Relationship: 2401: Decreased, ALDH1A activity leads to Decreased, atRA concentration

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

Key Event Relationship Description

All-trans retinoic acid (atRA) is the active metabolite of vitamin A in developing mammals and its physiological levels is tightly regulated by enzymatic pathways. This KER is particularly relevant for mammalian embryogenesis/fetal development stages.

atRA is synthesized from dietary vitamin A (retinol) by a two-step oxidation pathway (Chatzi et al, 2013; Kedishvili, 2016): 1) retinol dehydrogenase (RDH10) metabolizes retinol to retinaldehyde (reversible step), 2) retinaldehyde dehydrogenase ALDH1A (ALDH1A1, ALDH1A2, ALDH1A3) metabolizes retinaldehyde to RA (irreversible step). All three isoenzymes can carry out the second (irreversible step) to produce atRA, but ALDH1A2 is the most active form during development (Kedishvili, 2016). Thus, inhibition of ALDH1A2 during development will decrease atRA concentrations.

Evidence Supporting this KER

Evidence showing that retinaldehyde dehydrogenases is responsible for the irreversible oxidation of retinal to retinoic acid was provided by several studies in the 1960s, using calf and rat livers (Dmitrovskii, 1961; Dunagin Jr et al, 1964; Elder & Topper, 1962; Futterman, 1962; Lakshmanan et al, 1964; Mahadevan et al, 1962), as reviewed by (Kedishvili, 2016). The identification of the three isoenzymes ALDH1A1 (RALDH1), ALDH1A2 (RALDH2), ALDH1A3 (RALDH3) followed during 1980-1990 (Kedishvili, 2016). It is now considered canonical knowledge that the three retinaldehyde dehydrogenases are responsible for the in vivo biosynthesis of retinoic acid from retinal (Marchitti et al, 2008; Napoli, 2012).

Embryogenesis/fetal development in mammals

Of the three isoenzymes, ALDH1A2 is the most active form during early development in mammals. This is evidenced in mice ablated for Aldh1a2 (Raldh2^-/-), which are incapable of producing atRA and present with severe developmental defects (Niederreither et al, 1999). Conversely, mice lacking Aldh1a1 or Aldh1a3 survive fetal development, with phenotypes presenting postnatally (Dupé et al, 2003; Fan et al, 2003; Molotkov & Duester, 2003). Thus, the biological plausibility that inhibition of ALDH1A2 will lead to decreased atRA in cells and tissues during development is strong.

The empirical evidence for linkage is strong and widely accepted. The enzymatic activity of ALDH1A2 and capacity to oxidize retinal has been proven in vitro (see KE 1880). In vivo, the strongest evidence comes from the Aldh1a2-deficient mice that fail to synthesize retinoic acid during embryogenesis (Niederreither et al, 1999). Additionally, ovary culture with the potent ALDH1A2 inhibitor WIN18,446 results in failure to upregulate the atRA-regulated gene Stra8 in oocytes, resulting in germ cell loss (Rosario et al, 2020). Additional evidence for this relationship using WIN18,466 also comes from in vivo studies looking at spermatogenesis; inhibition of ALDH1A2 via WIN18,466 results in loss of atRA expression and halted spermatogenesis in diverse species such as mice, rabbits and zebrafish (Amory et al, 2011; Paik et al, 2014; Pradhan & Olsson, 2015).

There are redundant pathways for atRA synthesis (e.g. ALDH isoforms) which may buffer a decrease in atRA concentrations caused by reduced ALDH1A activity, complicating the prediction of changes to atRA concentration. There is also tissue-specific expression of various components of the atRA synthesis pathways, which introduces additional variability in atRA concentration outcomes depending on biological context.

Quantitative Understanding of the Linkage

The distribution of retinoic acid in cells and tissues are highly variable, as has been shown across species including chicken (Maden et al, 1998), frogs (Chen et al, 1994), mice (Kane et al, 2005; Obrochta et al, 2014) and rats (Bhat, 1997), as well as serum/plasma from humans (Kane et al, 2008; Miyagi et al, 2001; Napoli et al, 1985).

The exact relationship between ALDH1A2 inhibition and resulting atRA concentrations in mammalian ovaries is unclear. The ALDH1A2 inhibitor WIN18,446 inhibits enzyme activity in vitro with an IC(50) of 0.3 μM (Amory et al, 2011), and a dose of only 0.01 µM is sufficient to significantly reduce expression of Stra8 in cultured mouse fetal ovaries and with actual loss of oocytes from 2 µM (Rosario et al, 2020).

Since atRA must be enzymatically synthesized by ALDH1A enzymes (in this case ALDH1A2), the temporal and linear relationship between the two KEs are essential.

Retinoic acid status is regulated by complex feedback loops. For instance, atRA induces expression of retinoid enzymes to promote synthesis of retinyl esters, but simultaneously atRA induces expression of its own catabolizing CYP26 enzymes (Kedishvili, 2013; Kedishvili, 2016; Teletin et al, 2017).

References

Amory JK, Muller CH, Shimshoni JA, Isoherranen N, Paik J, Moreb JS, Amory Sr DW, Evanoff R, Goldstein AS, Griswold MD (2011) Suppression of spermatogenesis by bisdichloroacetyldiamines is mediated by inhibition of testicular retinoic acid biosynthesis. J Androl 32: 111-119

Bhat PV (1997) Tissue concentrations of retinol, retinyl esters, and retinoic acid in vitamin A deficient rats administered a single dose of radioactive retinol. Can J Physiol Pharmacol 75: 74-77

Chatzi C, Cunningham TJ, Duester G (2013) Investigation of retinoic acid function during embryonic brain development using retinaldehyde-rescued Rdh10 knockout mice. Dev Dyn 242: 1056-1065

Chen Y, Huang L, Solursh M (1994) A concentration gradient of retinoids in the early Xenopus laevis embryo. Dev Biol 161: 70-76

Dmitrovskii AA (1961) Oxidation of vitamin A aldehyde to vitamin A acid catalyzed by aldehyde oxidase. Biokhimiya 26: 126

Dunagin Jr PE, Zachman RD, Olson JA (1964) Identification of free and conjugated retinoic acid as a product of retinal (vitamin A aldehyde) metabolism in the rat in vivo. Biochim Biophys Acta 90: 432-434

Dupé V, Matt N, Garnier JM, Chambon P, Mark M, Ghyselinck NB (2003) A newborn lethal defect due to inactivation of retinaldehyde dehydrogenase type 3 is prevented by maternal retinoic acid treatment. Proc Natl Acad Sci U S A 100: 14036-14041

Elder TD, Topper YJ (1962) The oxidation of retinene (vitamin A1 aldehyde) to vitamin A acid by mammalian steroid-sensitive aldehyde dehydrogenase. Biochim Biophys Acta 64: 430

Fan X, Molotkov A, Manabe SI, Donmoyer CM, Deltour L, Foglio MH, Cuenca AE, Blaner WS, Lipton SA, Duester G (2003) Targeted disruption of Aldh1a1 (Raldh1) provides evidence for a complex mechanism of retinoic acid synthesis in the developing retina. Mol Cell Biol 23: 4637-4648

Futterman S (1962) Enzymatic oxidation of vitamin A aldehyde to vitamin A acid. J Biol Chem 237: 677-680

Kane MA, Chen N, Sparks S, Napoli JL (2005) Quantification of endogenous retinoic acid in limited biological samples by LC/MS/MS. Biochem J 388: 363-369

Kane MA, Folias AE, Napoli JL (2008) HPLC/UV quantitation of retinal, retinol, and retinyl esters in serum and tissues. Anal Biochem 378: 71-79

Kedishvili NY (2013) Enzymology of retinoic acid biosynthesis and degradation. J Lipid Res 54: 1744-1760

Kedishvili NY (2016) Retinoic Acid Synthesis and Degradation. Subcell Biochem 81: 127-161

Lakshmanan MR, Vaidyanathan CS, Cama HR (1964) Oxidation of vitamin A1 aldehyde and vitamin A2 aldehyde to the corresponding acids by aldehyde oxidase from different species. Biochem J 90: 569-573

Maden M, Sonneveld E, van der Saag PT, Gale E (1998) The distribution of endogenous retinoic acid in the chick embryo: implications for developmental mechanisms. Development 125: 4133-4144

Mahadevan S, Murthy SK, Ganguly J (1962) Enzymic oxidation of vitamin A aldehyde to vitamin A acid by rat liver. Biochem J 85: 326-331

Marchitti SA, Brocker C, Stagos D, Vasiliou V (2008) Non-P450 aldehyde oxidizing enzymes: the aldehyde dehydrogenase superfamily. Expert Opin Drug Metab Toxicol 4: 697-720

Miyagi M, Yokoyama H, Shiraishi H, Matsumoto M, Ishii H (2001) Simultaneous quantification of retinol, retinal, and retinoic acid isomers by high-performance liquid chromatography with a simple gradiation. J Chromatogr B Biomed Sci Appl 757: 365-368

Molotkov A, Duester G (2003) Genetic evidence that retinaldehyde dehydrogenase Raldh1 (Aldh1a1) functions downstream of alcohol dehydrogenase Adh1 in metabolism of retinol to retinoic acid. J Biol Chem 278: 36085-36090

Napoli JL (2012) Physiological insights into all-trans-retinoic acid biosynthesis. Biochim Biophys Acta 1821: 152-167

Napoli JL, Pramanik BC, Williams JB, Dawson MI, Hobbs PD (1985) Quantification of retinoic acid by gas-liquid chromatography-mass spectrometry: total versus all-trans-retinoic acid in human plasma. J Lipid Res 26: 387-392

Niederreither K, Subbarayan V, Dollé P, Chambon P (1999) Embryonic retinoic acid synthesis is essential for early mouse post-implantation development. Nat Genet 21: 444-448

Obrochta KM, Kane MA, Napoli JL (2014) Effects of diet and strain on mouse serum and tissue retinoid concentrations. PLoS One 9: e99435

Paik J, Haenisch M, Muller CH, Goldstein AS, Arnold S, Isoherranen N, Brabb T, Treuting PM, Amory JK (2014) Inhibition of retinoic acid biosynthesis by the bisdichloroacetyldiamine WIN 18,446 markedly suppresses spermatogenesis and alters retinoid metabolism in mice. J Biol Chem 289: 15104-15117

Pradhan A, Olsson PE (2015) Inhibition of retinoic acid synthesis disrupts spermatogenesis and fecundity in zebrafish. Gen Comp Endocrinol 217-218: 81-91

Rosario R, Stewart HL, Walshe E, Anderson RA (2020) Reduced retinoic acid synthesis accelerates prophase I and follicle activation. Reproduction 160: 331-341

Teletin M, Vernet N, Ghyselinck NB, Mark M (2017) Roles of Retinoic Acid in Germ Cell Differentiation. Curr Top Dev Biol 125: 191-225

Relationship: 2477: Decreased, atRA concentration leads to Disrupted, meiotic initiation in oocyte

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

Key Event Relationship Description

All-trans retinoic acid (atRA) is the active metabolite of vitamin A and is involved in regulating a large number of developmental processes (Bushue & Wan, 2010; Ghyselinck & Duester, 2019). atRA is produced in spatial and temporal gradients, and these patterns are maintained by regulated expression of the synthesis and degradation enzymes of the atRA pathway (Kedishvili, 2013).The presence of atRA in the fetal ovaries induces germ cells to enter meiosis (Spiller et al, 2017). The initiation of meiosis at this time during fetal life is critical for maintenance of the germ line throughout development and establishment of the oocyte pool at birth. If atRA is not present at the correct time and at sufficient concentration, meiotic initiation is either delayed or prevented from occurring, ultimately disrupting germ cell development.

All-trans retinoic acid (atRA) is the active metabolite of vitamin A and is involved in regulating a large number of developmental processes (Bushue & Wan, 2010; Ghyselinck & Duester, 2019). atRA is produced in spatial and temporal gradients, and these patterns are maintained by regulated expression of the synthesis and degradation enzymes of the atRA pathway (Kedishvili, 2013).

The presence of atRA in the fetal ovaries induces germ cells to enter meiosis (Spiller et al, 2017). The initiation of meiosis at this time during fetal life is critical for maintenance of the germ line throughout development and establishment of the oocyte pool at birth. If atRA is not present at the correct time and at sufficient concentration, meiotic initiation is either delayed or prevented from occurring, ultimately disrupting germ cell development.

Evidence Supporting this KER

The majority of evidence for this KER comes from rodent studies. In pregnant rats, depletion of vitamin A, the precursor of atRA, leads to an inability of ovarian germ cells to initiate meiosis (Li & Clagett-Dame, 2009).  Further studies in mice have produced strong evidence that atRA acts as a meiosis-inducing factor in oogonia of the ovaries, although there are some conflicting data depending on which techniques are used (Griswold et al, 2012; Spiller & Bowles, 2022). Evidence for the same mechanisms in human fetal ovaries is less substantiated and there may be species differences, particularly the manner in which atRA is made available (reviewed by (Jørgensen & Rajpert-De Meyts, 2014). In humans, evidence to support the KER comes from studies using explanted ovary culture.

In mammalian germ cells, the initiation and progression of meiosis is critically dependent on the expression of Stimulated by retinoic acid gene 8 (Stra8). In mice, deleting Stra8 leads to infertility in both males and females due to meiotic failure (Anderson et al, 2008; Baltus et al, 2006; Mark et al, 2008). What regulates the temporal expression of Stra8, and other factors (such as Rec8 and Dazl) in the germ cells is not completely clear, but there is strong evidence to support an important role for atRA (Bowles et al, 2006; Feng et al, 2021; Griswold et al, 2012; Koubova et al, 2014; Soh et al, 2015).

In the fetal mouse ovary, entry into meiosis, preceded by Stra8 expression, occurs in an overlapping anterior-to-posterior wave from E12.5 (Bowles et al, 2006; Menke et al, 2003). Stra8 is also expressed in rat oogonia at comparative developmental stages to the mouse (Liu et al, 2020). atRA can similarly upregulate Stra8 in vitro, but this is restricted to pluripotent cell lines(Feng et al, 2021; Oulad-Abdelghani et al, 1996; Wang et al, 2016). Culture of mouse skin-derived stem cells with atRA stimulates the formation of functioning gametes and improves oogonia-like cells entry into meiosis (Dyce et al, 2018; Miyauchi et al, 2017). Stra8 expression cannot be induced by atRA in non-pluripotent cell lines, nor in somatic cells in vivo (Feng et al, 2021).

Exposure of pre-meiotic tammar (marsupial) ovaries to atRA induces Stra8 expression and oogonial meiotic entry (Hickford et al, 2017). Culturing fetal mouse ovaries in the presence of atRA increases the number of meiotic oocytes (Livera et al, 2000) and the same phenomenon is observed in cultured human fetal ovaries (Jørgensen et al, 2015). 

In mouse ovaries lacking the atRA synthesizing enzyme ALDH1A1, the onset of germ cell meiosis is delayed (Bowles et al, 2016). This supports a previous study showing that atRA derived from the ovary (rather than mesonephros) is sufficient to initiate meiosis in mice (Mu et al, 2013). In humans, the local synthesis of atRA by ALDH1A enzymes within the ovary may also be involved in meiotic regulation (Childs et al, 2011; Le Bouffant et al, 2010). In two recent studies looking at mouse ovaries lacking all known atRA synthesizing enzymes (Chassot et al, 2020) or RA receptors (Vernet et al, 2020), expression of Stra8 was delayed, albeit some meiosis was still observed in these mice.

Animal models

In vitro/ex vivo

Mouse deletion models for the atRA synthesis enzymes Aldh1a1, Aldh1a2 and Aldh1a3 showed decreased expression of Stra8 in double (Aldh1a2/3) and triple (Aldh1a1/2/3) knockouts, although ultimately some germ cells were observed undergoing meiosis in these ovaries, suggesting that atRA is not essential for meiotic onset or progression(Chassot et al, 2020; Kumar et al, 2011). Similarly, transgenic mice lacking the three atRA nuclear receptors (RAR-α, -β, -γ) showed reduced levels of Stra8, although ultimately some germ cells were observed undergoing meiosis and were capable of producing live offspring (Vernet et al, 2020). Whether or not these models led to impaired fertility (such as sub-fertility) has not been elucidated and the size of their oocyte pools were not determined. In addition, the completeness of the genetic deletions in these models is not clear (discussed in (Spiller & Bowles, 2022)).

Gain of function mouse ovary models for CYP26A1 and CYP26B1 show that CYP26B1 can prevent oocytes from entering meiosis (as assessed by failure to induce Stra8 expression), whereas CYP26A1 does not have the same effect despite being a potent atRA degrading enzyme. This suggests that CYP26B1 works by additional mechanism(s) other than RA degradation (Bellutti et al, 2019).

Quantitative Understanding of the Linkage

The quantitative knowledge pertaining to this KER is very limited as little is known about 1) the levels of endogenous atRA produced in the ovaries in different mammals and 2) the levels of atRA required to achieve meiotic initiation.

In vitro and ex vivo, it has been conclusively shown that low levels (as low as 1uM) of exogenous atRA can induce germ cells to enter meiosis in mice (Bowles et al, 2010) and rats (Livera et al, 2000) and, similarly, that it is necessary to achieve meiosis in in vitro-derived oocytes via primordial germ cells (PGCs)/PGC-like cells (PGCLCs) (Miyauchi et al, 2017). Yet, its exact role in vivo is under debate.

Whilst the relative levels of endogenous atRA produced by the ovary (for any species) remains unknown, similarly, the quantitative relationship between atRA levels and induction of meiosis also remains unclear. As such, the quantitative understanding of how much atRA needs to be reduced to prevent germ cells to enter meiosis in vivo is rated low.

The time-scale for this KER is relatively short, limited to just a couple of days in e.g. mouse models. The induction of meiosis occurs shortly after the germ cells have colonized the ovary and occurs asynchronously (Bullejos & Koopman, 2004) (in mice this begins at E13.5 and is completed for all germ cells 2 days later at E15.5). Proliferation is halted and cells progress through leptonema, zygonema, pachynema, and arrest in diplonema of prophase I prior to birth (Zamboni, 1986). Time and duration of oogenesis varies between species, with rats the shortest duration of only 1-2 days, with other mammals such as pigs, cows, monkeys and humans lasting months (Peters, 1970).

The rat model of vitamin A deficiency (VAD) revealed severe defects to meiosis induction when Vitamin A was restricted/removed from the diet at E10.5, which is just 3 days prior to normal meiotic induction (Li & Clagett-Dame, 2009). Shorter time-frames have not been assessed to date, nor has rescue of VAD during later embryonic time-points been attempted.

No modulating factors are currently known to alter the quantitative relationship between the two KEs.

During development, retinoic acid homeostasis is regulated by feedback loops, as both too much and too little RA can have deleterious effects on the embryo or fetus. The availability of atRA is regulated locally by maintaining a balance between synthesis (ALDH1 enzymes) and metabolism (CYP26 enzymes) (Kedishvili, 2013; Niederreither & Dollé, 2008; Roberts, 2020; Teletin et al, 2017).

The expression of Aldh1a2 and Cyp26a1 can act as part of a negative feedback loop in response to changes in RA levels. Exogenous atRA suppresses expression of Aldh1a2 (Niederreither et al, 1997) whereas blocking atRA signalling increases expression of Aldh1a2. Although Cyp26 expression does not require atRA, addition of atRA greatly increases the expression of Cyp26a1, and conversely, reduced levels of atRA reduces Cyp26a1 expression (de Roos et al, 1999; Hollemann et al, 1998; Ross & Zolfaghari, 2011; Sirbu et al, 2005). Negative feedback loops also extend to the enzymes that convert retinol to all-trans retinaldehyde as well as other related enzymes (Feng et al, 2010; Strate et al, 2009), including Ski, which seem to have cell-type specific roles (Melling et al, 2013; Niederreither & Dollé, 2008).

References

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Aoki T, Takada T (2012) Bisphenol A modulates germ cell differentiation and retinoic acid signaling in mouse ES cells. Reprod Toxicol 34: 463-470

Baltus AE, Menke DB, Hu YC, Goodheart ML, Carpenter AE, de Rooij DG, Page DC (2006) In germ cells of mouse embryonic ovaries, the decision to enter meiosis precedes premeiotic DNA replication. Nat Genet 38: 1430-1434

Bellutti L, Abby E, Tourpin S, Messiaen S, Moison D, Trautmann E, Guerquin MJ, Rouiller-Fabre V, Habert R, Livera G (2019) Divergent Roles of CYP26B1 and Endogenous Retinoic Acid in Mouse Fetal Gonads. Biomolecules 9: 536

Bowles J, Feng CW, Miles K, Inseson J, Spiller CM, Koopman P (2016) ALDH1A1 provides a source of meiosis-inducing retinoic acid in mouse fetal ovaries. Nat Commun 7: 10845

Bowles J, Feng CW, Spiller CM, Davidson TL, Jackson A, Koopman P (2010) FGF9 suppresses meiosis and promotes male germ cell fate in mice. Dev Cell 19: 440-449

Bowles J, Knight D, Smith C, Wilhelm D, Richman J, Mamiya S, Yashiro K, Chawengsaksophak K, Wilson MJ, Rossant J, Hamada H, Koopman P (2006) Retinoid signaling determines germ cell fate in mice. Science 312: 596-600

Bullejos M, Koopman P (2004) Germ cells enter meiosis in a rostro-caudal wave during development of the mouse ovary. Mol Reprod Dev 68: 422-428

Bushue N, Wan YJY (2010) Retinoid pathway and cancer therapeutics. Adv Drug Deliv Rev 62: 1285-1298

Chassot AA, Le Rolle M, Jolivet G, Stevant I, Guigonis JM, Da Silva F, Nef S, Pailhoux E, Schedl A, Ghyselinck NB, Chaboissier MC (2020) Retinoic acid synthesis by ALDH1A proteins is dispensable for meiosis initiation in the mouse fetal ovary. Sci Adv 6: eaaz1261

Childs AJ, Cowan G, Kinnell HL, Anderson RA, Saunders PTK (2011) Retinoic Acid signalling and the control of meiotic entry in the human fetal gonad. PLoS One 6: e20249

de Roos K, Sonneveld E, Compaan B, ten Berge D, Durston AJ, van der Saag PT (1999) Expression of retinoic acid 4-hydroxylase (CYP26) during mouse and Xenopus laevis embryogenesis. Mech Dev 82: 205-211

Dyce PW, Tenn N, Kidder GM (2018) Retinoic acid enhances germ cell differentiation of mouse skin-derived stem cells. J Ovarian Res 11: 19

Feng CW, Burnet G, Spiller CM, Cheung FKM, Chawengsaksophak K, Koopman P, Bowles J (2021) Identification of regulatory elements required for Stra8 expression in fetal ovarian germ cells of the mouse. Development 148: dev194977

Feng L, Hernandez RE, Waxman JS, Yelon D, Moens CB (2010) Dhrs3a regulates retinoic acid biosynthesis through a feedback inhibition mechanism. Dev Biol 338: 1-14

Ghyselinck NB, Duester G (2019) Retinoic acid signaling pathways. Development 146: dev167502

Griswold MD, Hogarth CA, Bowles J, Koopman P (2012) Initiating meiosis: the case for retinoic acid. Biol Reprod 86: 35

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Hollemann T, Chen Y, Grunz H, Pieler T (1998) Regionalized metabolic activity establishes boundaries of retinoic acid signalling. EMBO J 17: 7361-7372

Jørgensen A, Nielsen JE, Perlman S, Lundvall L, Mitchell RT, Juul A, Rajpert-De Meyts E (2015) Ex vivo culture of human fetal gonads: manipulation of meiosis signalling by retinoic acid treatment disrupts testis development. Hum Reprod 30: 2351-2363

Jørgensen A, Rajpert-De Meyts E (2014) Regulation of meiotic entry and gonadal sex differentiation in the human: normal and disrupted signaling. Biomol Concepts 5: 331-341

Kedishvili NY (2013) Enzymology of retinoic acid biosynthesis and degradation. J Lipid Res 54: 1744-1760

Koubova J, Hu YC, Bhattacharyya T, Soh YQS, Gill ME, Goodheart ML, Hogarth CA, Griswold MD, Page DC (2014) Retinoic acid activates two pathways required for meiosis in mice. PLoS Genet 10: e1004541

Koubova J, Menke DB, Zhou Q, Capel B, Griswold MD, Page DC (2006) Retinoic acid regulates sex-specific timing of meiotic initiation in mice. Proc Natl Acad Sci U S A 103: 2474-2479

Kumar S, Chatzi C, Brade T, Cunningham TJ, Zhao X, Duester G (2011) Sex-specific timing of meiotic initiation is regulated by Cyp26b1 independent of retinoic acid signalling. Nat Commun 2: 151

Le Bouffant R, Guerquin MJ, Duquenne C, Frydman N, Coffigny H, Rouiller-Fabre V, Frydman R, Habert R, Livera G (2010) Meiosis initiation in the human ovary requires intrinsic retinoic acid synthesis. Hum Reprod 25: 2579-2590

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Relationship: 2481: Disrupted, meiotic initiation in oocyte leads to Decreased, ovarian reserve

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

Female mammals during fetal life, as this corresponds to initiation of meiosis prophase I in oocytes of the ovaries.

Key Event Relationship Description

The establishment of the primordial follicle pool is a multistep process that spans from early fetal life to reproductive maturity. This period of time varies greatly between species, lasting only a few weeks in mice and rats, but years in humans (Tingen et al, 2009). One important process is for the mitotic primordial germs cells to enter meiosis prior to cyst formation (Findlay et al, 2015; Tingen et al, 2009). Notably, in females there is a massive loss of oocytes between cyst formation and time of maturity, and the exact mechanisms behind this oocyte degradation is not well understood (Findlay et al, 2015; Sun et al, 2017).

Evidence Supporting this KER

It is well established that disruption to meiosis during oocyte development can lead to     sub-/infertility in females at reproductive age. There are numerous gene mutation in mice showing links between meiotic defects and fertility phenotypes, as well as associations to female fertility phenotypes in humans (Adelfalk et al, 2011).

Although the entry into meiosis is required for oocyte development, the relationship between meiotic entry and final oocyte reserve remains unclear. However, there are strong correlations between disrupted meiosis and infertility (or aneuploidy) in females (Handel & Schimenti, 2010). For instance, in mice, ablation of Stra8 prevents oocytes from entering meiosis in the fetal ovaries and mature females are infertile (Baltus et al, 2006; Zhou et al, 2008). Mutation in Atm, a gene involved in recombination during meiosis, results in complete loss of primary oocytes in mice, and greatly reduced follicle pool in humans (Adelfalk et al, 2011; Agamanolis & Greenstein, 1979; Aguilar et al, 1968; Xu et al, 1996). Other examples include Fanca and Fancd2 genes that are involved in recombination. Mutations to these genes lead to oocyte degeneration and subfertility in mice (Cheng et al, 2000; Houghtaling et al, 2003; Wong et al, 2003).

Mice with Lhx8 ablation display total loss of oocytes. Lhx8-/- mice maintain oocytes during fetal development, but loose the oocytes shortly after birth by autophagy, likely because the oocytes have failed to enter meiosis in utero (Choi et al, 2008; D'Ignazio et al, 2018). Fzr1 is a regulator of mitotic cell division. When conditionally ablated from the germ cells, female mice display premature ovarian failure by 5 months of age and are subfertile; oocytes are lost in utero during early meiotic prophase I (Holt et al, 2014).

CYP51 (lanosterol 14 α-demethylase) is expressed by fetal oocytes and is involved in meiotic regulation (Mu et al, 2018).  Inhibition of CYP51 activity reduces the formation of primordial follicles (Zhang et al, 2009) by disrupting entry into diplotene stage (Mu et al, 2018).

The mechanisms and outcomes of meiosis I disruption may vary significantly across species, making it challenging to generalize findings from animal models to humans. Also, the extent of disruption that is required to significantly affect the ovarian reserve remains uncertain, as there may be potential threshold effects influenced by genetic, epigenetic, and environmental factors.

Timing of disruption to meiosis I initiation in oocytes may influence to what extent the ovary reserve is ultimately affected.

Methods to quantify the ovarian reserve (e.g. follicle count, AMH levels) may not directly reflect the impact of meiotic disruption, leading to inconsistencies in observed effects.

Quantitative Understanding of the Linkage

The quantitative understanding of this KER remains poorly understood, not least because the quantification of actual oocyte numbers at various stages of development are very difficult to perform.

The ovarian follicle pool (ovarian reserve) refers to the final number of primordial follicles in the mature ovary and is established through a series of events. In most mammals, it is determined during gestation or just after birth and relies on i) how many germ cells were established during embryogenesis, ii) their proliferation during migration and early ovary development, iii) death rate during oogenesis and iv) formation of primordial follicles at nest breakdown (Findlay et al, 2015). The last two stages, which includes nest formation and breakdown, is largely influenced by the mitotic-meiotic transition, in that oocytes that have failed to enter meiosis may contribute to the cysts population, but only high quality oocytes in meiotic prophase are spared during cyst breakdown (Findlay et al, 2015). Thus, there is a response-response relationship between meiotic entry and final follicle pool, albeit the quantitative relationship is not that well understood.

The time-scale for oocyte mitotic-meiotic transition and subsequent nest breakdown varies between species, but generally takes place from mid gestation to around the time of birth. In mice, meiosis and nest formation is initiated from around E13, whereas in humans it initiates at around GW12-14 (Childs et al, 2012; Findlay et al, 2015; Grive & Freiman, 2015; Pepling, 2006; Tingen et al, 2009). Nest breakdown starts just before birth in mice and completes around postnatal day 5 (Grive & Freiman, 2015; Pepling, 2006). In humans, nest breakdown takes place during second trimester (Grive & Freiman, 2015; Tingen et al, 2009). 

Variations in genes involved in meiotic regulation (e.g., SYCP3, MSH5, DAZL) may influence sensitivity to disruptions, including species differences, as may variations in epigenetic status of oocytes.   

No (known) relevant feedback loop.

References

Adelfalk C, Ahmed EA, Scherthan H (2011) Reproductive Phenotypes of Mouse Models Illuminate Human Infertility. J Reproduktionsmed Endokrinol 8: 376-383

Agamanolis DP, Greenstein JI (1979) Ataxia-telangiectasia. Report of a case with Lewy bodies and vascular abnormalities within cerebral tissue. J Neuropathol Exp Neurol 38: 475-489

Aguilar MJ, Kamoshita S, Landing BH, Boder E, Sedgwick RP (1968) Pathological observations in ataxia-telangiectasia. A report of five cases. J Neuropathol Exp Neurol 27: 659-676

Baltus AE, Menke DB, Hu YC, Goodheart ML, Carpenter AE, de Rooij DG, Page DC (2006) In germ cells of mouse embryonic ovaries, the decision to enter meiosis precedes premeiotic DNA replication. Nat Genet 38: 1430-1434

Cheng NC, van de Vrugt HJ, van der Valk MA, Oostra AB, Krimpenfort P, de Vries Y, Joenje H, Berns A, Arwert F (2000) Mice with a targeted disruption of the Fanconi anemia homolog Fanca. Hum Mol Genet 9: 1805-1811

Childs AJ, Kinnell HL, He J, Anderson RA (2012) LIN28 is selectively expressed by primordial and pre-meiotic germ cells in the human fetal ovary. Stem Cells Dev 21: 2343-2349

Choi Y, Ballow DJ, Xin Y, Rajkovic A (2008) Lim homeobox gene, lhx8, is essential for mouse oocyte differentiation and survival. Biol Reprod 79: 442-449

D'Ignazio L, Michel M, Beyer M, Thompson K, Forabosco A, Schlessinger D, Pelosi E (2018) Lhx8 ablation leads to massive autophagy of mouse oocytes associated with DNA damage. Biol Reprod 98: 532-542

Dean A, van den Driesche S, Wang Y, McKinnell C, Macpherson S, Eddie SL, Kinnell HL, Hurtado-Gonzalez P, Chambers TJ, Stevenson K, Wolfinger E, Hrabalkova L, Calarrao A, Bayne RA, Hagen CP, Mitchell RT, Anderson RA, Sharpe RM (2016) Analgesic exposure in pregnant rats affects fetal germ cell development with inter-generational reproductive consequences. Sci Rep 6: 19789

Findlay JK, Hutt KJ, Hickey M, Anderson RA (2015) How Is the Number of Primordial Follicles in the Ovarian Reserve Established? Biol Reprod 93: 111

Grive KJ, Freiman RN (2015) The developmental origins of the mammalian ovarian reserve. Development 142: 2554-2563

Handel MA, Schimenti JC (2010) Genetics of mammalian meiosis: regulation, dynamics and impact on fertility. Nat Rev Genet 11: 124-136

Holt JE, Pye V, Boon E, Stewart JL, García-Higuera I, Moreno S, Rodríguez R, Jones KT, McLaughlin EA (2014) The APC/C activator FZR1 is essential for meiotic prophase I in mice. Development 141: 1354-1365

Houghtaling S, Timmers C, Noll M, Finegold MJ, Jones SN, Meyn MS, Grompe M (2003) Epithelial cancer in Fanconi anemia complementation group D2 (Fancd2) knockout mice. Genes Dev 17: 2021-2035

Mu X, Wen J, Chen Q, Wang Z, Wang Y, Guo M, Yang Y, Xu J, Wei Z, Xia G, Yang M, Wang C (2018) Retinoic acid-induced CYP51 nuclear translocation promotes meiosis prophase I process and is correlated to the expression of REC8 and STAG3 in mice. Biol Open 7: bio035626

Pepling ME (2006) From primordial germ cell to primordial follicle: mammalian female germ cell development. Genesis 44: 622-632

Sun YC, Sun XF, Dyce PW, Shen W, Chen H (2017) The role of germ cell loss during primordial follicle assembly: a review of current advances. Int J Biol Sci 13: 449-457

Tingen C, Kim A, Woodruff TK (2009) The primordial pool of follicles and nest breakdown in mammalian ovaries. Mol Hum Reprod 15: 795-803

Wong JCY, Alon N, Mckerlie C, Huang JR, Meyn MS, Buchwald M (2003) Targeted disruption of exons 1 to 6 of the Fanconi Anemia group A gene leads to growth retardation, strain-specific microphthalmia, meiotic defects and primordial germ cell hypoplasia. Hum Mol Genet 12: 2063-2076

Xu Y, Ashley T, Brainerd EE, Bronson RT, Meyn MS, Baltimore D (1996) Targeted disruption of ATM leads to growth retardation, chromosomal fragmentation during meiosis, immune defects, and thymic lymphoma. Genes Dev 10: 2411-2422

Zhang H, Xu B, Xie H, Zhou B, Quyang H, Ning G, Li G, Zhang M (2009) Lanosterol metabolic product(s) is involved in primordial folliculogenesis and establishment of primordial follicle pool in mouse fetal ovary. Mol Reprod Dev 76: 514-521

Zhang HQ, Zhang XF, Zhang LJ, Chao HH, Pan B, Feng YM, Li L, Sun XF, Shen W (2012) Fetal exposure to bisphenol A affects the primordial follicle formation by inhibiting the meiotic progression of oocytes. Mol Biol Rep 39: 5651-5657

Zhang XF, Zhang T, Han Z, Liu JC, Liu YP, Ma JY, Li L, Shen W (2015) Transgenerational inheritance of ovarian development deficiency induced by maternal diethylhexyl phthalate exposure. Reprod Fertil Dev 27: 1213-1221

Zhou Q, Nie R, Li Y, Friel P, Mitchell D, Hess RA, Small C, Griswold MD (2008) Expression of Stimulated by Retinoic Acid Gene 8 (Stra8) in Spermatogenic Cells Induced by Retinoic Acid: An In Vivo Study in Vitamin A-Sufficient Postnatal Murine Testes. Biol Reprod 79: 35-42

Relationship: 2525: Decreased, ovarian reserve leads to disrupted, ovarian cycle

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

Applicable for (mammalian) females during postnatal life. Although diminished ovary reserve was caused by disruption during fetal development, the link between reduced ovary reserve and irregular cycling occurs postnatally during reproductive ages.

Key Event Relationship Description

Reduced ovarian reserve, meaning the finite pool of primordial follicles containing the immature oocytes, is leading to ovarian cycle irregularities. Cycle irregularities include disturbances of the ovarian cycle like shorter cycle and prolonged estrus and/or ovulation problems like deferred ovulation and anovulation. This KER is considered canonical information.

Evidence Supporting this KER

All primordial follicles are formed during early development (fetal period in humans, perinatally in rodents) and can stay dormant for long periods of time (years in humans, months in rodents). The stock of primordial dormant follicles constitutes the ‘ovarian reserve’. In humans, millions of follicles are formed by mid-gestation (Wallace & Kelsey, 2010). Upon puberty, the hypothalamus-pituitary-ovary (HPO) axis matures enabling the primordial follicles to grow into maturity in a process called folliculogenesis, which serves two functions: i) secretion of steroid hormones that enable pregnancy, and ii) production of mature oocytes that are ovulated for possible fertilization.

Cohorts of primordial follicles continuously enter the growing pool. After puberty, growing antral follicles are recruited for final maturation during each menstrual cycle (estrus cycle in rodents) by gonadotropins secreted from the pituitary gland but only a limited number will reach maturity and ovulate oocytes (typically one in humans and 10-20 in rodents). The majority of follicles never reach maturity and instead die in a process called atresia. Therefore, the ovarian reserve is irreversibly depleted with age. When the reserve is depleted to a level that cannot faithfully maintain steroid production, fertility ceases. In humans, fertility ends in sterility at menopause when less than 1,000 follicles remain (Wallace & Kelsey, 2010).

Regular cycles are considered as an indicator of reproductive health and often used in animal studies as the earliest biomarker to reflect disruption of fertility and ovotoxicity (Hooser et al, 1994). OECD test guidelines 407, 416 and 443 include rodent cyclicity as an endpoint to assess reproductive toxicity (OECD, 2001; OECD, 2008; OECD, 2018). In humans, normal menstrual cycle lasts 28-35 days, and in rodents 4-6 days. When the ovarian reserve is depleted to a critically low level, like naturally during the perimenopausal period in humans, the variability of the cycle length increases with many of the cycles being anovulatory. The lower the ovarian reserve is, the lower the probability of growing follicles to exist at any given time point. Since the growing follicles produce steroid hormones that are essential for cyclicity, the lack of growing follicles leads to disturbances of the HPO axis. Therefore, the lower the ovarian reserve is, the less probable are regular cycles; reflected by regular menstruation in humans (O'Connor et al, 1998; O'Connor et al, 2001).

Supporting evidence exists. Anti-Müllerian hormone (AMH) is a growth factor secreted by growing follicles. Low levels of AMH correlate with longer cycle length (Harris et al, 2021). Other studies have connected AMH or antral follicle count (AFC), another ovarian reserve marker, to shorter cycles (Younis et al, 2020). A systematic review and meta-analysis have revealed in regularly cycling women, a shorter cycle is associated with lower ovarian reserve based on AMH or AFC (Younis et al, 2020). Young women diagnosed with premature ovarian failure have also reported shorter cycles (Guzel et al, 2017). In addition, it is well established in humans that diminishing ovarian reserve leads to perimenopause (a period or irregular cycles) and eventually menopause (complete cessation of cycles).

Stressors that are known to deplete the ovarian reserve include cancer treatments, which kill primordial follicles and disrupt folliculogenesis. Alkylating chemotherapy agents like cyclophosphamide and cisplatin are highly ovotoxic, as well as radiation therapy towards ovaries (Pampanini et al, 2020). Therapies based on high dose alkylators and radiation towards ovaries lead with high likelihood to amenorrhea, infertility and premature ovarian insufficiency in humans due to depletion of ovarian reserve and therefore constitute an indication for clinical fertility preservation (ESHRE_Guideline_Group_on_Female_Fertility_Preservation et al, 2020; Pampanini et al, 2020).

In mice, it has been shown that cisplatin induces primordial follicle loss in a dose-depended manner (Meirow et al, 1999). Cyclophosphamide can accelerate primordial follicle loss in mice and in human ovarian tissue (Kalich-Philosoph et al., 2013; Lande et al., 2017). In human xenografts in mice, the same compound decreased the primordial follicle density(Oktem & Oktay, 2007). These key studies indicate the effect of chemotherapy compounds on the size of the ovarian reserve (Meirow et al, 2010). In treated patients, although there is often no direct information on the size of the ovarian reserve, a rapid decrease of AMH levels has been observed following high risk chemotherapy. Patients receiving chemotherapy have also been shown to have an increased risk of premature ovarian failure. In addition, some studies have shown that the number of healthy follicles is significantly decreased and that of atretic follicles increased in human ovarian tissue following alkylating chemotherapy (Pampanini et al, 2019). These data establish a clear indication of diminished ovarian reserve following chemotherapy. Importantly, chemotherapy compounds also affect the menstrual cycle, with patients experiencing amenorrhea and irregular cycles (Jacobson et al., 2016; Meirow et al., 2010).

Another stressor known to affect the ovarian reserve is smoking. Cigarette smoke contains thousands of chemicals, several of which have been shown to be ovotoxic (Budani & Tiboni, 2017). Mice exposed in vivo to cigarette smoke have significantly fewer primordial follicles compared to the control group (Tuttle et al., 2009). Smoking women display reduced ovarian reserve markers and experience irregular cycles compared to non-smokers of the same age group (El-Nemr et al, 1998; Sharara et al, 1994).

Additional chemical insults affecting the ovarian reserve and causing menstrual cycle irregularities are presented in Table 1. These studies in animal models demonstrate how these chemicals directly target primordial follicles and cause menstrual cycle irregularities. Vinylcyclohexene diepoxide (VCD), metabolite of 4-vinylcyclohexene (VCH), is the most commonly used chemical in these studies and is often used to induce reproductive senescence in model organisms.

Table 1: In vivo studies demonstrating that effects on the KE upstream affect the KE downstream. VCD: vinylcyclohexene diepoxide, VCH: 4-vinylcyclohexene, BPA: bisphenol A, DEHP: bis(2-ethylhexyl) phthalate, B[a]P: Benzo[a]pyrene

As mentioned, several chemotherapy agents damage ovarian reserve and disrupt folliculogenesis. However, it has been shown that regular menses can resume upon treatment cessation (Jacobson et al, 2016). Therefore, in this case reduced ovarian reserve did not lead to permanent irregularities of ovarian cycle. In a systematic review and meta-analysis investigating the connection between the ovarian reserve and the length of the menstrual cycle, studies are mentioned where reduced ovarian reserve markers did not associate with irregular menstrual cycles (Younis et al, 2020). Several factors affect the impact of chemotherapy on ovarian health in humans, including the age at the treatment, size of ovarian reserve at treatment, and treatment regimen. However, late side effects of chemotherapy often include amenorrhea, premature ovarian insufficiency, and infertility.

Menstrual irregularities can be caused by factors other than reduced ovarian reserve. The most common factor affecting cyclicity is HPO axis dysregulation causing hypothalamic amenorrhea (Berga & Naftolin, 2012). Another example is the contraceptive pill that decreases gonadotropin secretion by the pituitary gland, leading to inhibition of folliculogenesis and amenorrhea. Changes in hormone levels produced by the pituitary gland have also been connected to shorter and anovulatory cycles (Mumford et al., 2012). Another factor affecting cyclicity is the thyroid gland function. Thyroid function disturbances, like hypo and hyperthyroidism have been connected to menstrual disturbances (Koutras, 1997).

Quantitative Understanding of the Linkage

poor

The timescale at which disruption in cyclicity occurs depends on the type of follicles that are affected, size of the reserve at the time of insult, and the extent of the damage. When a stressor targets selectively the ovarian reserve, it might take months (or years in humans) for the disruptions in cyclicity to be observed (Hoyer & Sipes, 1996). This delay was evident in some of the animal studies mentioned in Table 1 (Hooser et al., 1994; Lohff et al., 2006; Mayer et al., 2002).  

The size of the ovarian reserve at the time of stressor exposure is a factor that can affect the response-response relationship of this KER. Therefore, age can also be a modulating factor, as observed in the animal study mentioned in table 1, where even though all treated rats exhibited reduction in the ovarian reserve, irregular cycles were only observed in the adult ones but not the immature ones (Flaws et al., 1994). In addition, chemotherapy effects on fertility tend to be more severe with increasing age due to a smaller ovarian reserve (Jacobson et al, 2016).

Changes in hormones can affect menstrual/estrus cyclicity, without being connected to the size of the ovarian reserve. For instance, experiencing stress has been shown to affect the hypothalamus-pituitary-adrenal axis (HPA) activity. A high body mass index (BMI) has been shown to affect sex hormone-binding globulin (SHBG), free androgen index (FAI), testosterone, and insulin levels. Smoking, although it can also affect the reserve, can cause hypoestrogenism. Therefore, stress, obesity and smoking can affect menstrual cyclicity and influence the response-response relationship of this KER (Bae et al, 2018).

HPO axis regulates estrus/menstrual cycle, and is based on positive and negative feedback loops by ovarian steroids and peptide hormones, and hormones released by the hypothalamus and pituitary gland.

References

Bae J, Park SU, Kwon JW (2018) Factors associated with menstrual cycle irregularity and menopause. BMC Womens Health 18: 36

Berga S, Naftolin F (2012) Neuroendocrine control of ovulation. Gynecol Endocrinol 28 Suppl 1: 9-13

Budani MC, Tiboni GM (2017) Ovotoxicity of cigarette smoke: A systematic review of the literature. Reprod Toxicol 72: 164-181

El-Nemr A, Al-Shawaf T, Sabatini L, Wilson C, Lower AM, Grudzinskas JG (1998) Effect of smoking on ovarian reserve and ovarian stimulation in in-vitro fertilization and embryo transfer. Hum Reprod 13: 2192-2198

ESHRE_Guideline_Group_on_Female_Fertility_Preservation, Anderson RA, Amant F, Braat D, D'Angelo A, de Sousa Lopes SMC, Demeestere I, Dwek S, Frith L, Lambertini M, Maslin C, Moura-Ramos M, Nogueira D, Rodriguez-Wallberg K, Vermeulen N (2020) ESHRE guideline: female fertility preservation Hum Reprod Update 2020: hoaa052

Flaws JA, Doerr JK, Sipes IG, Hoyer PB (1994) Destruction of preantral follicles in adult rats by 4-vinyl-1-cyclohexene diepoxide. Reprod Toxicol 8: 509-514

Guzel Y, Aba YA, Yakin K, Oktem O (2017) Menstrual cycle characteristics of young females with occult primary ovarian insufficiency at initial diagnosis and one-year follow-up with serum amh level and antral follicle count. PLoS One 12: e0188334

Hannon PR, Peretz J, Flaws JA (2014) Daily exposure to Di(2-ethylhexyl) phthalate alters estrous cyclicity and accelerates primordial follicle recruitment potentially via dysregulation of the phosphatidylinositol 3-kinase signaling pathway in adult mice. Biol Reprod 90: 136

Harris BS, Steiner AZ, Jukic AM (2021) Ovarian Reserve Biomarkers and Menstrual Cycle Length in a Prospective Cohort Study. J Clin Endocrinol Metab 106: e3748-e3759

Hooser SB, Douds DP, DeMerell DG, Hoyer PB, Sipes IG (1994) Long-term ovarian and gonadotropin changes in mice exposed to 4-vinylcyclohexene. Reprod Toxicol 8: 315-323

Hoyer PB, Sipes IG (1996) Assessment of follicle destruction in chemical-induced ovarian toxicity. Annu Rev Pharmacol Toxicol 36: 307-331

Hu Y, Yuan DZ, Wu Y, Yu LL, Xu LZ, Yue LM, Liu L, Xu WM, Qiao XY, Zeng RJ, Yang ZL, Yin WY, Ma YX, Nie Y (2018) Bisphenol A Initiates Excessive Premature Activation of Primordial Follicles in Mouse Ovaries via the PTEN Signaling Pathway. Reprod Sci 25: 609-620

Jacobson MH, Mertens AC, Spencer JB, Manatunga AK, Howards PP (2016) Menses resumption after cancer treatment-induced amenorrhea occurs early or not at all. Fertil Steril 105: 765-772

Kalich-Philosoph, L., Roness, H., Carmely, A., Fishel-Bartal, M., Ligumsky, H., Paglin, S., Wolf, I., Kanety, H., Sredni, B., & Meirow, D. (2013). Cyclophosphamide triggers follicle activation and "burnout "; AS101 prevents follicle loss and preserves fertility. Science Translational Medicine, 5(185).

Koutras, D. A. (1997). Disturbances of menstruation in thyroid disease. Annals of the New York Academy of Sciences, 816, 280–284. 

Lande, Y., Fisch, B., Tsur, A., Farhi, J., Prag-Rosenberg, R., Ben-Haroush, A., Kessler-Icekson, G., Zahalka, M. A., Ludeman, S. M., & Abir, R. (2017). Short-term exposure of human ovarian follicles to cyclophosphamide metabolites seems to promote follicular activation in vitro. Reproductive BioMedicine Online, 34(1), 104–114. 

Lohff JC, Christian PJ, Marion SL, Arrandale A, Hoyer PB (2005) Characterization of cyclicity and hormonal profile with impending ovarian failure in a novel chemical-induced mouse model of perimenopause. Comp Med 55: 523-527

Lohff JC, Christian PJ, Marion SL, Hoyer PB (2006) Effect of duration of dosing on onset of ovarian failure in a chemical-induced mouse model of perimenopause. Menopause 13: 482-488

Mayer LP, Devine PJ, Dyer CA, Hoyer PB (2004) The follicle-deplete mouse ovary produces androgen. Biol Reprod 71: 130-138

Mayer LP, Pearsall NA, Christian PJ, Devine PJ, Payne CM, McCuskey MK, Marion SL, Sipes IG, Hoyer PB (2002) Long-term effects of ovarian follicular depletion in rats by 4-vinylcyclohexene diepoxide. Reprod Toxicol 16: 775-781

Meirow D, Biederman H, Anderson RA, Wallace WHB (2010) Toxicity of chemotherapy and radiation on female reproduction. Clin Obstet Gynecol 53: 727-739

Meirow D, Lewis H, Nugent D, Epstein M (1999) Subclinical depletion of primordial follicular reserve in mice treated with cyclophosphamide: clinical importance and proposed accurate investigative tool. Hum Reprod 14: 1903-1907

Mumford, S. L., Steiner, A. Z., Pollack, A. Z., Perkins, N. J., Filiberto, A. C., Albert, P. S., Mattison, D. R., Wactawski-Wende, J., & Schisterman, E. F. (2012). The utility of menstrual cycle length as an indicator of cumulative hormonal exposure. Journal of Clinical Endocrinology and Metabolism, 97(10). 

O'Connor KA, Holman DJ, Wood JW (1998) Declining fecundity and ovarian ageing in natural fertility populations. Maturitas 30: 127-136

O'Connor KA, Holman DJ, Wood JW (2001) Menstrual cycle variability and the perimenopause. Am J Hum Biol 13: 465-478

OECD. (2001) Test No. 416: Two-Generation Reproduction Toxicity, OECD Guidelines for the Testing of Chemicals, Section 4. OECD Publishing, Paris.

OECD. (2008) Test No. 407: Repeated Dose 28-day Oral Toxicity Study in Rodents. OECD Publishing, Paris.

OECD. (2018) Test No. 443: Extended One-Generation Reproductive Toxicity Study, OECD Guidelines for the Testing of Chemicals, Section 4. OECD Publishing, Paris.

Oktem O, Oktay K (2007) A novel ovarian xenografting model to characterize the impact of chemotherapy agents on human primordial follicle reserve. Cancer Res 67: 10159-10162

Pampanini V, Hassan J, Oliver E, Stukenborg JB, Damdimopoulou P, Jahnukainen K (2020) Fertility Preservation for Prepubertal Patients at Risk of Infertility: Present Status and Future Perspectives. Horm Res Paediatr 93: 599-608

Pampanini V, Wagner M, Asadi-Azarbaijani B, Oskam IC, Sheikhi M, Sjödin MOD, Lindberg J, Hovatta O, Sahlin L, Björvang RD, Otala M, Damdimopoulou P, Jahnukainen K (2019) Impact of first-line cancer treatment on the follicle quality in cryopreserved ovarian samples from girls and young women. Hum Reprod 34: 1674-1685

Sharara FI, Beatse SN, Leonardi MR, Navot D, Scott Jr RT (1994) Cigarette smoking accelerates the development of diminished ovarian reserve as evidenced by the clomiphene citrate challenge test. Fertil Steril 62: 257-262

Tuttle AM, Stämpfli M, Foster WG (2009) Cigarette smoke causes follicle loss in mice ovaries at concentrations representative of human exposure. Hum Reprod 24: 1452-1459

Wallace WHB, Kelsey TW (2010) Human ovarian reserve from conception to the menopause. PLoS One 5: e8772

Windham GC, Elkin EP, Swan SH, Waller KO, Fenster L (1999) Cigarette smoking and effects on menstrual function. Obstet Gynecol 93: 59-65

Xu C, Chen JA, Qiu Z, Zhao Q, Luo J, Yang L, Zeng H, Huang Y, Zhang L, Cao J, Shu W (2010) Ovotoxicity and PPAR-mediated aromatase downregulation in female Sprague-Dawley rats following combined oral exposure to benzo[a]pyrene and di-(2-ethylhexyl) phthalate. Toxicol Lett 199: 323-332

Younis JS, Iskander R, Fauser BMJM, Izhaki I (2020) Does an association exist between menstrual cycle length within the normal range and ovarian reserve biomarkers during the reproductive years? A systematic review and meta-analysis. Hum Reprod Update 26: 904-928

Relationship: 394: disrupted, ovarian cycle leads to decreased, Fertility

AOPs Referencing Relationship

Evidence Supporting Applicability of this Relationship

In many instances, human female reproductive toxicity of an agent is suspected based on studies performed in experimental animals. The neuroendocrinology, steroid biochemistry, and other physiologic events in the females of most small experimental species often used (mouse, rat, hamster) are similar in their susceptibility to disruption by toxicants (Massaro, 1997).

Although the assessment of the human ovarian cycle may have a variety of biomarkers distinct from those in rats, many of the underlying endocrine mechanisms associated with successful follicular development, ovulation, pregnancy, and parturition are homologous between the two (for review see (Bretveld et al., 2006). For this reason, a toxicant-induced perturbation of ovarian cycles in female rats suggest that a compound may function as a reproductive toxicant in human females.

Mice

• environmental air pollution (Mohallem et al., 2005)
• phthalates (DEHP)
• abortion rate of 100% in F0 dams in the 500-mg/kg/day was observed, in F1 females found that the total number of F2 embryos (exposed to DEHP only as germ cells) was not impaired. However, in the 0.05- and 5-mg DEHP groups, 28% and 29%, respectively, of the blastocysts were degenerated, compared with 8% of controls (Schmidt et al., 2012).
• Lamb et al. studied fertility effects of DEHP in mice (both sexes) and found that DEHP caused dose-dependent decreases in fertility. DBP exposure resulted in a reduction in the numbers of litters per pair and of live pups per litter and in the proportion of pups born alive at the 1.0% amount, but not at lower dose levels. A crossover mating trial demonstrated that female mice, but not males, were affected by DBP, as shown by significant decreases in the percentage of fertile pairs, the number of live pups per litter, the proportion of pups born alive, and live pup weight. DHP in the diet resulted in dose-related adverse effects on the numbers of litters per pair and of live pups per litter and proportion of pups born alive at 0.3, 0.6, and 1.2% DHP in the diet. A crossover mating study demonstrated that both sexes were affected. DEHP (at 0.1 and 0.3%) caused dose-dependent decreases in fertility and in the number and the proportion of pups born alive. A crossover mating trial showed that both sexes were affected by exposure to DEHP. These data demonstrate the ability of the continuous breeding protocol to discriminate the qualitative and quantitative reproductive effects of the more and less active congeners as well as the large differences in reproductive toxicity attributable to subtle changes in the alkyl substitution of phthalate esters (Lamb et al., 1987).

Rat phthalates (DEHP)

• female rats exposed to a high dose of DEHP (3,000 mg/kg/day) had irregular estrous cycles and a slight decline in pregnancy rate (Takai et al., 2009). At 1,000 mg/kg bw/day over a period of 4 weeks did not disturb female fertility or early embryo development.

• There was significant evidence that 5, 15, 50, and 400 mg /kg/day females differed from the control females in the relative amount of time spent in oestrous stages, however no changes were revealed in the number of females with regular cycles, cycle length, number of cycles, and in number of cycling females across the dose groups as compared to the control females The litter size (number of live pups) produced by the P0 generation was significantly reduced in the 400 mg/kg/day dose group (Blystone et al., 2010).

Human

Studies showing a correlation between decreased fertility and;

• professional activity (Olsen, 1994)
• phthalates (DEHP) In occupationally exposed women to high concentration of phthalates exhibit hypoestrogenic anovulary cycles and was associated with decreased pregnancy rate and higher miscarriage rates (Aldyreva,M.V.,Klimove,T.S.,Iziumova,A.S.,Timofeevskaia,L.A., 1975).
• smoking (Hull, North, Taylor, Farrow, & Ford, 2000)
• the use of certain drugs or radiation exposure (Dobson & Felton, 1983)

For the taxonomic applicability see also the Table 1.

Key Event Relationship Description

The ovarian cycle irregularities impact on reproductive capacity of the females that may result in impaired fertility:

1. Irregular cycles may reflect impaired ovulation. Extended vaginal estrus usually indicates that the female cannot spontaneously achieve the ovulatory surge of LH (Huang and Meites, 1975). The persistence of regular vaginal cycles after treatment does not necessarily indicate that ovulation occurred, because luteal tissue may form in follicles that have not ruptured. However, that effect should be reflected in reduced fertility. Conversely, subtle alterations of cyclicity can occur at doses below those that alter fertility (Gray et al., 1989).

2. Persistent or constant vaginal cornification (or vaginal estrus) may result from one or several effects. Typically, in the adult, if the vaginal epithelium becomes cornified and remains so in response to toxicant exposure, it is the result of the agent’s estrogenic properties (i.e., DES or methoxychlor), or the ability of the agent to block ovulation. In the latter case, the follicle persists and endogenous estrogen levels bring about the persistent vaginal cornification. Histologically, the ovaries in persistent estrus will be atrophied following exposure to estrogenic substances. In contrast, the ovaries of females in which ovulation has been blocked because of altered gonadotropin secretion will contain several large follicles and no corpora lutea. Females in constant estrus may be sexually receptive regardless of the mechanism responsible for this altered ovarian condition. However, if ovulation has been blocked by the treatment, an LH surge may be induced by mating (Brown-Grant et al., 1973; Smith, E.R. and Davidson, 1974) and a pregnancy or pseudopregnancy may ensue. The fertility of such matings is reduced (Cooper et al., 1994).

3. Significant delays in ovulation can result in increased embryonic abnormalities and pregnancy loss (Fugo and Butcher, 1966; Cooper et al., 1994).

4. Persistent diestrus indicates temporary or permanent cessation of follicular development and ovulation, and thus at least temporary infertility.

5. Prolonged vaginal diestrus, or anestrus, may be indicative of agents (e.g., polyaromatic hydrocarbons) that interfere with follicular development or deplete the pool of primordial follicles (Mattison and Nightingale, 1980) or agents such as atrazine that interrupt gonadotropin support of the ovary (Cooper et al., 1996). Pseudopregnancy is another altered endocrine state reflected by persistent diestrus. The ovaries of anestrous females are atrophic, with few primary follicles and an unstimulated uterus (Huang and Meites, 1975). Serum estradiol and progesterone are abnormally low.

6. Lengthening of the cycle may be a result of increased duration of either estrus or diestrus.

Evidence Supporting this KER

In females, normal reproductive function involves the appropriate interaction of central nervous system, anterior pituitary, oviducts, uterus, cervix and ovaries. During the reproductive years the ovary is the central organ in this axis. The functional unit within the ovary is the follicle which is composed of theca; granulosa cells and the oocyte. The somatic compartment synthesizes and secrets hormones (steroids and growth factors) necessary for the orchestration of the inter-relationship between the other parts of the reproductive tract and the central nervous system. Oestrus cycle is under strict hormonal control, therefore perturbations of hormonal balance lead to perturbations of normal cyclicity (change in number of cycles or duration of each phase) and/or ovulation problems leading to impaired female reproductive function. However, there are other mechanisms that might result in impaired fertility (e.g cellular maturation in ovary).

Many chemicals are found to interfere with reproductive function in the female. This interference is commonly expressed as a change in normal morphology of the reproductive tract or in ovarian cycle irregularities (disturbance in the duration of particular phases of the estrous cycle and/or ovulation problems). Monitoring estrous cyclicity provides a means to identify alterations in reproductive functions which are mediated through nonestrogenic as well as estrogenic mechanisms (Blasberg, Langan, & Clark, 1997), (Clark, Blasberg, & Brandling-Bennett, 1998). Adverse alteration in the nonpregnant female reproductive system have been observed at dose levels below those that result in reduced fertility or produce other overt effects on pregnancy or pregnancy outcomes. A disruption of cycling caused by xenobiotic treatment can induce a persistent estrus, a persistent diestrus, an irregular pattern with cycles of extended duration and ovulation problems. Common classes of chemicals have been shown to cause cycle irregularities in rats, humans, and non-human primates. Examples include the polychlorinated biphenyls (PCBs) and dioxins, which are associated with such irregularities in rats and humans (e.g (Li, Johnson, & Rozman, 1995) (Meerts et al., 2004), (Chao, Wang, Lin, Lee, & Päpke, 2007) and various agricultural pesticides, including herbicides, fungicides, and fumigants for review see (Bhattacharya & Keating, 2012),(Bretveld, Thomas, Scheepers, Zielhuis, & Roeleveld, 2006).

Table 1 Summary the empirical evidence supporting the KER.

It is known that exposure to 17-β-estradiol can disrupt the normal 4- to 5-day estrous cycle in adult female rats by inducing an extended period of diestrus consistent with pseudopregnancy within 5–7 days after the exposure (Gilmore & McDonald, 1969). This is due to the estrogen-dependent increase in prolactin that rescues ovarian corpora lutea and the subsequent synthesis and release of progesterone (Cooper, R. L., and Goldman, 1999). Significant evidence that the estrous cycle (or menstrual cycle in primates) has been disrupted should be considered an adverse effect (OECD, 2008).

Chemicals may be found to interfere with reproductive function in the female. This interference is commonly expressed as a change in normal morphology of the reproductive tract or a disturbance in the duration of particular phases of the estrous cycle. However, menstrual cyclicity is affected by many parameters such as age, nutritional status, stress, exercise level, certain drugs, and the use of contraceptive measures that alter endocrine feedback. In nonpregnant females, repetitive occurrence of the four stages of the estrous cycle at regular, normal intervals suggests that neuroendocrine control of the cycle and ovarian responses to that control are normal. Even normal, control animals can show irregular cycles. However, a significant alteration compared with controls in the interval between occurrence of estrus for a treatment group is cause for concern. Generally, the cycle will be lengthened or the animals will become acyclic. Therefore changes in cyclicity should be interpreted with caution and not judged adverse without a comprehensive consideration of additional relevant endpoints in a weight-of-evidence approach.

Inconsistencies

Two generation studies by Tyl et al with Butyl benzyl phthalate (BBP) did not observe effects in F0 females on any parameters of estrous cycling, mating, or gestation. However, F1 females carrying F2 litters at and reduced number of total and live pups/litter at birth, with no effects on pre- or postnatal survival (Tyl et al., 2004).

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