Antagonism, Androgen receptor

131983-72-7

PPDBOQMNKNNODG-UHFFFAOYNA-N

PPDBOQMNKNNODG-UHFFFAOYSA-N

Triticonazole

5-[(4-Chlorophenyl)methylene]-2,2-dimethyl-1-(1H-1,2,4-triazol-1-ylmethyl)cyclopentanol

DTXSID0032655

85509-19-9

FQKUGOMFVDPBIZ-UHFFFAOYSA-N

Flusilazole

NuStar

DTXSID3024235

133855-98-8

ZMYFCFLJBGAQRS-UHFFFAOYNA-N

ZMYFCFLJBGAQRS-UHFFFAOYSA-N

Epoxiconazole

DTXSID1040372

67747-09-5

TVLSRXXIMLFWEO-UHFFFAOYSA-N

Prochloraz

1H-Imidazole-1-carboxamide, N-propyl-N-[2-(2,4,6-trichlorophenoxy)ethyl]- BTS 40542-7877 N-propil-N-[2-(2,4,6-triclorofenoxi)etil]-1H-imidazol-1-carboxamida N-propyl-N-[2-(2,4,6-trichlorophenoxy)ethyl]-1H-imidazole-1-carboxamide N-Propyl-N-[2-(2,4,6-trichlorophenoxy)ethyl-1H-imidazole-1-carboxamide N-Propyl-N-[2-(2,4,6-trichlorphenoxy)ethyl]-1H-imidazol-1-carboxamid Plocloraz Prelude Sportak Sportake

DTXSID4024270

60207-90-1

STJLVHWMYQXCPB-UHFFFAOYNA-N

STJLVHWMYQXCPB-UHFFFAOYSA-N

Propiconazole

ppz 1H-1,2,4-Triazole, 1-[[2-(2,4-dichlorophenyl)-4-propyl-1,3-dioxolan-2-yl]methyl]- (.+-.)-1-[2-(2,4-dichlorophenyl)-4-propyl-1,3-dioxolan-2-yl-methyl]-1H-1,2,4-triazole (.+-.)-1-[2-(2,4-dichlorophenyl)-4-propyl-1,3-dioxolan-2-ylmethyl]-1H-1,2,4-triazole 1-[[2-(2,4-dichlorophenyl)-4-propyl-1,3-dioxolan-2-yl]methyl]-1H-1,2,4-triazole 1-[[2-(2,4-dichlorophenyl)-4-propyl-1,3-dioxolane-2-yl]methyl]-1H-1,2,4-triazole 1-[[2-(2,4-Dichlorphenyl)-4-propyl-1,3-dioxolan-2-yl]methyl]-1H-1,2,4-triazol 1-[[2-(2,4-diclorofenil)-4-propil-1,3-dioxolan-2-il]metil]-1H-1,2,4-triazol Bamper 25EC Banner Maxx Cane Sett Treatment Fertilome Liquid Systemic Fungicide Microban PZ Microban S 2140 Mycostat P Proconazole PROPICONAZOL Tilt Premium Wocosen Technical Wocosin Wocosin 50TK

DTXSID8024280

107534-96-3

PXMNMQRDXWABCY-UHFFFAOYNA-N

PXMNMQRDXWABCY-UHFFFAOYSA-N

Tebuconazole

1H-1,2,4-Triazole-1-ethanol, .alpha.-(2-(4-chlorophenyl)ethyl)-.alpha. +- 1H-1,2,4-Triazole-1-ethanol, α-[2-(4-chlorophenyl)ethyl]-α-(1,1-dimethylethyl)- (.+-.)-Tebuconazole 1-(4-Chlorophenyl)-4,4-dimethyl-3-(1,2,4-triazol-1-ylmethyl)pentan-3-ol 1H-1,2,4-Triazole-1-ethanol, α-[2-(4-chlorophenyl)ethyl]-α-(1,1-dimethylethyl)-, (.+-.)- 1H-1,2,4-Triazole-1-ethanol,α-[2-(4-chlorophenyl) ethyl]-α-(1,1-dimethylethyl)-, (.+-.)- BAY-HWG 1608 ETHANOL, α-[2-(4-CHLOROPHENYL)ETHYL]-α- (1,1-DIMETHYLETHYL)-1H-1,2,4-TRIAZOLE Ethyltrianol Etiltrianol Fenetrazole Folicur Microban S 2142 Microban TZ Preventol A 8 TEBUCONAZOL Tebuconazole Resp. HWG 1608 Terbutrazole α-[2-(4-Chlorophenyl)-ethyl]-α-(1,1-dimethylethyl)-1H-1,2,4-triazole-1-ethanol α-[2-(4-chlorophenyl)ethyl]-α-(1,1-dimethylethyl)-1H-1,2,4-triazole-1-ethanol α-tert-Butyl-α-(p-chlorophenethyl)-1H-1,2,4-triazole-1-ethanol

DTXSID9032113

13311-84-7

MKXKFYHWDHIYRV-UHFFFAOYSA-N

Flutamide

Propanamide, 2-methyl-N-[4-nitro-3-(trifluoromethyl)phenyl]- 4-Nitro-3-(trifluoromethyl)isobutyranilide 4'-Nitro-3'-trifluoromethylisobutyranilide Eulexin Flucinom Flutamid flutamida m-Propionotoluidide, α,α,α-trifluoro-2-methyl-4'-nitro- N-(Isopropylcarbonyl)-4-nitro-3-trifluoromethylaniline Niftholide Niftolide NSC 147834 NSC 215876

DTXSID7032004

427-51-0

UWFYSQMTEOIJJG-FDTZYFLXSA-N

Cyproterone acetate

3'H-Cyclopropa[1,2]pregna-1,4,6-triene-3,20-dione, 17-(acetyloxy)-6-chloro-1,2-dihydro-, (1β,2β)- 1,2α-Methylene-6-chloro-17α-acetoxy-4,6-pregnadiene-3,20-dione 1,2α-Methylene-6-chloro-pregna-4,6-diene-3,20-dione 17α-acetate 1,2α-Methylene-6-chloro-Δ4,6-pregnadien-17α-ol-3,20-dione acetate 17-acetate de 6-chloro-1-β,2-β-dihydro-17-hydroxy-3'H-cyclopropa[1,2]pregna-1,4,6-triene-3,20-dione 17-acetato de 6-cloro-1-β,2-β-dihidro-17-hidroxi-3'H-ciclopropa[1,2]pregna-1,4,6-trieno-3,20-diona 17α-Acetoxy-6-chloro-1α,2α-methylenepregna-4,6-diene-3,20-dione 3'H-Cyclopropa[1,2]pregna-1,4,6-triene-3,20-dione 3'H-Cyclopropa[1,2]pregna-1,4,6-triene-3,20-dione, 6-chloro-1β,2β-dihydro-17-hydroxy-, acetate 6-Chlor-1-β,2-β-dihydro-17-hydroxy-3'H-cyclopropa[1,2]pregna-1,4,6-trien-3,20-dion-17-acetat 6-Chloro-1,2α-methylene-17α-hydroxy-Δ6-progesterone acetate 6-Chloro-1,2α-methylene-6-dehydro-17α-hydroxyprogesterone acetate 6-Chloro-17-hydroxy-1α,2α-methylenepregna-4,6-diene-3,20-dione acetate 6-chloro-1-β,2-β-dihydro-17-hydroxy-3'H-cyclopropa[1,2]pregna-1,4,6-triene-3,20-dione 17-acetate Androcur Cyprostat Cyproterone 17-O-acetate Cyproterone 17α-acetate Cyproviron NSC 81430 Pregna-4,6-diene-3,20-dione, 6-chloro-17-hydroxy-1α,2α-methylene-, acetate

DTXSID5020366

50471-44-8

FSCWZHGZWWDELK-UHFFFAOYNA-N

FSCWZHGZWWDELK-UHFFFAOYSA-N

Vinclozolin

2,4-Oxazolidinedione, 3-(3,5-dichlorophenyl)-5-ethenyl-5-methyl- (.+-.)-Vinclozolin BAS 352-04F N-3,5-Dichlorophenyl-5-methyl-5-vinyl-1,3-oxazolidine-2,4-dione N-3,5-Dichlorophenyl-5-methyl-5-vinyloxazolidine-2,4-dione N-3,5-Dichlorphenyl-5-methyl-5-vinyl-1,3-oxazolidin-2,4-dion N-3,5-diclorofenil-5-metil-5-vinil-1,3-oxazolidina-2,4-diona Ornalin Ranilan Ronilan Ronilan 50WP

DTXSID4022361

PR:000004191

PR androgen receptor

CHEBI:50113

CHEBI androgen

PR:000001449

PR fibroblast growth factor receptor 2

PR:000007480

PR fibroblast growth factor 10

PR:000007489

PR fibroblast growth factor 2

UBERON:0011374

UBERON prepuce

UBERON:0001332

UBERON prepuce of penis

GO:0004882

GO androgen receptor activity

GO:0000060

GO protein import into nucleus, translocation

GO:0051101

GO regulation of DNA binding

GO:0060638

GO mesenchymal-epithelial cell signaling

GO:0001775

GO cell activation

GO:0035261

GO external genitalia morphogenesis

MP:0008940

MP delayed balanopreputial separation

WIKI decreased

WIKI disrupted

WIKI delayed

WIKI arrested

Mercaptobenzole

2016-11-29T18:42:26

Triticonazole

2020-05-16T11:02:07

2020-05-16T11:09:42

Flusilazole

2020-05-16T11:15:34

Epoxiconazole

2020-05-16T11:35:44

Prochloraz

2016-11-29T18:42:22

Propiconazole

2017-05-17T13:18:07

Tebuconazole

2017-05-17T13:17:14

Flutamide Flutamide is a selective androgen receptor (AR) antagonist (Simard et al 1986) that has been shown to induce shorter male AGD in rats after in utero exposure (Foster & Harris 2005; Hass et al 2007; Kita et al 2016; McIntyre et al 2001; Mylchreest et al 1999; Scott et al 2007; Welsh et al 2007).

2016-11-29T18:42:27

2025-08-14T05:22:07

Cyproterone acetate

2020-05-17T10:13:28

Vinclozolin

2020-05-14T11:28:31

WikiUser_17

mammals

10090

NCBI Mus musculus

60556

NCBI Oryctolagus sp.

9606

NCBI Homo sapiens

10141

NCBI guinea pig

10116

NCBI Rattus norvegicus

10141

NCBI Cavia porcellus

40674

https://www.ncbi.nlm.nih.gov/Taxonomy/Browser/wwwtax.cgi?command=show&mode=node&id=40674&lvl= Mammalia

Antagonism, Androgen receptor

Molecular

The androgen receptor (AR) and its function The AR is a ligand-activated transcription factor belonging to the steroid hormone nuclear receptor family (<a href="https://aopwiki.org/events/26#_ENREF_1" title="Davey, 2016 #250">Davey & Grossmann, 2016</a>). The AR has three domains: the N-terminal domain, the DNA-binding domain, and the ligand-binding domain, with the latter being most evolutionary conserved. Testosterone (T) and the more biologically active dihydrotestosterone (DHT) are endogenous ligands for the AR (<a href="#_ENREF_4" title="MacLean, 1993 #251">MacLean et al, 1993</a>; <a href="#_ENREF_5" title="MacLeod, 2010 #27">MacLeod et al, 2010</a>; <a href="#_ENREF_8" title="Schwartz, 2019 #252">Schwartz et al, 2019</a>). In teleost fishes, 11-ketotestosterone is the second main ligand (<a href="#" title="Schuppe et al, 2020">Schuppe et al, 2020</a>). Human AR mutations and mouse knock-out models have established a pivotal role for the AR in masculinization and spermatogenesis (<a href="#_ENREF_9" title="Walters, 2010 #254">Walters et al, 2010</a>). Apart from the essential role for AR in male reproductive development and function (<a href="#_ENREF_9" title="Walters, 2010 #254">Walters et al, 2010</a>), the AR is also expressed in many other tissues and organs such as bone, muscles, ovaries, and the immune system (<a href="#_ENREF_7" title="Rana, 2014 #253">Rana et al, 2014</a>).  AR antagonism as Key Event The main function of the AR is to activate gene transcription in cells. Canonical signaling occurs by ligands (androgens) binding to AR in the cytoplasm which results in translocation to the cell nucleus, receptor dimerization, and binding to specific regulatory DNA sequences (<a href="#_ENREF_2" title="Heemers, 2007 #255">Heemers & Tindall, 2007</a>). The gene targets regulated by AR activation depends on cell/tissue type and what stage of development activation occurs, and is, for instance, dependent on available co-factors. Apart from the canonical signaling pathway, AR can also initiate cytoplasmic signaling pathways with other functions than the nuclear pathway, for instance rapid change in cell function by ion transport changes (<a href="#_ENREF_3" title="Heinlein, 2002 #256">Heinlein & Chang, 2002</a>) and association with Src kinase to activate MAPK/ERK signaling and activation of the PI3K/Akt pathway (<a href="#" title="Leung & Sadar, 2017">Leung & Sadar, 2017</a>).

AR antagonism can be measured in vitro by transient or stable transactivation assays to evaluate nuclear receptor activation. There is already a validated test guideline for AR (ant)agonism adopted by the OECD, Test No. 458: Stably Transfected Human Androgen Receptor Transcriptional Activation Assay for Detection of Androgenic Agonist and Antagonist Activity of Chemicals (<a href="#_ENREF_13" title="OECD, 2016 #257">OECD, 2016</a>). This test guideline contains three different methods. More information on limitations, advantages, protocols, and availability, and description of cells are given in the test guideline. Besides these validated methods, other transiently or stably transfected reporter cell lines are available as well as yeast based systems (Campana et al, 2015; <a href="#_ENREF_10" title="Körner, 2004 #282">Körner et al, 2004</a>). AR nuclear translocation can be monitored by various assays (Campana et al 2015), for example by monitoring fluorescent rat AR movement in living cells (Tyagi et al 2020), with several human AR translocation assays being commercially available; e.g. Fluorescent AR Nuclear Translocation Assay (tGFP-hAR/HEK293) or Human Androgen NHR Cell Based Antagonist Translocation LeadHunter Assay. Additional information on AR interaction can be obtained employing competitive AR binding assays (Freyberger et al 2010, Shaw et al 2018), which can also inform on relative potency of the compounds, though not on downstream effect of the AR binding. The recently developed AR dimerization assay provides an assay with an improved ability to measure potential stressor-mediated disruption of dimerization/activation (<a href="#_ENREF_11" title="Lee, 2021 #288">Lee et al, 2021</a>).

Both the DNA-binding and ligand-binding domains of the AR are highly evolutionary conserved, whereas the transactivation domain show more divergence which may affect AR-mediated gene regulation across species (<a href="#_ENREF_1" title="Davey, 2016 #250">Davey & Grossmann, 2016</a>). Despite certain inter-species differences, AR function mediated through gene expression is highly conserved, with mutations studies from both humans and rodents showing strong correlation for AR-dependent development and function (<a href="#_ENREF_9" title="Walters, 2010 #254">Walters et al, 2010</a>).  This KE is applicable for both sexes, across developmental stages into adulthood, in numerous cells and tissues, and across mammalian taxa. It is, however, acknowledged that this KE most likely has a much broader domain of applicability extending to non-mammalian vertebrates. AOP developers are encouraged to add additional relevant knowledge to expand on the applicability to also include other vertebrates.

CL:0000255

CL eukaryotic cell

High Mixed

High

During development and at adulthood

High

Campana C, Pezzi V, Rainey WE (2015) Cell based assays for screening androgen receptor ligands. Semin Reprod Med 33: 225-234. <a name="_ENREF_2">Davey RA, Grossmann M (2016) Androgen Receptor Structure, Function and Biology: From Bench to Bedside. Clin Biochem Rev 37: 3-15</a> Freyberger A, Weimer M, Tran HS, Ahr HJ. Assessment of a recombinant androgen receptor binding assay: initial steps towards validation. Reprod Toxicol. 2010 Aug;30(1):2-8. doi: 10.1016/j.reprotox.2009.10.001. Epub 2009 Oct 13. PMID: 19833195. <a name="_ENREF_6">Heemers HV, Tindall DJ (2007) Androgen receptor (AR) coregulators: a diversity of functions converging on and regulating the AR transcriptional complex. Endocr Rev 28: 778-808</a> <a name="_ENREF_7">Heinlein CA, Chang C (2002) The roles of androgen receptors and androgen-binding proteins in nongenomic androgen actions. Mol Endocrinol 16: 2181-2187</a> <a name="_ENREF_10">Körner W, Vinggaard AM, Térouanne B, Ma R, Wieloch C, Schlumpf M, Sultan C, Soto AM (2004) Interlaboratory comparison of four in vitro assays for assessing androgenic and antiandrogenic activity of environmental chemicals. Environ Health Perspect 112: 695-702</a> <a name="_ENREF_11">Lee SH, Hong KY, Seo H, Lee HS, Park Y (2021) Mechanistic insight into human androgen receptor-mediated endocrine-disrupting potentials by a stable bioluminescence resonance energy transfer-based dimerization assay. Chem Biol Interact 349: 109655</a> <a id="_ENREF_23" name="_ENREF_23">Leung, J. K., & Sadar, M. D. (2017). Non-Genomic Actions of the Androgen Receptor in Prostate Cancer. Frontiers in Endocrinology, 8. https://doi.org/10.3389/fendo.2017.00002</a> <a name="_ENREF_12">MacLean HE, Chu S, Warne GL, Zajac JD (1993) Related individuals with different androgen receptor gene deletions. J Clin Invest 91: 1123-1128</a> <a name="_ENREF_13">MacLeod DJ, Sharpe RM, Welsh M, Fisken M, Scott HM, Hutchison GR, Drake AJ, van den Driesche S (2010) Androgen action in the masculinization programming window and development of male reproductive organs. Int J Androl 33: 279-287</a> <a name="_ENREF_14">OECD. (2016) Test No. 458: Stably Transfected Human Androgen Receptor Transcriptional Activation Assay for Detection of Androgenic Agonist and Antagonist Activity of Chemicals. OECD Guidelines for the Testing of Chemicals, Section 4, Paris.</a> OECD (2022). Test No. 251: <a name="_Hlk148359154">Rapid Androgen Disruption Activity Reporter (RADAR) assay</a>. Paris: OECD Publishing doi:10.1787/da264d82-en. <a name="_ENREF_15">Rana K, davey RA, Zajac JD (2014) Human androgen deficiency: insights gained from androgen receptor knockout mouse models. Asian J Androl 16: 169-177</a> <a name="_ENREF_16">Satoh K, Ohyama K, Aoki N, Iida M, Nagai F (2004) Study on anti-androgenic effects of bisphenol a diglycidyl ether (BADGE), bisphenol F diglycidyl ether (BFDGE) and their derivatives using cells stably transfected with human androgen receptor, AR-EcoScreen. Food Chem Toxicol 42: 983-993</a> <a id="_ENREF_22" name="_ENREF_22">Schuppe, E. R., Miles, M. C., and Fuxjager, M. J. (2020). Evolution of the androgen receptor: Perspectives from human health to dancing birds. Mol. Cell. Endocrinol. 499, 110577. doi:10.1016/J.MCE.2019.110577 </a> <a name="_ENREF_17">Schwartz CL, Christiansen S, Vinggaard AM, Axelstad M, Hass U, Svingen T (2019) Anogenital distance as a toxicological or clinical marker for fetal androgen action and risk for reproductive disorders. Arch Toxicol 93: 253-272</a> Shaw J, Leveridge M, Norling C, Karén J, Molina DM, O'Neill D, Dowling JE, Davey P, Cowan S, Dabrowski M, Main M, Gianni D. Determining direct binders of the Androgen Receptor using a high-throughput Cellular Thermal Shift Assay. Sci Rep. 2018 Jan 9;8(1):163. doi: 10.1038/s41598-017-18650-x. PMID: 29317749; PMCID: PMC5760633. Tyagi RK, Lavrovsky Y, Ahn SC, Song CS, Chatterjee B, Roy AK (2000) Dynamics of intracellular movement and nucleocytoplasmic recycling of the ligand-activated androgen receptor in living cells. Mol Endocrinol 14: 1162-1174 <a id="_ENREF_21" name="_ENREF_21">Walters KA, Simanainen U, Handelsman DJ (2010) Molecular insights into androgen actions in male and female reproductive function from androgen receptor knockout models. Hum Reprod Update 16: 543-558</a> AOPWiki 2016-11-29T18:41:22

2026-03-20T11:42:44

Androgen receptor nuclear transcriptional activity in genital-tubercle tissues, reduced

AR transcriptional activity in GT tissues, reduced

Cellular

The event describes a decrease in androgen receptor (AR)-mediated gene transcription in genital tubercle (GT) tissue.  It reflects a state in which AR fails to be activated to as a result of reduced AR translocation to the nucleus, diminished DNA binding to androgen response elements, and/or lower recruitment of transcriptional co-regulators.   The biological compartment it is measured in are GT tissues, particularly mesenchymal and epithelial cells.  AR transcriptional activity is involved in differentiation of the GT by regulating genes that control outgrowth and elongation of the tubercle, formation for the penile erectile tissues, ventral urethral closure and patterning, and morphogenesis of external genitalia.

Direct evidence with well validated and widely used techniques that strongly correlate with AR activation state. <ul> <li>Nuclear AR localization, which is the amount of AR protein in the cell nucleus, can be measured via: <ul> <li>Immunohistochemistry</li> <li>Immunofluorescence</li> <li>Western blot</li> </ul> </li> </ul> <ul> <li>AR DNA binding, which measures androgen response elements on chromatin, can be measured via: <ul> <li>Chromatin Immunoprecipitation followed by qPCR or seq.</li> </ul> </li> </ul> Indirect evidence well validated and widely used techniques that strongly correlate with AR activation state. <ul> <li>Expression of AR target genes, which measure either mRNA or protein levels of AR-regulated genes, can be measured via: <ul> <li>RT-qPCR</li> <li>RNA-seq</li> <li>In situ hybridization</li> <li>Proteomics</li> </ul> </li> </ul>

Taxonomic Applicability Reduced AR nuclear transcriptional activity in genital-tubercle tissues has been measured mammals including humans and rodents (Veyssiere et al., 1985; Agras et al., 2006; Rodriguez et al., 2012).  It’s plausible for any species expressing AR in GT tissue developing into external genitalia. SeqAPASS results for taxonomic conservation within Mammalia is attached for AR (<a href="http://www.ncbi.nlm.nih.gov/protein/AAA51780.1" target="_blank">AAA51780.1</a>) as <a href="https://aopwiki.org/system/dragonfly/production/2025/12/17/8jzsq5oyu5_AR_SeqAPASS_Mammalia.xlsx">AR-SeqAPASS_Mammalia.xlsx</a>. Lifestage Applicability AR transcriptional activity in GT tissues is critical in embryonic and fetal periods.  AR signalling during this time-period influences genital tissue patterning and morphogenesis. <ul> <li>Gestation day 18 in rabbits (Veyssiere et al., 1985) </li> <li>Gestational day 15.5-17.5 (Miyagawa et al., 2009) </li> <li>Gestation day 14 in mice (Agras et al., 2006)</li> </ul> Sex Applicability The genital tubercle forms early in embryonic development in males and females.  AR nuclear transcription occurs in that tissue in both males and females although it is the dominant driver of GT differentiation in males and only occurs transiently and at lower levels in females (Sajjad et al., 2004).

UBERON:0005876

UBERON undifferentiated genital tubercle

High Unspecific

High

Development

High

Fetal to Parturition

High

Foetal

High

Embryo

High High High High

Agras, K., Willingham, E., Liu, B., & Baskin, L. S. (2006). Ontogeny of androgen receptor and disruption of its mRNA expression by exogenous estrogens during morphogenesis of the genital tubercle. The Journal of urology, 176(4), 1883-1888. Miyagawa, S., Satoh, Y., Haraguchi, R., Suzuki, K., Iguchi, T., Taketo, M. M., ... & Yamada, G. (2009). Genetic interactions of the androgen and Wnt/β-catenin pathways for the masculinization of external genitalia. Molecular endocrinology, 23(6), 871-880 Rodriguez Jr, E., Weiss, D. A., Ferretti, M., Wang, H., Menshenia, J., Risbridger, G., ... & Baskin, L. (2012). Specific morphogenetic events in mouse external genitalia sex differentiation are responsive/dependent upon androgens and/or estrogens. Differentiation, 84(3), 269-279. Sajjad, Y., Quenby, S., Nickson, P., Lewis-Jones, D. I., & Vince, G. (2004). Immunohistochemical localization of androgen receptors in the urogenital tracts of human embryos. Reproduction, 128(3), 331-339. Veyssiere, G., Berger, M., Jean-Faucher, C., De Turckheim, M., & Jean, C. (1985). Androgen receptor in genital tubercle of rabbit fetuses and newborns. Ontogeny and properties. Journal of steroid biochemistry, 23(4), 399-404. AOPWiki 2025-12-03T11:50:25

2025-12-18T11:35:20

Fibroblast growth factor 10, fibroblast growth factor receptor 2 isoform IIIb signaling in genital tissue, reduced

FGF10/FGFR2-IIIb signaling in genital tissue, reduced

Cellular

This event describes a decrease in Fibroblast growth factor (FGF) signaling mediated by changes to FGF10 and/or FGFR2-IIIb within genital tissues.  It reflects a state in which FGF10 ligand availability is reduced and/or FGFR2-IIIb expression or activation is diminished. The biological compartment it is measured in are mesenchymal and epithelial cells in genital tissues including GT, preputial lamina, and urethral epithelium (Wang et al., 2025). FGF10/FGFR2-IIIb signaling plays an essential role in epithelial morphogenesis during external genital development by influencing outgrowth of preputial epithelium, maintenance of epithelial cell processes, and coordination of mesenchymal epithelial interactions required from normal foreskin/clitoral hood formation. This signaling has been demonstrated to play the same role in other tissues, i.e., lung and palate, as well (Rice et al., 2004; Warburton et al., 2003).

Direct evidence with well validated and widely used techniques that measure ligand/receptor expression and activation. Expression of FGF10 and FGFR2-IIIb: <ul> <li>RT-qPCR</li> <li>In situ hybridation</li> <li>IHC/IF</li> <li>Western blotting</li> </ul> Receptor activation of FGFR2-IIIb by way of phosphorylation state: <ul> <li>ICH or Western blot</li> <li>Immunoprecipation followed by phospho-Western blotting</li> </ul> Indirect evidence well validated and widely used techniques that correlate to ligand/receptor expression and activation. Expression of genes induced by FGF signaling can be measured via: <ul> <li>RT-qPCR</li> <li>RNA-seq</li> <li>In situ hybridization</li> </ul>

Taxonomic Applicability FGF family and FGF receptors are present and functionally conserved across vertebrates however, their association with external genital development has not been established in non-mammals.  This association has been established in mice, humans, and guinea pigs (Wang et al., 2025; Beleza-Meireles et al., 2007; Gredler et al., 2015).  SeqAPASS results for taxonomic conservation is attached for FGF10 and FGFR2 as <a href="https://aopwiki.org/system/dragonfly/production/2025/12/18/8er4wjnt7c_FGF10_SeqAPASS.xlsx">FGF10_SeqAPASS.xlsx</a> and <a href="https://aopwiki.org/system/dragonfly/production/2025/12/18/49ir5rux5o_FGFR2_SeqAPASS.xlsx">FGFR2_SeqAPASS.xlsx</a> respectively. Lifestage Applicability The highest impact of FGF10/FGFR2-IIIb signaling on the development of GT and preputial tissues occurs in embryonic stages, during the period where these structures are established (Harada et al., 2015).  It’s plausible that there are residual effects of this signaling in epithelial maturation in the perinatal and postnatal periods based on its role in other tissues (Cui and Li 2013). Sex Applicability FGF10/FGFR2-IIIb signaling in genital and pre-cursor tissues occurs in both sexes during development.

UBERON:0005876

UBERON undifferentiated genital tubercle

High Unspecific

High

Embryo

High

Foetal

High

Fetal to Parturition

High

Development

High High High High

Beleza-Meireles, A., Lundberg, F., Lagerstedt, K., Zhou, X., Omrani, D., Frisén, L., & Nordenskjöld, A. (2007). FGFR2, FGF8, FGF10 and BMP7 as candidate genes for hypospadias. European Journal of Human Genetics, 15(4), 405-410. Cui, Y., & Li, Q. (2013). Expression and functions of fibroblast growth factor 10 in the mouse mammary gland. International journal of molecular sciences, 14(2), 4094-4105. Gredler, M. L., Seifert, A. W., & Cohn, M. J. (2015). Tissue-specific roles of Fgfr2 in development of the external Genitalia. Development, 142(12), 2203-2212. Harada, M., Omori, A., Nakahara, C., Nakagata, N., Akita, K., & Yamada, G. (2015). Tissue‐specific roles of FGF signaling in external genitalia development. Developmental Dynamics, 244(6), 759-773. Rice, Ritva, Bradley Spencer-Dene, Elaine C. Connor, Amel Gritli-Linde, Andrew P. McMahon, Clive Dickson, Irma Thesleff, and David PC Rice. "Disruption of Fgf10/Fgfr2b-coordinated epithelial-mesenchymal interactions causes cleft palate." The Journal of clinical investigation 113, no. 12 (2004): 1692-1700. Wang, S., & Zheng, Z. (2025). Differences in Formation of Prepuce and Urethral Groove During Penile Development Between Guinea Pigs and Mice Are Controlled by Differential Expression of Shh, Fgf10 and Fgfr2. Cells, 14(5), 348. Warburton, D., Bellusci, S., Del Moral, P. M., Kaartinen, V., Lee, M., Tefft, D., & Shi, W. (2003). Growth factor signaling in lung morphogenetic centers: automaticity, stereotypy and symmetry. Respiratory research, 4(1), 5. AOPWiki 2025-12-11T18:01:12

2025-12-18T11:43:02

Preputial epithelial morphogenesis, disrupted

Cellular

This state is characterized by a disruption of the normal cellular and tissue processes involved in the formation of the prepuce.  These processes include: <ul> <li>Cellular proliferation, survival, adhesion, and differentiation.</li> <li>Tissue formation, position, and presence/absence.</li> </ul> There are cellular and tissue measurements that take place in the preputial tissue, either the foreskin or clitoral hood.  The cell types the measurements take place in are epithelial and mesenchymal cells. When preputial epithelial morphogenesis occurs normally, the result is the appropriate formation of the prepuce, both inner mucosal and outer cutaneous layers, covering and protecting the glans.

Direct evidence with well validated and widely used techniques based on quantitative and qualitative histological measurements. <ul> <li>Hematoxylin and eosin staining</li> <li>Serial sectioning with stereo microscopy, imaging, morphometric analysis</li> <li>Ki-67 immunostaining and BrdU or EdU incorporation assays for proliferation</li> <li>TUNEL and cleaved caspase-3 for apoptosis</li> </ul>

Taxonomic Applicability Preputial epithelial morphogenesis is applicable in mammals, having been demonstrated/measured directly in mice and humans (Liu et al., 2018; Cunha et al., 2020) as well as other rodents.  It’s not applicable outside of Mammalia because true preputial structures and the associated morphogenesis are mammal specific. Lifestage Applicability This is a developmental process primarily applicable in embryonic and fetal stages during the period of external genital morphogenesis. Sex Applicability Preputial morphogenesis occurs in both males and females with male morphogenesis culminating in properly developed and attached foreskin and female morphogenesis culminating in properly developed and attached clitoral hood.  Concordantly, the phenotypic outcomes of disruption differ.

UBERON:0011374

UBERON prepuce

CL:0000066

CL epithelial cell

High Unspecific

High

Embryo

High

Foetal

High

Fetal to Parturition

High

Development

High High High

Liu, X., Liu, G., Shen, J., Yue, A., Isaacson, D., Sinclair, A., ... & Baskin, L. (2018). Human glans and preputial development. Differentiation, 103, 86-99. Cunha, G. R., Sinclair, A., Cao, M., & Baskin, L. S. (2020). Development of the human prepuce and its innervation. Differentiation, 111, 22-40. AOPWiki 2025-12-15T18:13:37

2025-12-18T15:33:39

Male preputial separation, failed/delayed

Male PPS, failed/delayed

Tissue

This state is characterized by failure or delay in the natural detachment of the prepuce from the glans penis. The biological compartment this is measured in is the glans penis, glans-prepuce adhesion interface, and preputial epithelium. Normal preputial separation serves protective, mechanical, immunological and erogenous functions (Paraboschi and Garriboli 2020).  Biological consequences of a failure in this separation fall into the same areas.

Gross anatomical observations are a direct, standardized, and reproducible strategy and can be conducted periodically.  Binary scoring and ordinal scales can be employed to describe degree of separation and timing.

Taxonomic Applicability Delayed/failed preputial separation is limited to mammals because the process is unique to mammalian foreskin anatomy.  It is well documented in rats and mice as a developmental milestone (Gray et al., 2001).  It also occurs in humans but isn’t understood as a standardized developmental marker (Cunha et al., 2020). Lifestage Applicability Preputial separation is a maturation event that takes place postnatally.  It is an indicator of the onset of puberty in rodents, typically occuring 4-5 weeks after birth in mice and 6-7 weeks after birth in rats.  In humans, it is a variable and ongoing process starting in infancy with significant physiological shifts coinciding with puberty.  In all cases, if separation has not occurred by adulthood, it has failed permanently. Sex Applicability Male preputial separation refers to the detachment of the foreskin from glans penis thereby limiting the applicability to the male.

UBERON:0001332

UBERON prepuce of penis

High Male

High

Development

High

1 to < 3 months

High High High High

Cunha, G. R., Sinclair, A., Cao, M., & Baskin, L. S. (2020). Development of the human prepuce and its innervation. Differentiation, 111, 22-40. Federal Insecticide, Fungicide, and Rodenticide Act, 7 U.S.C. §§ 136–136y (2023). Gray Jr, L. E., Ostby, J., Furr, J., Wolf, C. J., Lambright, C., Parks, L., ... & Guillette, L. (2001). Effects of environmental antiandrogens on reproductive development in experimental animals. Apmis, 109(S103), S302-S319. OECD (2001), Test No. 416: Two-Generation Reproduction Toxicity, OECD Guidelines for the Testing of Chemicals, Section 4, OECD Publishing, Paris, <a href="https://doi.org/10.1787/9789264070868-en">https://doi.org/10.1787/9789264070868-en</a>. OECD (2025), Test No. 443: Extended One-Generation Reproductive Toxicity Study, OECD Guidelines for the Testing of Chemicals, Section 4, OECD Publishing, Paris, <a href="https://doi.org/10.1787/9789264185371-en">https://doi.org/10.1787/9789264185371-en</a>. Paraboschi, I., Garriboli, M. (2020). Medical and Surgical Uses of the Prepuce. In: Normal and Abnormal Prepuce. Springer, Cham. https://doi.org/10.1007/978-3-030-37621-5_8 Toxic Substances Control Act, as amended by the Frank R. Lautenberg Chemical Safety for the 21st Century Act, 15 U.S.C. §§ 2601–2697 (2023 U.S. Environmental Protection Agency. (2013, June 14). Endocrine disruptor screening program; final policies and procedures for screening Safe Drinking Water Act substances. Federal Register. <a href="https://www.federalregister.gov/documents/2013/06/14/2013-14228/endocrine-disruptor-screening-program-final-policies-and-procedures-for-screening-safe-drinking">https://www.federalregister.gov/documents/2013/06/14/2013-14228/endocrine-disruptor-screening-program-final-policies-and-procedures-for-screening-safe-drinking</a> U.S. Environmental Protection Agency. Final List of Initial Pesticide Active Ingredients and Pesticide Inert Ingredients to be Screened Under the Federal Food, Drug, and Cosmetic Act [Document ID EPA-HQ-OPPT-2004-0109-0080]. Regulations.gov. <a href="https://www.regulations.gov/document/EPA-HQ-OPPT-2004-0109-0080">https://www.regulations.gov/document/EPA-HQ-OPPT-2004-0109-0080</a> U.S. Environmental Protection Agency, Office of Prevention, Pesticides and Toxic Substances. (1998). Health Effects Test Guidelines: OPPTS 870.3800—Reproduction and fertility effects. <a href="https://www.epa.gov/test-guidelines-pesticide-registration/series-870-health-effects-test-guidelines">https://www.epa.gov/test-guidelines-pesticide-registration/series-870-health-effects-test-guidelines</a> AOPWiki 2025-12-15T18:15:47

2026-02-06T16:55:39

<upstream-id>ba82ea90-e914-402e-948c-c2611a79634b</upstream-id> <downstream-id>51864830-ea4f-4d13-b4e2-8bfa9e05dcb1</downstream-id> The androgen receptor (AR) is a ligand-activated nuclear transcription factor belonging to the nuclear receptor superfamily. Under normal developmental conditions, androgens (principally testosterone and its more potent metabolite dihydrotestosterone, DHT) bind to the AR in target tissues, triggering a conformational change, dissociation from heat shock proteins, receptor dimerization, nuclear translocation, and binding to androgen response elements (AREs) in the promoter regions of androgen-responsive genes (Brinkmann et al., 1999; Heinlein & Chang, 2002). This sequence of events constitutes AR-mediated transcriptional activation. In the genital tubercle, AR-dependent transcription contributes to a program of gene expression necessary for androgen-dependent GT outgrowth and masculinization during fetal development. The GT is the embryological precursor of the penis (and glans clitoris in females), and its masculinization depends in part on sustained AR signaling during a defined developmental window (Welsh et al., 2008; Blaschko et al., 2012). Downstream targets of AR-mediated transcription in the GT include components of the FGF signaling pathway (Petiot et al. 2005). AR antagonism, whether by exogenous antiandrogens (e.g., flutamide, vinclozolin, procymidone, finasteride-class compounds active via AR) or endocrine disrupting chemicals with antiandrogenic activity, competitively occupies the ligand-binding domain (LBD) of the AR without inducing the transcriptionally competent conformation. The antagonist-bound AR fails to productively recruit coactivators and fails to drive ARE-dependent transcription. The consequence in GT tissue is a reduction in the transcriptional output that normally promotes GT growth, mesenchymal-epithelial signaling, and patterning gene expression. Persistent AR antagonism during the critical developmental window in the GT is therefore causally linked to a reduction in AR transcriptional activity in GT mesenchymal and epithelial cells, which in turn compromises the androgen-driven gene expression program required for normal masculinization.

Evidence for this KER was assembled through a combination of expert knowledge and AI-assisted literature search and synthesis. Specifically, a combination of Claude (Anthropic) using Sonnet 4.6, and EPA AI, using GPT5, was used to identify, retrieve, and summarize relevant primary and secondary literature, and to draft the initial content of this KER page. All citations generated through this process were subsequently reviewed and verified by the KER author against primary sources and DOI resolution checks, prior to inclusion. Users of this KER are advised that AI-assisted evidence assembly may introduce selection bias or gaps in coverage that differ from a fully systematic human-conducted review, and independent verification of the evidence base is encouraged.  A copy of the initial prompt is attached to AOP619. A review of primary experimental literature using the following databases and search strategies was employed: PubMed/MEDLINE searches were conducted (literature through early 2025) using the following terms and combinations: "androgen receptor antagonism genital tubercle," "AR transcriptional activity fetal genitalia," "antiandrogen hypospadias," "flutamide androgen response element," "DHT transcription GT," "vinclozolin androgen receptor," "AR coactivator recruitment antiandrogen," "androgen responsive genes penis development," and "dihydrotestosterone genital tubercle gene expression." Additional sources consulted include: OECD Test Guideline documentation (TG 421, TG 422, TG 441, and the Hershberger assay TG 441); U.S. EPA EDSP (Endocrine Disruptor Screening Program) documentation; and review articles on male reproductive development and AR biology. Screening criteria prioritized studies that directly measured AR transcriptional readouts (ARE-reporter activity, androgen-responsive gene mRNA/protein levels) in GT tissue or GT-relevant cell lines in the context of AR ligand antagonism, or that linked AR antagonism to altered GT gene expression during the male programming window.

The quantitative relationship between the degree of AR antagonism and the magnitude of reduction in AR transcriptional activity in GT tissues is biologically grounded in competitive receptor pharmacology but is incompletely characterized with GT-tissue-specific empirical data. The available evidence is summarized below by subsection.

The biological rationale for this KER is grounded in the canonical mechanism of nuclear receptor pharmacology and developmental androgen biology. The AR ligand-binding domain (LBD) accommodates either agonist ligands (testosterone, DHT) or antagonist ligands (e.g., flutamide, bicalutamide, vinclozolin metabolites, hydroxyflutamide). Agonist binding induces Helix-12 repositioning within the LBD, creating a surface that recruits p160 coactivators (SRC-1, SRC-2/GRIP1, SRC-3) via their LXXLL motifs; this coactivator complex recruits general transcription machinery and histone acetyltransferases, ultimately enabling transcription of androgen-responsive genes (Heinlein & Chang, 2002). Chromatin immunoprecipitation studies confirm that agonist-bound AR recruits coactivators and RNA polymerase II to androgen response elements at target gene enhancers and promoters, whereas antagonist-bound AR instead recruits corepressors to the promoter (Shang et al., 2002). In contrast, antagonist binding can prevent productive AR transcriptional activation. Structural studies of bicalutamide bound to a mutant AR LBD provide evidence that antagonist binding makes direct contacts with Helix-12 residues in a manner that would disrupt the AF-2 coactivator-binding groove, preventing productive coactivator surface formation, as seen in a mutant complex (Bohl et al., 2005). Masiello et al., 2002 showed that bicalutamide-liganded AR translocates to the nucleus and binds DNA but fails to stimulate AR N/C-terminal interaction or recruit SRC-1 or SRC-2 coactivator proteins, resulting in a transcriptionally inactive receptor complex on DNA. In the GT specifically, AR is expressed in mesenchymal cells of the developing phallus from early embryonic stages, particularly in the bilateral mesenchyme flanking the urethral plate from approximately E14.5 in mice (Miyagawa et al., 2009; Matsushita et al., 2018; Blaschko et al., 2012). Downstream androgen-responsive genes in the GT include those governing cell proliferation, apoptosis suppression, and mesenchymal-epithelial inductive signaling (e.g., Fgf10, Bmp4, Wnt5a regulation downstream of androgen signaling; Seifert et al., 2008). The direct mechanistic chain from AR occupancy by an antagonist to failure of ARE-driven transcriptional activation in GT mesenchyme is therefore strongly supported by the combined understanding of AR structural biology and GT developmental biology.

AR transactivation assays provide direct empirical evidence that antiandrogens reduce AR-driven transcriptional output. Cell-based ARE-reporter assays demonstrate concentration-dependent and competitive inhibition of DHT-stimulated AR transcriptional activity by classical AR antagonists, including vinclozolin metabolites M1 and M2 (Kelce et al., 1994) and bicalutamide (Masiello et al., 2002). These assays show hat AR occupancy by an antagonist can suppresses ARE-dependent reporter gene activity in a manner that is competitive with DHT. Welsh et al. (2008) demonstrated that exposure of pregnant rats to the antiandrogen flutamide (100 mg/kg/day) during various programming windows (E15.5 to E21.5) produced dose-window-dependent suppression of GT masculinization endpoints in male offspring, including failure of urethral fold fusion resulting in hypospadias, absence of os bone ossification, and reduction of phallus length to near-female dimensions. These morphological outcomes serve as functional surrogates for impaired androgen action in GT tissue during the programming window, providing in vivo evidence that AR antagonism disrupts the androgen-dependent program governing GT development, though androgen-responsive transcription levels in GT tissue was not directly measured in this study. Rider et al. (2009) demonstrated that perinatal exposure to a mixture of antiandrogens in rats produced additive suppression of androgen-responsive endpoints in male offspring, including AGD and GT morphology, consistent with additive suppression of AR transcriptional activity in GT tissues. Kelce et al. (1994) provided foundational evidence that the developmental toxicity of vinclozolin is mediated through its antiandrogenic metabolites M1 and M2, which competitively displace DHT from the AR with binding Ki values of approximately 92 µM (92,000 nM) for M1 and 9.7 µM (9,700 nM) for M2, reflecting their comparatively lower AR-binding affinity relative to pharmaceutical antiandrogens such as hydroxyflutamide (Ki approximately 175 nM). M1 and M2 also inhibit DHT-induced AR transcriptional activity in AR transactivation assays, with M2 acting at concentrations approximately 2-fold less potent than hydroxyflutamide (Wong et al., 1995). Zheng et al. (2015) show that substantial AR antagonism delivered during the prenatal “programming” window is sufficient to change GT gene expression. Operationally, they used flutamide at 120 mg/kg to pregnant mice during E12.5–E16.5 windows (e.g., E14.5–E15.5) and detected significant transcriptional changes in the GT. Elmelund et al. (2025) show that continuous exposure to 3 or 6 mg/kg/day prenatal flutamide, from GD7 to GD17/19/21, changed Ar and Esr1 expression (bulk RT‑qPCR and spatial RNAscope), with region‑specific increases/decreases depending on gestational day. Clinical evidence: Males with complete androgen insensitivity syndrome (CAIS) carry loss-of-function mutations in the AR gene; despite normal or elevated circulating androgens, the AR is unable to drive transcription, and these individuals develop female-appearing external genitalia despite XY karyotype (Hughes et al., 2012; Gottlieb et al., 2012). This extreme phenotype provides the most compelling human evidence that loss of AR transcriptional activity in GT-equivalent tissues (genital folds, GT precursor) abolishes masculinization entirely. Partial AIS (PAIS) cases, in which AR function is partially retained, demonstrate a spectrum of GT phenotypes correlating with residual AR transcriptional activity, further supporting a quantitative relationship between AR transcriptional output and GT masculinization (Quigley et al. 1995; Hughes et al. 2012). Note: Some evidence in this section rely on in vivo studies where GT transcriptional activity is inferred from downstream morphological and gene expression endpoints rather than directly measured via ARE-reporter assays in intact fetal GT tissue.  The directional inference is well supported but direct transcriptomic quantification of AR target genes in GT tissue from antiandrogen-treated fetuses remains a data gap. The biological plausibility component is well supported by structural and mechanistic studies.

Several uncertainties and gaps should be noted: <ol> <li>Direct measurement of AR transcriptional activity specifically within GT mesenchymal cells in intact fetal tissue, following pharmacological AR antagonism at defined doses, is technically challenging. Most in vivo evidence relies on surrogate endpoints (AGD, GT morphology, anogenital index) or on measurement of downstream gene expression rather than a direct ARE-reporter readout within GT tissue. Transcriptomic profiling of GT tissue from antiandrogen-treated fetuses is limited in the literature, representing a data gap.</li> <li>The distinction between AR antagonism as the sole mechanism and combined effects on androgen biosynthesis (e.g., some phthalates reduce testosterone production as a primary mechanism, with AR antagonism being secondary or absent) can complicate attribution of reduced GT transcriptional activity specifically to AR antagonism versus ligand depletion.</li> <li>Species differences in AR LBD amino acid sequence may affect binding affinity of specific antiandrogens, introducing uncertainty in cross-species extrapolation of specific chemical potencies, even though the overall mechanism is conserved.</li> <li>AR coregulator expression profiles in GT mesenchyme during the critical window are not fully characterized; variation in coregulator availability could modulate the transcriptional response to a given degree of AR occupancy by antagonist, representing a potential source of inter-individual or inter-strain variability.</li> </ol>

<div> <table class="table table-bordered table-fullwidth"> <thead> <tr> <th>Modulating Factor (MF)</th> <th>MF Specification</th> <th>Effect(s) on the KER</th> <th>Reference(s)</th> </tr> </thead> <tbody> <tr> <td>5-alpha-reductase 2 (SRD5A2) activity</td> <td>SRD5A2 converts testosterone to DHT in GT mesenchyme</td> <td>Higher SRD5A2 activity increases local DHT, requiring higher antagonist concentration to achieve equivalent AR occupancy and transcriptional suppression; SRD5A2 deficiency reduces DHT and sensitizes AR to inhibition at lower androgen-to-antagonist ratios</td> <td>Deslypere et al. (1992); Kim et al. (2002); Blaschko et al. (2012)</td> </tr> <tr> <td>Gestational timing of exposure</td> <td>The male programming window (MPW) in rats is approximately E15.5 to E18.5; earlier or later exposure has reduced impact on GT transcriptional response as qualified by phenotypic and protein level evidence</td> <td>Antiandrogen timing, whether during windows of increased or decreased AR-responsiveness  have consequences for GT gene expression</td> <td>Welsh et al. (2008); van den Driesche et al. (2012); Seifert et al. (2012)</td> </tr> <tr> <td>Dose/concentration of antagonist</td> <td>Competitive antagonism is dose-dependent; the magnitude of AR transcriptional suppression depends on the ratio of antagonist to agonist concentrations at the receptor</td> <td>Higher antagonist-to-androgen ratios produce greater suppression of AR transcriptional activity; at subthreshold doses effects may be absent or minimal</td> <td>Kelce et al. (1997); Gray et al. (2022); Rider et al. (2009)</td> </tr> <tr> <td>AR coactivator availability</td> <td>Higher coactivator availability generally increases AR transcriptional output; lower availability decreases it. This can be assessed by coactivator expression or recruitment measures. Effects are cell‑ and promoter‑context dependent.</td> <td>Coactivator abundance modulates the magnitude of transcriptional decrease elicited by AR antagonists. Higher coactivator levels can, in some contexts, permit greater residual AR-driven transcription in the presence of antagonists, whereas lower levels tend to enhance suppression</td> <td>Shang et al. (2002); Heinlein & Chang (2002)</td> </tr> <tr> <td>Combined/mixture antiandrogen exposure</td> <td>Co‑exposure to multiple chemicals that reduce androgen signaling produces cumulative (approximately additive) suppression of androgen‑dependent endpoints </td> <td>Mixture exposures can suppress AR transcription at individual chemical concentrations that would be sub-effective alone</td> <td>Rider et al. (2009)</td> </tr> <tr> <td>Species / AR sequence variation</td> <td>Differences in AR amino‑acid sequence, especially within the ligand‑binding domain LBD, can alter ligand binding and efficacy for specific antagonists.</td> <td>Species/AR sequence variation modulates the magnitude of the decrease in AR‑dependent transcription elicited by AR antagonists at a given exposure. For a given ligand, lower AR affinity in one species (or variant) can attenuate suppression; higher affinity can amplify it.</td> <td>Kelce et al. (1997); Bohl et al. (2005); Hosokawa et al. (1993)</td> </tr> </tbody> </table> </div>

Cell models (hours scale): In HepARE-Luc cells, agonist-driven AR reporter activity begins to rise by ~6 h and is routinely quantified at 24 h; flutamide co-exposure reduces DHT-driven transcription over this 24 h window (e.g., ~50% at 1 μM; ~85% at 5 μM with 0.3 nM DHT) (Agrawal et al., 2022). Likewise, the AR-CALUX assay defines antagonism from 24 h concentration–response data with DHT held at its EC50 (~1 nM), again emphasizing an hours-to-24 h detection window for transcriptional inhibition (van Tongeren et al., 2022). Developing genital tubercle (days scale): In mice, prenatal flutamide during a discrete programming window (within E13.5–E16.5) alters expression of multiple GT genes (22 of 88 assayed at E15.5), including marked down-regulation of Indian Hedgehog (Ihh); these measurements are made over gestational days rather than minutes or hours (Zheng et al., 2015). In rats, continuous in utero exposure to low-dose flutamide (3–6 mg/kg/day from GD7) produces prenatal antiandrogenic phenotypes by GD21 and region-specific changes in Ar/Esr1 expression in the GT, again mapping effects across gestational days (Elmelund et al., 2025). The broader developmental context for timing sensitivity (the “programming window”) is established in rats using windowed in utero flutamide exposures, with masculinization endpoints affected when exposure occurs in early/mid windows but not late, underscoring day- rather than hour-scale sensitivity (Welsh et al., 2008).

Systemic level: Pure AR antagonists reduce androgenic negative feedback at the hypothalamo–pituitary level, provoking rises in LH and testicular androgens that can counteract peripheral antagonism; this limitation is overcome by combining with an LHRH agonist to block gonadotropin secretion (Labrie, 1984; Séguin et al., 1981; Simard et al., 1986) . Mechanistically, antiandrogens prevent androgen-stabilized AR from activating target genes by blocking the ligand-induced conformational changes needed for transcriptional activation (Kelce & Wilson, 1997). Together, these points support the inference that HPG-axis compensation (higher testosterone/DHT) could partially offset AR antagonism in genital tubercle if antagonist exposure is insufficient relative to the increased androgen drive. Local GT level: At the level of the developing genital tubercle, defined negative/positive feedback circuits that directly regulate AR’s own transcriptional output have not been delineated; rather, available evidence points to pathway cross-talk and relay downstream of AR. In mice, prenatal flutamide during the critical programming window alters expression of multiple GT transcripts in Hedgehog, FGF, Wnt, and BMP pathways and markedly reduces Ihh; conditional deletion of Ihh demasculinizes the penis, indicating a mesenchymal–epithelial relay of androgen signals rather than a mapped AR-centric feedback loop (Zheng et al., 2015). In rats, continuous prenatal flutamide at 3–6 mg/kg/day produces region-specific changes in Ar and Esr1 expression within the GT alongside prenatal hypospadias, consistent with local steroid-signaling interplay but without defining a closed feedback or feedforward circuit controlling AR transcription (Elmelund et al., 2025). These transcriptional and spatial effects occur within defined gestational windows when androgen signaling programs masculinization, reinforcing the importance of timing but not yet establishing an intra-GT feedback architecture for AR transcription (Welsh et al., 2008; Zheng et al., 2015).

High Male Low Female

Moderate

Development

High

Fetal to Parturition

High

Foetal

High

Embryo

High High High High

Taxonomic Applicability This is most empirically supported in rodents (rat, mouse), for which direct in vivo developmental studies with AR antagonists exist (Welsh et al., 2008; Rider et al., 2009; Zheng et al., 2015; Matsushita et al., 2018; Hashimoto et al., 2019; Rodriguez et al., 2012). It is biologically plausible across all therian mammals given the high conservation of AR structure and function, the conserved role of androgens in GT/phallus masculinization, and the clinical evidence from humans with androgen insensitivity (Hughes et al., 2012; Gottlieb et al., 2012). Lifestage Applicability In rats the masculinization programming window occurs during E15.5–E19.5; only antiandrogen exposure within this fetal window induces hypospadias/cryptorchidism and reduces penile length in males, and AGD reductions map to this same period (Welsh et al., 2008).  In mice, a prenatal AR-dependent window (approximately E13.5–E16.5) governs urethral tube closure and stromal patterning; a subsequent neonatal window controls growth of the glans via AR/ERα balance, such that either AR disruption or ERα activation can cause micropenis (Zheng et al., 2015). Mouse reviews and summaries place male-specific GT patterning and urethral canalization around E16.5 (Matsushita et al., 2018; Hashimoto et al., 2019). Although many penile features in mice differentiate postnatally, specification of penile identity is prenatal and androgen-dependent (Rodriguez et al., 2012).  The KER is not applicable to postnatal or adult life stages in the context of GT masculinization, as the GT has already differentiated by parturition. Sex Applicability Male (empirically supported and of regulatory significance for GT masculinization). Early-stage embryonic applicability to bipotential GT in both sexes is biologically plausible but less studied and of less regulatory significance.

Agrawal, H., Thakur, K., Mitra, S., Mitra, D., Keswani, C., Sircar, D., ... & Roy, P. (2022). Evaluation of (Anti) androgenic Activities of Environmental Xenobiotics in Milk Using a Human Liver Cell Line and Androgen Receptor-Based Promoter-Reporter Assay. ACS omega, 7(45), 41531-41547. Blaschko, S. D., Cunha, G. R., & Baskin, L. S. (2012). Molecular mechanisms of external genitalia development. Differentiation, 84(3), 261-268. <a href="https://doi.org/10.1016/j.diff.2012.06.003">https://doi.org/10.1016/j.diff.2012.06.003</a> Bohl, C. E., Gao, W., Miller, D. D., Bell, C. E., & Dalton, J. T. (2005). Structural basis for antagonism and resistance of bicalutamide in prostate cancer. Proceedings of the National Academy of Sciences, 102(17), 6201-6206. <a href="https://doi.org/10.1073/pnas.0500381102">https://doi.org/10.1073/pnas.0500381102</a> Brinkmann, A. O., Blok, L. J., de Ruiter, P. E., Doesburg, P., Steketee, K., Berrevoets, C. A., & Trapman, J. (1999). Mechanisms of androgen receptor activation and function. Journal of Steroid Biochemistry and Molecular Biology, 69(1-6), 307-313. <a href="https://doi.org/10.1016/s0960-0760(99)00049-7">https://doi.org/10.1016/s0960-0760(99)00049-7</a> Deslypere, J. P., Young, M., Wilson, J. D., & McPhaul, M. J. (1992). Testosterone and 5 alpha-dihydrotestosterone interact differently with the androgen receptor to enhance transcription of the MMTV-CAT reporter gene. Molecular and Cellular Endocrinology, 88(1-3), 15-22. <a href="https://doi.org/10.1016/0303-7207(92)90004-p">https://doi.org/10.1016/0303-7207(92)90004-p</a> Elmelund, E., Draskau, M. K., Berg, M., Strand, I. W., Black, J. R., Axelstad, M., ... & Svingen, T. (2025). Androgen receptor antagonist flutamide modulates estrogen receptor alpha expression in distinct regions of the hypospadiac rat penis. Frontiers in Endocrinology, 16, 1654965. Gottlieb, B., Beitel, L. K., Nadarajah, A., Paliouras, M., & Trifiro, M. (2012). The androgen receptor gene mutations database: 2012 update. Human Mutation, 33(5), 887-894. <a href="https://doi.org/10.1002/humu.22046">https://doi.org/10.1002/humu.22046</a> Gray, L. E., Jr., Furr, J., Lambright, C. S., Sampson, H., Hannas, B. R., & Wilson, V. S. (2022). Quantification of the uncertainties in extrapolating from in vitro androgen receptor antagonism to in vivo Hershberger assay endpoints and adverse reproductive development in male rats. Toxicological Sciences, Volume 176, Issue 2, August 2020, Pages 297–311, <a href="https://doi.org/10.1093/toxsci/kfaa067">https://doi.org/10.1093/toxsci/kfaa067</a> Haraguchi, R., Suzuki, K., Murakami, R., Sakai, M., Kamikawa, M., Kengaku, M., Sekine, K., Kawano, H., Kato, S., Ueno, N., & Yamada, G. (2000). Molecular analysis of external genitalia formation: the role of fibroblast growth factor (Fgf) genes during genital tubercle formation. Development, 127(11), 2471-2479. <a href="https://doi.org/10.1242/dev.127.11.2471">https://doi.org/10.1242/dev.127.11.2471</a> Hashimoto, D., Hyuga, T., Acebedo, A. R., Alcantara, M. C., Suzuki, K., & Yamada, G. (2019). Developmental mutant mouse models for external genitalia formation. Congenital Anomalies, 59, 74–80. <a class="inline-flex" href="https://doi.org/10.1111/cga.12319" style="box-sizing: border-box; border: 0px solid rgb(229, 231, 235); --tw-border-spacing-x: 0; --tw-border-spacing-y: 0; --tw-translate-x: 0; --tw-translate-y: 0; --tw-rotate: 0; --tw-skew-x: 0; --tw-skew-y: 0; --tw-scale-x: 1; --tw-scale-y: 1; --tw-pan-x: ; --tw-pan-y: ; --tw-pinch-zoom: ; --tw-scroll-snap-strictness: proximity; --tw-gradient-from-position: ; --tw-gradient-via-position: ; --tw-gradient-to-position: ; --tw-ordinal: ; --tw-slashed-zero: ; --tw-numeric-figure: ; --tw-numeric-spacing: ; --tw-numeric-fraction: ; --tw-ring-inset: ; --tw-ring-offset-width: 0px; --tw-ring-offset-color: #fff; --tw-ring-color: rgba(59,130,246,.5); --tw-ring-offset-shadow: 0 0 #0000; --tw-ring-shadow: 0 0 #0000; --tw-shadow: 0 0 #0000; --tw-shadow-colored: 0 0 #0000; --tw-blur: ; --tw-brightness: ; --tw-contrast: ; --tw-grayscale: ; --tw-hue-rotate: ; --tw-invert: ; --tw-saturate: ; --tw-sepia: ; --tw-drop-shadow: ; --tw-backdrop-blur: ; --tw-backdrop-brightness: ; --tw-backdrop-contrast: ; --tw-backdrop-grayscale: ; --tw-backdrop-hue-rotate: ; --tw-backdrop-invert: ; --tw-backdrop-opacity: ; --tw-backdrop-saturate: ; --tw-backdrop-sepia: ; --tw-contain-size: ; --tw-contain-layout: ; --tw-contain-paint: ; --tw-contain-style: ; color: rgb(17, 24, 39); text-decoration: underline; font-weight: 500; display: inline-flex;" target="_blank">https://doi.org/10.1111/cga.12319</a> Heinlein, C. A., & Chang, C. (2002). Androgen receptor (AR) coregulators: an overview. Endocrine Reviews, 23(2), 175-200. <a href="https://doi.org/10.1210/edrv.23.2.0460">https://doi.org/10.1210/edrv.23.2.0460</a> Hosokawa S, Murakami M, Ineyama M, Yamada T, Yoshitake A, Yamada H, Miyamoto J (1993) The affinity of procymidone to androgen receptor in rats and mice. J Toxicol Sci 18:83–93 Hughes, I. A., Davies, J. D., Bunch, T. I., Pasterski, V., Mastroyannopoulou, K., & MacDougall, J. (2012). Androgen insensitivity syndrome. The Lancet, 380(9851), 1419-1428. <a href="https://doi.org/10.1016/S0140-6736(12)60071-3">https://doi.org/10.1016/S0140-6736(12)60071-3</a> Kelce, W. R., Monosson, E., Gamcsik, M. P., Laws, S. C., & Gray, L. E. Jr. (1994). Environmental hormone disruptors: evidence that vinclozolin developmental toxicity is mediated by antiandrogenic metabolites. Toxicology and Applied Pharmacology, 126(2), 276-285. <a href="https://doi.org/10.1006/taap.1994.1117">https://doi.org/10.1006/taap.1994.1117</a> Kelce, W. R., & Wilson, E. M. (1997). Environmental antiandrogens: developmental effects, molecular mechanisms, and clinical implications. Journal of Molecular Medicine, 75(3), 198-207. <a href="https://doi.org/10.1007/s001090050104">https://doi.org/10.1007/s001090050104</a> Kim, K. S., Liu, W., Cunha, G. R., Russell, D. W., Huang, H., Shapiro, E., & Baskin, L. S. (2002). Expression of the androgen receptor and 5 alpha-reductase type 2 in the developing human fetal penis and urethra. Cell and Tissue Research, 307(2), 145-153. <a href="https://doi.org/10.1007/s004410100464">https://doi.org/10.1007/s004410100464</a> Labrie, F. (1984). A new approach in the hormonal treatment of prostate cancer: complete instead of partial blockade of androgens. International journal of andrology, 7(1), 1-4. Masiello, D., Cheng, S., Bubley, G. J., Lu, M. L., & Balk, S. P. (2002). Bicalutamide functions as an androgen receptor antagonist by assembly of a transcriptionally inactive receptor. Journal of Biological Chemistry, 277(29), 26321-26326. <a href="https://doi.org/10.1074/jbc.M203310200">https://doi.org/10.1074/jbc.M203310200</a> Matsushita, S., Suzuki, K., Murashima, A., et al. (2018). Regulation of masculinization: Androgen signalling for external genitalia development. Nature Reviews Urology, 15, 358–368. <a class="inline-flex" href="https://doi.org/10.1038/s41585-018-0008-y" style="box-sizing: border-box; border: 0px solid rgb(229, 231, 235); --tw-border-spacing-x: 0; --tw-border-spacing-y: 0; --tw-translate-x: 0; --tw-translate-y: 0; --tw-rotate: 0; --tw-skew-x: 0; --tw-skew-y: 0; --tw-scale-x: 1; --tw-scale-y: 1; --tw-pan-x: ; --tw-pan-y: ; --tw-pinch-zoom: ; --tw-scroll-snap-strictness: proximity; --tw-gradient-from-position: ; --tw-gradient-via-position: ; --tw-gradient-to-position: ; --tw-ordinal: ; --tw-slashed-zero: ; --tw-numeric-figure: ; --tw-numeric-spacing: ; --tw-numeric-fraction: ; --tw-ring-inset: ; --tw-ring-offset-width: 0px; --tw-ring-offset-color: #fff; --tw-ring-color: rgba(59,130,246,.5); --tw-ring-offset-shadow: 0 0 #0000; --tw-ring-shadow: 0 0 #0000; --tw-shadow: 0 0 #0000; --tw-shadow-colored: 0 0 #0000; --tw-blur: ; --tw-brightness: ; --tw-contrast: ; --tw-grayscale: ; --tw-hue-rotate: ; --tw-invert: ; --tw-saturate: ; --tw-sepia: ; --tw-drop-shadow: ; --tw-backdrop-blur: ; --tw-backdrop-brightness: ; --tw-backdrop-contrast: ; --tw-backdrop-grayscale: ; --tw-backdrop-hue-rotate: ; --tw-backdrop-invert: ; --tw-backdrop-opacity: ; --tw-backdrop-saturate: ; --tw-backdrop-sepia: ; --tw-contain-size: ; --tw-contain-layout: ; --tw-contain-paint: ; --tw-contain-style: ; color: rgb(17, 24, 39); text-decoration: underline; font-weight: 500; display: inline-flex;" target="_blank">https://doi.org/10.1038/s41585-018-0008-y</a> Petiot, A., Perriton, C. L., Dickson, C., & Cohn, M. J. (2005). Development of the mammalian urethra is controlled by Fgfr2-IIIb. Development, 132(10), 2441-2450. <a href="https://doi.org/10.1242/dev.01778">https://doi.org/10.1242/dev.01778</a> Quigley, C. A., De Bellis, A., Marschke, K. B., el-Awady, M. K., Wilson, E. M., & French, F. S. (1995). Androgen receptor defects: historical, clinical, and molecular perspectives. Endocrine Reviews, 16(3), 271-321. <a href="https://doi.org/10.1210/edrv-16-3-271">https://doi.org/10.1210/edrv-16-3-271</a> Rider, C. V., Wilson, V. S., Howdeshell, K. L., Hotchkiss, A. K., Furr, J. R., Lambright, C. R., & Gray, L. E. Jr. (2009). Cumulative effects of in utero administration of mixtures of "antiandrogens" on male rat reproductive development. Toxicologic Pathology, 37(1), 100-113. <a href="https://doi.org/10.1177/0192623308329478">https://doi.org/10.1177/0192623308329478</a> Rodriguez, E., Weiss, D. A., Ferretti, M., Wang, H., Menshenina, J., Risbridger, G., Handelsman, D., Cunha, G., & Baskin, L. (2012). Specific morphogenetic events in mouse external genitalia sex differentiation are responsive/dependent upon androgens and/or estrogens. Differentiation, 84(3), 269–279. <a class="inline-flex" href="https://doi.org/10.1016/j.diff.2012.07.003" style="box-sizing: border-box; border: 0px solid rgb(229, 231, 235); --tw-border-spacing-x: 0; --tw-border-spacing-y: 0; --tw-translate-x: 0; --tw-translate-y: 0; --tw-rotate: 0; --tw-skew-x: 0; --tw-skew-y: 0; --tw-scale-x: 1; --tw-scale-y: 1; --tw-pan-x: ; --tw-pan-y: ; --tw-pinch-zoom: ; --tw-scroll-snap-strictness: proximity; --tw-gradient-from-position: ; --tw-gradient-via-position: ; --tw-gradient-to-position: ; --tw-ordinal: ; --tw-slashed-zero: ; --tw-numeric-figure: ; --tw-numeric-spacing: ; --tw-numeric-fraction: ; --tw-ring-inset: ; --tw-ring-offset-width: 0px; --tw-ring-offset-color: #fff; --tw-ring-color: rgba(59,130,246,.5); --tw-ring-offset-shadow: 0 0 #0000; --tw-ring-shadow: 0 0 #0000; --tw-shadow: 0 0 #0000; --tw-shadow-colored: 0 0 #0000; --tw-blur: ; --tw-brightness: ; --tw-contrast: ; --tw-grayscale: ; --tw-hue-rotate: ; --tw-invert: ; --tw-saturate: ; --tw-sepia: ; --tw-drop-shadow: ; --tw-backdrop-blur: ; --tw-backdrop-brightness: ; --tw-backdrop-contrast: ; --tw-backdrop-grayscale: ; --tw-backdrop-hue-rotate: ; --tw-backdrop-invert: ; --tw-backdrop-opacity: ; --tw-backdrop-saturate: ; --tw-backdrop-sepia: ; --tw-contain-size: ; --tw-contain-layout: ; --tw-contain-paint: ; --tw-contain-style: ; color: rgb(17, 24, 39); text-decoration: underline; font-weight: 500; display: inline-flex; font-family: __Inter_f367f3, __Inter_Fallback_f367f3; font-size: 16px; font-style: normal; font-variant-ligatures: normal; font-variant-caps: normal; letter-spacing: normal; orphans: 2; text-align: start; text-indent: 0px; text-transform: none; widows: 2; word-spacing: 0px; -webkit-text-stroke-width: 0px; white-space: normal; background-color: rgb(249, 250, 251);" target="_blank">https://doi.org/10.1016/j.diff.2012.07.003</a> Séguin, C., Cusan, L., Belanger, A., Kelly, P. A., Labrie, F., & Raynaud, J. P. (1981). Additive inhibitory effects of treatment with an LHRH agonist and an antiandrogen on androgen-dependent issues in the rat. Molecular and Cellular Endocrinology, 21(1), 37-41. Seifert, A. W., Bouldin, C. M., Choi, K. S., Harfe, B. D., & Bhatt, D. L. (2008). Multiphasic and tissue-specific roles of sonic hedgehog in cloacal septation and external genitalia development. Development, 135(23), 3777-3787. Shang, Y., Myers, M., & Brown, M. (2002). Formation of the androgen receptor transcription complex. Molecular Cell, 9(3), 601-610. <a href="https://doi.org/10.1016/s1097-2765(02)00471-9">https://doi.org/10.1016/s1097-2765(02)00471-9</a> van den Driesche, S., Walker, M., McKinnell, C., Scott, H. M., Eddie, S. L., Mitchell, R. T., Seckl, J. R., Drake, A. J., Smith, L. B., Anderson, R. A., & Sharpe, R. M. (2012). Proposed role for COUP-TFII in regulating fetal Leydig cell steroidogenesis, perturbation of which leads to masculinization disorders in rodents. PLoS ONE, 7(5), e37064. <a href="https://doi.org/10.1371/journal.pone.0037064">https://doi.org/10.1371/journal.pone.0037064</a> van Tongeren, T. C., Carmichael, P. L., Rietjens, I. M., & Li, H. (2022). Next generation risk assessment of the anti-androgen flutamide including the contribution of its active metabolite hydroxyflutamide. Frontiers in Toxicology, 4, 881235. Welsh, M., Saunders, P. T., Fisken, M., Scott, H. M., Hutchison, G. R., Smith, L. B., & Sharpe, R. M. (2008). Identification in rats of a programming window for reproductive tract masculinization, disruption of which leads to hypospadias and cryptorchidism. Journal of Clinical Investigation, 118(4), 1479-1490. <a href="https://doi.org/10.1172/JCI34241">https://doi.org/10.1172/JCI34241</a> Wilson, V. S., Lambright, C., Furr, J., Ostby, J., Wood, C., Humphrey, S., & Gray, L. E. Jr. (2004). Phthalate ester-induced gubernacular lesions are associated with reduced insl3 gene expression in the fetal rat testis. Toxicology Letters, 146(3), 207-215. <a href="https://doi.org/10.1016/j.toxlet.2003.09.012">https://doi.org/10.1016/j.toxlet.2003.09.012</a> Wong, C., Kelce, W. R., Sar, M., & Wilson, E. M. (1995). Androgen receptor antagonist versus agonist activities of the fungicide vinclozolin relative to hydroxyflutamide. Journal of Biological Chemistry, 270(34), 19998-20003. <a href="https://doi.org/10.1074/jbc.270.34.19998">https://doi.org/10.1074/jbc.270.34.19998</a> Zheng, Z., Armfield, B. A., & Cohn, M. J. (2015). Timing of androgen receptor disruption and estrogen exposure underlies a spectrum of congenital penile anomalies. Proceedings of the National Academy of Sciences, 112(52), E7194-E7203. AOPWiki 2025-12-16T14:54:43

2026-04-08T13:13:56

<upstream-id>51864830-ea4f-4d13-b4e2-8bfa9e05dcb1</upstream-id> <downstream-id>9ab14690-ce1b-4c6d-8c50-5a53040c33fa</downstream-id> This KER captures an AR‑dependent regulatory relationship in the genital tubercle in which reduced AR transcriptional activity diminishes Fgfr2‑IIIb and Fgf10 signalling in genital tissues. AR functions as a ligand‑activated nuclear transcription factor. Upon binding 5α‑dihydrotestosterone (DHT), AR undergoes a ligand‑induced conformational change that stabilizes the receptor and supports recruitment of transcriptional co‑activators (Furutani et al., 2002). In GT organ culture, AR antagonism with flutamide down‑regulates Fgfr2‑IIIb transcripts, and co‑addition of DHT rescues Fgfr2‑IIIb expression, demonstrating AR‑dependent control of Fgfr2‑IIIb in this tissue (Petiot et al., 2005). An in silico stereotypic androgen response element (ARE) is present in the Fgfr2 promoter, suggesting possible direct transcriptional regulation of Fgfr2 by AR in this tissue (Petiot et al., 2005). AR antagonism also reduces Fgf10 transcripts in GT organ culture in a dose‑dependent fashion, although direct versus indirect regulation of Fgf10 by AR remains unresolved (Petiot et al., 2005). FGF10, produced by GT mesenchyme, is the primary ligand for the FGFR2‑IIIb isoform in the adjacent urethral epithelium and surface ectoderm; global loss of Fgf10 or Fgfr2‑IIIb and tissue‑specific deletion of Fgfr2 demonstrate that this axis drives urethral epithelial proliferation, maturation/stratification, tubulogenesis, and prepuce morphogenesis, and that its disruption causes hypospadias (Petiot et al., 2005; Gredler et al., 2015). When AR activity is reduced by pharmacological antagonism in GT organ culture, Fgfr2‑IIIb and Fgf10 mRNA are downregulated in a dose‑dependent manner, and Fgfr2‑IIIb downregulation is rescued by DHT (Petiot et al., 2005). This reduction in FGF10/FGFR2‑IIIb signaling is sufficient to arrest epithelial progenitor cell proliferation and disrupt stratification and maturation of the urethral plate epithelium, ultimately impairing urethral morphogenesis (Petiot et al., 2005; Gredler et al., 2015).

Evidence for this KER was assembled through a combination of expert knowledge and AI-assisted literature search and synthesis. Specifically, a combination of Claude (Anthropic) using Sonnet 4.6, and EPA AI, using GPT5, was used to identify, retrieve, and summarize relevant primary and secondary literature, and to draft the initial content of this KER page. All citations generated through this process were subsequently reviewed and verified by the KER author against primary sources and DOI resolution checks, prior to inclusion. Users of this KER are advised that AI-assisted evidence assembly may introduce selection bias or gaps in coverage that differ from a fully systematic human-conducted review, and independent verification of the evidence base is encouraged.  A copy of the initial prompt is attached to AOP619. A review of primary experimental literature using the following databases and search strategies was employed: PubMed and Google Scholar using the following search terms and combinations: "FGF10 FGFR2 androgen receptor genital tubercle," "FGFR2-IIIb androgen response element," "flutamide FGF10 FGFR2 hypospadias," "AR transcriptional activity external genitalia FGF signaling," "FGF10 FGFR2 urethral development masculinization," and "hypospadias FGF8 FGF10 FGFR2 androgen." No formal date restriction was applied. Priority was given to primary experimental studies in rodent genetic and pharmacological models, followed by human genetic and histological studies. Knockout mouse studies, ex vivo culture experiments, in situ hybridization studies, and transcriptome analyses were included. Review articles were used to corroborate mechanistic frameworks.

AR is a member of the nuclear steroid hormone receptor superfamily and functions as a ligand‑inducible transcription factor. Upon binding DHT, the AR-DHT complex undergoes an agonist‑induced conformational change, dimerizes, translocates to the nucleus, and binds androgen response elements (AREs) composed of two palindromic half‑sites separated by a three‑nucleotide spacer to drive transcription (Gelmann, 2002; Quigley et al., 1995). In developing external genital tissues, testosterone is locally converted to DHT by 5α‑reductase type 2; in the human fetal penis, SRD5A2 is strongly expressed in the stroma along the ventral urethral seam while AR is enriched in the urethral epithelium (Kim et al., 2002). Loss of AR function (e.g., complete androgen insensitivity) leads to female‑appearing external genitalia, and 5α‑reductase type 2 deficiency causes undervirilization and hypospadias, establishing that DHT-AR‑dependent transcription is required for normal external genital development (Quigley et al., 1995; Kim et al., 2002). Within the genital tubercle (GT), AR antagonism (flutamide) down‑regulates Fgfr2‑IIIb transcripts in a dose‑dependent manner with rescue by DHT, and reduces Fgf10 transcripts at higher antagonist doses, demonstrating AR‑dependent control of the FGF10-FGFR2‑IIIb axis during the developmental period when these genes are expressed (Petiot et al., 2005). The Fgfr2 promoter contains a stereotypic ARE‑like motif identified in silico between nucleotides 1193-1198 within the region 1041-1610 upstream of the transcription start site, providing a plausible direct molecular link between AR transcriptional activity and Fgfr2‑IIIb expression in urethral epithelium; however, GT‑specific AR-DNA binding or reporter evidence has not yet been shown, so direct regulation remains putative (Petiot et al., 2005). Whether AR regulation of Fgf10 in the GT is direct or involves intermediate signals remains unresolved; by analogy, in the prostate FGF10 is a mesenchymal growth cue required for epithelial proliferation and budding, but neonatal prostate data indicate FGF10 is not directly regulated by testosterone (Thomson & Cunha, 1999; Donjacour et al., 2003). FGF10, produced by GT mesenchyme, acts as a major ligand for the epithelial FGFR2‑IIIb isoform, forming a locally acting epithelial-mesenchymal signaling axis (Ohuchi et al., 2000). Genetic loss‑of‑function and tissue‑specific deletion studies show that reduced FGF10/FGFR2‑IIIb signaling is sufficient to arrest proliferation of urethral epithelial progenitors, disrupt epithelial stratification and maturation, and produce hypospadias; endodermal FGFR2 is required for urethral epithelial maturation and ectodermal FGFR2 for ventral prepuce closure and maintenance of a closed urethral tube (Petiot et al., 2005; Gredler et al., 2015). At the cellular level, FGFR2 signaling in GT epithelia couples G1/S progression with columnar morphogenesis and cell‑adhesion organization, linking the FGF axis to proliferation‑coupled epithelial maturation required for urethral tubulogenesis (Gredler et al., 2015). Taken together, these findings support the biological plausibility of a causal chain in which reduced AR transcriptional activity during the developmental window diminishes FGF10–FGFR2‑IIIb signaling in GT tissues, thereby compromising urethral epithelial progenitor maintenance and stratification required for urethral tubulogenesis and disrupting ectodermal FGFR2‑dependent ventral prepuce closure and external preputial lamina formation—prenatal defects that plausibly predispose to persistent ventral tethering and abnormal postnatal preputial separation (Petiot et al., 2005; Gredler et al., 2015; Harada et al., 2015).

Pharmacological AR antagonism in ex vivo GT organ culture (mouse). Petiot et al. (2005) showed that flutamide produced a clear, dose‑dependent decrease in Fgfr2‑IIIb transcripts in the urethral plate epithelium of cultured GTs, with no change at 10-5 M, progressively reduced at 10⁻⁴ M and 5×10⁻⁴ M, and markedly diminished/undetectable at 10⁻³ M; importantly, co‑addition of DHT rescued Fgfr2‑IIIb expression and normalized morphology, confirming AR specificity (Petiot et al., 2005). Male and female GTs were cultured separately and showed no differences in response to flutamide (Petiot et al., 2005). AR antagonism also down‑regulates Fgf10 in GT organ culture. In the same study, flutamide reduced Fgf10 transcripts in a dose‑dependent fashion in both male and female GTs, indicating that androgen‑dependent regulation of the mesenchymal ligand is operative during this developmental window (Petiot et al., 2005). ARE in the Fgfr2 promoter (molecular plausibility for a direct transcriptional link). Petiot et al. (2005) identified a stereotypic ARE‑like motif within 1041 to 1610 bp upstream of the Fgfr2 transcription start site, consistent with direct AR‑dependent transcriptional regulation of Fgfr2‑IIIb; they noted, however, that intermediate regulatory steps cannot be excluded and direct AR-DNA binding in GT was not shown. Disruption of androgen signaling in vivo alters urethral plate cell behavior. Using lineage tracing, Seifert et al. (2008) showed that flutamide‑treated males failed to undergo urethral septation/internalization and instead retained urethral plate cells to the ventral margin, mimicking untreated females; flutamide‑treated females showed normal urethral positioning. These data confirm that androgen‑dependent signals are required for the morphogenetic cell behaviors distinguishing male from female urethral development and align with the FGFR2 endodermal deletion phenotype (Seifert et al., 2008; Gredler et al., 2015). Network context and tissue‑specific roles of FGF signaling in GT. Tissue‑specific Fgfr knockouts demonstrate that mesenchymal FGF signaling is required for early GT outgrowth, ectodermal signaling for urethral tube formation, and endodermal signaling for epithelial stratification/adhesion during later stages, placing the FGF10-FGFR2‑IIIb axis precisely within the epithelial-mesenchymal mechanics of urethral tubulogenesis (Harada et al., 2015). Computational corroboration.  A multicellular agent‑based model of GT development recapitulated urethral tube closure as an emergent, androgen‑dependent property that is quantitatively sensitive to SHH‑ and FGF10‑driven mesenchymal proliferation and to endodermal apoptosis; androgen insufficiency or delayed androgenization produced feminization or incomplete closure, respectively, providing in silico corroboration of the AR→FGF10/FGFR2‑IIIb linkage’s impact on urethrogenesis (Leung et al., 2016).

<ol> <li>Directionality of Fgf10 regulation: In GT organ culture, flutamide dose‑dependently reduces Fgf10 in both sexes, but whether AR regulates Fgf10 directly or via intermediates is unresolved; in neonatal prostate, FGF10 is not directly regulated by testosterone, supporting an indirect route (Petiot et al., 2005; Thomson & Cunha, 1999; Donjacour et al., 2003).</li> <li>Possible ARE‑independent regulation of Fgfr2: A putative AR recognition hexamer is present upstream of Fgfr2 and DHT rescues flutamide‑suppressed Fgfr2‑IIIb in GT organ culture, but GT‑specific AR-DNA binding and transactivation have not been demonstrated (no ChIP or promoter‑reporter assays), so direct regulation remains unproven (Petiot et al., 2005).</li> <li>Species extrapolation: Most mechanistic data are from mouse/rat; the timing and endocrine control of the masculinization programming window differ between rodents and humans, and human genetic/IHC data are correlative, limiting direct inference of timing/magnitude/cell‑type specificity of AR→FGF regulation in humans (Welsh et al., 2008; Sharpe, 2020; Kim et al., 2002).</li> <li>Tissue‑compartment specificity: AR is present in both GT mesenchyme and urethral epithelium (mouse), and in human fetal penis AR is enriched in urethral epithelium while 5α‑reductase type 2 is concentrated in ventral stroma at the urethral seam; whether AR‑driven Fgfr2 and/or Fgf10 transcription occurs cell‑autonomously in specific GT compartments in vivo remains to be resolved (Petiot et al., 2005; Kim et al., 2002).</li> </ol>

The precise in vivo response-response relationship between the degree of AR activity reduction and the magnitude of FGF10/FGFR2‑IIIb signaling decrease is not yet quantified; available dose-response data derive from ex vivo GT cultures and semi‑quantitative in situ hybridization rather than absolute measures of FGF signaling output (e.g., receptor phosphorylation). Petiot et al. (2005) demonstrated a clear dose-response relationship between flutamide concentration and reduction of Fgfr2‑IIIb transcripts in mouse GT organ culture after 48 h: Fgfr2‑IIIb expression was maintained at 10⁻⁵ M flutamide, progressively reduced at 10⁻⁴ M and 5×10⁻⁴ M, and markedly diminished/undetectable at 10⁻³ M; co‑treatment with DHT (5×10⁻⁶ M) rescued Fgfr2‑IIIb expression, confirming AR specificity (Petiot et al., 2005). Fgf10 transcripts persisted at 10⁻⁵ M but showed progressive down‑regulation at 10⁻⁴-10⁻³ M, including loss of detectable signal at 10⁻³ M in female GT; DHT rescue was shown for Fgfr2‑IIIb (not for Fgf10) (Petiot et al., 2005). A multicellular agent‑based model of GT development indicated that partial reductions in androgen signaling can produce graded urethral‑closure outcomes—from mild to severe hypospadias—depending on the timing and magnitude of perturbation, providing in silico corroboration of graded AR‑dependent control over FGF‑sensitive epithelial behaviors (Leung et al., 2016).

In mouse GT organ culture, AR antagonism produces detectable reductions in Fgfr2‑IIIb (and, at higher doses, Fgf10) within 48 h, and DHT rescues Fgfr2‑IIIb within that same window, indicating an hours‑to‑days timescale for AR‑dependent FGF transcript changes in GT tissue (Petiot et al., 2005; Furutani et al., 2002). Overall, the KER operates during the androgen‑dependent sexual differentiation phase of GT development. In mouse, the critical programming window for penile masculinization is approximately E14.5-E17.5, while urethral septation/internalization and preputial morphogenesis continue through late gestation to birth (P0) (Amato et al., 2022; Seifert et al., 2008). In rat, the masculinization programming window spans GD15.5-18.5; suppression of androgen action within this window produces the greatest feminization/hypospadias, with much smaller effects outside it (Welsh et al., 2008; Sharpe, 2020). By analogy, the presumptive human window is ~8-14 gestational weeks, although direct human GT mechanistic data are limited (Sharpe, 2020).

Petiot et al. investigated whether FGFR2‑IIIb regulates AR in a positive feedback loop. Immunohistochemistry at E16.5 showed that AR distribution in Fgfr2‑IIIb null embryos was indistinguishable from wild type, indicating that FGF10/FGFR2‑IIIb signaling does not regulate AR expression in GT at this stage; thus, a downstream‑to‑upstream feedback loop was not detected.  Consistently, conditional deletion of Shh at E11.5 did not alter Fgfr2 mRNA levels, arguing against a Shh→Fgfr2 feedback at E14.5 (Gredler et al., 2015). Conversely, Sonic hedgehog (SHH) signaling interacts with androgen pathways during GT masculinization. SHH is expressed in the urethral plate epithelium and signals via Gli2 in adjacent mesenchyme; Gli2 mutants show reduced expression of androgen‑responsive, sexually dimorphic genes (e.g., Mafb, Fkbp5) and fail to be masculinized by exogenous androgens despite normal testicular testosterone, indicating hedgehog signaling is required to maintain androgen responsiveness in GT mesenchyme (Miyagawa et al., 2011). In GT, SHH also lies upstream of mesenchymal Fgf10 (in contrast to limb, where FGFR2‑IIIb acts upstream of Shh), providing a route by which hedgehog signaling can reinforce the mesenchymal ligand side of the FGF10-FGFR2‑IIIb axis (Petiot et al., 2005; Revest et al., 2001). Taken together, current data support a potential feed‑forward influence wherein SHH/Gli2 enhances androgen responsiveness and promotes Fgf10 expression, indirectly supporting AR‑dependent FGF signaling in GT; direct SHH‑dependent regulation of Fgfr2 has not been observed in GT (Gredler et al., 2015).

High Male Moderate Female High Unspecific

High

Fetal

High

Foetal

High

Embryo

High

Fetal to Parturition

High High Low

Taxonomy: Empirical applicability is strongest in mice, where genetic loss‑of‑function and pharmacological perturbation studies establish that reduced AR activity diminishes the FGF10-FGFR2‑IIIb axis in GT tissues and produces urethral/preputial defects (Petiot et al., 2005; Gredler et al., 2015). In rats, most evidence comes from in vivo anti‑androgen exposure models within the masculinization programming window (MPW), which feminize urethral development and induce hypospadias, consistent with diminished AR‑dependent control of pathways governing urethrogenesis (Welsh et al., 2008; Seifert et al., 2008; Sinclair et al., 2016). In humans, evidence is indirect and correlative: AR and 5α‑reductase type 2 are present in the right fetal tissues to support local DHT production (Kim et al., 2002), FGFR2 variants have been reported in familial hypospadias (Beleza‑Meireles et al., 2007), FGF10 SNPs associate with risk in a large case-control cohort (Carmichael et al., 2013), and postnatal foreskin from hypospadiac boys shows altered FGF/FGFR2 immunostaining patterns (Haid et al., 2020). Cross‑species extrapolation is biologically plausible given conserved epithelial-mesenchymal FGF signaling in external genital development, but differences in distal urethral morphogenesis and endocrine timing across mammals should be noted (Amato et al., 2022; Sharpe, 2020).  Mammalia, given conserved androgen‑responsive epithelial-mesenchymal signaling during external genital development, with caveats on interspecies differences in distal urethral morphogenesis and endocrine control (Amato et al., 2022; Sharpe, 2020) Life Stage: This KER operates during the androgen‑dependent phase of external genital development. In mouse, the critical programming window for penile masculinization is approximately E14.5-E17.5, with urethral septation/internalization and preputial morphogenesis continuing through late gestation to birth (Seifert et al., 2008; Amato et al., 2022). In rat, the MPW is GD15.5-18.5, during which suppression of androgen action has maximal effects on urethral development (Welsh et al., 2008; Sharpe, 2020). Evidence does not support activity of this specific regulatory relationship in postnatal life; AR‑dependent FGF regulation in adult tissues (e.g., prostate) is context‑specific and not directly informative for prenatal GT (Donjacour et al., 2003). Sex: The KER has greater downstream morphogenetic consequence for males. In organ culture, both male and female GTs show dose‑dependent down‑regulation of Fgfr2‑IIIb and Fgf10 with flutamide, and DHT rescues Fgfr2‑IIIb, indicating that the AR→FGF10/FGFR2‑IIIb linkage operates in both sexes at the molecular level (Petiot et al., 2005). In vivo, AR antagonism during the programming window feminizes male urethral development, whereas females show little change in urethral position, supporting primary applicability to males with moderate applicability to females (Seifert et al., 2008).

Amato, C. M., Yao, H. H.-C., & Zhao, F. (2022). One tool for many jobs: Divergent and conserved actions of androgen signaling in male internal reproductive tract and external genitalia. Frontiers in Endocrinology, 13, 910964. <a href="https://doi.org/10.3389/fendo.2022.910964" target="_blank">https://doi.org/10.3389/fendo.2022.910964</a> (PMID: 35846302) Beleza-Meireles, A., Lundberg, F., Lagerstedt, K., Zhou, X., Omrani, D., Frisén, L., & Nordenskjöld, A. (2007). FGFR2, FGF8, FGF10 and BMP7 as candidate genes for hypospadias. European Journal of Human Genetics, 15(4), 405-410. <a href="https://doi.org/10.1038/sj.ejhg.5201777" target="_blank">https://doi.org/10.1038/sj.ejhg.5201777</a> (PMID: 17264867) Carmichael, S. L., Ma, C., Choudhry, S., Lammer, E. J., Witte, J. S., & Shaw, G. M. (2013). Hypospadias and genes related to genital tubercle and early urethral development. Journal of Urology, 190(5), 1884-1892. <a href="https://doi.org/10.1016/j.juro.2013.05.061" target="_blank">https://doi.org/10.1016/j.juro.2013.05.061</a> (PMID: 23714460; PMC: PMC4103581) Donjacour, A. A., Thomson, A. A., & Cunha, G. R. (2003). FGF-10 plays an essential role in the growth of the fetal prostate. Developmental Biology, 261(1), 39-54. <a href="https://doi.org/10.1016/S0012-1606(03)00250-1" target="_blank">https://doi.org/10.1016/S0012-1606(03)00250-1</a> (PMID: 12941620) Furutani, T., Watanabe, T., Tanimoto, K., Hashimoto, T., Koutoku, H., Kudoh, M., Shimizu, Y., Kato, S. and Shikama, H. (2002). Stabilization of androgen receptor protein is induced by agonist, not by antagonists. Biochem. Biophys. Res. Commun. 294, 779-784. Gelmann, E. P. (2002). Molecular biology of the androgen receptor. Journal of Clinical Oncology, 20(13), 3001-3015. <a href="https://doi.org/10.1200/JCO.2002.10.018" target="_blank">https://doi.org/10.1200/JCO.2002.10.018</a> (PMID: 12089231) Gredler, M. L., Seifert, A. W., & Cohn, M. J. (2015). Tissue-specific roles of Fgfr2 in development of the external genitalia. Development, 142(12), 2203-2212. <a href="https://doi.org/10.1242/dev.119891" target="_blank">https://doi.org/10.1242/dev.119891</a> (PMID: 26081573; PMC: PMC4483768) Haid, B., Pechriggl, E., Nägele, F., Dudas, J., Webersinke, G., & Rammer, M. (2020). FGF8, FGF10 and FGF receptor 2 in foreskin of children with hypospadias: An analysis of immunohistochemical expression patterns and gene transcription. Journal of Pediatric Urology, 16(1), 41.e1-41.e10. <a href="https://doi.org/10.1016/j.jpurol.2019.10.007" target="_blank">https://doi.org/10.1016/j.jpurol.2019.10.007</a> (PMID: 31676182) Harada, M., Omori, A., Nakahara, C., Nakagata, N., Akita, K., & Yamada, G. (2015). Tissue-specific roles of FGF signaling in external genitalia development. Developmental Dynamics, 244(6), 759-773. <a href="https://doi.org/10.1002/dvdy.24277">https://doi.org/10.1002/dvdy.24277</a> Kim, K. S., Liu, W., Cunha, G. R., Russell, D. W., Huang, H., Shapiro, E., & Baskin, L. S. (2002). Expression of the androgen receptor and 5 alpha-reductase type 2 in the developing human fetal penis and urethra. Cell and Tissue Research, 307(2), 145-153. Leung, M. C. K., Hutson, M. S., Seifert, A. W., Spencer, R. M., & Knudsen, T. B. (2016). Computational modeling and simulation of genital tubercle development. Reproductive Toxicology, 64, 151-161. <a href="https://doi.org/10.1016/j.reprotox.2016.05.005" target="_blank">https://doi.org/10.1016/j.reprotox.2016.05.005</a> (PMID: 27181558) Miyagawa, S., Moon, A., Haraguchi, R., Inoue, C., Harada, M., Nakahara, C., Suzuki, K., Nakagata, N., Ng, R. C., Akita, K., Yamada, G. (2011). The role of sonic hedgehog-Gli2 pathway in the masculinization of external genitalia. Endocrinology, 152(7), 2894-2903. (PMID: 21586556) Ohuchi, H., Hori, Y., Yamasaki, M., Harada, H., Sekine, K., Kato, S., & Itoh, N. (2000). FGF10 acts as a major ligand for FGF receptor 2 IIIb in mouse multi-organ development. Biochemical and Biophysical Research Communications, 277(3), 643-649. <a href="https://doi.org/10.1006/bbrc.2000.3721">https://doi.org/10.1006/bbrc.2000.3721</a> Petiot, A., Perriton, C. L., Dickson, C., & Cohn, M. J. (2005). Development of the mammalian urethra is controlled by Fgfr2-IIIb. Development, 132(10), 2441-2450. <a href="https://doi.org/10.1242/dev.01778" target="_blank">https://doi.org/10.1242/dev.01778</a> (PMID: 15843416) Quigley, C. A., De Bellis, A., Marschke, K. B., El-Awady, M. K., Wilson, E. M., & French, F. S. (1995). Androgen receptor defects: historical, clinical, and molecular perspectives. Endocrine reviews, 16(3), 271-321. Revest, J. M., Spencer-Dene, B., Kerr, K., De Moerlooze, L., Rosewell, I., & Dickson, C. (2001). Fibroblast growth factor receptor 2-IIIb acts upstream of Shh and Fgf4 and is required for limb bud maintenance but not for the induction of Fgf8, Fgf10, Msx1, or Bmp4. Developmental Biology, 231(1), 47-62. <a href="https://doi.org/10.1006/dbio.2000.0144">https://doi.org/10.1006/dbio.2000.0144</a> Seifert, A. W., Harfe, B. D., & Cohn, M. J. (2008). Cell lineage analysis demonstrates an endodermal origin of the distal urethra and perineum. Developmental Biology, 318(1), 143-152. <a href="https://doi.org/10.1016/j.ydbio.2008.03.017" target="_blank">https://doi.org/10.1016/j.ydbio.2008.03.017</a> (PMID: 18439576; PMC: PMC3047571) Seifert, A. W., Zheng, Z., Ormerod, B. K., & Cohn, M. J. (2010). Sonic hedgehog controls growth of external genitalia by regulating cell cycle kinetics. Nature Communications, 1, Article 23. <a href="https://doi.org/10.1038/ncomms1020">https://doi.org/10.1038/ncomms1020</a> Sharpe, R. M. (2020). Androgens and the masculinization programming window: Human-rodent differences. Biochemical Society Transactions, 48(4), 1725-1735. <a href="https://doi.org/10.1042/BST20200200" target="_blank">https://doi.org/10.1042/BST20200200</a> (PMID: 32830844) Sinclair, A. W., Cao, M., Pask, A., Baskin, L., & Cunha, G. R. (2017). Flutamide-induced hypospadias in rats: A critical assessment. Differentiation, 94, 37-57. <a href="https://doi.org/10.1016/j.diff.2016.12.001" target="_blank">https://doi.org/10.1016/j.diff.2016.12.001</a> (PMID: 28043016) Thomson, A. A., & Cunha, G. R. (1999). Prostatic growth and development are regulated by FGF10. Development, 126(16), 3693-3701. Welsh, M., Saunders, P. T., Fisken, M., Scott, H. M., Hutchison, G. R., Smith, L. B., & Sharpe, R. M. (2008). Identification in rats of a programming window for reproductive tract masculinization, disruption of which leads to hypospadias and cryptorchidism. The Journal of clinical investigation, 118(4), 1479-1490. AOPWiki 2025-12-16T14:54:51

2026-05-07T15:54:39

<upstream-id>9ab14690-ce1b-4c6d-8c50-5a53040c33fa</upstream-id> <downstream-id>71f737c5-2a8f-4a4b-b9d9-585e6994255b</downstream-id> This KER describes the mechanistic link between reduced paracrine FGF10–FGFR2‑IIIb signaling in the developing genital tubercle (KEupstream) and disruption of preputial epithelial morphogenesis (KEdownstream). FGF10 produced by genital mesenchyme signals to the epithelial FGFR2‑IIIb isoform in adjacent surface ectoderm and urethral endoderm, supporting epithelial progenitor proliferation, stratification/maturation, and the ventral outgrowth and fusion of the preputial swellings around the developing glans and urethral tube (Ohuchi et al., 2000; Petiot et al., 2005; Gredler et al., 2015; Harada et al., 2015). Reduction of this axis, whether by genetic loss of FGF10 or FGFR2‑IIIb, tissue‑specific deletion of FGFR2 in ectoderm or endoderm, or upstream pathway perturbations that lower ligand availability or receptor expression/activation (e.g., AR antagonism reducing FGFR2‑IIIb and FGF10 in GT organ culture; SHH signaling that positions mesenchymal FGF10 in GT; WNT/β‑catenin or ISL1 programs that modulate FGF10). impairs epithelial proliferation, stratification, and organization, resulting in failed ventral prepuce closure and hypospadias with ventral tethering phenotypes (Petiot et al., 2005; Gredler et al., 2015; Harada et al., 2015; Perriton et al., 2002; Haraguchi et al., 2000; Lin et al., 2008; Ching et al., 2018). The relationship is rooted in epithelial-mesenchymal crosstalk conserved across organs: FGF10, a principal ligand for epithelial FGFR2‑IIIb, activates this receptor and its canonical cascades (ERK/AKT/PLCγ) in epithelia, providing plausible mediators between receptor output and epithelial behaviors; in GT, reduced FGFR2‑IIIb is empirically linked to reduced proliferation and disordered epithelial organization, but cascade‑level causality remains to be shown (Ohuchi et al., 2000; Itoh & Ornitz, 2011; Gredler et al., 2015). In the genital tubercle, the spatial deployment of FGF10-FGFR2‑IIIb signaling contributes to ventral prepuce expansion and closure; ectodermal FGFR2 is specifically required for ventral fusion and for maintaining a closed urethral tube, whereas endodermal FGFR2 is required for urethral epithelial maturation (Gredler et al., 2015; Harada et al., 2015; Satoh et al., 2004; Yamada et al., 2006). Within the established GT signaling network, SHH from the urethral epithelium promotes mesenchymal FGF10, and WNT/ISL1 programs influence mesenchymal FGF10 and epithelial behaviors; thus, perturbations in these inputs can secondarily diminish FGF10–FGFR2‑IIIb signaling and disrupt preputial morphogenesis. (Perriton et al., 2002; Seifert et al., 2010; Lin et al., 2008; Ching et al., 2018; Suzuki et al., 2002).

Evidence for this KER was assembled through a combination of expert knowledge and AI-assisted literature search and synthesis. Specifically, a combination of Claude (Anthropic) using Sonnet 4.6, and EPA AI, using GPT5, was used to identify, retrieve, and summarize relevant primary and secondary literature, and to draft the initial content of this KER page. All citations generated through this process were subsequently reviewed and verified by the KER author against primary sources and DOI resolution checks, prior to inclusion. Users of this KER are advised that AI-assisted evidence assembly may introduce selection bias or gaps in coverage that differ from a fully systematic human-conducted review, and independent verification of the evidence base is encouraged.  A copy of the initial prompt is attached to AOP619. A review of primary experimental literature using the following databases and search strategies was employed: PubMed/MEDLINE and Google Scholar using the following search terms and Boolean combinations: "FGF10" AND "FGFR2" AND "prepuce" OR "preputial", "FGFR2-IIIb" AND "hypospadias" AND "genital tubercle", "FGF10 knockout" AND "external genitalia" AND "mouse", "FGFR2 conditional knockout" AND "prepuce", "FGF10 FGFR2 hypospadias human", "preputial morphogenesis" AND "fibroblast growth factor". Searches were conducted in April 2026 with no formal date restriction. Screening criteria prioritized: (1) primary experimental studies using genetic loss-of-function models (null mutants and conditional knockouts); (2) pharmacological studies (AR antagonism) that implicate this pathway; (3) human tissue studies (immunohistochemistry, sequencing) in hypospadias patients; and (4) cross-species comparative studies. Reviews were used to triangulate but not as primary evidence sources.

In the genital tubercle (GT), FGF10 is produced by genital mesenchyme and FGFR2‑IIIb is transcribed in adjacent surface ectoderm and urethral epithelium, positioning a paracrine axis across the mesenchyme–epithelium boundary where the preputial folds arise (Gredler et al., 2015; Satoh et al., 2004; Yamada et al., 2006; Suzuki et al., 2002). FGF10 is a major ligand for epithelial FGFR2‑IIIb across multiple organs, and high‑affinity signaling requires heparan‑sulfate proteoglycans as cofactors (Ohuchi et al., 2000; Ornitz, 2015). FGFR2b activates canonical cascades in epithelia (MAPK/ERK, PI3K/AKT, PLCγ), providing plausible mediators between receptor output and epithelial behaviors (Ornitz, 2015). In the genital tubercle, epithelial FGFR2 deletion prolongs G1, reduces proliferation, and disrupts columnar morphogenesis/adhesion and stratification, linking diminished receptor output to the epithelial behaviors required for preputial fold expansion and fusion; assignment of these effects to specific cascades in GT epithelium has not yet been shown (Gredler et al., 2015). These pathway outputs map directly onto the epithelial behaviors needed for preputial morphogenesis. Preputial swellings elevate from the lateral GT at ~E13–E13.5 and must expand and fuse ventrally around the glans/urethral tube between ~E15.5 and E17.5 (Satoh et al., 2004; Yamada et al., 2006; Suzuki et al., 2002). When FGF10→FGFR2‑IIIb signaling is reduced, GT epithelia exhibit diminished proliferation and impaired columnar/adhesive organization, blunting lateral‑to‑ventral fold expansion and compromising midline fusion competence; consequently, the ventral prepuce fails to close, yielding ventral tethering and a persistent ventral groove by late gestation (Gredler et al., 2015; Harada et al., 2015). Genetic sufficiency data reinforce this linkage: global loss of FGFR2‑IIIb or FGF10 arrests urethral epithelial maturation and produces glans/prepuce malformations and severe hypospadias (Petiot et al., 2005). Tissue‑specific deletions show compartmental requirements, ectodermal FGFR2 is necessary for ventral prepuce closure and for maintaining a closed urethral tube, whereas endodermal FGFR2 is required for urethral epithelial proliferation, stratification, and maturation, indicating that reduced signaling in ectoderm predominantly blocks fold fusion, while reduced signaling in endoderm weakens the epithelial template on which closure depends (Gredler et al., 2015; Harada et al., 2015). Finally, pathway context helps explain sensitivity and modifiers: SHH from the urethral epithelium promotes mesenchymal FGF10 and GT growth, and WNT/β‑catenin and ISL1 programs regulate mesenchymal FGF10 and epithelial behaviors; perturbations in these inputs can secondarily depress FGF10– FGFR2‑IIIb signaling and disrupt preputial morphogenesis (Perriton et al., 2002; Seifert et al., 2010; Lin et al., 2008; Ching et al., 2018). Because FGF10 signals via FGFR2‑IIIb and high‑affinity signaling requires heparan‑sulfate proteoglycans, disturbances in HS biosynthesis or sulfation are plausible modulators of this KER, although GT‑specific HS loss‑of‑function data were not identified (Ohuchi et al., 2000; Ornitz, 2015)

Genetic null models (mouse) FGFR2‑IIIb−/− and FGF10−/−. In FGFR2‑IIIb null embryos, early GT outgrowth is grossly normal, but by E13.5 the ventral urethral plate develops a precocious proximal opening and a furrow along the urethral seam; by E15.5–E17.5, the penile urethra fails to internalize and ventral prepuce does not form, yielding severe hypospadias. FGF10 null males and females similarly exhibit severe proximal hypospadias with arrested epithelial maturation. Concordant phenotypes in FGF10 and FGFR2‑IIIb nulls provide strong genetic evidence that the FGF10–FGFR2‑IIIb pair is essential for urethral tube internalization and ventral prepuce development (Petiot et al., 2005). Tissue-specific (conditional) knockout models (mouse) Ectoderm vs endoderm roles. Conditional deletion of FGFR2 in the surface ectoderm disrupts ventral prepuce closure and maintenance of a closed urethral tube, producing severe hypospadias and ventral tethering; mislocalization of preputial swellings is evident from the onset of prepuce development (≈E13.0) and progresses to hypospadic orifices. In contrast, endodermal FGFR2 deletion causes milder hypospadias with impaired maturation and stratification of the urethral epithelium. These data demonstrate compartment‑specific sufficiency of FGFR2 signaling for preputial epithelial morphogenesis (Gredler et al., 2015; Harada et al., 2015). Endocrine pharmacology linking upstream perturbation to the FGF axis and downstream morphology <ul> <li>Ex vivo GT organ culture: Flutamide produced a clear, dose‑dependent reduction of FGFR2‑IIIb transcripts after 48 h (10⁻⁵ M no effect; 10⁻⁴–10⁻³ M progressive loss), with co‑treatment DHT (5×10⁻⁶ M) rescuing FGFR2‑IIIb; FGF10 was also reduced at higher flutamide doses in both male and female GTs. These findings show AR‑dependent control over the FGF10–FGFR2‑IIIb axis in GT tissue (Petiot et al., 2005).</li> <li>In vivo alignment: AR antagonism during the programming window feminizes male urethral development and yields ventral tethering/failed ventral closure, consistent with the preputial defects seen with ectodermal FGFR2 loss; while in vivo FGF10/FGFR2‑IIIb transcripts were not quantified, the morphological outcomes align with reduced epithelial FGF signaling (Seifert et al., 2008; Sinclair et al., 2016). Note: Gredler’s comment on flutamide lowering FGFR2 in GT transcriptionally cites Petiot’s ex vivo data (Gredler et al., 2015; Petiot et al., 2005).</li> </ul> Cross-species comparative and organ culture manipulation (mouse ↔ guinea pig) Species differences and causal tuning by FGF/SHH level. In guinea pig (which normally maintains an open urethral groove), GT expression of Shh, FGF8, FGF10, FGFR2, and Hoxd13 is >4‑fold lower than in mouse. In organ culture, Hedgehog and FGF pathway inhibitors induced/maintained a urethral groove and restrained preputial development in mouse GT, whereas recombinant SHH or FGF10 proteins promoted preputial development in guinea pig GT. These gain‑ and loss‑of‑function manipulations support that the level of FGF10–FGFR2‑IIIb (and SHH) signaling causally influences whether preputial morphogenesis proceeds versus biasing toward an open groove configuration (Wang & Zheng, 2025). Human tissue and genetic evidence (translation; indirect) <ul> <li>Protein-level alterations in prepuce/foreskin. In hypospadiac foreskin, immunohistochemistry demonstrates altered or reduced staining for FGF10/FGFR2 relative to controls (with some studies including FGF8); some cohorts report significant protein differences without concordant mRNA changes by qPCR, suggesting post‑transcriptional regulation or altered localization (Haid et al., 2020; Emaratpardaz et al., 2024).</li> <li>Severity correlation. In preputial tissue from boys with hypospadias, reduced FGF10 protein levels (IHC/Western) have been reported, with greater reduction in more severe cases, supporting a link between diminished upstream FGF10 and preputial pathology (Yamada et al., 2025).</li> <li>Genetic associations. Rare FGFR2 variants have been identified in familial hypospadias (Swedish cohort), and population‑level FGF10 SNPs are associated with increased hypospadias risk in a large, diverse cohort, supporting human plausibility that perturbations of the FGF10–FGFR2‑IIIb axis can contribute to preputial/urethral developmental defects (Beleza‑Meireles et al., 2007; Carmichael et al., 2013). For anatomical translation of epithelial compartments to human foreskin/prepuce, see keratin boundary mapping in fetal tissues (Kurzrock et al., 1999).</li> </ul>

Human protein vs mRNA discordance and postnatal sampling In hypospadiac foreskin, immunohistochemistry shows altered/lower FGF8, FGF10, and FGFR2 protein patterns versus controls (p < 0.01), yet qPCR detects no significant mRNA differences in the same samples; cases were postnatal surgical specimens (mean age ≈25 months for patients, ≈77 months for controls), not embryonic tissues. This limits inference about whether reduced prenatal FGF10/FGFR2‑IIIb signaling is causal versus secondary to altered tissue architecture or post‑transcriptional regulation (Haid et al., 2020). A separate pediatric foreskin study also reports lower FGF8/FGF10/FGFR2 IHC signals in hypospadias (p < 0.05), further supporting protein‑level differences with uncertain transcriptional basis (Emaratpardaz et al., 2024). Quantitative response–response gap in vivo Ex vivo GT culture shows a clear dose‑response for transcript loss after 48 h of AR antagonism (FGFR2‑IIIb, and FGF10 at higher doses), and compartmental sufficiency is defined genetically. However, graded in vivo measurements linking partial reductions in FGF10/FGFR2‑IIIb signaling to incremental failures of ventral prepuce closure are limited (semi‑quantitative rather than absolute readouts of receptor signaling; few time‑resolved, compartment‑specific quantifications) (Petiot et al., 2005; Gredler et al., 2015). Cross‑species endpoint definition Because guinea pig normally retains an open urethral groove and lacks full ventral prepuce enclosure, “disrupted preputial epithelial morphogenesis” should be scored as shifts in prepuce‑vs‑groove balance in that species. Applying a mouse‑style “failed ventral enclosure” endpoint across species could misclassify outcomes and obscure true cross‑species concordance (Wang & Zheng, 2025).

<table class="Table"> <thead> <tr> <td> Modulating Factor </td> <td> Details </td> <td> Effect on KER </td> <td> References </td> </tr> </thead> <tbody> <tr> <td> Heparan sulfate proteoglycans (HSPGs) </td> <td> HSPGs are required cofactors that promote high‑affinity FGF10–FGFR2‑IIIb complex formation and signaling; HS chain length and sulfation pattern modulate ligand–receptor binding and signaling strength. </td> <td> Disruption of HS biosynthesis/sulfation is expected to attenuate FGF10–FGFR2‑IIIb signaling and weaken preputial epithelial morphogenesis; GT‑specific HS loss‑of‑function data are not yet available. </td> <td> Ohuchi et al., 2000; Ornitz, 2015. </td> </tr> <tr> <td> Sex (androgen environment) </td> <td> Male and female GT explants show similar dose‑dependent reductions of FGFR2‑IIIb and FGF10 with flutamide ex vivo, but in vivo AR antagonism during the window feminizes male urethral development while females are largely unaffected. </td> <td> The molecular AR→FGF axis operates in both sexes, but reduced signaling has greater morphogenetic consequences in males. </td> <td> Petiot et al., 2005; Seifert et al., 2008; Sharpe, 2020. </td> </tr> <tr> <td> Sonic hedgehog (SHH) pathway activity </td> <td> SHH from the urethral epithelium promotes mesenchymal FGF10 and GT growth; conditional Shh loss reduces FGF10 (GT), and SHH/FGF manipulations in organ culture shift the balance between preputial outgrowth and persistence of a urethral groove. Shh does not appear to regulate FGFR2 transcription in GT. </td> <td> Reduced SHH can secondarily lower FGF10 and diminish the upstream KE, biasing development toward groove persistence and restrained preputial development. </td> <td> Perriton et al., 2002; Seifert et al., 2010; Gredler et al., 2015; Wang & Zheng, 2025. </td> </tr> <tr> <td> WNT/β‑catenin signaling </td> <td> β‑catenin is required in external genital epithelial tissues; WNT programs interface with epithelial adhesion/morphogenesis and with mesenchymal cues that include FGF10. </td> <td> Reduced WNT/β‑catenin signaling can impair epithelial behaviors needed for preputial fusion and may lower FGF10 support, weakening the upstream KE and exacerbating KEdownstream defects. </td> <td> Lin et al., 2008. </td> </tr> <tr> <td> ISL1 transcriptional program </td> <td> ISL1 controls mesenchymal expansion via regulation of Bmp4, FGF10, and Wnt5a in the developing external genitalia. </td> <td> Reduced ISL1 activity diminishes FGF10 and related cues, decreasing FGF10–FGFR2‑IIIb signaling and compromising preputial epithelial morphogenesis. </td> <td> Ching et al., 2018. </td> </tr> <tr> <td> Ligand redundancy at FGFR2‑IIIb </td> <td> FGFR2‑IIIb can bind multiple epithelial ligands (e.g., FGF7, FGF10); FGF10 is a major ligand across epithelia. </td> <td> Partial compensation by other ligands (e.g., FGF7) could blunt the impact of reduced FGF10 on this KER, potentially modifying effect size and penetrance. </td> <td> Ohuchi et al., 2000; Itoh & Ornitz, 2011. </td> </tr> <tr> <td> FGFR2 compartment specificity </td> <td> Ectodermal FGFR2 is required for ventral prepuce closure and maintenance of a closed urethral tube; endodermal FGFR2 supports urethral epithelial proliferation/stratification. </td> <td> The same reduction in upstream signaling produces different downstream severities depending on which epithelial compartment is most affected. </td> <td> Gredler et al., 2015; Harada et al., 2015. </td> </tr> </tbody> </table>

Direct, quantitative dose–response data that map graded reductions in FGF10/FGFR2‑IIIb signaling to graded severity of preputial epithelial morphogenesis defects remain limited. However, several semi‑quantitative relationships support a graded or threshold‑like linkage: Genetic dosage and compartment specificity (mouse). Homozygous loss of FGFR2‑IIIb (or FGF10) produces severe hypospadias with failure of ventral prepuce formation, whereas early GT outgrowth remains grossly normal, indicating a threshold requirement for epithelial FGFR2‑IIIb signaling during the period of preputial outgrowth/fusion (Petiot et al., 2005). Conditional deletions show graded, compartment‑specific consequences: endodermal FGFR2 loss yields milder hypospadias with impaired epithelial maturation, while ectodermal FGFR2 loss causes severe failure of ventral prepuce closure and ventral tethering, implying that the magnitude and site of receptor loss scale with the severity of preputial defects (Gredler et al., 2015; Harada et al., 2015). Temporal and morphometric context (mouse). The period of preputial elevation and ventral fusion (~E13–E17.5) aligns with the stages when epithelial FGFR2‑IIIb signaling is required; failure of ventral prepuce closure becomes apparent as development proceeds toward late gestation, consistent with a progressive deficit when upstream signaling is reduced (Satoh et al., 2004; Yamada et al., 2006; Suzuki et al., 2002; Gredler et al., 2015; Harada et al., 2015). Together, these findings support a semi‑quantitative picture in which partial attenuation of FGF10/FGFR2‑IIIb signaling can produce milder epithelial maturation defects, while more profound or compartment‑targeted reductions (especially in ectoderm) cross a threshold that results in failed ventral prepuce closure and ventral tethering. Nonetheless, the field lacks in vivo, compartment‑resolved measurements that link incremental decreases in ligand/receptor levels or receptor activation (e.g., pERK readouts) to graded preputial outcomes.

The temporal relationship is clear from mouse developmental staging. FGFR2‑IIIb transcripts are detected in the urethral plate epithelium beginning at E10.5 and persist through E16.5; expression appears in the paired preputial swellings as they emerge along the lateral edges of the tubercle at ~E13.5 (Petiot et al., 2005). Consistent with this, FGF signaling is first detected in GT mesenchyme at ~E11.5, is evident in mesenchyme, ectoderm, and endoderm by ~E13.5, and becomes predominantly epithelial by ~E14.5 (Harada et al., 2015). The preputial swellings initiate around E13.0–E13.5 and then grow laterally and ventrally to form the prepuce; by E15.5 the swellings cover the proximal glans, and by ~E17.0–E17.5 the prepuce normally surrounds the glans and penile shaft (Satoh et al., 2004; Suzuki et al., 2002; Petiot et al., 2005; Gredler et al., 2015; Wang & Zheng, 2025). In FGFR2‑IIIb null embryos, ventral defects in the urethral plate and preputial domain are evident by E13.5, and the ventral prepuce fails to form thereafter; by late gestation the urethra remains open with only lateral preputial tissue present (Petiot et al., 2005). In ectoderm‑specific FGFR2 deletion, mislocalization of the preputial swellings is already apparent at ~E13.0 and progresses to failed ventral prepuce closure (Gredler et al., 2015). Taken together, these data indicate that detectable disruption of preputial morphogenesis follows within roughly one to two embryonic days of the stage when preputial swellings normally initiate (~E13.0–E13.5), with the full morphogenetic outcome (failed ventral enclosure/ventral tethering) established over the ensuing embryonic days through ~E17.0–E17.5 (Petiot et al., 2005; Gredler et al., 2015).

SHH→FGF10 feed‑forward influence Evidence: In the genital tubercle (GT), SHH is produced by the urethral epithelium and promotes mesenchymal growth; Shh loss reduces mesenchymal FGF10 in GT, placing SHH upstream of the FGF10–FGFR2‑IIIb axis (Perriton et al., 2002; Seifert et al., 2010). In organ culture, inhibiting SHH/FGF signaling restrains preputial development and maintains a urethral groove in mouse GT, whereas adding SHH or FGF10 promotes preputial outgrowth in guinea pig GT, which normally exhibits an open groove (Wang & Zheng, 2025). Together, these findings support a feed‑forward influence in which SHH maintains/boosts FGF10 to drive epithelial FGFR2‑IIIb signaling needed for preputial morphogenesis. FGF→SHH feedback in GT (not demonstrated) Evidence: In limb, FGFR2‑IIIb acts upstream of Shh, illustrating a reciprocal logic in another organ (Revest et al., 2001). In GT, however, data argue against a similar feedback: conditional Shh deletion did not alter FGFR2 mRNA in GT, and early urethral Shh/FGF8 domains are established even in FGFR2‑IIIb nulls (though maturation subsequently fails), indicating that direct FGF→Shh feedback has not been shown in GT (Gredler et al., 2015; Petiot et al., 2005). Thus, a reciprocal SHH↔FGF loop (as in lung branching) is not established for genital preputial morphogenesis. SHH/FGF cooperation vs formal feedback loop Evidence: The combined gain‑ and loss‑of‑function organ culture data indicate that SHH and FGF10 act cooperatively to bias development toward preputial outgrowth versus a persistent urethral groove (Wang & Zheng, 2025). Whether this cooperation constitutes a bona fide feedback loop (with mutual regulation) or parallel, convergent inputs remains unresolved for GT.

High Male Moderate Female

High

Embryo

High

Foetal

High

Fetal

High Moderate Moderate Moderate Moderate

Taxonimic Applicability: In Mus musculus, global loss of FGF10 or FGFR2‑IIIb and tissue‑specific deletion of FGFR2 demonstrate that epithelial FGFR2‑IIIb signaling is required for urethral epithelial proliferation/stratification and ventral prepuce closure, with ectodermal deletion yielding severe failure of ventral enclosure and ventral tethering (Petiot et al., 2005; Gredler et al., 2015; Harada et al., 2015). In Rattus norvegicus, anti‑androgen exposures within the masculinization programming window (GD15.5–18.5) feminize urethral development and produce ventral fusion/prepuce defects, consistent with downstream consequences of reduced epithelial FGF signaling, although direct compartment‑specific FGFR2‑IIIb genetics in rat are limited (Welsh et al., 2008; Seifert et al., 2008; Sinclair et al., 2016; Sharpe, 2020). In Cavia porcellus, which normally retains an open urethral groove, organ‑culture manipulations show that inhibiting SHH/FGF restrains prepuce and maintains a groove in mouse GT, whereas adding SHH or FGF10 promotes prepuce outgrowth in guinea pig GT, indicating that the level of FGF10– FGFR2‑IIIb signaling causally biases development toward prepuce formation versus groove persistence, albeit with limited in vivo genetic confirmation in this species (Wang & Zheng, 2025). Together, these data provide a strong rodent‑wide rationale that reduced FGF10–FGFR2‑IIIb signaling in genital tissues leads to disrupted preputial epithelial morphogenesis. Human foreskin from boys with hypospadias shows altered protein‑level patterns for FGF10/FGFR2 (and FGF8) versus controls; cohort genetics implicate FGF10 SNPs (population‑level risk) and rare FGFR2 variants (familial cases). Direct mechanistic data in embryonic human GT during the sensitive window are not available, so support is correlative. Mammalia is plausible given conserved mesenchyme‑to‑epithelium FGF10→FGFR2‑IIIb signaling in external genital development, with caveats for species‑specific distal urethral/prepuce anatomy and timing (Amato et al., 2022; Haraguchi et al., 2000; Perriton et al., 2002; Seifert et al., 2010; Sharpe, 2020). Life Stage Applicability: In mouse, FGFR2‑IIIb is detectable in the urethral plate by ~E10.5, but preputial swellings initiate at ~E13–E13.5, cover the proximal glans by ~E15.5, and normally enclose the glans/shaft by ~E17–E17.5; loss of epithelial FGFR2 signaling produces defects detectable at or soon after preputial initiation and culminates in failed ventral prepuce closure by late gestation (Petiot et al., 2005; Satoh et al., 2004; Suzuki et al., 2002; Yamada et al., 2006; Gredler et al., 2015; Harada et al., 2015). In rat, sensitivity is concentrated within the fetal masculinization programming window (GD15.5–18.5), during which upstream anti‑androgen exposure yields ventral fusion/prepuce defects consistent with reduced epithelial FGF signaling (Welsh et al., 2008; Seifert et al., 2008; Sharpe, 2020). By analogy, the presumptive human window is ~8–14 gestational weeks, though direct fetal mechanistic data are limited (Sharpe, 2020). In guinea pig, organ‑culture experiments show that modulating SHH/FGF levels during embryonic stages shifts development toward preputial outgrowth versus persistent urethral groove, underscoring a level‑ and time‑dependent requirement for FGF10–FGFR2‑IIIb during the species‑specific embryonic window (Wang & Zheng, 2025). Sex Applicability: This KER operates in both sexes during the sexually indifferent stage of external genital development: in GT organ culture, AR antagonism produces similar dose‑dependent reductions of FGFR2‑IIIb and FGF10 in male and female explants, and DHT rescues FGFR2‑IIIb, indicating AR‑dependent control of the FGF10–FGFR2‑IIIb axis in both sexes (Petiot et al., 2005). Global loss of FGFR2‑IIIb or FGF10 yields severe external genital malformations consistent with hypospadias in embryos, with Petiot reporting effects across sexes, while tissue‑specific deletions of FGFR2 (characterized primarily in males) show that ectodermal FGFR2 is required for ventral prepuce closure and endodermal FGFR2 for urethral epithelial maturation (Petiot et al., 2005; Gredler et al., 2015). In vivo, however, androgen suppression within the programming window feminizes male urethral development whereas flutamide‑treated females show little change in urethral position, indicating that downstream morphogenetic consequences are more pronounced in males; this male‑biased sensitivity is consistent with higher androgen tone during the window rather than proven sex‑biased amplification of FGF10/FGFR2 expression per se (Seifert et al., 2008; Sharpe, 2020).

Amato, R., Yao, H.-H.-C., & Zhao, F. (2022). One tool for many jobs: Divergent and conserved actions of androgen signaling in male internal reproductive tract and external genitalia. Frontiers in Endocrinology, 13, 910964. https://doi.org/10.3389/fendo.2022.910964 Beleza-Meireles, A., Lundberg, F., Lagerstedt, K., et al. (2007). FGFR2, FGF8, FGF10 and BMP7 as candidate genes for hypospadias. European Journal of Human Genetics, 15(4), 405–410. https://doi.org/10.1038/sj.ejhg.5201777 Carmichael, S. L., Ma, C., Choudhry, S., Lammer, E. J., Witte, J. S., & Shaw, G. M. (2013). Hypospadias and genes related to genital tubercle and early urethral development. Journal of Urology, 190(5), 1884–1892. https://doi.org/10.1016/j.juro.2013.05.061 Ching, S. T., Infante, C. R., Du, W., Sharir, A., Park, S., Menke, D. B., & Klein, O. D. (2018). Isl1 mediates mesenchymal expansion in the developing external genitalia via regulation of Bmp4, Fgf10 and Wnt5a. Human Molecular Genetics, 27(1), 107–119. https://doi.org/10.1093/hmg/ddx388 Emaratpardaz, N., Turkyilmaz, Z., et al. (2024). Comparison of FGF-8, FGF-10, FGF-Receptor 2, androgen receptor, estrogen receptor-α and SS in healthy and hypospadiac children. Balkan Journal of Medical Genetics, 27(1), 21–29. https://doi.org/10.2478/bjmg-2024-0002 Gredler, M. L., Seifert, A. W., & Cohn, M. J. (2015). Tissue-specific roles of Fgfr2 in development of the external genitalia. Development, 142(12), 2203–2212. https://doi.org/10.1242/dev.119891 Haid, B., Pechriggl, E., Nägele, F., et al. (2020). FGF8, FGF10 and FGF receptor 2 in foreskin of children with hypospadias: An analysis of immunohistochemical expression patterns and gene transcription. Journal of Pediatric Urology, 16(1), 41.e1–41.e10. https://doi.org/10.1016/j.jpurol.2019.10.007 Harada, M., Omori, A., Nakahara, C, Nakagata, N., Akita, K., & Yamada, G. (2015). Tissue-specific roles of FGF signaling in external genitalia development. Developmental Dynamics, 244(6), 759–773. https://doi.org/10.1002/dvdy.24277 Haraguchi, R., Suzuki, K., Murakami, R., et al. (2000). Molecular analysis of external genitalia formation: The role of fibroblast growth factor (Fgf) genes during genital tubercle formation. Development, 127(11), 2471–2479. Itoh, N., & Ornitz, D. M. (2011). Fibroblast growth factors: From molecular evolution to roles in development, metabolism and disease. Journal of Biochemistry, 149(2), 121–130. https://doi.org/10.1093/jb/mvq166 Kurzrock, E. A., Baskin, L. S., & Cunha, G. R. (1999). Ontogeny of the male urethra: Theory of endodermal differentiation. Differentiation, 64(2), 115–122. Lin, C., Yin, Y., Long, F., & Ma, L. (2008). Tissue-specific requirements of β-catenin in external genitalia development. Development, 135(16), 2815–2825. https://doi.org/10.1242/dev.020586 Ohuchi, H., Hori, Y., Yamasaki, M., Harada, H, Sekine, K., Kato, S., & Itoh, N. (2000). FGF10 acts as a major ligand for FGF receptor 2 IIIb in mouse multi-organ development. Biochemical and Biophysical Research Communications, 277(3), 643–649. https://doi.org/10.1006/bbrc.2000.3721 Ornitz, D. M. (2015). The fibroblast growth factor signaling pathway. WIREs Developmental Biology, 4(3), 215–266. https://doi.org/10.1002/wdev.176 Perriton, C. L., Powles, N., Chiang, C., Maconochie, M. K., & Cohn, M. J. (2002). Sonic hedgehog signaling from the urethral epithelium controls external genital development. Developmental Biology, 247(1), 26–46. https://doi.org/10.1006/dbio.2002.0668 Petiot, A., Perriton, C. L., Dickson, C., & Cohn, M. J. (2005). Development of the mammalian urethra is controlled by Fgfr2-IIIb. Development, 132(10), 2441–2450. https://doi.org/10.1242/dev.01778 Revest, J. M., Spencer-Dene, B., Kerr, K., De Moerlooze, L., Rosewell, I., & Dickson, C. (2001). Fibroblast growth factor receptor 2-IIIb acts upstream of Shh and Fgf4 and is required for limb bud maintenance but not for the induction of Fgf8, Fgf10, Msx1, or Bmp4. Developmental Biology, 231(1), 47–62. https://doi.org/10.1006/dbio.2000.0144 Seifert, A. W., Zheng, Z., Ormerod, B. K., & Cohn, M. J. (2010). Sonic hedgehog controls growth of external genitalia by regulating cell cycle kinetics. Nature Communications, 1, Article 23. https://doi.org/10.1038/ncomms1020 Sharpe, R. M. (2020). Androgens and the masculinization programming window: Human–rodent differences. Biochemical Society Transactions, 48(4), 1725–1735. https://doi.org/10.1042/BST20200200 Sinclair, A. W., Cao, M., Pask, A., Baskin, L. S., & Cunha, G. R. (2017). Flutamide-induced hypospadias in rats: A critical assessment. Differentiation, 94, 37–57. https://doi.org/10.1016/j.diff.2016.06.003 Satoh, Y., Haraguchi, R., Wright, T. J., Mansour, S. L., Partanen, J., Hajihosseini, M. K., Eswarakumar, V. P., Lonai, P., & Yamada, G. (2004). Regulation of external genitalia development by concerted actions of FGF ligands and FGF receptors. Anatomical Embryology, 208(6), 479–486. https://doi.org/10.1007/s00429-004-0419-9 Suzuki, K., Ogino, Y., Murakami, R., Satoh, Y., Bachiller, D., & Yamada, G. (2002). Embryonic development of mouse external genitalia: Insights into a unique mode of organogenesis. Evolution & Development, 4(2), 133–141. Welsh, M., Saunders, P. T. K., Fisken, M., et al. (2008). Identification in rats of a programming window for reproductive tract masculinization, disruption of which leads to hypospadias and cryptorchidism. Journal of Clinical Investigation, 118(4), 1479–1490. https://doi.org/10.1172/JCI34241 Wang, X., & Zheng, Z. (2025). Comparative modulation of SHH/FGF signaling determines preputial outgrowth versus urethral groove persistence in rodent genital tubercle organ culture. Cells, 14, 348. https://doi.org/10.3390/cells14050348 Yamada, G., Suzuki, K., Haraguchi, R., et al. (2006). Molecular genetic cascades for external genitalia formation: An emerging organogenesis program. Developmental Dynamics, 235(7), 1738–1752. https://doi.org/10.1002/dvdy.20829 Yamada, S., et al. (2025). Down-regulation of FGF10 in hypospadias prepuce associated with severity. Journal of Pediatric Surgery Open, 10, 100206. AOPWiki 2025-12-16T14:54:58

2026-06-04T15:18:21

<upstream-id>71f737c5-2a8f-4a4b-b9d9-585e6994255b</upstream-id> <downstream-id>8dc8298b-d39c-4971-a2a5-8c036379243b</downstream-id>

AOPWiki 2025-12-16T14:55:08

2025-12-16T14:55:08

Androgen receptor antagonism leads to delayed preputial seperation via reduced fibroblast growth factor in genital-tubercle tissues

AR agonism leads to delayed PPS via reduced FGF expression

Allie Always

Travis Karschnik (General Dynamics Information Technology, Duluth, MN, USA.)

BY-SA

2.7

<a href="https://aopwiki.org/system/dragonfly/production/2026/03/27/kd0q21izj_AOP_Wiki_curation_assistant_master_prompt.docx">Master prompt</a> <div> This AOP was as part of an Environmental Protection Agency effort to develop AOPs that establish scientifically supported causal linkages between alternative endpoints measured using new approach methodologies (NAMs) and guideline apical endpoints measured in Tier 1 and Tier 2 test guidelines (U.S. EPA, 2024) employed by the Endocrine Disruptor Screening Program (EDSP).  A series of key events that represent significant, measurable, milestones connecting molecular initiation to apical endpoints indicative of adversity were identified based on scientific review articles and empirical studies.  Additionally, scientific evidence supporting the causal relationships between each pair of key events was assembled and evaluated.   </div>

The scope of the aforementioned EPA project was to develop AOP(s) relevant to apical endpoints observed in the test guidelines, based on mechanisms consistent with empirical studies. The literature used to support this AOP and its constituent pages began with the test guidelines and followed to primary, secondary, and/or tertiary works concerning the relevant underlying biology. KE and KER page creation and re-use was determined using Handbook principles where page re-use was preferred.

A large number of drugs and chemicals have been shown to antagonise the AR using various AR reporter gene assays. The AR is specifically targeted in AR-sensitive cancers, for example the use of the anti-androgenic drug flutamide in treating prostate cancer (<a href="#_ENREF_1" title="Alapi, 2006 #262">Alapi & Fischer, 2006</a>). Flutamide has also been used in several rodent in vivo studies showing anti-androgenic effects (feminization of male offspring) evident by e.g. short anogenital distance (AGD) in males (<a href="#_ENREF_4" title="Foster, 2005 #53">Foster & Harris, 2005</a>; <a href="#_ENREF_5" title="Hass, 2007 #76">Hass et al, 2007</a>; <a href="#_ENREF_8" title="Kita, 2016 #34">Kita et al, 2016</a>). QSAR models can predict AR antagonism for a wide range of chemicals, many of which have shown in vitro antagonistic potential (<a href="#_ENREF_17" title="Vinggaard, 2008 #263">Vinggaard et al, 2008</a>).

A delay or failure in preputial separation represents an apical endpoint in standard developmental and reproductive toxicity tests which are used for hazard identification, risk assessment, and regulatory decision-making, especially as it relates to chemicals with potential endocrine-disrupting activity. International Regulatory Context PPS is included in several OECD Test Guidelines that are adopted by OECD member countries as well as several non-OECD countries that adhere to Mutual Acceptance of Data (MAD) e.g., Argentina, Brazil, India). <ul> <li>OECD Test No. 443 - Extended One-Generation Reproductive Toxicity Study (EOGRTS)</li> <li>OECD Test No. 416 - Two-Generation Reproductive Toxicity Study: <ul> <li>Requires daily evaluation of balano-preputial separation in male pups.  It is used for comprehensive assessment of developmental and reproductive effects inclusive of endocrine disruption indicators.</li> </ul> </li> </ul> OECD member countries and regions adhering to MAD use data on delayed PPS as a regulatory endpoint used for: <ul> <li> <ul> <li>Hazard identification and classification (e.g., under EU CLP No 1272/2008)</li> <li>Risk assessment supporting chemical registration/authorization decisions under REACH/ECHA.</li> </ul> </li> </ul> United States Regulatory Context <ul> <li>U.S. EPA Two-Generation Reproductive Toxicity Test Guideline (OPPTS 870.3800) <ul> <li>PPS is an endpoint used for evaluation under Federal Insecticide, Fungicide, and Rodenticide Act (FIFRA) and Toxic Substances Control Act (TSCA) to detect effects on the integrity and performance of the reproductive system.</li> </ul> </li> <li>EPA Endocrine Disruptor Screening Program (EDSP) Tier 1 and Tier 2 <ul> <li>Incorporates rodent pubertal assays (including male pubertal assay) that use age at preputial separation as a sensitive measure of disruption of androgen signaling. <ul> <li>EDSP is integrated into the Federal Food, Drug, and Cosmetic Act (FFDCA) in section 408(p) and the Safe Drinking Water Act in Section 1457.</li> </ul> </li> </ul> </li> </ul> EPA uses these tests to screen, identify, and prioritize chemicals for potential endocrine activity and hazard.  Regulatory decisions like restrictions, registrations, and risk mitigation occur under TSCA and FIFRA statutes.

adjacent

Not Specified

Not Specified adjacent

Not Specified

Not Specified adjacent

Not Specified

Not Specified adjacent

Not Specified

<div> <table class="table table-bordered table-fullwidth"> <thead> <tr> <th>Modulating Factor (MF)</th> <th>Influence or Outcome</th> <th>KER(s) involved</th> </tr> </thead> <tbody> <tr> <td> </td> <td> </td> <td> </td> </tr> </tbody> </table> </div>

AOPWiki 2025-12-03T11:42:47

2026-06-26T16:12:16