SNAPSHOT

Key Event Component

Evidence for Perturbation by Stressor

Overview for Molecular Initiating Event

Fibrates are ligands of PPARα (Staels et al. 1998).

Phthalates

MHEP (CAS 4376-20-9) directly binds in vitro to PPARα (Lapinskas et al. 2005) and activates this receptor in transactivation assays PPARα (Lapinskas et al. 2005), (Maloney and Waxman 1999), (Hurst and Waxman 2003), (Bility et al. 2004), (Lampen, Zimnik, and Nau 2003), (Venkata et al. 2006) ]. DEHP (CAS 117-81-7) has not been found to bind and activate PPARα (Lapinskas et al. 2005), (Maloney and Waxman 1999). However, the recent studies shown activation of PPARα (ToxCastTM Data).

Notably, PPARα are responsive to DEHP in vitro as they are translocated to the nucleus (in primary Sertoli cells) (Dufour et al. 2003), (Bhattacharya et al. 2005). Expression of PPARα [mRNA and protein] has been reported to be also modulated by phthtalates: (to be up-regulated in vivo upon DEHP treatment (Xu et al. 2010) and down-regulated by Diisobutyl phthalate (DiBP) (Boberg et al. 2008)).

Perfluorooctanoic Acid (PFOA) is known to activate PPARα (Vanden Heuvel et al. 2006).

Organotin

Tributyltin (TBT) activates all three heterodimers of PPAR with RXR, primarily through its interaction with RXR (le Maire et al. 2009)

Domain of Applicability

Key Event Description

Biological state

The Peroxisome Proliferator Activated receptor α (PPARα) belongs to the Peroxisome Proliferator Activated receptors (PPARs; NR1C) steroid/thyroid/retinoid receptor superfamily of transcription factors.

Biological compartments

PPARα is expressed in high levels in tissues that perform significant catabolism of fatty acids (FAs), such as brown adipose tissue, liver, heart, kidney, and intestine (Michalik et al. 2006). The receptor is present also in skeletal muscle, intestine, pancreas, lung, placenta and testes (Mukherjee et al. 1997), (Schultz et al. 1999).

General role in biology

PPARs are activated by fatty acids and their derivatives; they are sensors of dietary lipids and are involved in lipid and carbohydrate metabolism, immune response and peroxisome proliferation (Wahli and Desvergne 1999), (Evans, Barish, & Wang, 2004). PAPRα is a also a target of hypothalamic hormone signalling and was found to play a role in embryonic development (Yessoufou and Wahli 2010).

Fibrates, activators of PPARα, are commonly used to treat hypertriglyceridemia and other dyslipidemic states as they have been shown to decrease circulating lipid levels (Lefebvre et al. 2006).

How it is Measured or Detected

Binding of ligands to PPARα is measured using binding assays in vitro and in silico, whereas the information about functional activation is derived from transactivation assays (e.g. transactivation assay with reporter gene) that demonstrate functional activation of a nuclear receptor by a specific compound. Binding of agonists within the ligand-binding site of PPARs causes a conformational change of nuclear receptor that promotes binding to transcriptional co-activators. Conversely, binding of antagonists results in a conformation that favours the binding of co-repressors (Yu and Reddy 2007), (Viswakarma et al. 2010). Transactivation assays are performed using transient or stably transfected cells with the PPARα expression plasmid and a reporter plasmid, respectively. There are also other methods that have been used to measure PPARα activity, such as the Electrophoretic Mobility Shift Assay (EMSA) or commercially available PPARα transcription factor assay kits, see Table 1. The transactivation (stable transfection) assay provides the most applicable OECD Level 2 assay (i.e. In vitro assays providing mechanistic data) aimed at identifying the initiating event leading to an adverse outcome (LeBlanc, Norris, and Kloas 2011). Currently no internationally validated assays for regulatory purposes are available.

Table 1 Summary of the chosen methods to measure the PPARα activation.

References

Bhattacharya, Nandini, Jannette M Dufour, My-Nuong Vo, Janice Okita, Richard Okita, and Kwan Hee Kim. 2005. “Differential Effects of Phthalates on the Testis and the Liver.” Biology of Reproduction 72 (3) (March): 745–54. doi:10.1095/biolreprod.104.031583.

Bility, Moses T, Jerry T Thompson, Richard H McKee, Raymond M David, John H Butala, John P Vanden Heuvel, and Jeffrey M Peters. 2004. “Activation of Mouse and Human Peroxisome Proliferator-Activated Receptors (PPARs) by Phthalate Monoesters.” Toxicological Sciences : An Official Journal of the Society of Toxicology 82 (1) (November): 170–82. doi:10.1093/toxsci/kfh253.

Dufour, Jannette M, My-Nuong Vo, Nandini Bhattacharya, Janice Okita, Richard Okita, and Kwan Hee Kim. 2003. “Peroxisome Proliferators Disrupt Retinoic Acid Receptor Alpha Signaling in the Testis.” Biology of Reproduction 68 (4) (April): 1215–24. doi:10.1095/biolreprod.102.010488.

Feige, Jérôme N, Laurent Gelman, Daniel Rossi, Vincent Zoete, Raphaël Métivier, Cicerone Tudor, Silvia I Anghel, et al. 2007. “The Endocrine Disruptor Monoethyl-Hexyl-Phthalate Is a Selective Peroxisome Proliferator-Activated Receptor Gamma Modulator That Promotes Adipogenesis.” The Journal of Biological Chemistry 282 (26) (June 29): 19152–66. doi:10.1074/jbc.M702724200.

Hurst, Christopher H, and David J Waxman. 2003. “Activation of PPARalpha and PPARgamma by Environmental Phthalate Monoesters.” Toxicological Sciences : An Official Journal of the Society of Toxicology 74 (2) (August): 297–308. doi:10.1093/toxsci/kfg145.

Kaya, Taner, Scott C Mohr, David J Waxman, and Sandor Vajda. 2006. “Computational Screening of Phthalate Monoesters for Binding to PPARgamma.” Chemical Research in Toxicology 19 (8) (August): 999–1009. doi:10.1021/tx050301s.

Lampen, Alfonso, Susan Zimnik, and Heinz Nau. 2003. “Teratogenic Phthalate Esters and Metabolites Activate the Nuclear Receptors PPARs and Induce Differentiation of F9 Cells.” Toxicology and Applied Pharmacology 188 (1) (April): 14–23. doi:10.1016/S0041-008X(03)00014-0.

Lapinskas, Paula J., Sherri Brown, Lisa M. Leesnitzer, Steven Blanchard, Cyndi Swanson, Russell C. Cattley, and J. Christopher Corton. 2005. “Role of PPARα in Mediating the Effects of Phthalates and Metabolites in the Liver.” Toxicology 207 (1): 149–163.

Le Maire, Albane, Marina Grimaldi, Dominique Roecklin, Sonia Dagnino, Valérie Vivat-Hannah, Patrick Balaguer, and William Bourguet. 2009. “Activation of RXR-PPAR Heterodimers by Organotin Environmental Endocrine Disruptors.” EMBO Reports 10 (4) (April): 367–73. doi:10.1038/embor.2009.8.

LeBlanc, GA, DO Norris, and W Kloas. 2011. “Detailed Review Paper State of the Science on Novel In Vitro and In Vivo Screening and Testing Methods and Endpoints for Evaluating Endocrine Disruptors” (178).

Lefebvre, Philippe, Giulia Chinetti, Jean-Charles Fruchart, and Bart Staels. 2006. “Sorting out the Roles of PPAR Alpha in Energy Metabolism and Vascular Homeostasis.” The Journal of Clinical Investigation 116 (3) (March): 571–80. doi:10.1172/JCI27989.

Maloney, Erin K., and David J. Waxman. 1999. “Trans-Activation of PPARα and PPARγ by Structurally Diverse Environmental Chemicals.” Toxicology and Applied Pharmacology 161 (2): 209–218.

Michalik, Liliane, Johan Auwerx, Joel P Berger, V Krishna Chatterjee, Christopher K Glass, Frank J Gonzalez, Paul A Grimaldi, et al. 2006. “International Union of Pharmacology. LXI. Peroxisome Proliferator-Activated Receptors.” Pharmacological Reviews 58 (4) (December): 726–41. doi:10.1124/pr.58.4.5.

Mukherjee, R, L Jow, G E Croston, and J R Paterniti. 1997. “Identification, Characterization, and Tissue Distribution of Human Peroxisome Proliferator-Activated Receptor (PPAR) Isoforms PPARgamma2 versus PPARgamma1 and Activation with Retinoid X Receptor Agonists and Antagonists.” The Journal of Biological Chemistry 272 (12) (March 21): 8071–6.

Schultz, R, W Yan, J Toppari, A Völkl, J A Gustafsson, and M Pelto-Huikko. 1999. “Expression of Peroxisome Proliferator-Activated Receptor Alpha Messenger Ribonucleic Acid and Protein in Human and Rat Testis.” Endocrinology 140 (7) (July): 2968–75. doi:10.1210/endo.140.7.6858.

Staels, B., J. Dallongeville, J. Auwerx, K. Schoonjans, E. Leitersdorf, and J.-C. Fruchart. 1998. “Mechanism of Action of Fibrates on Lipid and Lipoprotein Metabolism.” Circulation 98 (19) (November 10): 2088–2093. doi:10.1161/01.CIR.98.19.2088.

ToxCastTM Data. “ToxCastTM Data.” US Environmental Protection Agency. http://www.epa.gov/ncct/toxcast/data.html

Vanden Heuvel, John P, Jerry T Thompson, Steven R Frame, and Peter J Gillies. 2006. “Differential Activation of Nuclear Receptors by Perfluorinated Fatty Acid Analogs and Natural Fatty Acids: A Comparison of Human, Mouse, and Rat Peroxisome Proliferator-Activated Receptor-Alpha, -Beta, and -Gamma, Liver X Receptor-Beta, and Retinoid X Rec.” Toxicological Sciences : An Official Journal of the Society of Toxicology 92 (2) (August): 476–89. doi:10.1093/toxsci/kfl014.

Venkata, Nagaraj Gopisetty, Jodie a Robinson, Peter J Cabot, Barbara Davis, Greg R Monteith, and Sarah J Roberts-Thomson. 2006. “Mono(2-Ethylhexyl)phthalate and Mono-N-Butyl Phthalate Activation of Peroxisome Proliferator Activated-Receptors Alpha and Gamma in Breast.” Toxicology Letters 163 (3) (June 1): 224–34. doi:10.1016/j.toxlet.2005.11.001.

Viswakarma, Navin, Yuzhi Jia, Liang Bai, Aurore Vluggens, Jayme Borensztajn, Jianming Xu, and Janardan K Reddy. 2010. “Coactivators in PPAR-Regulated Gene Expression.” PPAR Research 2010 (January). doi:10.1155/2010/250126.

Wahli, Walter, and B Desvergne. 1999. “Peroxisome Proliferator-Activated Receptors: Nuclear Control of Metabolism.” Endocrine Reviews 20 (5) (October): 649–88. Wu, Bin, Jie Gao, and Ming-wei Wang. 2005. “Development of a Complex Scintillation Proximity Assay for High-Throughput Screening of PPARgamma Modulators.” Acta Pharmacologica Sinica 26 (3) (March): 339–44. doi:10.1111/j.1745-7254.2005.00040.x.

Xu, Chuan, Ji-An Chen, Zhiqun Qiu, Qing Zhao, Jiaohua Luo, Lan Yang, Hui Zeng, et al. 2010. “Ovotoxicity and PPAR-Mediated Aromatase Downregulation in Female Sprague-Dawley Rats Following Combined Oral Exposure to Benzo[a]pyrene and Di-(2-Ethylhexyl) Phthalate.” Toxicology Letters 199 (3) (December 15): 323–32. doi:10.1016/j.toxlet.2010.09.015.

Yessoufou, a, and W Wahli. 2010. “Multifaceted Roles of Peroxisome Proliferator-Activated Receptors (PPARs) at the Cellular and Whole Organism Levels.” Swiss Medical Weekly 140 (September) (January): w13071. doi:10.4414/smw.2010.13071.

Yu, Songtao, and Janardan K Reddy. 2007. “Transcription Coactivators for Peroxisome Proliferator-Activated Receptors.” Biochimica et Biophysica Acta 1771 (8) (August): 936–51. doi:10.1016/j.bbalip.2007.01.008.

Key Event Component

Key Event Component

Key Event Component

Key Event Component

Domain of Applicability

Key Event Description

Biological state

Testosterone (T) is a steroid hormone from the androgen group. T serves as a substrate for two metabolic pathways that produce antagonistic sex steroids.

Biological compartments

Testosterone is synthesized by the gonads and other steroidogenic tissues (e.g., brain, adipose), acts locally and/or is transported to other tissues via blood circulation. Leydig cells are the testosterone-producing cells of the testis.

General role in biology

Androgens, the main male sex steroids, are the critical factors responsible for the development of the male phenotype during embryogenesis and for the achievement of sexual maturation at puberty. In adulthood, androgens remain essential for the maintenance of male reproductive function and behaviour. Apart from their effects on reproduction, androgens affect a wide variety of non-reproductive tissues such as skin, bone, muscle, and brain (Heemers, Verhoeven, & Swinnen, 2006). Androgens, principally T and 5α-dihydrotestosterone (DHT), exert most of their effects by interacting with a specific receptor, the androgen receptor (AR), for review see (Murashima, Kishigami, Thomson, & Yamada, 2015). On the one hand, testosterone can be reduced by 5α-reductase to produce 5α dihydrotestosterone (DHT). On the other hand, testosterone can be aromatized to generate estrogens. Testosterone effects can also be classified by the age of usual occurrence, postnatal effects in both males and females are mostly dependent on the levels and duration of circulating free testosterone.

How it is Measured or Detected

Testosterone can be measured by immunoassays and by isotope-dilution gas chromatography-mass spectrometry in serum (Taieb et al., 2003), (Paduch et al., 2014). Testosterone levels are measured i.a. in: Fish Lifecycle Toxicity Test (FLCTT) (US EPA OPPTS 850.1500), Male pubertal assay (PP Male Assay) (US EPA OPPTS 890.1500), OECD TG 441: Hershberger Bioassay in Rats (H Assay).

References

Heemers, H. V, Verhoeven, G., & Swinnen, J. V. (2006). Androgen activation of the sterol regulatory element-binding protein pathway: Current insights. Molecular Endocrinology (Baltimore, Md.), 20(10), 2265–77. doi:10.1210/me.2005-0479

Murashima, A., Kishigami, S., Thomson, A., & Yamada, G. (2015). Androgens and mammalian male reproductive tract development. Biochimica et Biophysica Acta, 1849(2), 163–170. doi:10.1016/j.bbagrm.2014.05.020

Paduch, D. A., Brannigan, R. E., Fuchs, E. F., Kim, E. D., Marmar, J. L., & Sandlow, J. I. (2014). The laboratory diagnosis of testosterone deficiency. Urology, 83(5), 980–8. doi:10.1016/j.urology.2013.12.024

Taieb, J., Mathian, B., Millot, F., Patricot, M.-C., Mathieu, E., Queyrel, N., … Boudou, P. (2003). Testosterone measured by 10 immunoassays and by isotope-dilution gas chromatography-mass spectrometry in sera from 116 men, women, and children. Clinical Chemistry, 49(8), 1381–95.

Key Event Component

Domain of Applicability

Key Event Description

Biological state

Testosterone is a steroid hormone from the androgen group and is found in humans and other vertebrates.

Biological compartments

In humans and other mammals, testosterone is secreted primarily by the testicles of males and, to a lesser extent, the ovaries of females and other steroidogenic tissues (e.g., brain, adipose). It either acts locally /or is transported to other tissues via blood circulation. Testosterone synthesis takes place within the mitochondria of Leydig cells, the testosterone-producing cells of the testis. It is produced upon stimulation of these cells by Luteinizing hormone (LH) that is secreted in pulses into the peripheral circulation by the pituitary gland in response to Gonadotropin-releasing hormone (GnRH) from the hypothalamus. Testosterone and its aromatized product, estradiol, feed back to the hypothalamus and pituitary gland to suppress transiently LH and thus testosterone production. In response to reduced testosterone levels, GnRH and LH are produced. This negative feedback cycle results in pulsatile secretion of LH followed by pulsatile production of testosterone (Ellis, Desjardins, and Fraser 1983), (Chandrashekar and Bartke 1998).

General role in biology

Testosterone is the principal male sex hormone and an anabolic steroid. Male sexual differentiation depends on testosterone (T), dihydrotestosterone (DHT), and the expression of androgen receptors by target cells (Manson and Carr 2003). During the development secretion of androgens by Leydig cells is essential for masculinization of the foetus (Nef 2000). The foetal Leydig cells develop in utero. These cells become competent to produce testosterone in rat by gestational day (GD) 15.5, with increasing production thereafter. Peak steroidogenic activity is reached just prior to birth, on GD19 (Chen, Ge, and Zirkin 2009). Testosterone secreted by foetal Leydig cells is required for the differentiation of the male urogenital system late in gestation (Huhtaniemi and Pelliniemi 1992). Foetal Leydig cells also play a role in the scrotal descent of the testis through their synthesis of insulin-like growth factor 3 (Insl3), for review see (Nef 2000).

In humans, the first morphological sign of testicular differentiation is the formation of testicular cords, which can be seen between 6 and 7 weeks of gestation. Steroid-secreting Leydig cells can be seen in the testis at 8 weeks of gestation. At this period, the concentration of androgens in the testicular tissue and blood starts to rise, peaking at 14-16 weeks of gestation. This increase comes with an increase in the number of Leydig cells for review see (Rouiller-Fabre et al. 2009).

Adult Leydig cells, which are distinct from the foetal Leydig cells, form during puberty and supply the testosterone required for the onset of spermatogenesis, among other functions. Distinct stages of adult Leydig cell development have been identified and characterized. The stem Leydig cells are undifferentiated cells that are capable of indefinite self-renewal but also of differentiation to steroidogenic cells. These cells give rise to progenitor Leydig cells, which proliferate, continue to differentiate, and give rise to the immature Leydig cells. Immature Leydig cells synthesize high levels of testosterone metabolites and develop into terminally differentiated adult Leydig cells, which produce high levels of testosterone. With aging, both serum and testicular testosterone concentrations progressively decline, for review see (Nef 2000).

Androgens play a crucial role in the development and maintenance of male reproductive and sexual functions. Low levels of circulating androgens can cause disturbances in male sexual development, resulting in congenital abnormalities of the male reproductive tract. Later in life, this may cause reduced fertility, sexual dysfunction, decreased muscle formation and bone mineralisation, disturbances of fat metabolism, and cognitive dysfunction. Testosterone levels decrease as a process of ageing: signs and symptoms caused by this decline can be considered a normal part of ageing.

How it is Measured or Detected

OECD TG 456 [1] is the validated test guideline for an in vitro screen for chemical effects on steroidogenesis, specifically the production of 17ß-estradiol (E2) and testosterone (T). The testosterone syntheis can be measured in vitro cultured Leydig cells. The methods for culturing Leydig cells can be found in the Database Service on Alternative Methods to animal experimentation (DB-ALM): Leydig Cell-enriched Cultures [2], Testicular Organ and Tissue Culture Systems [3].

Testosterone synthesis in vitro cultured cells can be measured indirectly by testosterone radioimmunoassay or analytical methods such as LC-MS.

References

Chandrashekar, V, and A Bartke. 1998. “The Role of Growth Hormone in the Control of Gonadotropin Secretion in Adult Male Rats.” Endocrinology 139 (3) (March): 1067–74. doi:10.1210/endo.139.3.5816.

Ellis, G B, C Desjardins, and H M Fraser. 1983. “Control of Pulsatile LH Release in Male Rats.” Neuroendocrinology 37 (3) (September): 177–83. Huhtaniemi, I, and L J Pelliniemi. 1992. “Fetal Leydig Cells: Cellular Origin, Morphology, Life Span, and Special Functional Features.” Proceedings of the Society for Experimental Biology and Medicine. Society for Experimental Biology and Medicine (New York, N.Y.) 201 (2) (November): 125–40.

Manson, Jeanne M, and Michael C Carr. 2003. “Molecular Epidemiology of Hypospadias: Review of Genetic and Environmental Risk Factors.” Birth Defects Research. Part A, Clinical and Molecular Teratology 67 (10) (October): 825–36. doi:10.1002/bdra.10084.

Nef, S. 2000. “Hormones in Male Sexual Development.” Genes & Development 14 (24) (December 15): 3075–3086. doi:10.1101/gad.843800.

Rouiller-Fabre, Virginie, Vincent Muczynski, Romain Lambrot, Charlotte Lécureuil, Hervé Coffigny, Catherine Pairault, Delphine Moison, et al. 2009. “Ontogenesis of Testicular Function in Humans.” Folia Histochemica et Cytobiologica / Polish Academy of Sciences, Polish Histochemical and Cytochemical Society 47 (5) (January): S19–24. doi:10.2478/v10042-009-0065-4.

Key Event Component

Domain of Applicability

Key Event Description

Biological state

Steroidogenesis begins with the transport of cholesterol from intracellular stores into mitochondria. This process involves a series of protein-protein interactions involving cytosolic and mitochondrial proteins located at both the outer and inner mitochondrial membranes. In steroidogenic cells the cholesterol import to the mitochondrial inner membrane is crucial for steroid synthesis (Rone, Fan, and Papadopoulos 2009). This process is facilitated by the Scavenger Receptor Class B, type 1 (SR-B1) [more relevant for rodents, than for humans] that mediates the selective uptake of cholesterol esters from high-density lipoproteins. Steroidogenic acute regulatory protein (STAR) and the translator protein (TSPO) [former peripheral benzodiazepine receptor (PBR)] mediate cholesterol transport from the outer to the inner mitochondrial membrane. The conversion of cholesterol to pregnenolone is done by Cholesterol side-chain cleavage enzyme (P450scc), the start of steroidogenesis [reviewed in (Miller and Auchus 2011)].

Biological compartments

In mitochondria of steroidogenic tissues there are two specialized mechanisms related to hormone synthesis: one by which cholesterol is delivered to the mitochondria and the other by which specialized intra-mitochondrial enzymes participate in the synthesis of hormonal steroids.

General role in biology

Systemic steroid hormones are primarily formed by the gonads, adrenal glands, and during in utero development by the placenta. Some other organs like brain (Baulieu 1998), and heart (Kayes-Wandover and White 2000) have also been identified as steroid-producing tissues, mainly for local needs. The steroid hormones are indispensable for mammalian life. They are made from cholesterol via complex biosynthetic pathways that are initiated by specialized, tissue-specific enzymes in mitochondria. These hormones include glucocorticoids (cortisol, corticosterone) and mineralocorticoids (aldosterone) produced in the adrenal cortex, estrogens (estradiol), progestins (progesterone) and androgens (testosterone, dihydrotestosterone) produced in the gonads, and calciferols (1,25-dihydroxy vitamin D [1,25OH2D]) produced in the kidneys (Miller and Auchus 2011). Cholesterol is the precursor for the synthesis of steroid hormones in mitochondria. Steroidogenesis begins with the metabolism of cholesterol to pregnenolone facilitated by P450scc. The rate of steroid formation depends on the rate of cholesterol transport from intracellular stores to the inner mitochondrial membrane and the loading of P450scc with cholesterol (Miller and Auchus 2011). Interference with one or more of these reactions leads to reduced steroid production.

How it is Measured or Detected

This KE can be indirectly measured by:

1. Expression of the proteins involved in cholesterol transport by qPCR or Western blot.

3. Cholesterol transport to the mitochondrial inner membrane in intact cells:

• Indirectly as pregnenolone formation by cells. The pregnenolone concentration is assayed by commercially available radioimmunoassays and reflects the amount of cholesterol transported to the mitochondrial inner membrane (Charman et al. 2010).
• Filipin staining is one of the most widely used tools for studying intracellular cholesterol distribution. The fluorescent detergent filipin binds selectively to cholesterol (and not to cholesterol esters) (Schroeder, Holland, and Bieber 1971). Filipin can be only used for the qualitative analysis of cholesterol distribution, since its fluorescence intensity is not necessarily linearly related to cholesterol content.

The cholesterol transport can be measured in vitro cultured Leydig cells. The methods for culturing Leydig cells can be found in the Database Service on Alternative Methods to animal experimentation (DB-ALM): Leydig Cell-enriched Cultures [1] Testicular Organ and Tissue Culture Systems [2]

References

Albalat, Ricard, Frédéric Brunet, Vincent Laudet, and Michael Schubert. 2011. “Evolution of Retinoid and Steroid Signaling: Vertebrate Diversification from an Amphioxus Perspective.” Genome Biology and Evolution 3: 985–1005. doi:10.1093/gbe/evr084.

Baulieu, E E. 1998. “Neurosteroids: A Novel Function of the Brain.” Psychoneuroendocrinology 23 (8) (November): 963–87.

Charman, Mark, Barry E Kennedy, Nolan Osborne, and Barbara Karten. 2010. “MLN64 Mediates Egress of Cholesterol from Endosomes to Mitochondria in the Absence of Functional Niemann-Pick Type C1 Protein.” Journal of Lipid Research 51 (5) (May): 1023–34. doi:10.1194/jlr.M002345.

Kayes-Wandover, K M, and P C White. 2000. “Steroidogenic Enzyme Gene Expression in the Human Heart.” The Journal of Clinical Endocrinology and Metabolism 85 (7) (July): 2519–25. doi:10.1210/jcem.85.7.6663.

Miller, Walter L, and Richard J Auchus. 2011. “The Molecular Biology, Biochemistry, and Physiology of Human Steroidogenesis and Its Disorders.” Endocrine Reviews 32 (1) (February): 81–151. doi:10.1210/er.2010-0013.

Rone, Malena B, Jinjiang Fan, and Vassilios Papadopoulos. 2009. “Cholesterol Transport in Steroid Biosynthesis: Role of Protein-Protein Interactions and Implications in Disease States.” Biochimica et Biophysica Acta 1791 (7) (July): 646–58. doi:10.1016/j.bbalip.2009.03.001.

Schroeder, F, J F Holland, and L L Bieber. 1971. “Fluorometric Evidence for the Binding of Cholesterol to the Filipin Complex.” The Journal of Antibiotics 24 (12) (December): 846–9.

Steer, C. 1984. “Detection of Membrane Cholesterol by Filipin in Isolated Rat Liver Coated Vesicles Is Dependent upon Removal of the Clathrin Coat.” The Journal of Cell Biology 99 (1) (July 1): 315–319. doi:10.1083/jcb.99.1.315.

Key Event Component

Domain of Applicability

Key Event Description

Biological state

capability to produce offspring

Biological compartments

System

General role in biology

Fertility is the capacity to conceive or induce conception. Impairment of fertility represents disorders of male or female reproductive functions or capacity.

How it is Measured or Detected

As a measure, fertility rate, is the number of offspring born per mating pair, individual or population.

Regulatory Significance of the AO

Under REACH, information on reproductive toxicity is required for chemicals with an annual production/importation volume of 10 metric tonnes or more. Standard information requirements include a screening study on reproduction toxicity (OECD TG 421/422) at Annex VIII (10-100 t.p.a), a prenatal developmental toxicity study (OECD 414) on a first species at Annex IX (100-1000 t.p.a), and from March 2015 the OECD 443(Extended One-Generation Reproductive Toxicity Study) is reproductive toxicity requirement instead of the two generation reproductive toxicity study (OECD TG 416). If not conducted already at Annex IX, a prenatal developmental toxicity study on a second species at Annex X (≥ 1000 t.p.a.).

Under the Biocidal Products Regulation (BPR), information is also required on reproductive toxicity for active substances as part of core data set and additional data set (EU 2012, ECHA 2013). As a core data set, prenatal developmental toxicity study (EU TM B.31) in rabbits as a first species and a two-generation reproduction toxicity study (EU TM B.31) are required. OECD TG 443 (Extended One-Generation Reproductive Toxicity Study) shall be considered as an alternative approach to the multi-generation study.) According to the Classification, Labelling and Packaging (CLP) regulation (EC, 200; Annex I: 3.7.1.1): a) “reproductive toxicity” includes adverse effects on sexual function and fertility in adult males and females, as well as developmental toxicity in the offspring; b) “effects on fertility” includes adverse effects on sexual function and fertility; and c) “developmental toxicity” includes adverse effects on development of the offspring.

List of Adjacent Key Event Relationships

testosterone production, reduction ex vivo (Thompson, Ross, & Gaido, 2004)

List of Non Adjacent Key Event Relationships