 230–39.

Song, Yang, Zhen Chao Jia, Jin Yao Chen, Jun Xiang Hu, and Li Shi Zhang. 2014. “Toxic Effects of Atrazine on Reproductive System of Male Rats.” Biomedical and Environmental Sciences: BES 27 (4): 281–88.

Dasmahapatra, Asok K., Doris K. Powe, Thabitha P. S. Dasari, and Paul B. Tchounwou. 2020. “Assessment of Reproductive and Developmental Effects of Graphene Oxide on Japanese Medaka (Oryzias Latipes).” Chemosphere 259 (November): 127221.

Biswas, Subhasri, Soumyajyoti Ghosh, Anwesha Samanta, Sriparna Das, Urmi Mukherjee, and Sudipta Maitra. 2020. “Bisphenol A Impairs Reproductive Fitness in Zebrafish Ovary: Potential Involvement of Oxidative/nitrosative Stress, Inflammatory and Apoptotic Mediators.” Environmental Pollution  267 (December): 115692.

Mu, Xiyan, Suzhen Qi, Jia Liu, Hui Wang, Lilai Yuan, Le Qian, Tiejun Li, et al. 2022. “Environmental Level of Bisphenol F Induced Reproductive Toxicity toward Zebrafish.” The Science of the Total Environment 806 (Pt 1): 149992.

Zhao, Fan, Kun Li, Lijing Zhao, Jian Liu, Qi Suo, Jing Zhao, Hebin Wang, and Shuhua Zhao. 2014. “Effect of Nrf2 on Rat Ovarian Tissues against Atrazine-Induced Anti-Oxidative Response.” International Journal of Clinical and Experimental Pathology 7 (6): 2780–89.

Farombi, E. O., S. O. Abarikwu, A. C. Adesiyan, and T. O. Oyejola. 2013. “Quercetin Exacerbates the Effects of Subacute Treatment of Atrazine on Reproductive Tissue Antioxidant Defence System, Lipid Peroxidation and Sperm Quality in Rats.” Andrologia 45 (4): 256–65.

Abdel Aziz, Rabie L., Ahmed Abdel-Wahab, Fatma I. Abo El-Ela, Nour El-Houda Y. Hassan, El-Shaymaa El-Nahass, Marwa A. Ibrahim, and Abdel-Tawab A. Y. Khalil. 2018. “Dose- Dependent Ameliorative Effects of Quercetin and L-Carnitine against Atrazine- Induced Reproductive Toxicity in Adult Male Albino Rats.” Biomedicine & Pharmacotherapy = Biomedecine & Pharmacotherapie 102 (June): 855–64.

Adesiyan, Adebukola C., Titilola O. Oyejola, Sunny O. Abarikwu, Matthew O. Oyeyemi, and Ebenezer O. Farombi. 2011. “Selenium Provides Protection to the Liver but Not the Reproductive Organs in an Atrazine-Model of Experimental Toxicity.” Experimental and Toxicologic Pathology: Official Journal of the Gesellschaft Fur Toxikologische Pathologie 63 (3): 201–7.

Kalia, Sumiti, and M. P. Bansal. 2008. “Diethyl Maleate-Induced Oxidative Stress Leads to Testicular Germ Cell Apoptosis Involving Bax and Bcl-2.” Journal of Biochemical and Molecular Toxicology 22 (6): 371–81.

Kaur, Parminder, Sumiti Kalia, and Mohinder P. Bansal. 2006. “Effect of Diethyl Maleate Induced Oxidative Stress on Male Reproductive Activity in Mice: Redox Active Enzymes and Transcription Factors Expression.” Molecular and Cellular Biochemistry 291 (1-2): 55–61.

Boujbiha, Mohamed Ali, Khaled Hamden, Fadhel Guermazi, Ali Bouslama, Asma Omezzine, Abdelaziz Kammoun, and Abdelfattah El Feki. 2009. “Testicular Toxicity in Mercuric Chloride Treated Rats: Association with Oxidative Stress.” Reproductive Toxicology  28 (1): 81–89.

Rizzetti, Danize Aparecida, Caroline Silveira Martinez, Alyne Goulart Escobar, Taiz Martins da Silva, José Antonio Uranga-Ocio, Franck Maciel Peçanha, Dalton Valentim Vassallo, Marta Miguel Castro, and Giulia Alessandra Wiggers. 2017. “Egg White-Derived Peptides Prevent Male Reproductive Dysfunction Induced by Mercury in Rats.” Food and Chemical Toxicology: An International Journal Published for the British Industrial Biological Research Association 100 (February): 253–64.

Ibrahim, Ahmed Th A., Mahdi Banaee, and Antoni Sureda. 2019. “Selenium Protection against Mercury Toxicity on the Male Reproductive System of Clarias Gariepinus.” Comparative Biochemistry and Physiology. Toxicology & Pharmacology: CBP 225 (November): 108583.

Gomez, E., D. S. Irvine, and R. J. Aitken. 1998. “Evaluation of a Spectrophotometric Assay for the Measurement of Malondialdehyde and 4-Hydroxyalkenals in Human Spermatozoa: Relationships with Semen Quality and Sperm Function.” International Journal of Andrology 21 (2): 81–94.

Hsieh, Yao-Yuan, Chi-Chen Chang, and Chich-Sheng Lin. 2006. “Seminal Malondialdehyde Concentration but Not Glutathione Peroxidase Activity Is Negatively Correlated with Seminal Concentration and Motility.” International Journal of Biological Sciences 2 (1): 23–29.

Aitken, R. John, Jordana K. Wingate, Geoffry N. De Iuliis, and Eileen A. McLaughlin. 2007. “Analysis of Lipid Peroxidation in Human Spermatozoa Using BODIPY C11.” Molecular Human Reproduction 13 (4): 203–11.

Mihalas, Bettina P., Geoffry N. De Iuliis, Kate A. Redgrove, Eileen A. McLaughlin, and Brett Nixon. 2017. “The Lipid Peroxidation Product 4-Hydroxynonenal Contributes to Oxidative Stress-Mediated Deterioration of the Ageing Oocyte.” Scientific Reports 7 (1): 6247.

Abarikwu, S. O., E. O. Farombi, and A. B. Pant. 2011. “Biflavanone-Kolaviron Protects Human Dopaminergic SH-SY5Y Cells against Atrazine Induced Toxic Insult.” Toxicology in Vitro: An International Journal Published in Association with BIBRA 25 (4): 848–58.