Phosphoramide mustard induces autophagy markers and mTOR inhibition prevents follicle loss due to phosphoramide mustard exposure

Reproductive Toxicology 67 (2017) 65–78

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Reproductive Toxicology journal homepage: www.elsevier.com/locate/reprotox

Phosphoramide mustard induces autophagy markers and mTOR inhibition prevents follicle loss due to phosphoramide mustard exposure Jill A. Madden, Porsha Q. Thomas, Aileen F. Keating ∗ Department of Animal Science, Iowa State University, Ames, IA, 50011, United States

a r t i c l e

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Article history: Received 27 July 2016 Received in revised form 15 November 2016 Accepted 21 November 2016 Available online 22 November 2016 Keywords: Ovotoxicity Phosphoramide mustard Mammalian target of rapamycin Phosphatidylinositol-3 kinase

a b s t r a c t Phosphoramide mustard (PM) is an ovotoxic metabolite of cyclophosphamide. Postnatal day 4 Fisher 344 rat ovaries were exposed to vehicle control (1% DMSO) or PM (60 ␮M) ± LY294002 or rapamycin for 2 or 4 d. Transmission election microscopy revealed abnormally large golgi apparatus and electron dense mitochondria in PM-exposed ovaries prior to and at the time of follicle depletion. PM exposure increased (P < 0.05) mRNA abundance of Bbc3, Cdkn1a, Ctfr, Edn1, Gstp1, Nqo1, Tlr4, Tnfrsfla, Txnrd1 and decreased (P < 0.05) Casp1 and Il1b after 4d. PM exposure increased (P < 0.1) BECN1 and LAMP, decreased (P < 0.1) ABCB1 and did not alter ABCC1 protein. LY294002 did not impact PM-induced ovotoxicity, but decreased ABCC1 and ABCB1 protein. Rapamycin prevented PM-induced follicle loss. These data suggest that the mammalian target of rapamycin, mTOR, may be a gatekeeper of PM-induced follicle loss. © 2016 Elsevier Inc. All rights reserved.

1. Introduction The female ovarian reserve is established at birth and represents a finite number of follicles, comprised of the oocyte surrounded at initial stages of development by squamous granulosa cells. Throughout the lifespan of a woman, follicles gradually progress through development beginning at the primordial follicle stage and continuing through the primary and secondary follicle stages to eventually become surrounded by theca cells, form an antrum and ultimately be ovulated. However, less than 1% percent of follicles will complete the cycle to be ovulated. Follicles that are not ovulated will die, thus resulting in over 99% of mammalian follicles dying by the process of atresia [1]. Once the ovarian reserve is depleted of follicles, ovarian failure, or menopause, occurs. On average, the natural phenomenon of menopause occurs at the age of 51 [2], however, chemical exposures can deplete follicles leading to premature ovarian failure (POF). The chemotherapy drug cyclophosphamide (CPA) targets and depletes primordial follicles in mice [3,4] and antral follicles in rats [5,6] at concentrations relevant to human exposures [4,7]. The results from these studies coincide with the human side effects, such as amenorrhea, premature menopause and infertil-

∗ Corresponding author at: 2356J Kildee Hall, Department of Animal Science, Iowa State University, Ames, IA, 50011, United States. E-mail address: [email protected] (A.F. Keating). http://dx.doi.org/10.1016/j.reprotox.2016.11.014 0890-6238/© 2016 Elsevier Inc. All rights reserved.

ity, reported by women and young girls who have undergone CPA treatment [8,9]. CPA is a prodrug and must be metabolized in order to have anti-cancer effects. Hepatic metabolism by cytochrome P450 (CYP) enzymes initiates the biotransformation of CPA, which ultimately forms the active, anti-cancer and ovotoxic metabolite phosphoramide mustard (PM). Notably, PM can further biotransform to a volatile and ovotoxic metabolite, presumably chloroethylaziridine (CEZ), which also likely contributes to PM overall toxicity [10]. Previous studies suggest that oocytes are the main PM target, while in larger follicles, that the granulosa cells are the targeted cell type [4]. Using a neonatal rat ovary culture method, following PM exposure, granulosa cells of primary follicles stained TUNEL positive [4]. Interesting, caspase-3 staining in PMexposed ovaries was not different from control and no primordial follicles stained caspase-3 positive [4]. Additionally, a study investigating the effects of varying doses of CPA in vivo found an absence of TUNEL or caspase-3 positive primordial follicle, despite primordial follicle depletion induced by CPA [11], which again, encourages the possibility that another PCD form is occurring, especially in the primordial follicles, in response to PM exposure. Apoptosis is a type of programmed cell death (PCD) occurring in ovarian follicles undergoing atresia (follicular death). However, a number of studies now support that autophagy may be an important process in follicle depletion, working either independently or in tandem with apoptosis [12–14]. The autophagy proteins Beclin 1 (BECN1) and lysosome associated membrane protein (LAMP1) are involved in key steps of autophagosome and autolysosome assembly [15–17].

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An autophagosome is the double-membraned structure that forms around material targeted for destruction, transports that unwanted material, and merges with the lysosome for digestion forming the autolysosome [18,19]. A study evaluating PCD at birth, a time of significant follicle loss, found autophagy to be the active PCD pathway [13]. Autophagy was proposed to result from the nutrient deprivation experienced at birth due to the removal of the fetus from the placenta, a phenomenon that has also been observed in other tissues [13,20]. Nutrient deprivation is sensed by the key response element mammalian target of rapamycin (mTOR) leading to the induction of autophagy via initiation of the autophagosome-forming cascade [21]. mTOR lies downstream of the phosphatidylinositol-3 kinase (PI3 K) signaling pathway and the interaction of these two pathways mediates cell survival and viability as well as regulating autophagy [22]. Chemical exposures, such as cigarette smoke, have also been shown to induce autophagy-mediated follicular death in mice [14,23]. Compared to unexposed controls, cigarette smoke-exposed mice had reduced number of follicles, but did not have a higher level of terminal deoxynucleotide transferase dUTP nick end labeling (TUNEL), DNA laddering or caspase activity − all of which are considered classic signs of apoptosis [24]. However, cigarette smoke-exposed mice did have a higher number of autophagosomes in granulosa cells, increased BECN1 and LC3 protein levels and mitochondrial dysfunction [23]. Also, through the use of a neonatal rat ovary culture method, single low level exposures to 7,12dimethylbenz[a]anthracene (DMBA), a component of cigarette smoke, reduced follicle number and altered Becn1 mRNA and protein levels [25]. Recently, a temporal induction of autophagy proteins has been demonstrated in ovaries of mice exposed in utero to bisphenol A (BPA) [26] and 2,2 ,4,4 -tetrabromodiphenylether exposure also induced autophagy proteins, including BECN1, in rat ovaries as a mode of ovotoxicity [27]. Collectively, these studies suggest autophagy is an active process during the ovarian response to xenobiotics. Information regarding ovarian expression and function of phase III chemical biotransformation proteins in non-cancerous tissues remains scant. Multidrug resistance (MDR) to chemotherapeutics represents a challenge in terms of treatment but could be advantageous in protecting the ovary from the ovotoxic effects of chemotherapy. ATP binding cassette transporters are known to be involved in MDR and much of our knowledge in their ovarian function comes from studies of tumor biology [28–31]. We have reported alterations in ovarian phase I and II chemical biotransformation enzymes during PM exposure [32], however, the involvement of phase III biotransformation during PM exposure is unknown. ABCB1 [33,34] have recently been demonstrated to be regulated by PI3 K/mTOR signaling. Additionally, multidrug resistance gene 1 (Mdr1/Abcc1) is also regulated by mTOR signaling and can be modified by rapamycin treatment [35]. Amplification of the catalytic subunit of PI3 K is associated with resistance to chemotherapy in ovarian cancer patients [36], and this could be potentially due to alterations in PI3 K signaling, autophagy and phase III drug metabolism. We investigated the effect of the phase III enzymes ABCB1 and ABCC1 in this study due to intersect between their regulation and that of autophagy via mTOR. In this study we hypothesized that autophagy is an active ovarian response following PM exposure which contributes to PMinduced follicle loss and that phase III drug metabolism would also be activated in the ovary in response to PM exposure. We evaluated mRNA (Becn1) and/or protein (LAMP1) abundance of autophagy cellular components along with the presence of autophagosomes via transmission electron microscopy (TEM). To evaluate a functional role for autophagy in follicle death and survival, mTOR was inhibited using rapamycin and PI3 K was inhibited using LY294002

to evaluate the impacts on PM-induced follicle loss. Furthermore, we investigated the involvement of phase III chemical biotransformation in the ovary as a response to PM exposure and determined a role for ovarian PI3 K regulation of these proteins. 2. Materials and methods 2.1. Reagents Bovine serum albumin (BSA), ascorbic acid, transferrin, 2-␤mercaptoethanol, 30% acrylamide/0.8% bisacrylamide, ammonium persulphate, glycerol, N’ N’ N’ N’-tetramethylethylenediamine (TEMED), Tris base, Tris HCL, sodium chloride, Tween-20, phosphatase inhibitor, protease inhibitor and rapamycin were purchased from Sigma Aldrich Inc. (St. Louis, MO). Dulbecco Modified Eagle Medium: nutrient mixture F-12 (Ham) 1× (DMEM/Hams F12), Albumax, penicillin (5000U/ml) Hanks Balanced Salt Solution (without CaCl2 , Mg Cl2 , or MgSO4 ) were obtained from Invitrogen Co. (Grand Island, NY). Millicell-CM filter inserts and 48-well cell culture plates were obtained from Millipore (Billerica, MA) and Corning Inc. (Corning, NY), respectively. RNeasy Mini kit, QIA Shredder kit, RNeasy Mini Elute kit, Quantitect TM SYBR Green PCR kit, RT2 First Strand kit, RT2 SYBR Green Mastermix, and the Stress and Toxicity RT2 Profiler PCR arrays were purchased from Qiagen Inc. (Valencia, CA). RNAlater was obtained from Ambion Inc. (Grand Island, NY). 2-(4-morpholinyl)-8-phenyl-4H-1-benzopyran-4-one (LY294002) was purchased from A.G. Scientific, Inc. (San Diego, CA). Recombinant mouse kit ligand was obtained from R & D Systems Inc. (Minneapolis, MN). PM was obtained from the National Institutes of Health National Cancer Institute (Bethesda, MA). All primers were obtained from the DNA facility of the Iowa State University office of biotechnology (Ames, IA). Ponceau S was purchased from Fisher Scientific (Waltham, MA). SignalFireTM ECL Reagent, anti-LAMP1, and the HRP-linked secondary anti-rabbit ® antibody was purchased from Cell Signaling Technology (Danvers, MA). Anti-BECN1 antibody was purchased from Santa Cruz (Dallas, TX) and donkey anti-rabbit FITC labeled secondary antibody was obtained from Southern Biotech (Birmingham, AL). Anti-ABCC1 and anti-ABCB1 primary antibodies were purchased from Cell Signaling (Beverly, MA). 2.2. Animals The Iowa State University Institutional Animal Care and Use Committee’s approved all experimental procedures. Fisher 344 (F344) rats were housed one per plastic cage and maintained in a controlled environment (22 ± 2 ◦ C; 12 h light/12 h dark cycles). The animals were provided a standard diet with ad libitum access to food and water, and allowed to give birth. 2.3. In vitro ovarian cultures Following euthanasia, ovaries were removed from female postnatal day (PND) 4 F344 rats [37]. Each ovary was removed, trimmed of oviduct and excess tissue and placed onto a MillicellCM membrane floating on 250 ␮l of previously 37 ◦ C equilibrated DMEM/Hams F12 medium containing 1 mg/ml BSA, 1 mg/ml Albumax, 50 ␮g/ml ascorbic acid, 5 U/ml penicillin and 27.5 ␮g/ml transferrin per well in a 48-well plate. To prevent dehydration, a drop of medium was placed on top of each ovary. In order to eliminate CEZ as a confounding factor, PM-treated ovaries were cultured in a separate incubator from other treatments [10]. Ovaries were treated with vehicle control (DMSO; 1%) or PM (60 ␮M) for 1–4 days as indicated in specific experiments. Ovaries were also treated with vehicle control (DMSO; 1%) or PM (60 ␮M) ± kit ligand (400 ng/ml) for 4 d for protein isolation. As we have previously reported, PM

J.A. Madden et al. / Reproductive Toxicology 67 (2017) 65–78

metabolizes into a volatile cytotoxic compound that also negatively impacts the ovary [10]. The PM dose, duration and time of media change and PM addition, used is based upon our previous work and was found to be effective at induction of follicle loss [10,38]. The relevance to cancer patients is that we utilize an endpoint at which we observe approximately 50% follicle loss and we evaluate gene changes prior to and at the time of follicle loss. This represents a bioassay approach since it is impossible to know the amount of PM reaching the human ovary during treatment and how this varies per individual on varying regimens and dosage. 2.4. RNA isolation and quantitative real-time polymerase chain reaction (qRT-PCR) Following 2 or 4 d of in vitro culture, total RNA was isolated (n = 3; 10 ovaries per pool) using an RNeasy Mini kit according to the manufacturer’s instructions, concentrated via RNeasy Mini Elute kit and quantified with a NanoDrop (␭ = 260/280 nm; ND 1000; Nanodrop Technologies Inc., Wilmington, DE). 2.4.1. RT2 PCR array Total RNA (250 ng) was reverse transcribed to cDNA using the RT2 first-strand kit and applied to the Stress and Toxicity Profiler PCR Array according to the manufacturers protocol. The PCR conditions were a 10 min hold at 95 ◦ C and 40 cycles of denaturing at 95 ◦ C for 15 s and a combined annealing and extension for 1 min at 60 ◦ C. Genes of interest were normalized to the housekeeping genes recommended by company-provided analysis software, hypoxanthine phosphoribosyltransferase (Hprt) and ribosomal protein, large, P1 (Rplp1). The mRNA level of these two genes did not change across treatments. The online SABiosciences RT2 ProfilerTM PCR Array Data Analysis quantified the changes in mRNA levels using the 2−Ct method [39,40]. 2.4.2. Quantitative PCR analysis Total RNA (200 ng) was reverse transcribed to cDNA utilizing the Superscript III One-Step RT-PCR (Qiagen). cDNA was diluted (1:20) in RNase-free water. Diluted cDNA (2 ␮l) were amplified on an Eppendorf PCR Master cycler using Quantitect SYBR Green PCR kit (Qiagen). Primers for Becn1 and Gapdh were designed by Primer 3 Input Version (0.4.0) [41]. The regular cycling program consisted of a 15-min hold at 95 ◦ C and 45 cycles of denaturing at 95 ◦ C for 15 s, annealing at 58 ◦ C for 15 s, and extension at 72 ◦ C for 20 s at which point data were acquired. There was no difference in Gapdh mRNA expression between treatments, thus each sample was normalized to Gapdh before quantification. Quantification of fold-change in gene expression was performed using the 2−Ct method [40,42]. 2.5. Protein isolation and western blot analysis PND4 ovaries (n = 3; 10 ovaries per pool) were homogenized in 200 ␮l of ice-cold tissue lysis buffer and protein quantified using a standard BCA protocol on a 96-well assay plate. Total protein (15 ␮g) was separated on a 10% SDS-PAGE and transferred to a nitrocellulose membrane. Prior to 1 h blocking in 5% milk, membranes were stained with Ponceau S to confirm equal protein loading. The membrane was then incubated at 4 ◦ C with rabbit antiBECN1 (1:250; Santa Cruz); anti-ABCB1 (1:100) or ABCC1 (1:100). Following ∼60 h of incubation, donkey anti-rabbit secondary antibody was applied and rocked with membrane for 1 h at room temperature. Autoradiograms were visualized on X-ray films in a dark room following 10 min incubation of membranes with 1× SignalFireTM ECL reagent. Densitometry of the appropriate sized bands was measured using Image Studio Lite Version 3.1 (LI-COR

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Biosciences, Lincoln, NE) which eliminates background noise. Values were normalized to Ponceau S stained total protein. 2.6. Immunofluorescence staining Ovaries were placed in 4% paraformaldehyde for 2 h, washed and stored in 70% ethanol, paraffin embedded and serially sectioned (5 ␮M). Two sections per ovary (n = 3) were deparaffinized, blocked with 5% BSA and incubated with anti-LAMP1 primary antibody (1:50 dilution) at 4 ◦ C overnight. Secondary FITC-labelled antibody was applied for 1 h, followed by incubation with the nuclear stain, Hoechst (30 min; 1:1000 dilution/5 mg/ml; Invitrogen). Slides were repeatedly rinsed with PBS, cover-slipped, and stored in the dark (4 ◦ C) until visualization. Primary antibody was not added to immunonegative ovarian sections. Immunofluorescence was visualized on a Leica DMI300 B fluorescent microscope at ␭ = 488 and 633 nm for FITC and Hoechst, respectively. All images were captured using a 10× objective lens. Protein level of LAMP1 was quantified using OpenLab by setting an intensity threshold and measuring the number of pixels above that threshold in controland PM-treated ovaries (n = 3/treatment). The number of pixels in each section was normalized to the total area of that section. 2.7. Histological evaluation of ovarian morphology PND4 ovaries (n = 5) were exposed on alternate days to PM (60 ␮M) and collected from culture after 1, 2, 3 or 4 days. The ovaries were fixed with 2% glutaraldehyde (w/v) and 2% paraformaldehyde (w/v) in 0.1 M sodium cacodylate buffer, pH 7.2, for at least 48 h at 4 ◦ C. Samples were washed in buffer and then fixed in 1% osmium tetroxide in 0.1 M cacodylate buffer for 1 h at room temperature The samples were then dehydrated in a graded ethanol series, cleared with ultra-pure acetone, infiltrated and embedded using a modified EPON epoxy resin (Embed 812; Electron Microscopy Sciences, Ft. Washington, PA). Resin blocks were polymerized for 48 h at 70 ◦ C. Thick and ultrathin sections were made using a Leica UC6 ultramicrotome (Leeds Precision Instruments, Minneapolis, MN). Thick sections were stained with epoxy resin stain (Electron Microscopy Sciences, Ft. Washington, PA) and imaged with a Zeiss Axioplan II light microscope with a MRC AxioCam and Axiovision software (Carl Zeiss INC, Thornwood, NY). Ultrathin sections were collected onto copper grids and images were captured using a JEM 2100 200 kV scanning and transmission electron microscope (Japan Electron Optic Laboratories Inc., Peabody, MA). 2.8. Inhibition of mTOR using rapamycin Ovaries (n = 6 for follicle counting) were treated with vehicle control media (1% DMSO); PM (60 ␮M); rapamycin (1 ␮M); or PM (60 ␮M) + rapamycin (1 ␮M) on alternate days for 4d. The rapamycin concentration administered was adapted from a prior study [43]. 2.9. Inhibition of PI3 K using LY294002 Ovaries (n = 4–6 for follicle counting; 10 ovaries per pool, n = 3 for protein isolation) were treated on alternate days with vehicle control media (1% DMSO); PM (60 ␮M); LY294002 (20 ␮M); or PM (60 ␮M) + LY294002 (20 ␮M). The concentration of LY294002 using the PND4 ovary culture method was previously determined to be effective [44]. 2.10. Histological evaluation of follicle numbers Ovaries were placed in 4% paraformaldehyde for 2 h, washed and stored in 70% ethanol, paraffin embedded, and serially sec-

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tioned (5 ␮M). Every 6th section was mounted and stained with hematoxylin and eosin. Healthy oocyte-containing follicles were identified and counted in every 6th section. Unhealthy follicles were distinguished from healthy follicles by the appearance of pyknotic bodies and intense eosinophilic staining of oocytes. Healthy follicles were classified and enumerated as previously described [45]. Slides were blinded to prevent counting bias. 2.11. Statistical analysis Treatment comparisons for follicle count experiments were performed using Analysis of Variance (ANOVA). Quantitative RT-PCR, Western blot and immunofluorescence staining data were analyzed by t-test comparing treatment with control raw data at each individual time point. All statistical analysis was performed using Prism 5.04 software (GraphPad Software). Statistical significance was defined as P < 0.05 and a trend for a significant difference from control was designated at P < 0.1. 3. Results 3.1. Evaluation of changes in genes involved in stress and toxicity pathways following PM exposure A stress and toxicity RT2 Profiler PCR array was used to investigate the mRNA profile of PND4 rat ovaries (n = 3, 10 ovaries per pool) which had been exposed to PM (60 ␮M) for 2 or 4 d (Table 1). Following 2d of PM exposure, B2 m mRNA levels decreased (P < 0.05), as well as there being a trend for a decrease (P < 0.1) in Tnfa and a trend (P < 0.1) for an increase in Hspa1b and Sqstml, relative to control treated ovaries. After 4 d, PM increased (P < 0.05) Bbc3, Cdkn1a, Cftr, Edn1, Gstp1, Nqo1, Tlr4, Tnfrsf1a, Txnrd1 mRNA levels compared to control, while decreased (P < 0.05) Casp1 and Il1b mRNA levels were observed. In addition, 4 d of PM exposure induced a trend (P < 0.1) for increased mRNA levels for the following genes: Cd40lg, Fas, Gadd45a and Mre11a as well as a trend (P < 0.1) for a decrease in levels of Adm, Epo, Mmp9, Tnf, and Vegfa. The results of this experiment are summarized in Table 1. 3.2. Investigation of ovarian damage by TEM In order to determine histological signs of ovarian damage, PND4 ovaries were exposed to PM (60 ␮M) on alternate days and collected after days 1, 2, 3 and 4 of culture for TEM. PM-treated ovaries were compared against primordial (Fig. 1A) and small primary (Fig. 1B) follicles from control-treated ovaries. After 1 d, PM-exposed ovaries exhibited dark, electron dense structures in the oocyte cytoplasm of primordial follicles (Fig. 1C) and large double-membraned structures in the granulosa cells of the small primary follicles (Fig. 1D) which appeared to have engulfed mitochondria. Multilamellar bodies were seen in both control and PM-treated ovaries after 2d, with a greater abundance apparent visually in PM-treated ovaries (Fig. 1E,F). Some oocytes of small primary follicles following 2 d of PM exposure appeared non-viable with global vacuolization (Fig. 1F), while other small primary follicles had little signs of damage. Similar to observations after 1 d of PM exposure, the granulosa cells of small primary follicles contained large double-membraned structures after 2 d (Fig. 1F). After 3 d of PM exposure, dead primordial follicles were evident by the lack of definition between the granulosa cell layer and the oocyte cytoplasm as well as small potential autophagosomes displacing the oocyte nucleus (Fig. 1G,H). Like the earlier time points, the cytoplasm of granulosa cells continued to have large areas of vacuolization, which had increased electron density. Following 4 d of PM exposure, in addition to apparent degradation, abnormal golgi apparatus were evident across follicle types (Fig. 1G).

Fig. 1. Histological evaluation of small preantral follicles following PM exposure by TEM. PND4 rat ovaries were treated with 1% DMSO (vehicle control, CT) or PM (60 ␮M). Following 1, 2, 3 and 4 d of culture, ovaries were fixed and processed for TEM (A) 1d CT primordial follicle; (B) 1d CT small primary follicle; (C) 1d PM primordial (D) 1d PM small primary (E) 2d PM primordial (F) 2d PM small primary (G) 3d primordial (H) 3d small primary (I) 4d primordial (J) 4d small primary; Hollow arrow = autophagosomes; G = abnormal golgi apparatus; Thin black arrow = double membrane; Thin dashed black arrow = electron dense structures; Solid thick black arrow = multilamellar structures; Arrow head = global vacuolization of oocyte.

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Table 1 Effect of PM exposure on ovarian expression of stress and toxicity genes. PND4 rat ovaries were treated with 1% DMSO (vehicle control) or PM (60 ␮M) for 2 or 4 d. Following RNA isolation, mRNA levels were quantified with an RT2 Profiler PCR array. Values represent fold-change to a control value of 1, normalized to Hprt1 and Rplp1. * = different from control, P < 0.05; † = P < 0.1. 2d mRNA

4d mRNA

Gene Name

Symbol

P-value

Fold Change

P-value

Fold Change

Adrenomedullin Aquaporin 1 Aquaporin 2 (collecting duct) Aquaporin 4 Aryl hydrocarbon receptor nuclear translocator Activating transcription factor 4 (tax-responsive enhancer element B67) Activating transcription factor 6 ATG12 autophagy related 12 homolog (S. cerevisiae) ATG7 autophagy related 7 homolog (S. cerevisiae) Ataxia telangiectasia mutated homolog (human) Bcl-2 binding component 3 Beclin 1, autophagy related BH3 interacting domain death agonist BCL2/adenovirus E1 B interacting protein 3-like Calreticulin Carbonic anhydrase 9 Caspase 1 Caspase 7 Chemokine (C C motif) ligand 12 CD40 ligand Cyclin-dependent kinase inhibitor 1A Cystic fibrosis transmembrane conductance regulator homolog (human) CHK1 checkpoint homolog (S. pombe) CHK2 checkpoint homolog (S. pombe) C-reactive protein, pentraxin-related Damage specific DNA binding protein 2 DNA-damage inducible transcript 3 DnaJ (Hsp40) homolog, subfamily C, member 3 Endothelin 1 Erythropoietin Fas (TNF receptor superfamily, member 6) Ferritin, heavy polypeptide 1 Growth arrest and DNA-damage-inducible, alpha Growth arrest and DNA-damage-inducible, gamma Glutamate-cysteine ligase, catalytic subunit Glutamate cysteine ligase, modifier subunit Growth factor receptor bound protein 2 Glutathione reductase Glutathione S-transferase pi 1 Heme oxygenase (decycling) 1 Heat shock protein 90, alpha (cytosolic), class A member 1 Heat shock protein 90, beta, member 1 Heat shock 70 kD protein 1 B (mapped) Heat shock protein 4 Heat shock protein 4-like Heat shock protein 5 HUS1 checkpoint homolog (S. pombe) Interferon gamma Interleukin 1 alpha Interleukin 1 beta Interleukin 6 Lactate dehydrogenase A Myeloid cell leukemia sequence 1 Matrix metallopeptidase 9 MRE11 meiotic recombination 11 homolog A (S. cerevisiae) Nibrin Nuclear factor of activated T-cells 5 NAD(P)H dehydrogenase, quinone 1 Poly (ADP-ribose) polymerase 1 Poly (ADP-ribose) polymerase 2 Peroxiredoxin 1 Poliovirus receptor RAD17 homolog (S. pombe) RAD51 homolog (RecA homolog, E. coli) (S. cerevisiae) RAD9 homolog (S. pombe) Receptor (TNFRSF)-interacting serine-threonine kinase 1 Receptor-interacting serine-threonine kinase 3 Serpin peptidase inhibitor, clade E, member 1 Solute carrier family 2 (facilitated glucose transporter), member 1 Solute carrier family 5 (sodium/myo-inositol cotransporter), member 3

Adm Aqp1 Aqp2 Aqp4 Arnt Atf4 Atf6 Atg12 Atg7 Atm Bbc3 Becn1 Bid Bnip3l Calr Car9 Casp1 Casp7 Ccl12 Cd40lg Cdkn1a Cftr Chek1 Chek2 Crp Ddb2 Ddit3 Dnajc3 Edn1 Epo Fas Fth1 Gadd45a Gadd45g Gclc Gclm Grb2 Gsr Gstp1 Hmox1 Hsp90aa1 Hsp90b1 Hspa1b Hspa4 Hspa4l Hspa5 Hus1 Ifng Il1a Il1b Il6 Ldha Mcl1 Mmp9 Mre11a Nbn Nfat5 Nqo1 Parp1 Parp2 Prdx1 Pvr Rad17 Rad51 Rad9 Ripk1 Ripk3 Serpine1 Slc2a1 Slc5a3

0.419797 0.438229 0.628950 0.409614 0.402638 0.388710 0.366526 0.196181 0.232411 0.310347 0.121654 0.210899 0.467470 0.420711 0.476559 0.249993 0.454259 0.225881 0.966183 0.151793 0.103088 0.121441 0.195285 0.184901 0.306373 0.234869 0.253815 0.162845 0.085487 0.420016 0.152999 0.201300 0.117923 0.165395 0.213655 0.200845 0.129836 0.157221 0.169424 0.230743 0.135410 0.219337 0.078083† 0.162369 0.247055 0.156053 0.125403 0.112426 0.475107 0.535699 0.377604 0.209343 0.136931 0.649328 0.237973 0.207718 0.104889 0.190228 0.151388 0.152741 0.290861 0.220251 0.221535 0.194399 0.227146 0.366649 0.173003 0.332730 0.165728 0.128854

−1.11 −1.92 −1.41 1.9 1.98 2.31 2.26 2.46 2.06 1.87 7.5 2.73 1.41 1.58 1.18 2.84 1.36 2.18 −1.33 3.04 21.76 5.79 3.35 4 11.03 2.32 2.18 5.36 4.46 2.1 4.34 5.13 6.36 3.96 7.21 3.93 4.41 3.58 5.27 5.25 3.66 3.58 7.89 4.05 12.07 3.32 4.87 5.94 1.33 1.21 3.52 4.15 4.2 1.17 3.22 2.99 5.86 5.66 4.63 4.12 1.75 2.07 2.36 2.5 3.45 2.08 2.08 2.09 4.15 5.79

0.054799† 0.836907 0.867642 0.480284 0.912393 0.572265 0.497833 0.222277 0.229800 0.171512 0.024119* 0.314192 0.854024 0.218922 0.740587 0.147251 0.021000* 0.345033 0.110543 0.079118† 0.009860* 0.038517* 0.219069 0.134936 0.150496 0.433278 0.677588 0.279235 0.007401* 0.077783† 0.091059† 0.287588 0.056880† 0.327235 0.161268 0.235398 0.296947 0.192857 0.002306* 0.586164 0.206438 0.449016 0.167280 0.628763 0.113168 0.493379 0.386687 0.298985 0.356756 0.039169* 0.290754 0.163127 0.103449 0.064778† 0.095880† 0.486507 0.112377 0.034212* 0.351783 0.360298 0.701133 0.388302 0.472564 0.104578 0.158361 0.396183 0.120046 0.902335 0.513878 0.110675

−2.44 1.02 −1.09 1.27 −1.03 −1.16 1.17 1.23 1.4 1.44 5.22 1.29 −1.05 1.15 1.03 1.4 −1.53 1.16 −1.84 1.28 15.04 2.8 1.35 1.74 1.78 1.09 1.06 1.16 1.48 −1.86 1.65 1.21 2.23 1.28 1.4 1.45 1.12 1.14 1.41 −1.15 1.16 1.13 2.15 1.06 1.67 1.11 1.29 1.52 −1.23 −1.93 −1.9 −1.28 1.42 −2.58 1.37 1.09 1.38 2.43 1.26 1.27 −1.42 1.21 1.09 1.41 1.15 1.18 −1.59 −1.11 −1.25 1.69

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Table 1 (Continued) 2d mRNA

4d mRNA

Gene Name

Symbol

P-value

Fold Change

P-value

Fold Change

Solute carrier family 9 (sodium/hydrogen exchanger), member 3 Sequestosome 1 Toll-like receptor 4 Tumor necrosis factor (TNF superfamily, member 2) Tumor necrosis factor receptor superfamily, member 10b Tumor necrosis factor receptor superfamily, member 1a Tumor protein p53 Thioredoxin 1 Thioredoxin-like 4B Thioredoxin reductase 1 Unc-51 like kinase 1 (C. elegans) Vascular endothelial growth factor A X-box binding protein 1 Xeroderma pigmentosum, complementation group C Actin, beta Beta-2 microglobulin Hypoxanthine phosphoribosyltransferase 1 Lactate dehydrogenase A Ribosomal protein, large, P1

Slc9a3 Sqstm1 Tlr4 Tnf Tnfrsf10b Tnfrsf1a Tp53 Txn1 Txnl4b Txnrd1 Ulk1 Vegfa Xbp1 Xpc Actb B2m Hprt1 Ldha Rplp1

0.265134 0.092462† 0.642777 0.067945† 0.463041 0.312884 0.179527 0.399887 0.359633 0.385092 0.319923 0.221746 0.298901 0.120776 0.474897 0.018168* 0.202150 0.663564 0.218626

4.73 6.93 −1.45 −2.22 −1.19 2.15 2.17 1.56 1.82 1.71 2.66 2.14 2.4 7.93 −1.9 −2.68 −1.22 −1.06 1.22

0.502139 0.478914 0.023985* 0.092015† 0.205793 0.035641* 0.219701 0.320947 0.205090 0.029960* 0.293571 0.052465† 0.498726 0.510219 0.150776 0.123497 0.149469 0.170170 0.166232

1.2 1.17 1.21 −1.47 −1.11 1.25 −1.07 1.08 1.3 1.32 1.23 −1.81 1.16 1.16 −1.18 −1.23 1.13 −1.26 −1.1

Fold-changes in bold refer to those with a P-value < 0.05.

Fig. 2. Detection of autophagosomes by TEM in large preantral follicles following PM exposure. PND4 rat ovaries were treated with 1% DMSO (vehicle control, CT) or PM (60 ␮M). Following 1, 2, 3 and 4 d of culture, ovaries were fixed and processed for TEM (A) 3d CT; (B) 4d CT; (C) 3d PM and (D) 4d PM. Black double-ended arrows indicate the width of the granulosa cell layer; Hollow arrows = autophagosomes; G = abnormal golgi apparatus.

Large primary and secondary follicles, were evident after 3 and 4 d, respectively, in control ovaries (Fig. 2A,B), however, these two follicle types were difficult to distinguish in PM-exposed ovaries because, although the oocyte had grown, the granulosa cell layer was minute (Fig. 2D), about 2 ␮M, compared to the control, which was over 10 ␮M in width (Fig. 2C). Also, in contrast to control, the mitochondria appeared much darker and electron-dense in the large follicles of PM-treated ovaries (Fig. 2C, D). Analogous to the PM-treated primordial and small primary follicles, abnor-

mally large golgi apparatus and autophagosomes were present in the oocyte cytoplasm and granulosa cells, respectively (Fig. 2C,D). 3.3. Determination of increased abundance of autophagy signaling molecules We evaluated the mRNA abundance of Becn1 in ovaries exposed to PM for 2 or 4d. There was a trend (P < 0.1) for increased Becn1 mRNA (Fig. 3A) and protein (Fig. 3B,C) abundance after 4 and 2 days of PM exposure, respectively. We had difficulty with identifying

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addition of rapamycin did not impact the effects of PM on large primary and secondary follicles, relative to control treated ovaries (Fig. 6C and D). Compared to control, rapamycin treatment alone reduced (P < 0.05) secondary follicle number with no effect of treatment on any other follicle stage (Fig. 6D). 3.5. Involvement of phase III biotransformation proteins in ovarian response to PM exposure Total ovarian protein homogenate contained ABCC1 (Fig. 7A) and ABCB1 (Fig. 7B). No impact of PM exposure on total ovarian ABCC1 protein level was observed (Fig. 7A), however, protein abundance of ABCB1 was decreased (P = 0.05) by PM exposure at the time of follicle loss which occurs at day 4 (Fig. 7B). 3.6. Potential for PI3 K regulation of ovarian phase III biotransformation proteins Utilizing protein isolated from ovaries cultured in media containing the PI3 K inhibitor, LY294002 (20 ␮M) or the PI3 K activator, kit ligand, Western blotting demonstrated that in the absence of PI3 K signaling, ABCC1 protein was essentially absent (P < 0.05) from the ovary compared to control treatment (Fig. 8A). Interestingly, Kit Ligand treatment, did not impact ABCC1 protein levels and no difference from control treated ovaries was observed (Fig. 8A). Similarly to ABCC1, lack of PI3 K signaling in cultured ovaries reduced (P < 0.05) ABCB1 protein abundance, though to a lesser extent than that of ABCC1 (Fig. 8B). Kit Ligand treatment had no impact of ABCB1 protein abundance (Fig. 8B). 4. Discussion Fig. 3. Impact of PM exposure on ovarian Becn1 mRNA and protein abundance. PND4 rat ovaries (n = 3; 10 ovaries per pool) were treated with 1% DMSO (vehicle control; CT) or PM (60 ␮M). Following 2 or 4 d of culture, mRNA or protein was isolated and Becn1 levels evaluated. (A) mRNA values represent fold-change ± SEM relative to a control value of 1, normalized to Gapdh. (B) Representative Western blot of 2 d treated ovaries normalized to Ponceau S and (B) data quantification. Values represent signal intensity ± SEM. # = P < 0.1 difference from CT.

LAMP1 via Western blotting, thus, immunostaining was employed to investigate any potential effect of PM on LAMP1 protein, and relative to control-treated ovaries (Fig. 4A,D) increased LAMP1 protein staining signal was observed 4d post-PM exposure (Fig. 4B-F). 3.4. Evaluation of mechanisms of PM-induced ovotoxicity To gain insight into the mechanisms behind autophagy induced by PM exposure, we manipulated pathways that are associated with cell survival using pharmaceutical approaches. PND4 ovaries (n = 5 6) were treated with vehicle control (1% DMSO), PM (60 ␮M), rapamycin (1 ␮M), PM (60 ␮M) + rapamycin (1 ␮M), LY294002 (20 ␮M), or PM (60 ␮M) + LY294002 (20 ␮M) on alternate days for 4 d and then processed for histology. Relative to control treated ovaries, there were reduced numbers of all follicle stages due to PM exposure (Fig. 5 and 6). The addition of LY294002 resulted in an increased (P < 0.05) number of primordial follicles (Fig. 5A) and a trend (P < 0.10) for increased small primary follicles (Fig. 5B), and a reduction (P < 0.05) in large primary and secondary follicles (Fig. 5C,D), when compared to control ovaries. Surprisingly, ovarian follicle depletion by PM in the presence of LY294002 was not different from ovaries treated solely with PM (Fig. 5A-D). PM-induced primordial and small primary follicle loss was prevented in the presence of rapamycin (Fig. 6A and B). Interestingly, PM + rapamycin treated ovaries tended to have increased (P = 0.05) numbers of primordial follicles than control ovaries (Fig. 6A). The

PM is the ovotoxic and antineoplastic metabolite of the chemotherapy drug CPA [3,4]. CPA is used to treat a variety of cancers, but due it its ability to deplete primordial follicles [3], CPA increases a female patient’s risk of reduced fertility or POF [46], which represents a major concern for female cancer survivors. Although compromised fertility via follicle depletion is well documented as a side effect of CPA, little is known about the mechanisms by which this ovotoxicant elicits its detrimental effects on the ovary. A prior study, using the same neonatal rat ovary culture system as in our experiments, investigated apoptosis as a PCD mechanism active during PM-induced follicle loss and found that granulosa cells of primary follicles were positively identified during TUNEL and cleaved caspase-3 staining following PM exposure [4]. However, the number of follicles that were positive for cleaved caspase-3 did not differ from control and no primordial follicles stained positive for this apoptosis marker [4]. In addition, use of a caspase-3 inhibitor did not prevent PM-induced follicle loss leading the authors to conclude that PM-induced follicle loss occurs via a caspase-independent pathway [4]. Autophagy, a caspase-independent PCD process, represents an alternative atretic route outside of apoptosis. There is mounting evidence in favor of the involvement of autophagy in both ovarian follicular death and survival. Studies investigating mechanisms responsible for follicle death at birth and puberty, times of large natural follicle loss, have found the classic markers of autophagy plentiful, with little to no signs of apoptosis [13,47]. Furthermore, ovarian induction of autophagy genes was found in murine ovaries following exposure to cigarette smoke [14,23]. Also, we have demonstrated altered mRNA levels of autophagy genes in neonatal rat ovaries exposed to low levels of DMBA [25], suggesting autophagy as a contributing factor to both the ovotoxicity of these compounds and the removal of damaged and/or dead follicles.

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Fig. 4. Localization and effect of PM on ovarian LAMP1 protein. PND4 rat ovaries (n = 3; 2 sections per ovary) were treated with 1% DMSO (vehicle control; CT) or PM (60 ␮M). Following 2 or 4 d of culture, ovaries were fixed in formalin, and immunofluorescence staining was performed using a primary antibody directed against LAMP1: (A) 2d CT; (B) 2d PM; (C) 4d CT; (D) 4d PM; (E) Enlarged 2d PM; and (F) Enlarged 4d PM. LAMP1 is represented in red and the Hoechst nuclear stain is in blue. Quantification of the number of LAMP1 foci was performed on (G) 2d and (H) 4d PM treated ovaries.

Interestingly, autophagy has also been suggested to act as a pro-survival mechanism in the ovary, particularly in the perinatal ovary and its establishment of the primordial follicle pool from the female germ cells [48]. Specifically, PND1 Atg7−/− knockout mice had no apparent germ cells and Becn1+/− had less than half the number of germ cells compared to control, thus supporting the functional requirement of these autophagy proteins for normal follicular development and the consideration of autophagy as more than just a PCD process. Separation of the functions of autophagy in follicles of differing stages of development represents a challenge but is a direction that should yield valuable information in terms of biological changes during folliculogenesis. The ovarian culture method is perfectly suited to the study of autophagy involvement in ovotoxicity, since there is lack of the functional circulatory system by which immune cells could remove such debris. The lack of a concomitant increase in unhealthy or dead follicles mirroring the loss of healthy follicles after PM exposure supports that there is some mechanism in place for this removal. Utilizing an RT-PCR array approach, two days after PM exposure, we identified a tendency for increased abundance of Heat shock 70 kD protein 1 B (Hspa1b) which has been associated with the ovarian stress response in pig ovaries [49] as well as increased Sequestosome 1/p62, which is implicated as an autophagic receptor [50] and is increased in expression by exposure to other ovotoxicants

[27]. Further, we discovered a tendency towards reduced Tumor necrosis factor (Tnf) mRNA abundance. Tnf is associated with cell death [51,52] and negatively impacts ovarian function [53,54], thus a decrease during PM exposure may represent an effort by the ovary to minimize loss of cellular viability. Interestingly, these molecular alterations occur prior to observed PM-induced follicle loss [10], thus may be attractive candidates as initiating molecular events during PM-induced ovotoxicity. Bcl-2 binding component 3 (Puma), which has been shown to be involved in mitochondria-specific autophagy. In addition, a striking increase in cyclin-dependent kinase inhibitor 1A (p21) was observed in PM-exposed ovaries compared to control, also linked to the induction of autophagy [55]. In contrast, a decrease in mRNA level for caspase-1 after PM exposure was noted, in agreement with the previous study reporting PM-induced follicle depletion is caspase-independent [4]. Endothelin 1 (Edn1) [56], Cystic fibrosis transmembrane conductance regulator homolog (Cftr) [57], NAD(P)H dehydrogenase, quinone 1 (Nqo1) [58] were increased by PM exposure and associated with the process of autophagy and likely represent the ovarian response to PM exposure as a protective mechanism. Toll-like receptor 4 (Tlr4), the classic receptor for lipopolysaccharide (LPS), was increased in response to PM. The actions of the ligand for TLR4, LPS, has negative reproductive consequences, observed as primordial follicle loss [59,60] as

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Fig. 5. Impact of PI3 K inhibition during PM-induced ovotoxicity. PND4 rat ovaries were cultured and treated on alternate days with 1% DMSO (vehicle control, CT), PM (60 ␮M), LY (20 ␮M) or PM (60 ␮M) + LY (20 ␮M). Following 4 d of culture, healthy follicles were classified and counted: (A) Primordial Follicles; (B) Small Primary Follicles; (C) Large Primary Follicles; (D) Secondary Follicles. Values represent mean ± SE total follicles counted/ovary, n = 4–6. Different letters represent significant difference between treatments P < 0.05. * = P < 0.10 difference CT.

Fig. 6. Effect of mTOR inhibition on PM-induced ovotoxicity. PND4 rat ovaries were cultured and treated on alternate days with 1% DMSO (vehicle control, CT), PM (60 ␮M), Rapa (1 ␮M) or PM (60 ␮M) + Rapa (1 ␮M). Following 4 d of culture, healthy follicles were classified and counted: (A) Primordial Follicles; (B) Small Primary Follicles; (C) Large Primary Follicles; (D) Secondary Follicles. Values represent mean ± SE total follicles counted/ovary, n = 4–6. Different letters represent significant difference between treatments P < 0.05. * = P < 0.10 difference from CT.

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Fig. 7. Effect of PM treatment on ABCC1 and ABCB1 protein abundance in neonatal cultured ovaries. PND4 ovaries were cultured for 4 d in media containing vehicle control (CT; DMSO) or PM (60 ␮M). Total protein was isolated and Western blotting performed to quantify (A) ABCC1 or (B) ABCB1 protein abundance. Barchart indicates protein relative densitometric values for CT (C)- or PM (P)-treated ovaries; values represent mean ± standard error (n = 3 pools per treatment; 10 ovaries per pool). † indicates difference between treatments, P = 0.06.

Fig. 8. Effect of PI3 K inhibition on ABCC1 ovarian protein level. PND4 ovaries were cultured in media containing vehicle control, LY294002 (LY; 20 ␮M) or kit ligand (KL; 400 ng/ml) for 4 d. Total protein was isolated and (A) ABCC1 or (B) ABCB1 protein abundance determined by Western blotting. Barchart indicates relative protein densitometric values in CT-, LY-, or KL-treated ovaries; values represent mean ± standard error (n = 3 pools per treatment; 10 ovaries per pool). * indicates difference between treatments, P < 0.05.

well as effects on the endocrine system [61]. Interestingly, TLR4 can active PI3 K/AKT signaling [62], and has been identified to be linked to autophagy regulation [63]. Despite reduced Tnf two days after PM exposure, Tnf expression tended to be increased 4 days after PM exposure along with increased Tumor necrosis factor receptor superfamily, member 1a (Tnfrsf1a). Interestingly, this receptor is expressed in granulosa cells, thus may represent a response to altered Tnf, or PM exposure, or both. Also, there was a tendency for increased TNF receptor superfamily, member 6 (Fas). Interestingly, both Thioredoxin reductase 1 (Txnrd1) and Glutathione S-transferase pi 1 (Gstp1) were increased by PM exposure, both of whom regulate the redox state of the cell. Txnrd1 inhibition induced autophagy in colon cancer cells [64]. We have previously noted increased Gstp1 in response to PM exposure [32] and glutathionylation of PM represents a potential detoxification mechanism [32,65,66]. Reduction of interleukin 1 beta (Il1␤) by PM may also represent an ovotoxic mechanism since Il1␤ has been identi-

fied as a pro-survival factor in the ovary [67] and is also reduced during autophagy induction [68], thus increased Il1␤ during PM exposure may represent a protective mechanism within the ovary to prevent follicle damage and loss. Although present on the PCR array as an internal control, we observed that PM reduced mRNA abundance of Beta-2 microglobulin (B2m). In a previous study, we reported that inhibition of PI3 K dramatically affected the expression of beta-actin, thus chemical stressors can alter traditional internal control gene expression. Four days post-PM exposure, when follicle loss is observable [10], there were trends within the ovary for increased Adrenomedullin (Adm). ADM is altered in the ovary across the estrous cycle [69] and ovarian function includes oocyte-cumulus cell communication [70]. Interestingly, ADM interacts with EDN1 [71], which was also increased by PM exposure. CD40 ligand (Cd40lg) tended to be increased by PM exposure, and may represent a new marker of ovarian damage. We noted a tendency for increased MRE11 meiotic recombination 11 homolog A

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(Mre11a), a DNA damage response component [72]. Growth arrest and DNA-damage-inducible, alpha (Gadd45) was increased by PM exposure, also a DNA repair response element [73]. We have characterized PM-induced DNA damage as a mechanism of ovotoxicity [38,74], thus Mre11a and Gadd45 can be considered part of the ovarian DNA damage repair response. There were also tendencies for increased Erythropoietin (Epo), which has been identified as a modulator and potential therapeutic avenue for altering mTOR expression in disease states [75]. Finally, both Matrix metallopeptidase 9 (Mmp9) and Vascular endothelial growth factor A (Vegf) tended toward a PM-induced increase. Mmp9 has roles in ovarian steroidogenesis [76] and autophagy [77] while Vegf induces granulosa cell proliferation [78] and angiogenesis [79]. Finally, although not an overwhelming increase, we did observe a tendancy toward increased BECN1 and an increase in LAMP1 in a temporal response to PM. Taken together, the results from the PCR array support that mechanisms involved in autophagy are activated in the ovary post-PM exposure but also should be interpreted carefully as some of the observed alterations may simply be due to an altered cell population since follicle loss is occurring at the four day post-PM timepoint. Ovarian histology was examined using TEM following a time course of PM exposure. Since larger primary and secondary follicles develop after approximately 3–4 days in culture, determination of the impacts of PM on follicles of differing maturation stages (primordial, small primary, large primary and secondary) was possible. In small preantral (primordial and small primary) follicles, electron dense structures along with engulfed mitochondria in doublemembraned structures were noted. As the time of PM exposure progressed, vacuolization of oocytes was observed. The presence of abnormally large Golgi apparatus also became apparent, potentially suggesting a cellular response in protein modification in response to PM. In large preantral follicles (large primary and secondary), many of the same morphological alterations were seen. A major observation incompatible with follicle viability and growth is the reduction in the width of the granulosa cell layer, from about 2 to 10 ␮M in width. These data are in agreement with the PM-induced gap between the oocyte and granulosa cell that we have previously reported [10]. More importantly, PM-induced cellular component changes supportive of autophagy occurrence were observed prior to the observation of follicle loss (day 4; [10]), supporting that they are ovotoxic changes that occur due to PM exposure and contribute to ovotoxicity. Our findings coincide with a previous in vivo study, which found ovarian autophagy induced in response to whole-body cigarette exposure [14,23]. Similarly, the majority of autophagosomes were observed in the granulosa cells and these structures appeared to also contain mitochondria [14]. Collectively, these studies are among the first to report that autophagy is an active ovarian process that can be initiated in response to xenobiotic exposures. However, additional studies are required to further characterize the autophagy pathway and to ensure that the ovotoxicant-induced flux is occurring in induction of the autophagy pathway, opposed to a build-up of autophagy vesicles, due to lack of their degradation. Our evidence collected to this point strongly supported autophagy as an ovotoxic mechanism induced by PM, thus, we used pharmaceutical manipulation of pathways reported in the literature to regulate autophagy [43,80–83]. By administering PM in combination with LY294002, which was expected to inhibit autophagy via the PI3 K pathway [80–82], we hypothesized that if autophagy was prevented, that PM-induced follicle depletion would be at a higher level than in ovaries treated with PM alone. Surprisingly, there was no impact of PI3 K inhibition on PM-induced follicle loss. Interestingly, this result underscores the difference in mechanisms of ovotoxicity induced by chemical exposure. Increased and decreased primordial follicle loss was observed in

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ovaries treated with DMBA or 4-vinylcyclohexene diepoxide (VCD), respectively, during LY294002 treatment, relative to DMBA and VCD alone [44]. Additionally, this result contrasts with the findings that alteration of the PI3 K pathway through use of an immunomodulatory compound AS101 prevented follicle loss induced by CPA [11]. An in vivo study found that CPA increased phosphorylation of proteins in the PI3 K pathway, including mTOR and when the PI3 K pathway was perturbed, via the drug AS101, CPA-induced follicle loss was reduced [11]. How AS101 specifically impacts the PI3 K pathway remains unclear, since it is an immunomodulatory compound [91], and exposure was systemic rather than ovary-specific. It is interesting that many of the genes identified to be changed by PM exposure in our PCR array data are involved in processes associated with inflammation and/or immune response, thus the findings that AS101 altered CPA induced follicle loss may involve alterations to these genes identified in this study, however, this is speculative. Additionally, in our study characterizing follicle loss induced by PM [10], we did not observe evidence to support accelerated recruitment of primordial follicles into the growing pool as a mode of ovotoxicity. Our next step was to activate autophagy via the use of rapamycin, which inhibits mTOR, a negative regulator of autophagy [22,84,85]. We expected that if autophagy is an ovotoxic mechanism induced by PM, accelerated PM-induced follicle loss would be evident. In fact, the opposite was the case; PM-induced follicle loss was prevented in primordial and small primary follicles during mTOR inhibition. mTOR is a downstream component of PI3 K signaling and is activated and inhibited through phosphorylation at different sites [86]. If PM is working through mTOR to induce autophagy, it is unsurprising that no ovotoxic impact of PM during PI3 K inhibition was noted, since mTOR protein modification may have been unaltered simply by PI3 K inhibition in this experimental paradigm. We were unable to directly assess PMinduced mTOR alteration due to difficulties with the antibodies available for mTOR, and this is an area which we are actively pursuing. Interestingly, follicles cultured in rapamycin were previously noted to evade atresia [87] and rapamycin rescued poor developmental capability of aged porcine oocytes [88]. These studies indicate that pro-survival autophagy could be regulated independently from pro-death associated autophagy, which is likely a factor exemplified here with PM. Both the PI3 K and mTOR pathways have gained recent attention in ovarian physiology and toxicology, as two pathways suggested to play a key role in follicle activation and influence ovotoxicity [44,89,90]. Manipulation of this the mTOR pathway, shows promising results as a potential therapeutic to alleviate CPA/PM-induced infertility. Whether the follicles that were treated with rapamycin and PM are developmentally viable is a question for further research. Based on these largely unexpected findings, we next examined whether the phase III drug metabolism proteins ABCB1 and ABCC1 were also induced in response to PM exposure and regulated by PI3 K signaling. We have previously reported increased Abcb1 mRNA abundance in response to PM exposure [32]. In addition, we have demonstrated that phase I (Cyp2e1) and phase II chemical biotransformation proteins (Ephx1, Gstm1 and Gstp1), as well as transcriptional regulators of chemical biotransformation (Ahr and Nrf2) are downstream of PI3 K signaling [92–94]. While we anticipated that these drug transporters would be increased in response to PM, there was no impact on ABCC1 protein level, and PM decreased ABCB1 protein abundance, despite increased Abcb1 mRNA [32]. This reduction could be in response to binding the GSH conjugate of PM, transporting from the cell and resulting in a turnover of and thereby reduced ABCB1 protein level, however, this remains for further investigation. We have previously demonstrated that GSH supplementation protected the ovary from PM exposure [32], thus this hypothesis has merit. We have shown

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previously that the PI3 K pathway regulates enzymes involved in both phase I and phase II chemical biotransformation and in this study, we determined that this is also the case for phase III chemical biotransformation proteins. However, these results should be interpreted with the changes in ovarian histology that occur during both PI3 K inhibition and PM-induced ovotoxicity in mind, and determining ovarian localization of these proteins would reflect whether these findings are due to direct PI3 K actions or simply due to loss of the cellular fraction within the ovary in which these proteins are expressed. We have little information regarding ovarian phase III drug transport in non-carcinogenic samples, thus these data are informative and add to our understanding of the ovarian response to a toxic exposure. In summary, our results support that PM induces ovotoxicity, potentially through autophagy induction and we made the serendipitous observation that rapamycin prevented PM-induced follicle loss. We have discovered potential involvement of ABCB1 in the ovarian response to PM and that PI3 K may regulate the phase III chemical biotransformation proteins ABCB1 and ABCC1. This data lay the foundation for future experiments to confirm the ovarian protective mechanisms underway in response to chemotherapeutic treatment. Conflict of interest statement The authors do not have any conflicts of interest. Grant support The project described was partially supported by award number R00ES016818 to AFK. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of Environmental Health Sciences or the National Institutes of Health. References [1] A.N. Hirshfield, Development of follicles in the mammalian ovary, Int. Rev. Cytol. 124 (1991) 43–101. [2] H. Jick, J. Porter, Relation between smoking and age of natural menopause. report from the Boston collaborative drug surveillance program, Boston university medical center, Lancet 1 (8026) (1977) 1354–1355. [3] D.R. Plowchalk, D.R. Mattison, Phosphoramide mustard is responsible for the ovarian toxicity of cyclophosphamide, Toxicol. Appl. Pharmacol. 107 (3) (1991) 472–481. [4] P. Desmeules, P.J. Devine, Characterizing the ovotoxicity of cyclophosphamide metabolites on cultured mouse ovaries, Toxicol. Sci. 90 (2) (2006) 500–509. [5] J. Jarrell, E.V. Lai, R. Barr, A. McMahon, L. Belbeck, G. O’Connell, Ovarian toxicity of cyclophosphamide alone and in combination with ovarian irradiation in the rat, Cancer Res. 47 (9) (1987) 2340–2343. [6] D.P, P.B. Hoyer, Endocrinology and toxicology: the female reproductive system, in: M.J. Derelanko, M.A. Hollinger (Eds.), Handbook of Toxicology, CRC Press, 2002, pp. 573–596. [7] R.F. Struck, D.S. Alberts, K. Horne, J.G. Phillips, Y.M. Peng, D.J. Roe, Plasma pharmacokinetics of cyclophosphamide and its cytotoxic metabolites after intravenous versus oral administration in a randomized, crossover trial, Cancer Res. 47 (10) (1987) 2723–2726. [8] J.E. Sanders, C.D. Buckner, D. Amos, W. Levy, F.R. Appelbaum, K. Doney, R. Storb, K.M. Sullivan, R.P. Witherspoon, E.D. Thomas, Ovarian function following marrow transplantation for aplastic anemia or leukemia, J. Clin. Oncol. 6 (5) (1988) 813–818. [9] M.E. Suarez-Almazor, E. Belseck, B. Shea, G. Wells, P. Tugwell, Cyclophosphamide for treating rheumatoid arthritis, Cochrane Database Syst. Rev. 4 (2000) (CD001157). [10] J.A. Madden, P.B. Hoyer, P.J. Devine, A.F. Keating, Involvement of a volatile metabolite during phosphoramide mustard-induced ovotoxicity, Toxicol. Appl. Pharmacol. 277 (1) (2014) 1–7. [11] L. Kalich-Philosoph, H. Roness, A. Carmely, M. Fishel-Bartal, H. Ligumsky, S. Paglin, I. Wolf, H. Kanety, B. Sredni, D. Meirow, Cyclophosphamide triggers follicle activation and burnout; AS101 prevents follicle loss and preserves fertility, Sci. Transl. Med. 5 (185) (2013) (185ra62). [12] M.L. Escobar, O.M. Echeverria, R. Ortiz, G.H. Vazquez-Nin, Combined apoptosis and autophagy, the process that eliminates the oocytes of atretic follicles in immature rats, Apoptosis: Int. J. Progr. Cell Death 13 (10) (2008) 1253–1266.

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