The ovarian DNA damage repair response is induced prior to phosphoramide mustard-induced follicle depletion, and ataxia telangiectasia mutated inhibition prevents PM-induced follicle depletion

Toxicology and Applied Pharmacology 292 (2016) 65–74

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The ovarian DNA damage repair response is induced prior to phosphoramide mustard-induced follicle depletion, and ataxia telangiectasia mutated inhibition prevents PM-induced follicle depletion Shanthi Ganesan, Aileen F. Keating ⁎ Department of Animal Science, Iowa State University, Ames, IA 50011, USA

a r t i c l e

i n f o

Article history: Received 15 July 2015 Revised 16 December 2015 Accepted 17 December 2015 Available online 19 December 2015 Keywords: PM Ovary DNA repair Apoptosis PKC delta ATM inhibitor

a b s t r a c t Phosphoramide mustard (PM) is an ovotoxic metabolite of cyclophosphamide and destroys primordial and primary follicles potentially by DNA damage induction. The temporal pattern by which PM induces DNA damage and initiation of the ovarian response to DNA damage has not yet been well characterized. This study investigated DNA damage initiation, the DNA repair response, as well as induction of follicular demise using a neonatal rat ovarian culture system. Additionally, to delineate specific mechanisms involved in the ovarian response to PM exposure, utility was made of PKC delta (PKCδ) deficient mice as well as an ATM inhibitor (KU 55933; AI). Fisher 344 PND4 rat ovaries were cultured for 12, 24, 48 or 96 h in medium containing DMSO ± 60 μM PM or KU 55933 (48 h; 10 nM). PM-induced activation of DNA damage repair genes was observed as early as 12 h postexposure. ATM, PARP1, E2F7, P73 and CASP3 abundance were increased but RAD51 and BCL2 protein decreased after 96 h of PM exposure. PKCδ deficiency reduced numbers of all follicular stages, but did not have an additive impact on PM-induced ovotoxicity. ATM inhibition protected all follicle stages from PM-induced depletion. In conclusion, the ovarian DNA damage repair response is active post-PM exposure, supporting that DNA damage contributes to PM-induced ovotoxicity. © 2015 Elsevier Inc. All rights reserved.

1. Introduction The female gamete, the oocyte, is produced by the ovary, wherein they become integrated into follicular structures. Females are born with a finite number of oocytes, which once depleted, cannot be replaced (Hirshfield, 1991). Approximately 11% of women suffer infertility due to unknown etiology (Chandra et al., 2013) and a number of environmental exposures are known to impact ovarian function, including cigarette smoke, occupational chemicals, pesticides and chemotherapeutic agents, potentially hastening entry into premature ovarian senescence (Hoyer and Sipes, 1996; Hoyer and Keating, 2014). Cyclophosphamide (CPA) used in treatment of cancer and autoimmune diseases causes premature ovarian failure and rapid amenorrhea in treated individuals by destroying primordial and antral follicles (Brunner et al., 2006; Hudson, 2010). CPA targets primordial follicles in mice (Plowchalk and Mattison, 1992) and antral follicles in rats (Ataya et al., 1990). Phosphoramide mustard (PM), an ovotoxic CPA metabolite, destroys rapidly dividing cells by covalently binding to DNA, inducing DNA–DNA, DNA-protein cross links and DNA double strand ⁎ Corresponding author at: Department of Animal Science, Iowa State University, Ames, IA 50011, USA. E-mail addresses: [email protected] (S. Ganesan), [email protected] (A.F. Keating).

http://dx.doi.org/10.1016/j.taap.2015.12.010 0041-008X/© 2015 Elsevier Inc. All rights reserved.

breaks (DSB) (Helleday et al., 2008). Unfortunately, PM exposure causes dose- and time-dependent primordial and primary ovarian follicle loss following induction of DNA damage in mouse and rat oocytes (Petrillo et al., 2011). DSBs and DNA replication blocking lesions are both apoptotic signals and cells are equipped in their detection, prompting a protein cascade, which finally results in cell cycle arrest to facilitate DNA repair (Zhou and Elledge, 2000). If DNA repair fails, or the cell is overwhelmed by too many DNA lesions, cellular sensors initiate apoptosis (Friedberg, 2003). Proteins involved in DNA damage recognition include ataxia telangiectasia mutated (ATM), Ataxia telangiectasia and rad3-related (ATR) and DNA-dependent protein kinase (DNA-PK), which phosphorylate a multitude of proteins inducing a collective DNA damage response (DDR), in which p53 and BRCA1/2 play important roles (Roos and Kaina, 2006). ATM is implicated in three crucial functions: regulation and stimulation of DSB repair, activation of cell cycle checkpoints, and signaling for apoptosis induction; three key nodes in making the decision between survival and death following genotoxic exposures (Roos and Kaina, 2012). P73, a member of the P53 protein family, is a structural homolog of P53 and is involved in the cellular response to stress (Urist et al., 2004). P73 can induce the pro-apoptotic genes Bbc3 (aka Puma) and Bax (Melino et al., 2004) The E2F1 transcription factor can induce p73 transcription and also binds to the p73 promoter during DNA damage

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Table 1 Determination of genes involved in the ovarian response to PM exposure. 24 h

48 h

Gene symbol

Gene description

P-value

Fold-change

P-value

Fold-change

Abl1 Apex1 Atm Atrx Bard1 Bax Bbc3 Blm Brca1 Brca2 Cdc25a Cdc25c Cdkn1a Chek1 Chek2 Csnk2a2 Dclre1a Ddb2 Ddit3 Ercc1 Ercc2 Exo1 Fanca Fancc Fancd2 Fancg Fen1 Gadd45a Gadd45g Hus1 Lig1 Mbd4 Mgmt Mif Mlh1 Mlh3 Mpg Mre11a Msh2 Msh3 Nbn Nthl1 Ogg1 Parp1 Parp2 Pcna Pms1 Pms2 Pold3 Pole Polh Poli Ppm1d Ppp1r15a Prkdc Pttg1 Rad1 Rad17 Rad18 Rad21 Rad50 Rad51 Rad51c Rad51l1 Rad52 Rad9 Rev1 Rnf8 Rpa1 Sirt1 Smc1a Smc3 Sumo1 Terf1

C-abl oncogene 1, receptor tyrosine kinase APEX nuclease (multifunctional DNA repair enzyme) 1 Ataxia telangiectasia mutated homolog (human) Alpha thalassemia/mental retardation syndrome X-linked (RAD54 homolog, S. cerevisiae) BRCA1 associated RING domain 1 Bcl2-associated X protein Bcl-2 binding component 3 Bloom syndrome, RecQ helicase-like Breast cancer 1 Breast cancer 2 Cell division cycle 25 homolog A (S. pombe) Cell division cycle 25 homolog C (S. pombe) Cyclin-dependent kinase inhibitor 1A CHK1 checkpoint homolog (S. pombe) CHK2 checkpoint homolog (S. pombe) Casein kinase 2, alpha prime polypeptide DNA cross-link repair 1A, PSO2 homolog (S. cerevisiae) Damage specific DNA binding protein 2 DNA-damage inducible transcript 3 Excision repair cross-complementing rodent repair deficiency, complementation group 1 Excision repair cross-complementing rodent repair deficiency, complementation group 2 Exonuclease 1 Fanconi anemia, complementation group A Fanconi anemia, complementation group C Fanconi anemia, complementation group D2 Fanconi anemia, complementation group G Flap structure-specific endonuclease 1 Growth arrest and DNA-damage-inducible, alpha Growth arrest and DNA-damage-inducible, gamma HUS1 checkpoint homolog (S. pombe) Ligase I, DNA, ATP-dependent Methyl-CpG binding domain protein 4 O-6-methylguanine-DNA methyltransferase Macrophage migration inhibitory factor MutL homolog 1 (E. coli) MutL homolog 3 (E. coli) N-methylpurine-DNA glycosylase MRE11 meiotic recombination 11 homolog A (S. cerevisiae) MutS homolog 2 (E. coli) MutS homolog 3 (E. coli) Nibrin Nth (endonuclease III)-like 1 (E. coli) 8-Oxoguanine DNA glycosylase Poly (ADP-ribose) polymerase 1 Poly (ADP-ribose) polymerase 2 Proliferating cell nuclear antigen Postmeiotic segregation increased 1 (S. cerevisiae) PMS2 postmeiotic segregation increased 2 (S. cerevisiae) Polymerase (DNA-directed), delta 3, accessory subunit Polymerase (DNA directed), epsilon Polymerase (DNA directed), eta Polymerase (DNA directed), iota Protein phosphatase 1D magnesium-dependent, delta isoform Protein phosphatase 1, regulatory (inhibitor) subunit 15A Protein kinase, DNA activated, catalytic polypeptide Pituitary tumor-transforming 1 RAD1 homolog (S. pombe) RAD17 homolog (S. pombe) RAD18 homolog (S. cerevisiae) RAD21 homolog (S. pombe) RAD50 homolog (S. cerevisiae) RAD51 homolog (RecA homolog, E. coli) (S. cerevisiae) Rad51 homolog c (S. cerevisiae) RAD51-like 1 (S. cerevisiae) RAD52 homolog (S. cerevisiae) RAD9 homolog (S. pombe) REV1 homolog (S. cerevisiae) Ring finger protein 8 Replication protein A1 Sirtuin (silent mating type information regulation 2 homolog) 1 (S. cerevisiae) Structural maintenance of chromosomes 1A Structural maintenance of chromosomes 3 SMT3 suppressor of mif two 3 homolog 1 (S. cerevisiae) Telomeric repeat binding factor (NIMA-interacting) 1

0.31 0.13 0.01⁎ 0.24 0.29 0.005⁎ 0.012⁎

1.14 1.26 7.42 0.88 1.12 1.28 3.05 1.03 1.02 1.03 1.20 1.13 2.96 1.08 1.06 1.11 1.05 1.11 1.30 1.66 1.31 1.28 1.12 1.23 0.91 1.23 1.22 1.25 1.46 1.06 1.23 1.01 1.30 1.13 1.17 1.23 1.16 1.10 1.17 1.19 1.38 1.32 1.17 1.17 1.03 1.13 1.11 1.04 1.29 1.09 1.36 1.06 0.98 1.25 0.96 0.99 1.21 1.30 1.10 1.15 1.05 0.91 1.17 0.94 1.14 1.24 0.94 1.14 1.18 1.05 1.03 1.01 1.06 1.12

0.52 0.39 0.01⁎ 0.81 0.08⁎⁎

1.08 0.88 5.65 0.97 1.20 1.21 2.32 0.93 1.15 0.97 1.11 1.23 4.26 1.15 1.05 1.06 1 1.21 1.17 1.05 0.95 1.34 0.96 0.93 1.18 0.93 0.97 1.14 1.10 1.24 0.90 0.92 1.36 0.81 1.30 0.95 0.75 0.92 0.95 1.19 1.00 1.16 0.92 1.05 0.98 1.04 1.06 1.08 1.21 1.09 1.30 1.02 1.03 0.99 0.90 1.19 1.17 1.06 1.38 1.07 1.01 0.94 0.98 1.18 0.77 1.26 1.24 0.98 0.95 0.96 0.98 0.98 0.97 1.15

0.49 0.78 0.74 0.11 0.39 0.008⁎ 0.34 0.26 0.28 0.71 0.44 0.07⁎⁎ 0.14 0.08⁎⁎ 0.04⁎ 0.35 0.04⁎ 0.60 0.46 0.22 0.21 0.03⁎ 0.63 0.06⁎⁎ 0.89 0.04⁎ 0.46 0.35 0.20 0.23 0.60 0.49 0.44 0.05 0.054⁎⁎ 0.12 0.26 0.67 0.38 0.51 0.56 0.01⁎ 0.37 0.12 0.71 0.94 0.48 0.58 0.88 0.17 0.25 0.47 0.02⁎ 0.84 0.71 0.12 0.78 0.27 0.42 0.57 0.48 0.01⁎ 0.67 0.74 0.63 0.59 0.45

0.25 0.007⁎ 0.93 0.37 0.95 0.35 0.21 0.001⁎ 0.70 0.86 0.39 0.91 0.38 0.28 0.99 0.54 0.06⁎⁎ 0.56 0.52 0.33 0.81 0.81 0.08⁎⁎ 0.33 0.32 0.21 0.66 0.01⁎ 0.07⁎⁎ 0.12 0.56 0.06⁎⁎ 0.63 0.90 0.44 0.85 0.08⁎⁎ 0.34 0.42 0.93 0.94 0.64 0.34 0.045⁎ 0.65 0.24 0.99 0.75 0.94 0.15 0.14 0.10 0.67 0.056⁎⁎ 0.31 0.89 0.74 0.92 0.24 0.05 0.31 0.23 0.86 0.55 0.78 0.96 0.73 0.85 0.03⁎

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Table 1 (continued) 24 h

48 h

Gene symbol

Gene description

P-value

Fold-change

P-value

Fold-change

Topbp1 Tp53 Tp53bp1 Ung Wrn Wrnip1 Xpc Xrcc1 Xrcc2 Xrcc6 Actb B2m Hprt1 Ldha Rplp1

Topoisomerase (DNA) II binding protein 1 Tumor protein p53 Tumor protein p53 binding protein 1 Uracil-DNA glycosylase Werner syndrome Werner helicase interacting protein 1 Xeroderma pigmentosum, complementation group C X-ray repair complementing defective repair in Chinese hamster cells 1 X-ray repair complementing defective repair in Chinese hamster cells 2 X-ray repair complementing defective repair in Chinese hamster cells 6 Actin, beta Beta-2 microglobulin Hypoxanthine phosphoribosyltransferase 1 Lactate dehydrogenase A Ribosomal protein, large, P1

0.02⁎ 0.10 0.25 0.04⁎ 0.07 0.15 0.07 0.14 0.03⁎

1.24 1.11 1.34 1.13 1.11 1.32 1.23 1.24 1.24 1.10 1.13 1.16 1.12 1.07 1

0.89 0.87 0.63 0.77 0.19 0.81 0.30 0.95 0.79 0.65 0.58 0.22 0.85 0.21 0

1.03 1.05 0.93 1.07 0.82 1.03 0.90 1.00 1.02 1.04 1.05 1.27 0.97 0.08 1

0.53 0.44 0.15 0.35 0.58 0

Values represent fold-change ± SEM relative to a control value of 1 (10 ovaries per pool), normalized to Rplp1. Genes with PM-induced changes in abundance are indicated in bold. ⁎ Different from control, P b 0.05. ⁎⁎ Different from control, P b 0.1.

(Irwin et al., 2000; Lissy et al., 2000). The pro-apoptotic actions of P73 involve mitochondrial translocation of BAX, under the direction of P73-induced BBC3 (Yoon et al., 2015). Protein kinase c (PKC) is a serine–threonine kinase that participates in cellular processes such as growth, differentiation and apoptosis (Hug and Sarre, 1993). PKC delta (PKCδ) represents a PKC subfamily and is expressed in the rat ovary (Peters et al., 2000). The catalytic fragment of PKCδ interacts with P73 thereby phosphorylating the transactivation and DNA binding domains of P73 to induce its apoptotic function (Ren et al., 2002). However, PKCδ also enhances proliferation and survival of murine mammary cells (Grossoni et al., 2007), thus indicating that PKCδ has both apoptotic and anti-apoptotic properties. PKCδ has been detected in male germ cells (Um et al., 1995) and PKCδ deficient mice are fertile and develop normally (Leitges et al., 2001). DNA damage can lead to activation of the mitochondrial apoptosis pathway (Roos and Kaina, 2006), in which voltage-dependent anion channel (VDAC) regulates the release of apoptotic proteins, such as cytochrome c (CYCS), from the mitochondria (Igosheva et al., 2010). CYCS release is also regulated by the Bcl-2 family protein, BAX, which interacts with VDAC to increase pore size thus promoting CYCS release, while the anti-apoptotic protein BCL-xL produces the converse effect (Wu et al., 2010). Although it is recognized that infertility ensues from PM treatment, little is known about the earliest mechanistic events that either contribute to ovotoxicity or that could be targeted to provide protection from PM exposure. Our previous evidence using spontaneously immortalized granulosa cells indicated that PM exposure resulted in DNA adduct formation, followed by increased expression of members of the DNA repair response; γH2AX, ATM, PARP1, PRKDC, XRCC6, BRCA1 and RAD51 (Ganesan and Keating, 2015), thus, we hypothesized that similar events would be observed in ex vivo cultured whole ovaries from neonatal rats. We also hypothesized that ATM and PKCδ are important cellular sensors of ovarian DNA damage. This study was designed to investigate the temporal pattern of ovarian cellular signaling events post-PM exposure, using whole neonatal ovarian culture methods, with inclusion of an ATM inhibitor and use of an available mouse strain deficient in PKCδ. 2. Methods and materials 2.1. Reagents Phosphoramide mustard (National 170 Cancer Institute, Bethesda, MD), Bovine serum albumin (BSA), ascorbic acid, transferrin, 2-βmercaptoethanol, 30% acrylamide/0.8% bisacrylamide, ammonium persulfate, glycerol, N′N′N′N′-tetramethylethylenediamine (TEMED), Tris

base, Tris HCL, sodium chloride, and Tween-20 were purchased from Sigma-Aldrich Inc. (St. Louis, MO). Dulbecco's Modified Eagle Medium: nutrient mixture F-12 (Ham) 1× (DMEM/Ham's F12), Albumax, penicillin (5000 U/ml), Hanks' Balanced Salt Solution (without CaCl2, MgCl2 or MgSO4) from Invitrogen Co. (Carlsbad, CA). Millicell-CM filter inserts and 48 well cell culture plates were obtained from Millipore (Bedford, MA) and Corning Inc. (Corning, NY) respectively. RNeasy Mini kit, QIA shredder kit, RNeasy Min Elute kit, RT2 First Strand Kit, RT2 SYBR Green Mastermix, Quantitect™ SYBR Green PCR kit and DNA damage RT2 profiler PCR array kit for rats were purchased from Qiagen Inc. (Valencia, CA). All the primers were purchased from Iowa State University DNA facilities. All primary antibodies with the exception of anti-caspase 3 were purchased from Abcam (anti-ATM (ab78); antiPARP1 (ab6079); anti-RAD51 (ab1837); anti-E2F7 (ab56022); antiP73 (ab40658); anti-BCL2 (ab7973); Cambridge, MA). Anti-caspase 3 primary antibody was purchased from Santa Cruz Biotechnology (sc-22139; Santa Cruz, CA). RNA later was obtained from Ambion Inc. (Austin, TX). The polyclonal goat anti-rabbit secondary were obtained from Pierce Biotechnology (Rockford, IL). Ponceau S was from Fisher Scientific. ECL plus chemical luminescence detection kit was obtained from GE Healthcare, Amersham (Buckinghamshire, UK). KU-55933 was obtained from SelleckChem (Houston, TX).

2.2. Ovary culture Ovaries were collected from PND4 female F344 rats or PKC delta wild type and deficient mice and cultured as described by Devine et al. (2002). Briefly, PND4 female pups were euthanized, ovaries removed, trimmed of oviduct and other excess tissues and placed onto a membrane floating on 250 μl of DMEM/Ham's F12 medium containing 1 mg/ml BSA, 1 mg/ml Albumax, 50 μg/ml ascorbic acid, 5 U/ml penicillin and 27.5 μg/ml transferrin per well in a 48 well plate previously equilibrated to 37 °C. Cultured PND4 rat ovaries were cultured in medium containing vehicle control (DMSO) ± PM (60 μM) and maintained at 37°C with 5% CO2 for 12, 24, 48, or 96 h and medium replaced every 2 d. This concentration was based on our previous studies in which we demonstrated PM-induced follicle loss (Madden et al., 2014). PND4 ovaries from PKC delta wild type and deficient mice were cultured in medium containing vehicle control (DMSO) ± PM (10 μM) and maintained at 37°C with 5% CO2 for 6 d with replacement of media and treatment on alternate days. This concentration was based on the work of others (Desmeules and Devine, 2006; Petrillo et al., 2011). A drop of medium was placed on top of each ovary to prevent it from drying. Control-treated ovaries were maintained in a separate incubator from

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those exposed to PM, due to liberation of a volatile, ovotoxicant from PM (Madden et al., 2014). 2.3. Ovarian follicle counts PND4 female F344 rats ovaries were cultured in medium containing vehicle control (DMSO) ± PM (60 μM) ± KU-55933 (10 nM) and maintained at 37 °C with 5% CO2 for 96 h. The ATM inhibitor KU-55933 concentration was chosen based on our observations that primordial follicular viability was unaffected (data not presented). Ovaries from PKC delta wild type and deficient mice (Gift from Dr. Kanthasamy, Iowa State University) were cultured in vehicle control (DMSO) ± PM (10 μM) for 144 h. The chosen PM concentration of 10 μM was previously characterized to cause follicle depletion from day 2 onward after a single exposure in CD-1 mouse ovaries (Desmeules and Devine, 2006). Following incubation, ovaries were placed in 4% paraformaldehyde for 2 h and transferred to 70% ethanol. Ovaries were embedded in paraffin following standard histological procedures. Tissue sections (5 μM) were cut and stained with hematoxylin and eosin and every 6th section were mounted as described previously (Igawa et al., 2009). Follicle populations were classified as previously described (Keating et al., 2008) and healthy follicles were counted. 2.4. RNA isolation and RT2 profiler PCR array RNA was isolated using an RNeasy Mini kit and the concentration determined using a ND-1000 Spectrophotometer (λ = 260/280 nm; NanoDrop Technologies, Inc., Wilmington, DE). Three biological replicates were used to perform a RT2 profiler DNA damage PCR array. The PCR array contained 96-wells, each containing a gene-specific primer set, therefore one plate tested 96 genes per each sample. Total ovarian RNA (1 μg) was reverse transcribed to cDNA using an RT2 first-strand kit, combined with an appropriate RT2 SYBR Green Mastermix before being aliquoted into the wells of the RT2 profiler PCR array. The regular cycling program consisted of 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. Each gene was normalized to ribosomal protein, large, P1 (Rplp1), as recommended by the company-provided analysis software. There was no effect of PM exposure on ovarian mRNA levels of either of these two housekeeping genes. The SA Biosciences RT2 Profiler PCR Array Data Analysis software quantified the changes in mRNA levels using the 2−ΔΔCt method. 2.5. cDNA amplification Total RNA (1 μg) was reverse transcribed to cDNA utilizing the Superscript III One-Step qRT-PCR kit. cDNA was diluted (1:20) in RNasefree water and amplified on an Eppendorf PCR Master cycler using a Quantitect SYBR Green PCR kit. Primers for Atm, Prkdc, Xrcc6, Parp1, Rad51, Brca1 and β-actin were designed by Primer 3 Input Version (0.4.0) as previously described (Ganesan and Keating, 2015). 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. Primers were validated to produce a single product by analysis of a melt curve post amplification. Each sample was normalized to β-actin before quantification. Quantification of fold-change in gene expression was performed using the 2−ΔΔCt method (Livak and Schmittgen, 2001; Pfaffl, 2001).

and protein concentration was measured using a BCA protocol. Protein was stored at −80°C until use. SDS-PAGE was used to separate proteins which were then transferred to a nitrocellulose membrane. Membranes were blocked for 1 h in 5% milk in Tris-buffered saline containing Tween-20, followed by incubation with anti-PARP1 antibody (1:200), anti-ATM antibody (1:100), anti-RAD 51 antibody (1:500), anti-E2F1 antibody (1:100), anti-P73 antibody (1:200), anti-BCl2 antibody (1:200) and anti-CASP3 antibody (1:50) overnight at 4°°C. Following three washes in TTBS (1×), membranes were incubated with an appropriate species-specific secondary antibody (1:2000) for 1 h at room temperature. Membranes were washed three times in TTBS, once in TBS and then incubated in enhanced chemiluminescence detection substrate for 5 min and exposed to X-ray film. Densitometry of the appropriate sized bands was measured using Carestream molecular imaging software version 5.0 (Carestream Health Inc., Rochester, NY) which eliminates background noise. Proteins of interest were normalized to Ponceau S measurement by dividing the protein of interests densitometric value by a densitometric value obtained by measuring the entire protein staining per lane on the Ponceau S stained membrane. 2.7. Statistical analysis Treatment comparisons for follicle count experiments were performed using one-way analysis of variance. Quantitative RT-PCR and western blot data were analyzed by t-test comparing treatment with control raw data at each individual time point. RT2 Profiler PCR array data were analyzed using the online SA Biosciences software which quantified the changes in mRNA levels using the 2−ΔΔCt method. All other statistical analysis was performed using Prism 5.04 software (GraphPad Software). Statistical significance was defined as P b 0.05, with a trend for a difference considered at P b 0.1. For graphical purposes, protein expression is presented as a mean ± SE, relative to the respective control. 3. Results 3.1. Impact of PM exposure on abundance of genes encoding proteins involved in the ovarian response to DNA damage Fisher 344 PND4 rat ovaries were cultured for 24 or 48 h in medium containing vehicle control ±60 μM PM. RNA was isolated and used to perform RT2 Profiler PCR array for genes involved in the response to DNA damage. After 24 h of PM exposure, of the 89 genes tested, fourteen genes had increased (P b 0.05) gene expression, and four genes were trending (P b 0.1) toward being increased, compared with controltreated ovaries. After 48 h of PM exposure, five genes had increased (P b 0.05) abundance and 1 gene was decreased in response to PM exposure. Seven genes had a tendency (P b 0.1) for increased abundance relative to control-treated ovaries (Table 0.71). Interestingly, four

2.6. Protein isolation and western blot Total protein was isolated from cultured ovaries (n = 3; 10 ovaries per pool via homogenization in tissue lysis buffer containing protease and phosphatase inhibitors as previously described (Ganesan et al., 2013). Briefly, homogenized samples were placed on ice for 30 min, followed by two rounds of centrifugation at 10,000 rpm for 15 min

Fig. 1. Effect of PM exposure on ovarian DSBs repair gene mRNA abundance. Fisher 344 PND4 rat ovaries were cultured for 12 h in medium containing vehicle control ±60 μM PM. RNA was isolated and used to perform qRT-PCR. Values are expressed as mean fold change relative to control treated ovaries ± SE; n = 3 (10 ovaries per pool). Statistical significance was defined as * = P b 0.05.

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69

Fig. 2. Impact of PM exposure on transcription factor and apoptotic gene mRNA level. Fisher 344 PND4 rat ovaries were cultured for (A) 24, (B) 48 or (C) 96 h, in medium containing vehicle control ±60 μM PM. RNA was isolated and used to perform qRT-PCR. Values are expressed as mean fold change relative to control treated ovaries ± SE; n = 3 (10 ovaries per pool). Statistical significance was defined as * = P b 0.05.

genes had increased abundance (P b 0.05) at both PM-exposure time points: Atm, Bbc3, Mgmt, and Pold3 (Table 1). In addition to genes present on the PCR array, primers were designed to quantify changes in genes that were of interest but were either absent from the array or the array was not performed at the time point of interest (12 h). Following 12 h of PM exposure, Atm (3.4 fold ± 0.7); Parp1 (1.6 ± 0.5 fold); Prkdc (0.9 ± 0.3 fold); Xrcc6 (1.4 ± 0.3 fold); Rad51 (1.9 ± 0.5 fold) and Brca1 (1.9 ± 0.6 fold) mRNA abundance were increased (P b 0.05) compared to control (Fig. 1). Following 24 h of PM exposure, E2f1 (1.65 ± 0.15 fold), Mdm2 (0.7 ± 0.21 fold) and p73 (3.06 ± 0.7 fold) mRNA levels were increased (P b 0.05) compared to control. In contrast, Vdca1 (0.4 ± 0.1 fold) mRNA abundance was decreased (P b 0.05) compared to control. There was no impact of PM exposure on Cycs mRNA level (Fig. 2A). After 48 h of PM exposure, E2f1 (1.4 ± 0.2 fold); p73 (90 ± 1 fold); Mdm2 (0.7 ± 0.2 fold); Vdca1 (0.4 ± 0.2 fold) and Cycs (2.5 ± 0.5 fold) mRNA levels were increased (P b 0.05) compared to control-treated ovaries (Fig. 2B). 96 h post-PM exposure, increased (P b 0.05) levels of E2f1 (1.3 ± 0.2 fold); p73 (1.4 ± 0.5 fold); Mdm2 (1 ± 0.07 fold); Vdca1

(0.3 ± 0.06 fold) and Cycs (0.7 ± 0.1 fold) mRNA were observed compared to control (Fig. 2C).

3.2. Effect of exposure to PM on abundance of DNA damage response proteins Total ovarian protein was isolated following 96 h of PM exposure and western blot analysis performed. The DNA damage repair proteins ATM (PM: 1.6 ± 0.3; CT: 1.4 ± 0.02); PARP1 (PM: 1.8 ± 0.03; CT: 1.5 ± 0.02) were increased (P b 0.05) compared to ovaries cultured in control media. In contrast, RAD51 (PM: 1.4 ± 0.02; CT: 1.5 ± 0.02) protein abundance was decreased (P b 0.05) after PM exposure (Fig. 3A).The transcription factors E2F7 (PM: 1.5 ± 0.06; CT: 0.8 ± 0.04); P73 (PM: 1.9 ± 0.1; CT: 1.3 ± 0.03) and apoptotic proteins were increased (P b 0.05) 96 h post-PM exposure and CASP3 (PM: 1.5 ± 0.02); CT: 1.3 ± 0.01) was increased (P b 0.05), relative to control-treated ovaries. In contrast, the anti-apoptotic protein BCL2 (PM: 0.9 ± 0.05; CT: 1.2 ± 0.02) was decreased (P b 0.05) after PM exposure compared to control ovaries (Fig. 3B).

Fig. 3. PM exposure effects on DNA damage response protein level. Following 96 h of PM exposure, protein was isolated and used to perform western blot analysis for (A) DNA damage response proteins, (B) transcription factor proteins, (C) pro-apoptotic CASP3 and (D) anti-apoptotic BCL-2. Results were normalized to Ponceau S densitometric staining and expressed as mean raw data ± SE; n = 3 (10 ovaries per pool). Statistical significance was defined as * = P b 0.05.

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3.3. Involvement of ATM protein in ovotoxicity induced by PM exposure Ovaries were cultured in medium containing vehicle control (DMSO) ± PM (60 μM) ± KU-55933 (10 nM) for 96 h, and follicles counted. Primordial follicle number (PM: 279.7 ± 7.8; CT: 568 ± 26.2; AI: 578 ± 49.1; PM + AI: 536 ± 9.5) was reduced (P b 0.05) after PM treatment. There was no impact of ATM inhibition on primordial follicle number. Interestingly, in the presence of ATM inhibition, primordial follicles were protected from PM-induced depletion (Fig. 4A). PM exposure reduced (P b 0.05) small primary follicle number (PM: 40.3 ± 8; CT: 128.7 ± 5.8; AI: 124 ± 4.7; PM + AI: 69 ± 7.2), compared to all other groups. There was no impact of ATM inhibition on small primary follicle number, however, partially prevention (P b 0.05) of PM-induced destruction of small primary follicles was observed (PM + AI: 69 ± 7.2; PM: 40.3 ± 8) (Fig. 4B). A similar pattern was observed for both large primary and secondary follicles. PM exposure depleted (P b 0.05) large primary follicles number (PM: 13 ± 3.2; CT: 32.6 ± 2.3; AI: 51.6 ± 2.9; PM + AI: 32.3 ± 4.4). Interestingly, ATM inhibition increased large primary follicle numbers (AI: 51.6 ± 2.9; PM: 13 ± 3.2; CT: 32.6 ± 2.3; PM + AI: 32.3 ± 4.4), compared to all other groups (Fig. 4C), and prevented PM-induced depletion of large primary follicles (PM + AI: 32.3 ± 4.4; PM: 13 ± 3.2). Secondary follicles numbers were reduced (P b 0.05) by PM exposure (PM: 2.6 ± 1.4; CT: 20.6 ± 3.8; AI: 18.6 ± 2.1). ATM inhibition

alone did not impact secondary follicle number (CT: 20.6 ± 3.8; AI: 18.6 ± 2.1), but partially protected secondary follicles from PMinduced ovotoxicity (PM: 2.6 ± 1.4; PM + AI 12.3 ± 1.7) (Fig. 4D). 3.4. Impact of PKC delta deficiency on PM-induced ovotoxicity PKC delta (PKCδ) wild type (WT) and deficient (KO) mouse ovaries were cultured with vehicle control (DMSO) ± PM (10 μM) for 6 d. PKC delta deficiency alone reduced (P b 0.05) ovarian follicle number, relative to the WT mice; primordial (WT: 193 ± 39.4; KO: 84 ± 22.3), small primary (WT: 59.2 ± 14.2; KO: 24 ± 3.9), large primary (WT: 28 ± 4.6; KO: 18.4 ± 5.1) and secondary (WT: 6.6 ± 1.4; KO: 0.4 ± 0.4) follicles were all lower than those contained in ovaries from WT mice (Fig. 5A). The percentage of PM-induced follicle loss was calculated in both genotypes and no impact of PKCδ deficiency on PM-induced follicle loss was observed (Fig. 5B). 4. Discussion PM, an alkylating agent, causes both dose- and time-dependent ovarian primordial and primary follicle loss following DNA damage in mice and rat oocytes (Petrillo et al., 2011). In this study, we explored mechanisms involved with PM-induced ovarian follicle loss and the response to DNA damage in neonatal cultured rat ovaries and have

Fig. 4. ATM inhibition effect on ovotoxicity induced by PM. Ovaries were cultured in medium containing vehicle control (DMSO) ± PM (60 μM) ± KU-55933 (10 nM) for 96 h. Following culture, follicles were classified and counted. Total number of primordial (A), small primary (B), large primary (C) and secondary (D) follicles are presented as mean raw data ± SE; n = 5. Different letters indicates the statistical differences from control treated ovaries at P b 0.05. (E) Histological ovarian sections of the control treated (CT), PM treated (PM), ATM inhibitor treated (AI) and PM plus ATM inhibitor treated (PM + AI) are presented.

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summarized our findings in Fig. 6. Using this system, we previously demonstrated that PM caused depletion of all follicle types from day 4 onwards (Madden et al., 2014). We chose to perform the analyses described in this study on days prior to and at the time of observed follicle loss (12–96 h). Two approaches were employed to determine transcriptional effects of PM exposure on genes involved in the ovarian response to DNA damage. We first used a commercially available PCR array to survey impacts of PM exposure on a wide number of genes at either 24 or 48 h postexposure. A dynamic response in terms of gene activation was observed in that genes that were altered after 24 h included Atm, Bax, Cdkn1a, Ddit3, Ercc1, Fancc, Gadd45g, Lig1, Nthl1, Pold3, Rad21, Rpa1, Ung and Xrcc2. After 48 h of exposure, genes that remained elevated included Atm, Cdkn1a, Nthl1, and Pold3, however, a number of genes returned to control levels and genes that were increased at 48 h but not 24 h were Bard1, Gadd45a, Mif, Mlh3, Rad18 and Terf1. The sole gene for which PM exposure decreased abundance was Mpg. We chose to also examine a number of genes after 12 h of PM exposure by traditional PCR that we have previously determined to be induced by PM exposure in rat ovarian granulosa cells (Ganesan and Keating, 2015) and found that Atm, Parp1, Prkdc, Xrcc6, Rad51 and Brca1 were elevated in mRNA abundance, and with the exception of Atm, all were returned to control level by the 24 h time point. Thus, a temporal ovarian response to PM exposure was evident. Activation of these genes support that DNA DSBs were induced by PM exposure as seen in other experimental paradigms (Durocher and Jackson, 2001; Ganesan and Keating, 2015). The data also indicates that PM-induced ovarian DNA damage was occurring prior to activation of the ovarian DNA repair mechanism and, furthermore, support that PM induces DNA damage as an upstream event of PM-induced follicle loss and that the ovary activates DNA repair genes to protect against PMinduced ovotoxicity. We chose a number of proteins representative of the DNA damage response (ATM, PARP1, RAD51), apoptosis (CASP3 and BCL-2) as well as those that regulate these physiological processes (E2F7 and P73) and quantified mRNA expression 24, 48 and 96 h post-PM exposure as well as protein abundance after 96 h of PM exposure. As a reminder PM-induced follicle loss is observed at the 96 h time point, and proteins that are involved in the ovarian response to PM exposure are anticipated to be increased at this time of exposure. ATM and PARP1 were increased by PM exposure, while RAD51 was decreased. The decreased in RAD51 was unexpected and could indicate that examining RAD51 protein abundance at an earlier time point would be beneficial, as we previously observed in cultured PM exposed rat granulosa cells (Ganesan and Keating, 2015). The increase in ATM and PARP1, however, further support that the ovary is mounting a protective response to an environmental exposure that has caused an assault on the germline DNA. Unsurprisingly, increased pro-apoptotic CASP3 and decreased anti-apoptotic BCL-2 were observed after PM exposure. Additionally, PM exposure increased protein levels of the transcriptional regulators, E2F7 and P73. Upon phosphorylation of P53 by ATM, P53 becomes stabilized and blocks cell proliferation by up-regulation of P21 (CDKN1a), which triggers G1-S arrest (Roos and Kaina, 2006). In this study, Cdkn1a gene was increased after 24 h of PM exposure but no alterations in p53 mRNA abundance were evident. Low levels of P53 during DSBs are sufficient to propel the transcription of the Cdkn1a gene to result in cellcycle arrest (Roos and Kaina, 2006). In addition, both ATM and ATR phosphorylate checkpoint kinases 1 and 2 (CHK1 and CHK2) to activate the transcriptional regulator E2F1 upon DNA damage and E2F1 then can induce apoptosis through a P53-independent pathway via activation of P73 (Phillips et al., 1997). In human cancer cell lines, E2F1 was activated by etoposide which in turn stimulated the transcription of p73 gene (Urist et al., 2004). In this study, both E2f1 and p73 mRNA levels were increased after 24 h and E2F7 and P73 proteins were elevated after 96 h of PM exposure,

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potentially indicating that PM induces apoptosis via P73. MDM2 degrades P53 using a negative feedback loop resulting in a decrease in exogenous P53 in transient transfection experiments or in endogenous P53 induced by DNA damage (Wang et al., 2001). The transcriptional level of Mdm2 was increased after 24 h while there were no changes in p53 mRNA expression which might be due to increased P73. We did not examine the P53 protein abundance in this study and complete understanding of P53 involvement cannot be achieved by simply

Fig. 5. Impact of PKCδ deficiency on PM-induced ovotoxicity. Ovaries from PKCδ wild type (WT) or knockout (KO) mice were cultured with vehicle control (DMSO) ± PM (10 μM) for 6 d Following culture, follicles were counted between (A) treatments and strains and (B) the percentage reduction in follicle stage number calculated. The total number of primordial, small primary, large primary and secondary follicles are expressed as mean raw data ± SE; n = 5. Statistical significance from the respective control was defined as * = P b 0.05. Different letters indicates the statistical differences in follicle number between strains at P b 0.05. (C) Histological ovarian sections of the wild type control treated (WT CT), wild type PM treated (WT PM), knockout control treated (KO CT) and knockout PM treated (KO PM) are presented.

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examining mRNA abundance, thus, future studies will include analysis of P53 and MDM2 protein abundance and phosphorylation to determine post-translational abundance due to PM exposure. During P73-mediated apoptosis, transcriptional up regulation of Bbc3 resulted in increased BAX mitochondrial translocation and CYCS release (Melino et al. 2004). We observed that PM increased Bbc3 and Bax gene expression after 24 h of exposure, but decreased BCL2 protein level after 96 h. This observation is reminiscent of a study in which docetaxel/doxorubicin chemotherapy decreased BCL2 in human breast cancer cells (Buchholz et al., 2003). BCL-2 inhibits apoptosis by sequestering the CASP cysteine proteases or through preventing the release of mitochondrial apoptogenic factors such as CYSC into the cytoplasm (Tsujimoto, 1998). This study indicates that PM activates Bbc3 and Bax and decreases BCL2 to elicit mitochondrial apoptotic cell death. VDAC1 is a key player in mitochondria-mediated apoptosis, participating in the release of mitochondrial pro-apoptotic CYCS to the cytosol (Zaid et al., 2005). We found that PM increased both Vdac1 and Cycs mRNA level after 24 h in rat ovaries. These data are similar to a study that demonstrated increased Vdac1 expression after endostatin exposure which stimulated apoptosis in endothelial cells (Yuan et al., 2008). CASP3 protein observed herein to be increased after 96 h by PM exposure, the time point at which PM-induced ovotoxicity is occurring (Madden et al., 2014), supporting the involvement of CASP3 in PM-induced follicle loss. Increased CASP3 has also been demonstrated during apoptosis induced by another ovotoxicant, 7,12dimethylbenz[a]anthracene (Tsai-Turton et al., 2007; Ganesan et al., 2013). Since Atm mRNA expression was increased at the earliest time point examined (12 h) and was sustained at an elevated level over the time course of PM exposure, we investigated a functional role for ATM during

PM exposure using the ATM inhibitor KU-55933. Interestingly, though no apparent impact of ATM inhibition on primordial, small primary or secondary follicles was observed, lack of ATM increased the number of large primary follicles, potentially indicating a role for ATM in viability and/or maturation of this follicular subtype. We originally expected that absence of ATM would result in enhanced follicle loss induced by PM, however, the opposite was the case. Lack of ATM protected all follicle types PM-induce follicle depletion. This result was similar to a previous study that showed that ATM inhibitor improved oocyte survival after doxorubicin exposure by activation of c-AbI–Tap63 pathway (Soleimani et al., 2011). As a testament to its importance in maintaining cell viability, blocking ATM activity caused induction of Atm transcription, regulated by p73 and E2f1 (Khalil et al., 2012). These results are in correlation with our findings in the ovary, and suggest that ATM serves to eliminate cells with excessive DNA damage (Herzog et al., 1998). It remains unclear as to whether the retained follicles are in fact healthy or contain germline DNA damage. Thus, an interesting follow up study will be to determine if increased oocyte or granulosa cell DNA damage is evident in ovaries treated with both the ATM inhibitor and PM, work which is underway by our group. PKCδ, a member of the novel PKC subfamily, is actively involved in cell apoptosis in response to genotoxic and oxidative stress (Cross et al., 2000). PKCδ phosphorylates P73 resulting in P73 accumulation (Ren et al., 2002). Interestingly, PKCδ expression has recently been demonstrated to be positively regulated by ATM (Yu et al., 2015). Since P73 is increased by PM, we exposed PKCδ wild type and deficient mice to PM in order to deduce any involvement of PKCδ in PM-induced ovotoxicity. The absence of PKCδ reduced the numbers of all stage follicles compared to wild type ovaries, suggesting that PKCδ has importance in oogenesis or folliculogenesis. When these mice were exposed to PM, the number

Fig. 6. Working model of PM-induced ovotoxicity. 1) PM induces DNA double strand breaks resulting in 2) activation of ATM. 3) ATM halts the cell cycle and recruits DNA repair proteins to the site of the break. 4) If the DNA damage is too great for repair or the cell becomes overwhelmed by repeated exposures, E2F7 is activated and drawn to the P73 promoter, P73 protein level increased resulting in increased expression of Bbc3 and Bax, mitochondrial translocation of BAX, leakage of cytochrome c, activation of the executioner caspase 3, and resultant follicle loss. 5) Inhibition of ATM prevents follicle loss, likely due to lack of a coordinated cellular response to destroy follicles containing DNA damage.

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of healthy follicles was reduced by PM in both the wild type and deficient mice. However, based upon the percentage reduction in follicle number induced by PM relative to the control stage within genotype, there was no effect of PKCδ deficiency on PM-induced ovotoxicity. These results indicate that PKCδ alone is important for ovarian follicle survival and potentially ovarian follicle endowment, but that there was no apparent influence of PKCδ deficiency on PM-induced ovotoxicity. PKCδ has been demonstrated to enhance proliferation and survival of murine mammary cells (Grossoni et al., 2007) and human breast cancer cell lines (McCracken et al., 2003) indicating a role for PKCδ in cell viability. As summarized in Fig. 6, the ovary induces increased levels of mRNA and proteins involved in the response to DNA damage following PM exposure, suggesting that induction of DNA damage is an initiating event of PM-induced ovotoxicity. However, DNA repair components were reduced and cell death mechanisms were increased at the time of follicle loss. PM-induced apoptosis appears P53-independent but is potentially occurring through the E2F1-P73 dependent mitochondrial apoptotic pathway. Importantly, we have demonstrated that inhibition of ATM prevented PM-induced oocyte elimination, which may not necessarily represent a benefit to the germline, and is an area that requires further examination. Although fertility is reportedly unaffected by PKCδ deficiency (Leitges et al., 2001), there may be impacts on either ovarian endowment or folliculogenesis, however there was no impact of PKCδ deficiency on PM-induced ovotoxicity. The data shed additional light on potential targets for amelioration of PM-induced infertility that occurs as a side effect to anti-neoplastic therapy. Funding This work was partially supported by the ‘National Institute of Environmental Health Sciences at the National Institutes of Health’ (grant number R00ES016818). Conflict of interest statement There are no conflicts of interest to disclose. Transparency document The Transparency document associated with this article can be found in the online version. References Ataya, K.M., Pydyn, E.F., Ramahi-Ataya, A.J., 1990. The effect of “activated” cyclophosphamide on human and rat ovarian granulosa cells in vitro. Reprod. Toxicol. (Elmsford, N.Y.) 4, 121–125. Brunner, H.I., Bishnoi, A., Barron, A.C., Houk, L.J., Ware, A., Farhey, Y., Mongey, A.B., Strife, C.F., Graham, T.B., Passo, M.H., 2006. Disease outcomes and ovarian function of childhood-onset systemic lupus erythematosus. Lupus 15, 198–206. Buchholz, T.A., Davis, D.W., McConkey, D.J., Symmans, W.F., Valero, V., Jhingran, A., Tucker, S.L., Pusztai, L., Cristofanilli, M., Esteva, F.J., Hortobagyi, G.N., Sahin, A.A., 2003. Chemotherapy-induced apoptosis and Bcl-2 levels correlate with breast cancer response to chemotherapy. Cancer J. (Sudbury, Mass.) 9, 33–41. Chandra, A., Copen, C.E., Hervey Stephen, E., 2013. In: Services, H.A.H. (Ed.)Infertility and Impaired Fecundity in the United States, 1982–2010: Data From the National Survey of Family Growth 20782. Centers for Disease Control and Prevention, Hyattsville, MD, pp. 1–17. Cross, T.G., Scheel-Toellner, D., Henriquez, N.V., Deacon, E., Salmon, M., Lord, J.M., 2000. Serine/threonine protein kinases and apoptosis. Exp. Cell Res. 256, 34–41. Desmeules, P., Devine, P.J., 2006. Characterizing the ovotoxicity of cyclophosphamide metabolites on cultured mouse ovaries. Toxicol. Sci. 90, 500–509. Devine, P.J., Sipes, I.G., Skinner, M.K., Hoyer, P.B., 2002. Characterization of a Rat in Vitro Ovarian Culture System to Study the Ovarian Toxicant 4-Vinylcyclohexene Diepoxide. Toxicol. Appl. Pharmacol. 184 (2), 107–115. Durocher, D., Jackson, S.P., 2001. DNA-PK, ATM and ATR as sensors of DNA damage: variations on a theme? Curr. Opin. Cell Biol. 13, 225–231. Friedberg, E.C., 2003. DNA damage and repair. Nature 421, 436–440. Ganesan, S., Keating, A.F., 2015. Phosphoramide mustard exposure induces DNA adduct formation and the DNA damage repair response in rat ovarian granulosa cells. Toxicol. Appl. Pharmacol. 282 (3), 252–258 (Feb 1).

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