FOLLICLE PROTECTIVE ACTION OF TAMOXIFEN IN DOG OVARIAN EXPLANT TISSUE CULTURE IN PRESENCE OF CYCLOPHOSPHAMIDE By Puja Basu A DISSERTATION Michigan State University in partial fulfillment of the requirements for the degree of Comparative Medicine and Integrative Biology – Doctor of Philosophy Submitted to 2020 ABSTRACT FOLLICLE PROTECTIVE ACTION OF TAMOXIFEN IN DOG OVARIAN EXPLANT TISSUE CULTURE IN PRESENCE OF CYCLOPHOSPHAMIDE By Puja Basu Women exposed to cytotoxic chemotherapy are at risk of amenorrhea and premature ovarian failure due to the side effects of treatment. Few options are available to women who wish to preserve their germline, including embryo cryopreservation and ovary cryopreservation, however these procedures are expensive, invasive, often requires delay of treatment and the presence of a partner. Most importantly all these options fail to protect ovarian function. Thus, there is a clear need to investigate an alternate strategy to protect the ovary from the cytotoxic effects so as to maintain oocyte quality as well as endogenous hormone production to support follicular growth and for maintenance of pregnancy. Recent studies in rodents have demonstrated that tamoxifen treatment prevented follicular loss and apoptosis from chemotherapeutic drugs and radiation- mediated injury. However, rodent models may not predict clinical outcomes. There is a need for a clinical model which would reflect clinical efficacy in humans. We evaluated the ovarian-sparing effect of tamoxifen on ovarian follicle reserve (primordial and primary follicles) of dogs. Ovarian cortical explant tissues from pre-pubertal dogs (N = 10) were collected at elective ovariohysterectomy and randomly assigned and treated for 72 hours with active metabolites of alkylating agent cyclophosphamide (4-hydroxycyclophosphamide; CTX) (0, 1, and 10 µM; C, CTXL, and CTXH) and tamoxifen (4-hydroxytamoxifen; TAM) (0 and 10 µM; C and TAM). Results presented as total normal follicle reserve based on histological morphology. High dose CTXH caused marked follicular loss (P < 0.05) whereas treatment with TAM decreased follicular loss (P < 0.05) from CTX in-vitro. TAM alone did not have an effect on morphologically normal reserve follicle counts (P > 0.05). This work also developed a dog ovarian cortical explant model to test efficacy and action of drugs in a model that share some genetic, physiological and environmental similarities with humans, have increased life spans and a propensity for tumors. Another goal of this study was to characterize anti-Mullerian hormone (AMH) in dogs with relation to puberty, age, numbers of reserve follicles, stage of estrus cycle and spay status to determine usefulness of AMH as an indicator of ovarian reserves and fertility in dogs. The data presented will show that spayed and intact female dogs can be successfully identified using a readily available human AMH ELISA. The results also demonstrated a significant negative relationship between AMH and advancing age and that there is a significant difference in serum AMH concentrations at anestrus and proestrus. Taken together, the results demonstrate that dog ovarian cortical explant culture system may be an alternative to test effects and actions of chemotherapeutic agent. TABLE OF CONTENTS LIST OF FIGURES KEY TO ABBREVIATIONS CHAPTER 1. INTRODUCTION AND LITERATURE REVIEW OVARY Anatomy and histology of dog ovaries and follicles Follicle stimulating hormone (FSH) in follicle development and senescence Anti-Mullerian hormone (AMH) in follicle development and senescence vii ix Production and function Clinical uses of AMH in humans Clinical uses of AMH in dogs Predicting ovarian reserves Effects of xenobiotics on ovaries 1 2 2 3 4 4 5 5 6 7 8 8 9 10 10 11 12 Metabolism of TAM Actions of TAM Current uses in humans Tamoxifen and ovarian function TAMOXIFEN GOALS OF THIS THESIS REFERENCES CHAPTER 2. DOG PRIMORDIAL AND PRIMARY FOLLICLES MAINTAIN VIABILITY IN IN-VITRO OVARIAN CORTICAL EXPLANT AGAROSE GEL CULTURE SYSTEM INTRODUCTION MATERIAL AND METHODS Collection of ovaries Processing of ovary Culture media In-vitro culture systems Processing ovary for histological sections Statistical analysis 17 18 21 21 22 22 22 23 24 25 25 Loss of morphologically normal follicles after 72 hours of culture 27 Agarose gel improved primordial and primary follicle survival 27 29 30 31 32 RESULTS DISCUSSION CONCLUSION APPENDIX Appendix 2.1: Hematoxylin and eosin staining protocol REFERENCES CHAPTER 3. PROTECTIVE ACTION OF TAMOXIFEN IN DOG OVARIAN EXPLANTS CULTURED IN PRESENCE OF CYCLOPHOSPHAMIDE 36 iv INTRODUCTION MATERIAL AND METHODS Collection of ovaries Processing ovaries for in-vitro culture Culture media In-vitro ovarian culture Processing ovaries for histology Apoptosis evaluation Statistical analysis 37 40 40 40 41 41 42 43 44 44 44 45 47 48 50 51 52 53 58 59 61 61 61 62 63 63 63 64 64 65 66 67 68 68 70 73 74 75 76 77 RESULTS Loss of follicles after 72 hours in in-vitro explant tissue culture Co-administering tamoxifen with cyclophosphamide protects dog ovarian reserves Apoptosis marker did not show recovery of follicles on coadministration of TAM-CTX Appendix 3.1: Hematoxylin and eosin staining protocol DISCUSSION CONCLUSION APPENDIX Animals Collection of blood samples and ovaries Processing ovaries for histology Serum AMH enzyme-linked immunosorbent assay Progesterone assay Statistical analysis REFERENCES CHAPTER 4. ANTI-MULLERIAN HORMONE AS AN INDICATOR OF OVARIAN RESERVES IN DOMESTIC DOGS INTRODUCTION MATERIAL AND METHODS RESULTS DISCUSSION CONCLUSION APPENDICES REFERENCES Commercially available serum AMH assay identified spayed dogs Serum AMH differed between prepubertal (Pre-P) and post pubertal (Post-P) dogs AMH varies with age Significant AMH variation is seen in dogs that are in baseline and proestrus Total follicle numbers (TFN) negatively corelate with advancing age AMH and TFN are not positively correlated Appendix 4.1: Hematoxylin and eosin staining protocol Appendix 4.2: VDL Endocrinology laboratory Progesterone assay ranges and their implication for breeding v CHAPTER 5. SUMMARY AND FUTURE DIRECTIONS SUMMARY FUTURE DIRECTIONS Optimize the ovarian cortical explant culture system Determine the mechanism of action of TAM Better characterization of circulating AMH in dogs REFERENCES 89 90 90 90 91 92 93 vi LIST OF FIGURES Figure 2.1. Bar graph of total normal primordial and primary follicles of dog ovarian tissue cultured for 0 hour and 72 hours in 1.0% agarose gel (1.0%AG), 1.5% agarose gel (1.5%AG) and cell inserts coated with fibronectin (INSRT). Data is presented as mean and standard error of mean. Different letters indicate significant differences (P < 0.05). Figure 2.2. Bar graph of morphological normal follicles of dog ovarian tissue cultured for 0 hour and 72 hours in 1.0% agarose gel (1.0%AG), 1.5% agarose gel (1.5%AG) and cell inserts coated with fibronectin (INSRT). Data is presented as mean and standard error of mean. Different letters indicate significant differences (P < 0.05). Figure 3.1. Primordial and primary follicles counted in ovarian sections at 0 hours and after 72 hours of culture at 38.5 °C in 5% CO2 on 1.5% agarose gel pads. Data is presented as mean and standard error of mean. Different letters indicate significant differences (P < 0.05). Figure 3.2. Effect of 4-hydroxytamoxifen (TAM; 10 µM) and/or 4-hydroxycycloophosphamide (CTXL; 1 µM and CTXH; 10 µM) on the number primordial follicles in cultured dog (N = 10) ovaries after 72 hours. Values are presented as mean ± SEM follicles counted (n=10). CTRL: control group. Different letters indicate significant differences (P < 0.05). Figure 3.3. Effect of 4-hydroxytamoxifen (TAM; 10 µM) and/or 4-hydroxycyclophosphamide (CTXL; 1 µM and CTXH; 10 µM) on the number primordial and primary follicles in cultured dog ovaries after 72 hours. Values are presented as mean ± SEM follicles counted (n=10). CTRL: control group. Different letters indicate significant differences (P < 0.05). Figure 3.4. Effect of 4-hydroxytamoxifen (TAM; 10 µM) and/or 4-hydroxycyclophosphamide (CTXL; 1 µM and CTXH; 10 µM) on the number of apoptotic Caspase-3 positive primordial and primary follicles in cultured dog ovaries after 72 hours. Values are presented as mean ± SEM follicles counted (n=10). CTRL: control group. Different letters indicate significant differences (P < 0.05). Figure 4.1. Serum AMH values in spayed and intact dogs ranging between 6 months to 6 years. Different letters indicate significant differences (P < 0.05) between the two groups. Figure 4.2. Serum AMH values in prepubertal (< 6 months) and post-pubertal 65 vii 26 26 45 46 47 48 66 67 dogs (> 6 months to 6 years). Different letters indicate significant differences (P < 0.05) between the two groups. Figure 4.3. Serum AMH values in dogs ranging from 1 month old to 14 years age Figure 4.4. Serum AMH values in dogs (n= 40, 6 months – 6 years) significantly varied in dogs that were baseline (n = 26) and in proestrus (n = 14). Different letters indicate significant differences (P < 0.05) between the two groups. Figure 4.5. Total follicle numbers for dogs (N = 55) aged between 1 month to 14 years. There is significant negative correlation (r = -0.059, P < 0.0001) of total follicle numbers with advancing age. Figure 4.6. Serum AMH and reserve follicles did not show positive correlation (r = - 0.36, P < 0.05) with advancing age (N = 55). 68 69 69 viii AMH CTX CYP DMBA GnRH TAM KEY TO ABBREVIATIONS Anti-Mullerian hormone 4-hydroxycyclophophamide Cytochromes P450s 7,12-dimethylebenz[α]anthracene Gonadotropin-releasing hormone 4-hydroxytamoxifen ix CHAPTER 1. INTRODUCTION AND LITERATURE REVIEW 1 OVARY Anatomy and histology of dog ovaries and follicles Dog ovaries are paired organs, that fulfill both an endocrine and exocrine function. The ovary can be classified into two zones, the cortex and the medulla (Fiori, 2000). The cortex in turn is covered by a surface epithelium and contains follicles in various stages of development, corpora lutea, interstitial cells, interstitial glands and stroma (Fiori, 2000; Junqueira and Carneiro, 2004; Priedkalns, 1982). Follicles are the basic functional unit in an ovary. Primordial follicles are described as a follicle containing a primary oocyte enclosed in a single flattened layer of granulosa cells. Primary follicles are comprised of a primary oocyte enclosed in a single cuboidal layer of granulosa cells exhibiting first signs of the formation of the zona pellucida around the oocyte. A secondary follicle comprising of a oocyte and enclosed by more than one layer of granulosa cells and may contain a cavity (antrum) (Gougeon, 1996). A follicle with multiple layers of cuboidal cells, the complete development of the thecae, the presence of a large cavity, and the formation of cells surrounding the oocyte (corona radiata, and the cumulus oophorus) are characteristic of tertiary follicles (Junqueira and Carneiro, 2004). Diagone et. al. measured dog oocyte’s nuclear and cytoplasm diameter in primordial (19.76 μm and 33.37 μm respectively), primary follicles (21.74 μm and 42.50 μm respectively) and secondary follicles (41.01 μm and 111.38 μm respectively) (Diagone et al., 2008) demonstrating the increase in size as follicles developed. Dog ovarian follicles, much like humans, are heterogeneously distributed within the ovarian cortex (Hayashi et al., 2015; Wakasa et al., 2016) and similar to women, there is a wide variation of follicles in dogs of the same age (Hollinshead et al., 2017). Even within the same dog, there is 2 bilateral asymmetry in the density and distribution between the right and left ovaries (Hayashi et al., 2015). Follicle stimulating hormone (FSH) in follicle development and senescence Under the influence of follicle stimulating hormone (FSH), and anti-Mullerian hormone (AMH), the follicles are recruited into the growing pool (Diagone et al., 2008). FSH is a hormone produced by the pituitary. FSH receptors are first expressed in primary follicles and in later stages of the developing follicles (Hsueh et al., 2015). FSH treatment in prepubertal intact, hypophysectomized or GnRH antagonist-treated rats containing pre-antral and smaller follicles resulted in development of these follicles up to the antral stages (McGee et al., 1997). Cyclical recruitment of follicles in women increases as a result of circulating FSH during each reproductive cycle, and as a result it also rescues a cohort of antral follicles from atresia due to the survival action of FSH (McGee and Hsueh, 2000). FSH is also thought to be responsible for recruitment of a follicle to emerge as a dominant follicle (Fauser and Van Heusden, 1997). The exact mechanism underlying the recruitment of these dominant follicles are unknown however it is believed that these follicles are more sensitive to FSH due to increased FSH and/or luteinizing hormone (LH) receptor expression and/or increased in local growth factor expression (Bao et al., 1997; Evans and Fortune, 1997; Xu et al., 1995). The influence of FSH on growing follicles is influenced by presence of AMH as has been demonstrated using AMH knock out (AMHKO) mice. Durlinger et. al. demonstrated that ovaries of 4-month old AMHKO mice contained more preantral and small antral follicles despite a low serum FSH concentration when compared to wildtype litter mates (Durlinger et al., 1999). 3 The secretion of FSH from the pituitary is under the influence of GnRH as well as by a complex steroid feedback mechanism (Mateos et al., 2002) and therefore affected by a woman’s age. As age increases there are decreased numbers of follicles producing estrogen, which positively signals the pituitary to produce more FSH, resulting in increasing levels of FSH (Wang et al., 2018). In practice, diminished ovarian reserves are suspected when FSH values are over 10 mIU/ml (Wang et al., 2018). Anti-Mullerian hormone (AMH) in follicle development and senescence Production and function AMH is a peptide hormone produced by ovarian granulosa cells (Fleming, Seifer et al. 2015) that has been measured in serum samples from human, cats, dogs and horses using commercially available enzyme-linked immunosorbent assay (ELSIA) kits. In the female, AMH is first produced in the pre-antral follicles and continues to be produced up to small antral follicle stages at which time it is maximally being expressed (Fleming et al., 2015). AMH is involved in the regulation of primordial follicle numbers (thus preventing premature exhaustion of ovarian reserves) and steroidogenesis by granulosa cells (Fleming et al., 2015; Visser and Themmen, 2005). AMH’s effect on primordial follicle numbers has been demonstrated in 2-day-old mice ovaries cultured for 4 days in the presence of AMH had 40% fewer growing follicles compared with untreated ovaries. (Carlsson et al., 2006; Durlinger et al., 1999). Later in 2006, Carlsson et. al. demonstrated the AMH suppressed initiation of the growth of primordial follicles in ovarian biopsy specimens collected from women (Carlsson et al., 2006). AMH concentrations reflect the size of ovarian reserve follicle pool in women (Gomez et al., 2016), and has been shown to correlate with the number of early stage antral follicles (Amer et al., 4 2013; Fleming et al., 2015). AMH remains high in women until roughly 30 years of age and then begins to decline significantly reflecting a major decline in ovarian follicular reserve until complete depletion at menopause (Fleming et al., 2015). Clinical uses of AMH in humans Serum AMH has been shown to be a successful fertility marker in humans through several studies (Amer et al., 2013; Fleming et al., 2015). AMH concentrations are well characterized in women across adolescent and reproductive ages, levels measured are objective and there is convenience of testing serum at any time throughout the menstrual cycle (Fleming et al., 2015; Gomez et al., 2016). This is in contrast to other reproductive biomarkers of fertility such as Follicle Stimulating Hormone (FSH). Not only multiple samples are needed to study FSH trends in women, it also suffers from low sensitivity (Fleming et al., 2015; Tremellen et al., 2005). Previous research in women has demonstrated that AMH provided much better sensitivity and accuracy for predicting ovarian reserve and a woman’s ability to conceive than FSH measurements (Fleming et al., 2015). When combined with antral follicle counts (AFC), it can help determine a woman’s success when undergoing in vitro fertilization (IVF) and other fertility treatments (Tremellen et al., 2005). Clinical uses of AMH in dogs Serum AMH is currently used in dogs to differentiate between spayed and intact animals (Place et al., 2011). Serum AMH has also been used to diagnose ovarian remnant syndrome (4.40 ± 1.09 ng/ml) in female dogs which had AMH concentrations similar to unspayed dogs (4.26 ± 0.82 ng/ml) (Turna Yilmaz et al., 2015). Male dogs with sertoli cells tumors have been shown to have 5 serum AMH levels higher than 22ng/ml (Holst and Dreimanis, 2015). However, there are no tests to determine ovarian follicular reserves and fertility in dogs. Predicting ovarian reserves With increasing survivorship in cancer survivors, the quality of life including reproductive ability, need to be addressed. Mammals are born with a fixed number of ovarian reserves. For women, an ovarian reserve is estimated to be maximum at gestation (20 weeks) with approximately 6-7 million. At birth, this number is reduced to 1-2 million oocytes, which is further reduced to 300,000 – 500,000 at puberty. An average woman may have 25,000 oocytes at age 37 and only 1,000 at 51 years (average age of menopause in USA)(Fleming et al., 2015). This rate of loss is accelerated in women undergoing cancer therapy (chemotherapy and radiation therapy). Cytotoxic hemotherapy, particularly cyclophosphamide, in premenopausal women has been associated with increased fertility failure, an increased risk of amenorrhea and causes premature ovarian failure in 70% of cases as well as early menopause (McLaughlin et al., 2016; Morgan et al., 2012). Over the years a number of endocrine and non-endocrine markers of fertility have been tested including FSH, Inhibin etc. However, in recent years, AMH along with antral follicle counts (AFC), have been used to determine a woman’s success in in vitro fertilization (IVF) and other fertility treatments (Tremellen et al., 2005). It has been established that in women, AMH decreases before FSH begins to rise and Inhibin begins to decline, making AMH an early marker of a limited ovarian reserve (Broekmans et al., 2006; Sowers et al., 2010). Total follicular counts have not been established for dogs. Furthermore, biomarkers to test ovarian reserves and fertility in dogs have not yet been established. 6 Effects of xenobiotics on ovaries Ovarian response to xenobiotics could be a result of direct action via biotransformation within the ovary, or indirectly through systemic circulation of bioactive xenobiotics. Many xenobiotics require to be metabolized or biotransformed into a chemically active intermediate to induce toxicity. Although, liver is the primary organ of metabolizing chemicals, ovaries have been shown to contain both phase I and phase II enzymes involved in metabolism and has been shown to metabolize xenobiotics (Petroff and Basu, 2017). For example, DMBA (7,12-dimethylebenz[α]anthracene) is metabolized in the ovary by CYP 1B1, microsomal epoxide hydrolase (mEH) and CYP 1A1/1B1 and transformed into ovotoxicant DMBA-3,4-diol-1,2- epoxide (Rajapaksa et al., 2007). The implication of ovarian metabolism and its effect on follicles and therefore on fertility has not been studied extensively in humans and are still rare in dogs. Chemotherapy, using cyclophosphamide, in premenopausal women has been associated with an increased risk of amenorrhea and premature ovarian failure in 70% of cases (McLaughlin et al., 2016; Morgan et al., 2012). In women, early menopause often may result in reduced quality of life, increased risk of osteoporosis, cardiovascular disease and depression (Letourneau et al., 2012; Morgan et al., 2012). Few options are available to women who wish to preserve their germline, including embryo cryopreservation and ovary cryopreservation. Embryo cryopreservation is a comparatively successful procedure, but these procedures are expensive, invasive, often requires delay of treatment and the presence of a partner. Most importantly all these options fail to protect ovarian function. Ovarian cryopreservation has been much more difficult to establish, with less than 24 live births since 2004 (Donnez et al., 2013; Ting and Petroff, 2015). Thus, there is a clear need to investigate an alternate strategy to protect the ovary from the cytotoxic effects so as to maintain oocyte quality as well as endogenous hormone production to 7 support follicular growth and for maintenance of pregnancy. Recent studies in rodents have demonstrated that tamoxifen treatment prevented follicular loss and apoptosis from chemotherapeutic drugs and radiation-mediated injury. However, rodent models may not predict clinical outcomes. There is a need for a clinical model which would reflect clinical efficacy in humans. TAMOXIFEN Tamoxifen is a selective estrogen receptor modulator (SERM) that has been approved for use in the prevention of breast cancer in premenopausal women, treatment of advanced breast cancer, and as an adjuvant therapy for postmenopausal women with estrogen receptor-positive (ER) mammary tumor due to its selective antagonist action on ER+ breast cancer (2016c; Gottardis and Jordan, 1988). Metabolism of TAM Tamoxifen is a prodrug that undergoes extensive primary and secondary metabolism to its active metabolites N-desmethyl-tamoxifen, 4-hydroxy-N-desmethyl-tamoxifen and 4- hydroxytamoxifen. The concentration and bioactivity of tamoxifen and its metabolites vary widely (Goetz et al., 2005; Jordan and O'Malley, 2007). One metabolite, 4-hydroxytamoxifen (TAM), has been considered to play an important role in tamoxifen’s anti-cancer effect. TAM has been shown to have a 100 fold greater affinity for estrogen receptors as well as a 30 to 100 fold greater potency in suppressing estrogen dependent cell proliferation compared to the prodrug (Jordan, 1988; Jordan and O'Malley, 2007; Robertson et al., 1982). 8 Actions of TAM The mechanism of TAM is not completely understood, but is suggested to be a combination of complex and intertwined relationships between the number of estrogen receptors, the proportion of alpha and beta estrogen receptors, presence of co-activators and co-repressors in the cell, ligands and presence of growth factors that make it a selective estrogen or an antiestrogen (Jordan, 1988; Jordan and O'Malley, 2007; Smith and O'Malley, 2004). Because of its ability to act as an estrogen agonist, it is also used in some countries for induction of ovulation in sub-fertile women (Dhaliwal et al., 2011; Jordan, 1988). Recent studies by Piasecka-Srader et. al. (2015) and Ting and Petroff (2010), using in-vivo and in-vitro rodent model, have shown that TAM protects ovarian follicles from apoptosis and decreases follicular loss (Piasecka-Srader et al., 2015; Ting and Petroff, 2010). Administration of cyclophosphamide reduced fertility, lessened neonatal survival and decreased primordial and primary follicles in 5-week old rats by as much as one third of the control animals. However, these effects were prevented when TAM was present (Ting and Petroff, 2010). A follow up study using cultured rat ovaries showed similar protective effect of TAM where treatment with TAM decreased follicular loss and apoptosis from 4-hydroxycyclophophamide (CTX) in vitro, suggesting that the protective effects of tamoxifen may be via a direct action on the ovaries as opposed to indirect actions via hepatic metabolism or alteration in gonadotropin release (Inal et al., 2005; Piasecka- Srader et al., 2015). A recent population based cohort study using 397 cancer survivors aged 22- 45 years showed that women who had used TAM had anti-Mullerian hormone levels 2.7 times higher than the women who had never taken tamoxifen, suggesting that TAM users had higher ovarian reserves (Shandley et al., 2016). 9 Current uses in humans The National Comprehensive Cancer Network’s Breast cancer guidelines recommends tamoxifen as the first-line endocrine therapy in pre-menopausal women, as well as those pre- menopausal women presenting with metastatic disease >12 months after completion of adjuvant endocrine therapy or those presenting with de novo metastatic breast cancer (NCCN, 2019). Because of its ability to act as an estrogen agonist, it is also used in some countries for induction of ovulation in sub-fertile women (Dhaliwal et al., 2011; Jordan, 1988). Tamoxifen and ovarian function Tamoxifen appears to exert its own effect on ovarian function independent of chemotherapy (Berliere et al., 2013). This was evidenced in a prospective three-year study in premenopausal women who were treated with tamoxifen as monotherapy or after adjuvant chemotherapy. Irrespective of (presence or absence of) chemotherapy, all patients recovered menses and premenopausal status (Follicle stimulating hormone, FSH, <40 UI/l) after cessation of tamoxifen (Berliere et al., 2013). Depending on the time of tamoxifen administration in the estrus cycle, estrogen levels can increase to up to 3 – 10-fold (Jordan 1991, Jordan 1987) in presence of amenorrhea thus suggesting a false ovarian failure (Mourits et al., 2007). Interference of the normal feedback mechanism via the pituitary axis caused an FSH-driven ovarian steroidogenesis has been suggested as a mechanism (Mourits et al., 2007; Speroff et al., 1999). 10 GOALS OF THIS THESIS Recent studies in rodents have demonstrated that tamoxifen treatment prevented follicular loss and apoptosis from chemotherapeutic drugs and radiation-mediated injury. However, rodent models may not predict clinical outcomes. There is a need for a clinical model which would reflect clinical efficacy in humans. The goals of this study were three-fold. Our first aim was to establish a dog ovarian cortical explant tissue culture model that may be used for testing efficacy of drugs as there is growing interest in the use of dogs as translational models because of their increased life span, physiological similarities to humans, propensity for tumors, larger body size and shared environment. Our second aim was to model the effects of tamoxifen in dog ovarian cortical explant tissue culture model. This study showed that high dose CTXH caused marked follicular loss (P < 0.05) whereas treatment with TAM decreased follicular loss (P < 0.05) from CTX in-vitro. TAM alone did not have an effect on morphologically normal reserve follicle counts (P > 0.05). 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Mol Cell Endocrinol 234(1-2), 81-86. Wakasa, I., Hayashi, M., Abe, Y., Suzuki, H., 2016. Distribution of follicles in canine ovarian tissues and xenotransplantation of cryopreserved ovarian tissues with even distribution of follicles. Reprod Domest Anim. Wang, S., Zhang, Y., Mensah, V., Huber, W.J., 3rd, Huang, Y.T., Alvero, R., 2018. Discordant anti-mullerian hormone (AMH) and follicle stimulating hormone (FSH) among women undergoing in vitro fertilization (IVF): which one is the better predictor for live birth? J Ovarian Res 11(1), 60. Xu, Z., Garverick, H.A., Smith, G.W., Smith, M.F., Hamilton, S.A., Youngquist, R.S., 1995. Expression of follicle-stimulating hormone and luteinizing hormone receptor messenger ribonucleic acids in bovine follicles during the first follicular wave. Biol Reprod 53(4), 951- 957 16 CHAPTER 2. DOG PRIMORDIAL AND PRIMARY FOLLICLES MAINTAIN VIABILITY IN IN-VITRO OVARIAN CORTICAL EXPLANT AGAROSE GEL CULTURE SYSTEM 17 INTRODUCTION In comparison to rodents there is growing interest in the use of dogs as translational models because of their increased life span, physiological similarities to humans, propensity for tumors, larger body size and shared environment (Chastant-Maillard et al., 2010; Schiffman and Breen, 2015; Wakasa et al., 2016). Dogs are excellent genetic models as well as they share approximately 410 genetic disorders with humans (2016b; Antuofermo et al., 2007; Nicholas, 2003). More specifically, in reproductive research, the large size of the ovary, with increased surface area, increased numbers of granulosa cells and tissue availability for research make them an attractive alternative to rodent ovaries. Culture of canine ovaries can be used for understanding the mechanisms controlling follicular growth and differentiation, to study the effects of toxins on follicular numbers, growth and development, for in-vitro generation of competent oocytes capable of fertilization that could be used for preservation of genetic pools of endangered animals, distinguished guide-dogs (that are often spayed before their potential is realized) and may even pave the way for producing competent follicles in women and young girls undergoing cancer therapy (Fujihara et al., 2012; Songsasen et al., 2012; Songsasen et al., 2002). In-vitro culture methods include cell cultures, stem cell cultures, embryo cultures and organ/tissue cultures among others. Specifically, for the reproductive system, cell cultures include culturing of individual follicles (cortical strip culture, individual follicle culture, oocyte-cumulus cell culture, neonatal ovary culture and denuded oocyte culture), spermatogonia, or fertilized embryos and less commonly tissue cultures of ovaries and testes have been studied (Fujihara et al., 2012; Gohbara et al., 2010; Nagashima et al., 2015). Organ culture can be advantageous over cell culture methods because the follicles remain in-situ even as the tissue is cultured in-vitro 18 (Gohbara et al., 2010) thus maintaining follicle-follicle interactions and the influence of stromal cells and glands that are essential for ovarian function and at the same time isolating the ovary from its neuroendocrine inputs that have an impact on follicle growth and recruitment (Morgan and Spears, 2015). Ovaries can be anatomically divided in to a cortex and a medulla, wherein the thin cortex is composed of the ovarian follicles and supporting cells and the medulla contains the blood vessels, lymphatics and nerve supported by a thick connective tissue (Heide Schatten and M. Constantinescu, 2008). Dog ovaries have unique biology making them difficult to culture in- vitro (Songsasen et al., 2012; Songsasen and Wildt, 2007; Songsasen et al., 2011). Follicular loss is seen during the initial transport and processing due to ischemia (Hayashi et al., 2015; Wakasa et al., 2016) and can explain the approximately 20% morphologically abnormal follicles in ovaries processed within 1- 6 hours of collection (Fujihara et al., 2012). Further, dog ovarian follicles have been shown to degenerate extensively within 7 days of culture (Fujihara et al., 2012). Recent studies by Thongkittidilo et. al. (2016), Abdel-Ghani et. al. (2014), Fujihara et. al. (2012) and Songsasen et. al. (2012, 2010) have assessed the use of various growth factors, cytokines and supplements in maintaining tissue viability and follicular growth (Abdel-Ghani et al., 2014; Ackermann et al., 2016; Fujihara et al., 2012; Songsasen et al., 2012; Thongkittidilok et al., 2016). Eagle’s α-Minimum Essential Media (α-MEM) is a rich source of non-essential amino acids and has been used with moderate success in maintaining follicular growth and tissue viability (Fujihara et al., 2012; Nagashima et al., 2016; Thongkittidilok et al., 2016). Fujihara et. al. (2012) demonstrated the superiority of growing dog ovarian cortices in α-MEM culture media on 1.5% agarose gel block and protein free supplement of 0.1 % polyvinyl alcohol (Fujihara et al., 2012). Agarose gel saturated with culture media has shown to be superior while culturing mouse testicular tissue as it allows maintenance of normal cellular structure as well as promotes spermatogenesis 19 in vitro (Gohbara et al., 2010) acting perhaps by providing a better microenvironment for oxygen and nutrient uptake. Supplementing exogenous follicle stimulating factor (FSH) has been used to stimulate follicle growth (Shille et al., 1984). Fujihara and Songsasen reported better results when dog ovarian cortices were cultured in protein free media using 0.1% (w/v) polyvinyl alcohol (PVA) as opposed to those that were cultured in blood serum (Fujihara et al., 2012). Polyvinyl alcohol has been previously used to culture oocytes in cats (Johnston et al., 1993) and for maturing oocytes in a bovine study (Asada and Fukui, 2000). This success of sustaining follicles using PVA, Fujihara attributed to the lack of unknown factors that are often present in blood serum. Amino acids play a vital role in cell and tissue culture sustainability. For example, glutamine, taurine and glycine are known to stimulate the growth and cleavage of hamster embryos (McKiernan et al., 1995) and were included in culture media to provide a nitrogen source. Fibronectin is a component of the extracellular matrix and it is proposed that fibronectin coated cell inserts improve cell culture insert systems by providing protein substrates (2016a). Cell inserts have also been used for dog ovary culture with some success, demonstrating 50% survival after 7 days of culture (Abdel-Ghani et al., 2014). Dog ovarian follicles, much like in women, are heterogeneously distributed within the ovarian cortex (Wakasa et al., 2016) and therefore to attain maximum follicle numbers in the sections, neutral red (NR) was used to identify ovarian cortical fragments that contained viable follicles. Neutral red is a water soluble and non-toxic dye and has been previously used as a non-invasive tool to assess cell viability based on its ability to be taken up by lysosomes of living cells including ovarian follicles (Borenfreund and Puerner, 1985; Elliott and Auersperg, 1993; Langbeen et al., 2014; Repetto et al., 2008). Advantages of using neutral red include its quickness, easy of use, and 20 has shown to be non-toxic in bovine (Langbeen et al., 2014) and canine follicles (Songsasen, pers. comm.). The objective of this aim is to establish a dog cortical ovarian tissue explant culture model that would sustain more than 60% morphologically normal follicles, measured as percent of healthy follicles in the total population of counted follicles after 3 days of in-vitro culture. Our working hypothesis is that our culture media would be able to sustain over 60% of morphologically normal follicles. The overall goal of this study was to understand and optimize the in-vitro culture conditions that would sustain dog primordial and primary follicles. MATERIAL AND METHODS Collection of ovaries, culture media preparation, histological preparation and evaluation was performed as previously described by Fujihara (Fujihara et al., 2012) and Dr. Songsasen (Pers. Comm.) and are described in detail under each subsection. Collection of ovaries Ovaries were collected at elective ovariohysterectomy from clinically healthy animals aged 12 months or less (N = 7). Of the 7 dogs, one dog was in heat. Inclusion criteria were as follows: female, healthy dogs (as ascertained by attending veterinary surgeon), and not pregnant. Ovaries were collected in 1:10 v/v culture media (APPENDIX I). Connective tissue, bursa and oviduct were trimmed on-site before transporting to the laboratory. Ovaries were transported under sterile conditions on ice and processed within 3 hours from time of collection. 21 Processing of ovary Ovaries were transported to the laboratory and transferred from ice to a 100 mm diameter petri- dish and placed on stage-warmer maintained at 38.5°C. Strips of cortical tissue (~ 1.0 mm thick) were dissected from the surface of the ovary using a scalpel blade and scissors. Medial connective tissue was dissected, inside-out, to get cortical sections that were approximately 1 – 1.5 mm thickness. These sections were incubated in 0.05 µg/µl neutral red at 38.5°C in 5% CO2 for 20 minutes to identify sections with most follicles as seen under a field microscope (40x magnification). These pieces were further sectioned into pieces of 1 – 1.5 mm width giving a final approximate size of 1 – 1.5 mm3. Sections were washed multiple times using fresh culture media during processing. Culture media Ovaries were cultured in Eagle’s a-MEM media (Irvine scientific, USA), supplemented with 200 mM L-glutamine, 2.5 mM L-ascorbic acid, 0.3% Poly vinyl alcohol (PVA), 0.06 µl/ml 100x ITS, 50 units/ml penicillin and 50 µg/ml streptomycin. In addition, 10 ng/ml FSH (Folltropin®, Vetoquinol, USA) and 10 ng/ml EGF are added on the day of culture. All chemicals were purchased from Sigma-Aldrich, St. Louis, MO, USA unless otherwise indicated. In-vitro culture systems Three different culture systems for dog ovarian cortical explant cultures were compared based on previous studies on dog ovarian cortical in-vitro culture (Fujihara et al., 2012) and these included: 1.5% agarose gel (1.5%AG), 1.0% agarose gel (1.0%AG) and cell inserts coated with fibronectin (INSRT). 1.5% and 1.0% agarose gel were prepared in phosphate buffered saline. 22 Sections of both 1.5% and 1.0% agarose gel pads, measuring approximately 8 x 8 x 4 mm3 were soaked overnight in culture media at 38.5°C in 5% CO2 in a humidified environment. On day of the experiment (day 0), 24-well plates were pre-equilibrated for 1 hour at 38.5°C in 5% CO2 in a humidified environment, with one 8 x 8 mm 1.5% agarose gel pad per well (prepared and soaked overnight). Three sections of ovaries were placed on the previously soaked gel pad and appropriate media was added to the bottom edge of each gel such it surrounded (but did not submerge) the agarose gel pads and a drop of media placed over the ovarian sections to prevent drying. Half of the media was replaced with fresh media at 48 hours. For the third method, Millicell-CM filter inserts (4 µm) were incubated with 0.1 mg/ml Fibronectin for one hour at 38.5˚C and washed with culture media before use. On the day of culture, each pre-coated and incubated insert was placed in a 24-well plate that contained 400 µl culture media. The ovarian cortical sections were covered with a drop of culture media to prevent drying. Half of the culture media was replaced at 48 hours. Ovaries were cultured at 38.5°C in 5% CO2 in a humidified environment. Each experimental group was run in triplicate (3 wells) with 3 explants per well, culminating in 9 explants per culture system per dog. Processing ovary for histological sections Following 72 hours of culture, ovarian sections was fixed in 4 % paraformaldehyde for at least 24 hours at 4° C, before processing for histology. Ovarian cortical sections were embedded in paraffin following serial dehydration and rehydration in different grades of ethanol, clarified with d-Limonene (EK Industries, Joliet, IL, USA) and then embedded in paraffin. Paraffin blocks were refrigerated overnight before serial sectioning. Serial sections of 5 µm thickness were made and every 10th section was stained with hematoxylin-eosin stain (Appendix II) for morphological 23 evaluation. Follicle stages was determined by morphology. A follicle was evaluated as healthy if it had an intact oocyte with a visible nucleus, granulosa cells without a granular appearance and nuclei that are visible and not condensed. Deformed follicles exhibiting nuclear pyknosis, fragmentation or shrinking in the oocyte or granulosa cells were defined as apoptotic follicles (Fujihara et al., 2012). Follicles in section were classified as described by Gougeon (1996): (a) Primordial follicles comprising of a primary oocyte enclosed in a single flattened layer of granulosa cells, (b) primary follicle comprising of a primary oocyte enclosed in a single cuboidal layer of granulosa cells exhibiting first signs of the formation of the zona pellucida around the oocyte, (c) secondary follicle comprising of a oocyte and enclosed by more than one layer of granulosa cells and (d) antral follicles characterized by the formation of an antral cavity (Gougeon, 1996). For each tissue section (9 tissue sections/culture system/dog), the total number of morphologically normal primordial follicles and primary follicles and morphologically abnormal follicles were counted. Percent morphologically normal follicles were calculated as a percent of total morphologically healthy primordial and primary follicles in a culture system over the total number of follicles (morphologically normal and abnormal) counted in the same culture system. Statistical analysis Data presented as arithmetic mean and standard error of mean (SEM). P<0.05 was considered significant. Follicle counts from the 0-hour sections and the three culture techniques (1.5% AG, 1.0% AG and INSRT) were compared using Fisher’s exact test to test for differences in viable and apoptotic follicles. All statistical tests were performed in Prism 8 (GraphPad software Inc.). 24 RESULTS Loss of morphologically normal follicles after 72 hours of culture A total of 1779 ovarian cortical sections were examined which included 3830 primordial follicles and 747 primary follicles from 6 dogs for this entire study. Average numbers of primordial and primary follicles counted in fresh-fixed ovarian cortical sections (CTRL) were 229.5 ± 78.69. Average numbers for 1.0%AG, 1.5%AG and cell inserts (INSRT) were 132.5 ± 52.68, 151.0 ± 53.03 and 197.2 ± 77.28 respectfully (Figure 2.1). There was no significant difference in the means of the four groups. Percent morphologically normal follicles (Follicle %) were calculated as the total numbers of primordial and primary follicles divided by total follicles counted and expressed as percent. Percent morphologically normal follicles ranged from 94.98 ± 1.62 (n = 7, P < 0.05) in 0 hour control samples (CTRL) compared to 86.98 ± 4.21 (n = 7) in 1.0% AG, 84.52 ± 1.79 in 1.5%AG (n = 7) and 81.46 ± 3.82 in INSRT (n = 6) after 72 hours of culture. In the 72-hour tissue culture sections, there was also mild to moderate loss of normal tissue architecture with mild disarrangement of medial connective tissue and to a lesser extent the interstitial glands in the cortex. These changes included variable amounts of nuclear streaming, increased clear space between stromal connective tissue and loss of normal architecture of the interstitial glands. 25 s e l c i l l o f y r a m i r p d n a l a i d r o m i r p l a t o T 250 200 150 100 50 0 0 hour 1.0% AG 1.5%AG INSRT Figure 2.1. Bar graph of total normal primordial and primary follicles of dog ovarian tissue cultured for 0 hour and 72 hours in 1.0% agarose gel (1.0%AG), 1.5% agarose gel (1.5%AG) and cell inserts coated with fibronectin (INSRT). Data is presented as mean and standard error of mean. Different letters indicate significant differences (P < 0.05). 100 a b b b % e l c i l l o F 80 60 40 20 0 0 hour 1.0% AG 1.5% AG INSRT Figure 2.2. Bar graph of morphological normal follicles of dog ovarian tissue cultured for 0 hour and 72 hours in 1.0% agarose gel (1.0%AG), 1.5% agarose gel (1.5%AG) and cell inserts coated with fibronectin (INSRT). Data is presented as mean and standard error of mean. Different letters indicate significant differences (P < 0.05). 26 Agarose gel improved primordial and primary follicle survival When percent morphologically normal follicle percent was compared, both 1.0%AG and 1.5%AG proved superior (86.98 ± 4.21 and 84.52 ± 1.79 respectively, P < 0.05) to INSERT culture method (81.46 ± 3.82) for maintaining follicle morphology at 72 hours of culture, (Figure.2.2). There was no significant difference observed between 1.0% AG and 1.5% AG, however since the standard error of mean (SEM) of 1.5%AG was smaller and comparable to the 0-hour counts and 1.5% AG were easier to work with in the medium, it was decided to proceed with this method for performing TAM-CTX treatment protocol described in chapter 3. DISCUSSION We were able to sustain viability of a significant numbers of primordial and primary follicles with the ovarian cortical sections for 3 days, in part perhaps, to a faster collection and processing time in comparison to the work undertaken by Fujihara et. al. (Fujihara et al., 2012). We also demonstrated that 1.5% agarose gel was a better microenvironment for culturing dog ovaries compared to the cell inserts coated with fibronectin. Rodent models have been widely used for preclinical studies; however, they may not predict clinical outcomes. Only 10% of anti-cancer agents that enter clinical trials are approved for clinical use, highlighting the need for better preclinical models that would reflect clinical efficacy in humans. Dogs have unique and complex reproductive anatomy and physiology and up until recently it has been extremely difficult to develop in-vitro fertilized embryos in dogs (Fujihara et al., 2012; Nagashima et al., 2016). In an early study employing in-vivo matured oocyte to generate cleaved oocytes; the success rate averaged approximately 10.6% with the poor performance 27 attributed to suboptimal fertilization and poor quality embryo culture medium (Hase et al., 2000). More recently however, Nagashima and Songsasen have had better results; successfully cleaving over 78.8% of the in-vivo matured fertilized oocytes. (Nagashima et al., 2019). The rarity of successful studies of dog ovarian tissue culture models emphasizes the difficulty in sustaining viability. Fujihara and Songsasen demonstrated the superiority of agarose gel culture system that promoted overall survival of dog primordial follicles after 3 (40 %) and 9 (1%) days of incubation compared to in a 24-well cell culture plate (day 3 = 10 %, day 15 = 0 %) (Fujihara et al., 2012) perhaps because of the liquid/gas interface that facilitates nutrient exchange and allows a higher percent of oxygen to diffuse into the tissue (Fujihara et al., 2012; Gohbara et al., 2010). Beyond its usefulness in reproductive sciences, agarose gel culture has shown to be superior for growing and assembling a mechanically functional cartilage-like extracellular matrix (Buschmann et al., 1992), to enhance proliferation of mesenchymal stem cells (Suzawa et al., 2015) and allows increased neuronal differentiation from nervous stem cell (Park et al., 2015). At 0.8% agarose gel has been sufficiently rigid to support adherent cells as well as prevented cell damage while transporting cell lines over long distances (Yang et al., 2009). In this study we experimented using agarose gel at two concentrations, 1.0% and 1.5%, and choose to work with 1.5% agarose gel culture system due to its narrow range of SEM as well as ease of use compared to the 1.0% gel. In dogs, as well as in women, ovarian follicles are heterogeneously distributed within the ovarian cortex (Wakasa et al., 2016). Light and transmission electron microscopy have been historically used to assess follicle viability in human cryopreservation procedures previously, however, these techniques are invasive, destructive and therefore not preferred methods of viability assessment (Langbeen et al., 2014). Use of NR was not only non-invasive; its uptake is 28 temporary because viable follicles eliminate residual NR when follicles are cultured in NR-free media (Langbeen et al., 2014). Traditionally, researchers have used 3 to 4, 1 – 1.5 mm3 blocks of dog ovarian cortex for tissue culture to assess cyto-preservation and vitrification techniques (Abdel-Ghani et al., 2014; Fujihara et al., 2012; Thongkittidilok et al., 2016). It was ascertained that 9 sections of 1 – 1.5 mm3 blocks, collected randomly from both ovarian cortices of the dog will provide our experiment with appropriate robustness to test our hypothesis. To prevent double counting of follicles, primordial and primary follicles were counted on every 10th serial section, each of 5 µm thickness. This model was developed in our laboratory to serve as a basis for further research in reproductive toxicology since dogs may make better translational model than laboratory rodents. Primordial follicles are formed at or around the time of birth and the ovary contains the maximum number of follicles that are available for fertilization (Findlay et al., 2015). Additionally, primordial and primary follicles that have been shown to be extremely sensitive targets of cyclophosphamide and its metabolites. The study was designed to achieve the maximum benefit of the reserve follicle numbers without confounding influence of hormonal factors. CONCLUSION Our findings reflect previous reports of dog ovarian cortical tissue culture and show the superiority of using agarose gel system for promoting primordial and primary follicle survival. Although, the agarose gel system is superior over the other system tested here, there was a significant loss of viable follicles after 72-hours of culture suggesting that we are yet to harness the correct media and microenvironment to optimize and sustain viable follicles for longer periods. 29 APPENDIX 30 Appendix 2.1. Hematoxylin and eosin staining protocol Place the slides in a hot air oven for 25 minutes at 55-60 °C to drip out the excess paraffin. Proceed with the following steps Reagent d-Limonene Ethanol 100% Ethanol 90% Ethanol 80% Ethanol 70% Hematoxylin Tap water Acid 0.3% alcohol Tap water 2x 2x 1x 1x 1x 1x 1x 1x 1x Time 10 min each 5 min 1 min 1 min 1 min 2 min 2 min 2 dips 3 dips Notes 12-slide coplin staining jar were used for most of these reagents Made fresh every day Made fresh every day Made fresh every day Changed after every 4 times Standing tap water Prepared using commercial grade ethanol 200 proof (350 ml) + distilled water (150 ml) + concentrated hydrochloric acid (1.5 ml) Standing tap water 0.2% Ammonia water solution: Add concentrated Ammonium hydroxide (1ml) to distiller water (500 ml) Place the slide holder in a tray filled with gently running water. Place at an angle, so that the excess water runs off. Eosin stain changed after every second use. Made fresh every day Made fresh every day May use xylene. We have not used it. 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J Assist Reprod Genet 31(12), 1727-1736 McKiernan, S.H., Clayton, M.K., Bavister, B.D., 1995. Analysis of stimulatory and inhibitory amino acids for development of hamster one-cell embryos in vitro. Mol Reprod Dev 42(2), 188-199. Morgan, S., Spears, N., 2015. In Vitro Models of Ovarian Toxicity, in: Anderson, R.A., Spears, N. (Eds.), Cancer treatment and the ovary. Elsevier, Netherlands, pp. 79-90. Nagashima, J.B., Hansen, B.S., Songsasen, N., Travis, A.J., Place, N.J., 2016. Anti-Mullerian Hormone in the Domestic Dog during the Anestrus to Oestrous Transition. Reprod Domest Anim 51(1), 158-164. Nagashima, J.B., Sylvester, S.R., Nelson, J.L., Cheong, S.H., Mukai, C., Lambo, C., Flanders, J.A., Meyers-Wallen, V.N., Songsasen, N., Travis, A.J., 2015. Live Births from Domestic Dog (Canis familiaris) Embryos Produced by In Vitro Fertilization. PLoS One 10(12), e0143930. Nagashima, J.B., Travis, A.J., Songsasen, N., 2019. The Domestic Dog Embryo: In Vitro Fertilization, Culture, and Transfer. Methods Mol Biol 2006, 247-267. Nicholas, F.W., 2003. Online Mendelian Inheritance in Animals (OMIA): a comparative knowledgebase of genetic disorders and other familial traits in non-laboratory animals, Nucleic Acids Res. pp. 275-277. Park, K., Nam, Y., Choi, Y., 2015. An agarose gel-based neurosphere culture system leads to enrichment of neuronal lineage cells in vitro. In Vitro Cell Dev Biol Anim 51(5), 455-462. 34 Repetto, G., del Peso, A., Zurita, J.L., 2008. Neutral red uptake assay for the estimation of cell viability/cytotoxicity. Nat Protoc 3(7), 1125-1131. Schiffman, J.D., Breen, M., 2015. Comparative oncology: what dogs and other species can teach us about humans with cancer. Philos Trans R Soc Lond B Biol Sci 370(1673). Shille, V.M., Thatcher, M.J., Simmons, K.J., 1984. Efforts to induce estrus in the bitch, using pituitary gonadotropins. J Am Vet Med Assoc 184(12), 1469-1473. Songsasen, N., Comizzoli, P., Nagashima, J., Fujihara, M., Wildt, D.E., 2012. The domestic dog and cat as models for understanding the regulation of ovarian follicle development in vitro. Reprod Domest Anim 47 Suppl 6, 13-18. Songsasen, N., Wildt, D.E., 2007. Oocyte biology and challenges in developing in vitro maturation systems in the domestic dog. Anim Reprod Sci 98(1-2), 2-22. Songsasen, N., Woodruff, T.K., Wildt, D.E., 2011. In vitro growth and steroidogenesis of dog follicles are influenced by the physical and hormonal microenvironment. Reproduction 142(1), 113-122. Songsasen, N., Yu, I., Leibo, S.P., 2002. Nuclear maturation of canine oocytes cultured in protein- free media. Mol Reprod Dev 62(3), 407-415. Suzawa, Y., Kubo, N., Iwai, S., Yura, Y., Ohgushi, H., Akashi, M., 2015. Biomineral/Agarose Composite Gels Enhance Proliferation of Mesenchymal Stem Cells with Osteogenic Capability. Int J Mol Sci 16(6), 14245-14258. Thongkittidilok, C., Wildt, D.E., Songsasen, N., 2016. Responsiveness of intraovarian dog follicles in vitro to epidermal growth factor and vascular endothelial growth factor depends on ovarian donor age. Reproduction in Domestic Animals, n/a-n/a. Wakasa, I., Hayashi, M., Abe, Y., Suzuki, H., 2016. Distribution of follicles in canine ovarian tissues and xenotransplantation of cryopreserved ovarian tissues with even distribution of follicles. Reprod Domest Anim. Yang, L., Li, C., Chen, L., Li, Z., 2009. An agarose-gel based method for transporting cell lines. Curr Chem Genomics 3, 50-53. 35 CHAPTER 3. PROTECTIVE ACTION OF TAMOXIFEN IN DOG OVARIAN EXPLANTS CULTURED IN PRESENCE OF CYCLOPHOSPHAMIDE 36 INTRODUCTION Mammals are born with a fixed number of ovarian reserves. For women, the ovarian reserve is estimated to be maximum at gestation (20 weeks) with approximately 6-7 million. At birth, this number is reduced to 1-2 million oocytes, which is further reduced to 300,000 – 500,000 at puberty. An average woman may have 25,000 oocytes at age 37 and only 1,000 at 51 years (average age of menopause in USA)(Fleming et al., 2015). This rate of loss is accelerated in women undergoing cancer therapy (chemotherapy and radiation therapy). Cytotoxic hemotherapy, particularly cyclophosphamide, in premenopausal women has been associated with increased fertility failure, an increased risk of amenorrhea and causes premature ovarian failure in 70% of cases as well as early menopause (McLaughlin et al., 2016; Morgan et al., 2012). In women, early menopause often may result in reduced quality of life, increased risk of osteoporosis, cardiovascular disease and depression (Letourneau et al., 2012; Morgan et al., 2012). Few options are available to women who wish to preserve their germline, including embryo cryopreservation and ovary cryopreservation. Embryo cryopreservation is a comparatively successful procedure, but it is invasive, often requires that the patient delays treatment, that there is a current partner and most importantly it fails to protect ovarian function. Ovarian cryopreservation has been much more difficult to establish, with less than 24 live births since 2004 (Donnez et al., 2013; Ting and Petroff, 2015). Total follicular counts have not been established for dogs. According to the 2012 U.S Pet Ownership and Demographics sourcebook, there are a little over 43 million households owning pet dogs, with an average of 1.6 dogs per household (2016d). Over 4.2 million dogs (approximately 5,300/ 100,000 population rate) are annually diagnosed with cancer. It is estimated that one in four 37 intact dog over 4 years of age is expected to develop mammary neoplasia (Antuofermo et al., 2007), positioning valuable breeding dogs and working dogs at greater risk of developing cancers. Dog cancer treatment predominantly consists of chemotherapy, radiation therapy and surgical resection of tumors (Geschwind and Soulen, 2008; MacNeill, 2015). There are very few papers published regarding the effect of chemotherapy in dogs. Tamoxifen is a SERM that has been approved for use in the prevention of breast cancer in premenopausal women, treatment of advanced breast cancer, and as an adjuvant therapy for postmenopausal women with estrogen receptor-positive (ER) mammary tumor due to its selective antagonist action on ER+ breast cancer (2016c; Gottardis and Jordan, 1988). The mechanism of TAM action is not completely understood, but is suggested to be a combination of complex and intertwined relationships between the number of estrogen receptors, the proportion of alpha and beta estrogen receptors, presence of co-activators and co-repressors in the cell, ligands and presence of growth factors that make it a selective estrogen or an antiestrogen (Jordan, 1988; Jordan and O'Malley, 2007; Smith and O'Malley, 2004). Because of its ability to act as an estrogen agonist, it is also used in some countries for induction of ovulation in sub-fertile women (Dhaliwal et al., 2011; Jordan, 1988). Recent studies by Piasecka-Srader et. al. (2015) and Ting and Petroff (2010), using in-vivo and in-vitro rodent models, have shown that TAM protects ovarian follicles from apoptosis and decreases follicular loss (Piasecka-Srader et al., 2015; Ting and Petroff, 2010). Administration of cyclophosphamide reduced fertility, lessened neonatal survival and decreased primordial and primary follicles in 5-week old rats by as much as one third of the control animals. However, these effects were prevented when TAM was present (Ting and Petroff, 2010). A follow up study using cultured rat ovaries showed similar protective effect of TAM where treatment with TAM decreased 38 follicular loss and apoptosis from 4-hydroxycyclophophamide (CTX) in vitro, suggesting that the protective effects of tamoxifen may be via a direct action on the ovaries as opposed to indirect actions via hepatic metabolism or alteration in gonadotropin release (Inal et al., 2005; Piasecka- Srader et al., 2015). A recent population based cohort study using 397 cancer survivors aged 22- 45 years showed that women who had used TAM had anti-Mullerian hormone levels 2.7 times higher than the women who had never taken tamoxifen, suggesting that TAM users had higher ovarian reserves (Shandley et al., 2016). CTX, an alkylating agent is used in dogs for treatment of lymphoproliferative disorders, mammary carcinomas, thyroid carcinomas, hemangiosarcoma as well as an immunosuppressive in autoimmune disorders (Dobson, 2014). Most common side effects of CTX in dogs include myelosuppression, gastrointestinal cytotoxicity and less commonly hemorrhagic cystitis (Dobson, 2014; Wang S.L., 2015). In recent studies on rodents using CTX, apoptosis has been established as the mechanism by which ovarian and granulosa cells are damaged in a dose dependent manner (Donnez et al., 2013). Caspase -3 belongs to a family of highly conserved cystine proteases that are mediators in apoptotic cell death and has been previously shown to take part in normal follicular atresia in the ovaries of rats (Boone and Tsang, 1998), hen (Johnson and Bridgham, 2000), quail (Van Nassauw et al., 1999), mice and human (Matikainen et al., 2001). It is a reliable marker of apoptosis in ovarian follicles (Fenwick and Hurst, 2002). To our knowledge there is no known literature discussing the effects of CTX on fertility or ovarian reserves in dogs has been published. We aim to investigate the follicular protective property of tamoxifen in the dog ovarian explant culture model. We hypothesized that in the presence of CTX, an alkylating chemotherapeutic, TAM will promote a follicle protective effect on the primordial and primary follicles in the dog ovarian cortical explant model. 39 MATERIAL AND METHODS Collection of ovaries, culture media preparation, histological preparation and evaluation was performed as previously described by Fujihara ((Fujihara et al., 2012) and Dr. Songsasen (Pers. Comm.) and are described in detail under each subsection. Collection of ovaries Ovaries were collected at elective ovariohysterectomy from clinically healthy animals aged 6 months or less (N = 10). Inclusion criteria were as follows: healthy dogs (as ascertained by attending veterinary surgeon), dogs aged < 6 months with no prior history of heat (estrus), not displaying signs of heat at time of surgery and the absence of corpus luteum on ovaries. Ovaries were collected in 1:10 v/v culture media. Connective tissue, bursa and oviduct were trimmed on- site before transporting to the laboratory. Ovaries were transported under sterile conditions on ice and processed within 3 hours from time of collection. Processing ovaries for in-vitro culture Ovaries were transported to the laboratory and transferred from ice to a 100 mm diameter petri- dish and placed on stage-warmer maintained at 38.5°C. Strips of cortical tissue (~ 1.0 mm thick) were dissected from the surface of the ovary using a scalpel blade and scissors. Medial connective tissue was dissected, inside-out, to give cortical sections that were approximately 1 – 1.5 mm thickness. These sections were incubated in 0.05 µl/µl solution of neutral red at 38.5°C in 5% CO2 for 20 minutes to identify sections with most follicles as seen under a field microscope (40x magnification). These pieces were further sectioned into pieces of 1 – 1.5 mm width giving a final 40 approximate size of 1 – 1.5 mm3. Sections were washed multiple times using fresh culture media during processing. Culture media Ovaries were cultured in Eagle’s a-MEM media (Irvine scientific, USA), supplemented with 200 mM L-glutamine, 2.5 mM L-ascorbic acid, 0.3% Poly vinyl alcohol (PVA), 0.06 µl/ml 100x ITS (Insulin transferrin sodium selinate), 50 units/ml penicillin and 50 µg/ml streptomycin. In addition, 10 ng/ml FSH (Folltropin®, Vetoquinol, Fort Worth, TX, USA) and 10 ng/ml EGF are added on the day of culture. All chemicals were purchased from Sigma-Aldrich, St. Louis, MO, USA unless otherwise indicated. 4-hydroxycyclophosphamide (Mafosfamide; CTX) was Santa Cruz Biotechnology Inc. (Texas, USA) and 4-hydroxytamoxifen (TAM) was from Sigma-Aldrich. In-vitro ovarian culture 1.5% agarose gel pads, measuring approximately 8 x 8 x 4 mm3 were soaked overnight in treated culture media at 38.5°C in 5% CO2 in a humidified environment. On day of the experiment (day 0), 24-well plates were pre-equilibrated for 1 hour at 38.5°C in 5% CO2 in a humidified environment, with one 8 x 8 mm 1.5% agarose gel pad per well (prepared and soaked overnight). Ovaries were cultured at 38.5°C in 5% CO2 in a humidified environment. Ovaries were treated with vehicle (control, CRTL: 0.1% dimethyl sulfoxide plus ethanol), TAM (10 µM), and /or two doses of CTX (1 and 10 µM) for 72 hours (follicle counts and apoptosis). 200 µl of media was placed in each well such that it surrounded (but did not submerge) the 1.5% agarose gel. Half of the media was replaced with fresh media at 48 hours. Doses were based on human pharmacokinetics concentrations (Pollack et al., 1997; Pollack et al., 1990) as well as past work in 41 culture systems (Chen et al., 1997; Desmeules and Devine, 2006; Piasecka-Srader et al., 2015; Ramachandran et al., 2004; Ting and Petroff, 2010). Each treatment group was run in triplicate (3 wells) with 3 explants per well. The experiment was limited by the number of explant sections that were obtained per dog (both ovaries), with a maximum of approximately 75-80 1mm3 sections/dog. Processing ovaries for histology Following 72 hours of culture, ovarian sections was placed in 4% paraformaldehyde for at least 24 hours at 4° C, before processing for histology. Ovarian cortical sections were embedded in paraffin following serial dehydration and rehydration in different grades of ethanol, clarified with d-Limonene (EK Industries, Joliet, IL, USA) and then embedded in paraffin. Paraffin blocks were refrigerated overnight before serial sectioning. Serial sections of 5µm thickness were made and every 10th section was stained with hematoxylin-eosin stain (Appendix I) for morphological evaluation. Follicle stages was determined by morphology. A follicle was evaluated as healthy if it had an intact oocyte with a visible nucleus, granulosa cells without a granular appearance and nuclei that are visible and not condensed. Deformed follicles exhibiting nuclear pyknosis, fragmentation or shrinking in the oocyte or granulosa cells were defined as apoptotic follicles (Fujihara et al., 2012). Follicles in section were classified as described by Gougeon (1996): (a) Primordial follicles comprising of a primary oocyte enclosed in a single flattened layer of granulosa cells, (b) primary follicle comprising of a primary follicle enclosed in a single cuboidal layer of granulosa cells exhibiting first signs of the formation of the zona pellucida around the oocyte, (c) secondary follicle comprising of a oocyte and enclosed by more than one layer of granulosa cells and (d) antral follicles characterized by the formation of an antral cavity (Gougeon, 42 1996). For each section, the total number of morphologically normal primordial follicles, primary follicles and morphologically abnormal follicles was tabulated. The analysis was performed blind. Apoptosis evaluation Apoptosis of primordial and primary follicles was evaluated in three serial sections. These three sections represented the maximum number of follicles present on a slide in corresponding 10th sections (which were used for counting reserve follicles and assessing morphology). Caspase- 3 was performed by the Histology laboratory at the College of Natural Sciences at Michigan State University. In brief, un-stained charged slides were deparaffinized in Xylene and hydrated through descending grades of ethyl alcohol to distilled water. Slides were placed in Tris Buffered Saline pH 7.4 (Scytek Labs – Logan, UT) for 5 minutes for pH adjustment. Following TBS, Heat Induced Epitope Retrieval in Citrate Plus pH 6.0 (Scytek) performed in vegetable steamer at 100°C for 30 minutes. After citrate treatment, the slides were incubated for 10 minutes at room temperature and rinsed in several changes of distilled water. Endogenous peroxidase was blocked for 30 minutes at room temperature in a 3% hydrogen peroxide / methanol bath followed by running tap water and several rinses in distilled water; followed by Tris buffered saline + Tween 20 (Scytek) for 5 minutes. Following pretreatment standard Micro-polymer staining steps were performed at room temperature on the Biocare intelli-PATH automated staining instrument. All staining steps are followed by rinses in TBS Autowash Buffer + surfactant (Biocare – Concord, CA). After blocking for non-specific protein with Rodent Block M (Biocare) for 10 minutes; sections were incubated with primary antibody Caspase-3 (p17) (Abcam; ab2302) at 1:30 in Normal Antibody Diluent (Scytek) overnight at 4°C. Following this, rabbit on rodent HRP incubation for 20 minutes (Biocare). Reaction development utilized Biocare Romulin AEC Chromogen incubation of 10 43 minutes followed by counterstain in CATHE Hematoxylin 1:10 (Biocare) for 1 minute. The slides were allowed to air dry and mounted with synthetic mounting media. Granulosa cell or oocytes were considered caspase positive if there was positive cytoplasmic or nuclear staining, and results reported as total positive follicles in each treatment group in every dog (N = 8). Statistical analysis Data presented as arithmetic mean and standard error of mean. P<0.05 was considered to be significant. The 0 hour and 72-hour control groups (CTRL) were compared using Mann-Whitney non-parametric tests. Control and treatment groups were compared using two-way ANOVA followed by Tukey’s multiple comparison test to compare the means of each treatment group. Difference with a probability of P < 0.05 were considered statistically significant. All statistical tests were performed in Prism 8 (GraphPad software Inc.). RESULTS Loss of follicles after 72 hours in in-vitro explant tissue culture Follicle populations were predominantly primordial follicles, primary follicles and rarely secondary follicles. For this study, only primary and primordial follicles were counted and presented as total follicle numbers (TFN). There was a significant (P < 0.05) decline in the total numbers of morphologically normal follicles counted in CTRL group (146.40 ± 34.54) compared to 0-hour (310.5 ± 57.30) fixed ovarian sections with loss of approximately 50% of follicles in the first three days of culture (Figure. 3.1.). Other changes seen included mild to moderate loss of 44 normal tissue architecture with marked disarrangement and collapse of medial connective tissue and to a lesser extent the interstitial glands. 400 300 200 100 0 s e l c i l l o f y r a m i r p d n a l a i d r o m i r P a b 0 hour CTRL Figure 3.1. Primordial and primary follicles counted in ovarian sections at 0 hours and after 72 hours of culture at 38.5 °C in 5% CO2 on 1.5% agarose gel pads. Data is presented as mean and standard error of mean. Different letters indicate significant differences (P < 0.05). Co-administering tamoxifen with cyclophosphamide protects dog ovarian reserves Follicle populations were predominantly primordial follicles, primary follicles and rarely secondary follicles. For this study, only primordial and primary follicles were counted. Primordial follicles made up majority of the follicles. Active metabolite of CTX decreased the number of primordial and primary follicles after 72 hours of culture in a dose dependent manner, with the highest dose of CTX depleting reserves (presented as TNF) to approximately 70% of the controls (Figure. 3.2 and 3.3). TAM, on its own, was not protective, however when co-administered with CTX, it was associated with significantly greater follicular reserves. This is the first time this phenomenon has been documented in dogs, and is similar to what has been demonstrated in rodents (Piasecka-Srader et al., 2015; Ting and Petroff, 2010). Other changes seen included mild to severe 45 loss of normal tissue architecture with marked disarrangement and collapse of medial connective tissue and to a lesser extent the interstitial glands in a dose dependent manner in the CTXL and CTXH treatment groups as well as in TAM treated explants. These changes included variable amounts of nuclear streaming, increased clear space and discontinuity between stromal connective tissue and loss of normal architecture of the interstitial glands. s e l c i l l o f l a i d r o m i r P 150 100 50 0 a a , a , a , a , b CTRL TAM CTXL CTXH TAM+ CTXL TAM+ CTXH Figure 3.2. Effect of 4-hydroxytamoxifen (TAM; 10 µM) and/or 4-hydroxycycloophosphamide (CTXL; 1 µM and CTXH; 10 µM) on the number primordial follicles in cultured dog (N = 10) ovaries after 72 hours. Values are presented as mean ± SEM follicles counted (n=10). CTRL: control group. Different letters indicate significant differences (P < 0.05). 150 100 50 s e l c i l l o f l a i d r o m i r P 46 s e l c i l l o f y r a m i r p d n a l a i d r o m i r P 200 150 100 50 0 a a a a a b CTRL TAM CTXL CTXH TAM+ CTXL TAM+ CTXH Figure. 3.3. Effect of 4-hydroxytamoxifen (TAM; 10 µM) and/or 4-hydroxycyclophosphamide (CTXL; 1 µM and CTXH; 10 µM) on the number primordial and primary follicles in cultured dog ovaries after 72 hours. Values are presented as mean ± SEM follicles counted (n=10). CTRL: control group. Different letters indicate significant differences (P < 0.05). Apoptosis marker did not show recovery of follicles on coadministration of TAM-CTX Caspase-3 was localized in the granulosa cells as well as in oocytes of the primary and primordial follicles. Follicles were considered apoptotic, if one or more granulosa cells and/or oocyte stained positive for caspase-3. Although not statistically significant, changes in follicle counts that were positive for caspase-3 reflected the follicle count data seen at 72 hours, with a high dose CTX markedly inducing caspase-3 staining and recovery seen in ovaries that were co- treated with TAM (P = 0.99, Figure. 3.4). 47 % e l c i l l o f e v i t i s o p 3 - e s a p s a C 100 80 60 40 20 0 P = 0.93 a a a P = 0.99 a a a CTRL TAM CTXL CTXH TAM+ CTXL TAM+ CTXH Figure 3.4. Effect of 4-hydroxytamoxifen (TAM; 10 µM) and/or 4-hydroxycyclophosphamide (CTXL; 1 µM and CTXH; 10 µM) on the number of apoptotic Caspase-3 positive primordial and primary follicles in cultured dog ovaries after 72 hours. Values are presented as mean ± SEM follicles counted (n=10). CTRL: control group. Different letters indicate significant differences (P < 0.05). DISCUSSION The main objective of this study was to examine the effects of TAM when co-administered with CTX on cultured dog ovaries, specifically their actions on primordial and primary follicles. To our knowledge this is the first study that has shown the protective effects of TAM on dog ovarian follicles when it is co-administered with cyclophosphamide. CTX induced marked follicular loss via apoptosis, and while TAM alone did not alter basal apoptosis, ovaries treated with both TAM and CTXH ovaries had decreased apoptosis compared to CTX treated ovaries. Fujihara et. al. demonstrated the difficulty of maintaining morphologically normal primordial follicles. They demonstrated a loss of morphologically normal follicle numbers from 86% at time 48 of collection to approximately 40% after 72 hours of incubation (Fujihara et al., 2012). They however did not provide absolute follicle numbers making it difficult to compare data from this study. Piasecka-Srader et. al. experienced a decline of approximately 78% primordial follicles in an in-vitro study in ovaries of rodents (Piasecka-Srader et al., 2015; Ting and Petroff, 2010) that were cultured for 7 days using similar drugs and dosages as this study. The loss of morphologically normal ovaries by approximately 70% after 72 hours of culture in this study is accentuated perhaps in part due to the challenges in culturing dog ovarian explants. More recently, a retrospective study that studied the impact of tamoxifen on fertility in women showed that women who used tamoxifen had higher AMH levels and antral follicle counts, suggesting that these women had more ovarian reserves than the women who did not use tamoxifen ever (Shandley et al., 2017). In women tamoxifen is associated with risk of secondary endometrial cancer and life threatening thromboembolism (Smith, 2014). The most common side effects from use of tamoxifen are postmenopausal symptoms, including hot flashes, night sweats, vaginal dryness, discharge and irregular menses (Braems et al., 2011; Eigeliene et al., 2016; Jordan, 1988; Shandley et al., 2017; Smith, 2014). In dogs, estrogenic side effects have been reported, including vulval discharge and swelling, attractiveness to male dogs (Morris et al., 1993). This study had a few strengths. We demonstrated that tamoxifen had a similar protective action on dog ovaries as has been previously demonstrated in rodents. It is estimated that one in four intact dogs over 4 years of age is expected to develop mammary neoplasia (Antuofermo et al., 2007), positioning valuable breeding dogs and working dogs at greater risk of developing cancers. Not only does our study provide proof of concept that may help prevent loss of fertility in valuable breeding dogs; we also hope that this study’s results can help establish dogs as a translational 49 model for human mammary tumor studies since not only do dogs share physiological similarities to humans, but when compared to rodents, they also have increased life spans, a propensity for tumors, larger body size and shared environment (Chastant-Maillard et al., 2010; Schiffman and Breen, 2015; Wakasa et al., 2017). This study had a few limitations. Although we demonstrated a lowered apoptosis trend with use of TAM in presence of cyclophosphamide, we could not demonstrate a significant association between CTX and apoptosis in the dog ovarian follicle sections as has been demonstrated in rodents (Piasecka-Srader et al., 2015). This was perhaps because of the smaller sample sizes that were used for this part of the study. Another limitation of our study was our inability to demonstrate the mechanism by which TAM annulled the effect of CTX on dog ovaries after 72 hours of culture. Piesecka-Srader et al. demonstrated that TAM decreased the expression of multiple inflammation related genes, such as mediators of lipoxygenase and prostaglandin production, cytokine binding, second messenger signaling, tissue modelling and vasodilation (Piasecka-Srader et al., 2015). Another study looking at radiation induced follicular loss implicated enhancement of insulin like- growth factor -1 by tamoxifen (Mahran et al., 2013). It may be prudent to test for mechanism at earlier and different time points when the action of the chemotherapeutic is most established. The exact mechanism of TAM however remains elusive and perhaps is multifaceted. CONCLUSION To our knowledge this is the first study that examines the effects of coadministration of TAM with CTX in dogs and demonstrates that TAM has protective effects on ovarian follicles when it is co-administered with cyclophosphamide. 50 APPENDIX 51 Appendix 3.1. Hematoxylin and eosin staining protocol Place the slides in a hot air oven for 25 minutes at 55-60 °C to drip out the excess paraffin. Proceed with the following steps Reagent d-Limonene Ethanol 100% Ethanol 90% Ethanol 80% Ethanol 70% Hematoxylin Tap water Acid 0.3% alcohol Tap water 2x 2x 1x 1x 1x 1x 1x 1x 1x Time 10 min each 5 min 1 min 1 min 1 min 2 min 2 min 2 dips 3 dips Notes Use a 12-slide coplin staining jar for most of these reagents Made fresh every day Made fresh every day Made fresh every day Changed after every 4 staining cycles Standing tap water Add commercial grade ethanol (200 proof, 350 ml) + distilled water (150 ml) + concentrated hydrochloric acid (1.5 ml) Standing tap water 0.2% Ammonia water solution: Add concentrated Ammonium hydroxide (1ml) to distiller water (500 ml) Place the slide holder in a tray filled with gently running water. Place at an angle, so that the excess water runs off. Changed after every 2 cycles. made fresh every day made fresh every day May use xylene. We have not used it. DPX mountant in sufficien quantity to cover tissue sections and place a cover slip avoiding air bubbles. 52 Blueing agent 1x 2 min tap Running water Ethanol 70% Eosin Ethanol 80% Ethanol 90% Ethanol 100% d-Limonene Mounting 1x 1x 1x 1x 1x 2x 2x 8 min 3 min 2 min 5 min 10 sec 10 each 10 each sec sec REFERENCES 53 REFERENCES 2016a. U S Food and Drug Administration Home Page. http://www.fda.gov/. 2016b. U.S. Pet Ownership Statistics. https://www.avma.org/KB/Resources/Statistics/Pages/Market-research-statistics-US-pet- ownership.aspx. (Accessed December 2016). Antuofermo, E., Miller, M.A., Pirino, S., Xie, J., Badve, S., Mohammed, S.I., 2007. Spontaneous mammary intraepithelial lesions in dogs--a model of breast cancer. Cancer Epidemiol Biomarkers Prev 16(11), 2247-2256. Boone, D.L., Tsang, B.K., 1998. Caspase-3 in the rat ovary: localization and possible role in follicular atresia and luteal regression. Biol Reprod 58(6), 1533-1539. 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Chemotherapy-induced neutropenia is associated with 57 CHAPTER 4. ANTI-MULLERIAN HORMONE AS AN INDICATOR OF OVARIAN RESERVES IN DOMESTIC DOGS 58 INTRODUCTION All females are born with a limited number of ovarian germ cells the number of which vary within species and between individuals within the species (Hollinshead et al., 2017). Anti- Mullerian hormone (AMH) is a peptide hormone produced by ovarian granulosa cells (Fleming, Seifer et al. 2015) that has been detected in serum samples from human, cats, dogs and horses using commercially available enzyme-linked immunosorbent assay (ELISA) kits. In women, AMH is first produced in the pre-antral follicles and continues to be produced up to the small antral follicle stages at which time it is maximally being expressed (Fleming et al., 2015). AMH is responsible for the regulation of primordial follicle numbers (thus preventing premature exhaustion of ovarian reserves) and steroidogenesis by granulosa cells (Fleming et al., 2015; Visser and Themmen, 2005). AMH’s effects on primordial follicle numbers has been demonstrated in 2- day-old mice ovaries cultured for 4 days in the presence of AMH had 40% fewer growing follicles compared with untreated ovaries. (Carlsson et al., 2006; Durlinger et al., 1999). Later in 2006, Carlsson et. al. demonstrated the AMH supressed initiation of the growth of primordial follicles in ovarian biopsy specimens collected from women (Carlsson et al., 2006). AMH concentrations reflect the size of ovarian reserve follicle pool in women (Gomez et al., 2016), and has been shown to correlate with the number of early stage antral follicles (Amer et al., 2013; Fleming et al., 2015). Serum AMH has been shown to be a successful fertility marker in humans through several studies (Amer et al., 2013; Fleming et al., 2015). AMH concentrations are well characterized in women across adolescent and reproductive ages, levels measured are objective and there is convenience of testing serum at any time throughout the menstrual cycle (Fleming et al., 2015; Gomez et al., 2016). This is in contrast to other reproductive biomarkers of 59 fertility such as Follicle Stimulating Hormone (FSH). Not only multiple samples are needed to study FSH trends in women, it also suffers from low sensitivity (Fleming et al., 2015; Tremellen et al., 2005). Previous research in women has demonstrated that AMH provided much better sensitivity and accuracy for predicting ovarian reserve and a woman’s ability to conceive than FSH measurements (Fleming et al., 2015). When combined with antral follicle counts (AFC), it can help determine a woman’s success when undergoing in vitro fertilization (IVF) and other fertility treatments (Tremellen et al., 2005). AMH remains high in women until roughly 30 years of age and then begins to decline significantly reflecting a major decline in ovarian follicular reserve until complete depletion at menopause (Fleming et al., 2015). Serum AMH is currently used in dogs to differentiate between spayed and intact animals (Place et al., 2011). Serum AMH has also been used to diagnose ovarian remnant syndrome (4.40 ± 1.09 ng/ml) in female dogs which had AMH concentrations similar to unspayed dogs (4.26 ± 0.82 ng/ml) (Turna Yilmaz et al., 2015). Male dogs with sertoli cells tumors have been shown to have serum AMH levels higher than 22ng/ml (Holst and Dreimanis, 2015). However, there are no tests to determine ovarian follicular reserves and fertility in dogs. The aim of this study was to develop a similar model of ovarian reserve measurement in companion animals that can be used as a predictor of ovarian reserve and fertility in future canine clinical trials. The objective of this aim is to test circulating AMH concentration as a non-invasive measure of ovarian follicular reserve, so that it may serve as an indicator of ovarian function in dogs. 60 MATERIAL AND METHODS Animals Discarded ovaries were collected from clinically healthy mixed breed dogs (N=55) undergoing elective ovariohysterectomy from October 2016 to January 2018. Detailed reproductive history was generally not available for these dogs, however at the time of collection, age (1 -168 months), breed, weight and approximate stage in the reproductive cycle (as ascertained by attending veterinarian) was recorded based on physical examination (swollen vulva, serosanguinous discharge) and by examining macro ovarian morphology. Only dogs that were deemed clinically healthy were included in this study. Informed consent was obtained from all owners. Banked serum samples obtained from the Endocrine laboratory serum repository from age-matched spayed dogs were used as negative controls for serum AMH ELISA. Collection of blood samples and ovaries Blood (2ml -5ml) collected in red top tube during pre-operation site preparation, was allowed to clot at room temperature and transported on ice to the laboratory for further processing. Ovaries were collected in either 1:10 v/v culture media (previously described in chapter 2 and 3) if they were destined for in-vitro study or in 4% paraformaldehyde. Connective tissue, bursa and oviduct were trimmed on-site. Ovaries were transported under sterile conditions to the laboratory and processed within 4 hours. Clotted blood samples were spun at 3500 rpm for 10 minutes in a centrifuge. Serum was immediately separated into plastic tubes, frozen and maintained at - 20 degrees C until further analysis. 61 Processing ovaries for histology Strips of cortical tissue (~ 1.0 mm thick) were dissected from the surface of the ovary using a scalpel blade and scissors. Cortical sections were further sectioned into pieces of 1 – 1.5 mm width giving a final approximate size of 1 – 1.5 mm3 and allowed to fix in 4% paraformaldehyde for at least 24 hours at 4 ºC, before paraffin embedding. Nine sections per animal were sampled from ovarian cortical sections for analysis. Ovarian cortical sections were embedded in paraffin following serial dehydration and rehydration in different grades of ethanol. Paraffin blocks were refrigerated overnight before serial sectioning. Serial sections of 5µm thickness were made and every 10th section was stained with hematoxylin-eosin stain (Appendix I) for morphological evaluation. Follicle stages was determined by morphology. A follicle was evaluated as healthy if it had an intact oocyte with a visible nucleus, granulosa cells without a granular appearance and nuclei that are visible and not condensed. Deformed follicles exhibiting nuclear pyknosis, fragmentation or shrinking in the oocyte or granulosa cells were defined as apoptotic follicles (Fujihara et al., 2012). Follicles in section were classified as described by Gougeon (1996): (a) Primordial follicles are comprised of a primary oocyte enclosed in a single flattened layer of granulosa cells, (b) primary follicle comprising of a primary oocyte enclosed in a single cuboidal layer of granulosa cells exhibiting first signs of the formation of the zona pellucida around the oocyte, (c) secondary follicle comprising of an oocyte and enclosed by more than one layer of granulosa cells and (d) antral follicles characterized by the formation of an antral cavity (Gougeon, 1996). For each section, the total number of ‘healthy’ primordial follicles, primary follicles and apoptotic follicles were counted. Total number of primordial and primary follicles for each animal was tabulated. 62 Serum AMH enzyme-linked immunosorbent assay AMH was measured using a commercially available enzyme-linked immunosorbent assay (AMH Gen II ELISA, Beckman Coulter, Brea, CA). All samples were run in duplicates as per manufacturers instruction. Briefly, the assay is a two-site immunoassay, which utilizes two monoclonal antibodies labelled with biotin. Calibrators, controls and samples are incubated in microtitration wells coated with anti-AMH antibody. The wells are then treated with anti-AMH detection antibody labelled with biotin followed by streptavidin-horseradish peroxidase (HRP), followed by the addition of a stop solution. Extensive washing was performed after each incubation period. Enzymatic activity of the substrate was reflected by absorbance at 450 nm of HRP. The limit of quantification is 0.08 ng/ml. Intra assay coefficient of variation was 8.5%. Progesterone assay Progesterone assay was performed on serum samples (N = 40) at the Veterinary Diagnostic Laboratory’s endocrinology laboratory at Michigan State University and classified as baseline, proestrus or estrus as per laboratory standards (Appendix II). Serum from dogs ranging in age from 7 months up to 9 years were analyzed. Briefly, progesterone values between 0 to 2.9 nmol/L were classified as baseline (anestrous), progesterone values over 3 nmol/L but below 13 nmol/L were classified as proestrus and progesterone values between 13 nmol/L to 31 nmol/L were classified as estrous (Refsal, 2019). Statistical analysis Data are presented as arithmetic mean and standard error of mean. Serum AMH levels between spayed and intact dogs was compared using Wilcoxon-signed rank test. Mann-Whitney test was 63 used to analyze serum AMH levels between pre-pubertal and pubertal dogs, baseline and proestrus serum AMH levels. Pearson correlation test was used to determine the interaction between serum AMH, age and reserve follicles. P < 0.05 was considered to be significant. All statistical tests were performed in Prism 8 (GraphPad software Inc.). RESULTS Commercially available serum AMH assay identified spayed dogs As part of internal quality control, a pilot study was carried out to determine the feasibility of using a human based AMH ELISA using banked serum that were submitted for other tests. Sera from known intact (n=20, 0.569 ± 0.120 ng/ml) and spayed (n = 14, 0.005 ± 0.003 ng/ml) dogs showed significant (P < 0.05) differences. The dogs in both groups ranged in age from 1 – 11 years. In the follow up study, serum AMH measured in 20 spayed dogs (6 months – 6 years) was below the detection level (<0.08 ng/ml) of the assay kit. Circulating AMH in 28 intact dogs aged between 6 months to 6 years (assumed to be peak reproductive years) was 0.79 ± 0.106 ng/ml (P < 0.0001) (Figure. 4.1). 64 ) L m / g n ( H M A 1.0 0.8 0.6 0.4 0.2 0.0 a Spayed Intact Figure 4.1. Serum AMH values in spayed and intact dogs ranging between 6 months to 6 years. Different letters indicate significant differences (P < 0.05) between the two groups. Serum AMH differed between prepubertal (Pre-P) and post pubertal (Post-P) dogs Dogs were divided into two groups. Prepubertal dogs (n = 14) were classified as those younger than 6 months with no history of estrus (heat). Post-pubertal dogs (n = 28) were classified as dogs between 6 months – 6 years, with at least one known estrus cycle and assumed to be in prime reproductive age. Serum AMH in prepubertal dogs was 0.08 ± 0.05 ng/ml and in post-pubertal dogs was 0.79 ± 0.10 ng/ml were significantly different (P<0.0001) (Figure.4.2). 65 ) L m / g n ( H M A 1.0 0.8 0.6 0.4 0.2 0.0 a Prepubertal Postpubertal Figure 4.2. Serum AMH values in prepubertal (< 6 months) and post-pubertal dogs (> 6 months to 6 years). Different letters indicate significant differences (P < 0.05) between the two groups. AMH varies with age Serum AMH was determined for all dogs (N = 55) between ages 1 month and 14 years and ranged between 0 ng/ml to 2.4 ng/ml (Figure. 4.3). All dogs < 3 months (n = 7) had undetectable serum. Two out of four dogs aged 4.5 months and 5 months also had undetectable serum AMH. The youngest age at which serum was detected was 4.5 months (0.63 ng/ml). Circulating AMH was detecting in all dogs over 5 months of age except in two dogs in this study. One dog was a 4 year of Chihuahua-Pomeranian mix with a very low follicle count and the second dog was 14 years old Saluki with a known history of infertility for over two years (per owner). Average circulating AMH in dogs (n= 25) aged 1 – 6 years at 0.86 ± 0.11 ng/ml, ranging from 0 ng/ml to 2.4 ng/ml. Circulating AMH decreased with age (n = 13, 7 – 14 years), with average values of 0.64 ± 0.09 ng/ml (r = -0.53, P > 0.05). 66 ) L m g n ( / H M A 3 2 1 0 0 50 100 150 200 Age (months) Figure 4.3. Serum AMH values in dogs ranging from 1 month old to 14 years age. Significant AMH variation is seen in dogs that are in baseline and proestrus We classified 40 dogs as either baseline or in proestrus based on circulating progesterone levels under guidelines published by the endocrinology laboratory at Veterinary Diagnostic laboratory at Michigan State University. There was significant difference (P < 0.0001) in serum AHM in baseline dogs (n = 23, 1 – 6 years, 0.54 ± 0.04 ng/ml, range 0.19 – 0.9 ng/ml) and serum AMH in proestrus dogs (n = 14, 1 – 6 years, 1.28 ± 0.11 ng/ml, range 0.96 – 2.4 ng/ml) (Figure. 4.4). Circulating AMH of baseline dogs (n = 25, r = 0.53, P < 0.05) significantly corelated with age as opposed to AMH from dogs in proestrus (n = 14, P > 0.55). 67 ) L m / g n ( H M A 1.5 1.0 0.5 0.0 a Baseline Proestrus Figure 4.4. Serum AMH values in dogs (n= 40, 6 months – 6 years) significantly varied in dogs that were baseline (n = 26) and in proestrus (n = 14). Different letters indicate significant differences (P < 0.05) between the two groups. Total follicle numbers (TFN) negatively corelate with advancing age Primordial and primary follicles together represented as TFN showed significant negative correlation (r = -0.059, P < 0.0001) with advancing age (N = 55) (Figure.4.5). AMH and TFN are not positively correlated Serum AMH and reserve follicles did not show positive correlation (r = - 0.36, P < 0.05) with advancing age (N = 55). On the whole, the plotted data showed a strong negative correlation between circulating AMH and the total numbers of follicles counted with a large proportion of the data sets clustered in the lower left quadrant. We discuss these findings in the next section in detail. 68 800 600 400 200 s r e b m u n e l c i l l o f l a t o T 0 0 50 100 150 200 Age (months) Figure 4.5. Total follicle numbers for dogs (N = 55) aged between 1 month to 14 years. There is significant negative correlation (r = -0.059, P < 0.0001) of total follicle numbers with advancing age. ) L m / g n ( H M A 3 2 1 0 0 200 400 600 800 Total follicle numbers 69 Figure 4.6. Serum AMH and reserve follicles did not show positive correlation (r = - 0.36, P < 0.05) with advancing age (N = 55). DISCUSSION This is the first study to establish a relationship between serum AMH and age over a wide age group in dogs. We have also demonstrated a significant difference in circulating AMH levels during anestrous and proestrus. Further this study demonstrated a strong negative correlation between follicle reserves and advancing age. Sparse literature is available about the physiology and role of AMH in dogs. Current uses of AMH in veterinary medicine are limited to differentiating spay and neuter status, diagnosing ovarian remnant syndrome, sertoli cell tumors in dogs and granulosa cell tumors (GCT) in dogs and horses) since both Sertoli cells in male dogs and granulosa cells in female dogs can produce AMH (Almeida et al., 2011; Holst and Dreimanis, 2015). Almeida et. al. demonstrated that in GCT mares, serum AMH concentrations (1901.4 ± 1144.6 ng/ml) were higher compared to mares that were cycling (0.96 ± 0.08 ng/ml) or those which were pregnant (0.72 ± 0.05 ng/ml) (Almeida et al., 2011). AMH structure is highly conserved across species (Cate et al., 1986). Following favorable preliminary results that differentiated spayed and intact mature dog as previously shown (Axnér and Ström Holst, 2015; Hollinshead et al., 2017; Place et al., 2011), the current study measured circulating AMH in a wide range of dogs, from 1 month to 14 years old. The ELISA was performed on all test samples on the same day. None of the animals (n = 7) under the age of 4.5 months had measurable AMH. Two of four animals who were 4.5 and 5 months of age, respectively, also did not have detectable AMH levels. In women, AMH is secreted from granulosa cells in ovaries and production starts as soon as follicles are initially recruited (McGee and Hsueh, 2000) into the growing pool and reaching the highest levels of production in preantral and small antral follicles 70 (Axnér and Ström Holst, 2015; Mossa et al., 2017; Place et al., 2011; Yagci et al., 2016). In women AMH concentration has been shown to increase from birth to three months of age, peaking at an average age of 25, and thereafter declines steadily and becomes undetectable at menopause (Hagen et al., 2011; Kelsey et al., 2011; Nelson et al., 2011). Similar changes have been observed in cattle of different breeds such as Holstein, Maine-Anjou beef heifers and Bos indicus Nelore (Batista et al., 2016; Monniaux et al., 2012; Mossa et al., 2013). For example, AMH concentrations in Holstein female calves increase in the first 2 months, decrease at 5 months and are stable at 8-9 months suggesting that the AMH concentrations observed in heifers before puberty may be reflective of the changes seen in the growth pattern of small follicles as they are recruited from the reserve pool into the growth pool (Mossa et al., 2013). In the current study, average circulating AMH levels in all female dogs (n = 25, baseline and proestrus) from 1 to 6 years was 0.86 ± 0.11 ng/ml (which were significantly lower than those published by Hollinshead et al. (2.9 – 21.10 ng/ml), however their values were measured at estrus using a dog based AMH ELISA (Canine AMH ELISA; AnshLabsÒ) (Hollinshead et al., 2017). Hollinshead et al. measured serum AMH during estrous, however, serum AMH levels in all four breed categories (giant breeds small breeds, medium breeds and large breeds) overlapped significantly, with the reference intervals spanning 5.5 folds (Hollinshead et al., 2017). The current study demonstrates that there is a significant difference in circulating AMH levels during anestrus (baseline) and proestrus. It is possible that the higher levels seen by Hollinshead et. al. was in part due to the animals being in estrus. Another factor that could explain this variation is due to a possible difference in the AMH sequence that is used in the species specific ELISA kits (Hollinshead et al., 2017). However, establishing a true relationship would require parallel comparison of serum samples as significant variation has been demonstrated while measuring 71 human AMH in different settings (Freour et al., 2007; Li et al., 2012). AMH values in the current study were closer to the mean serum levels published by Yilmas et al. Circulating AMH values of un-spayed dogs using the Beckman and Coulter Gen II AMH ELISA (n = 10, 1.5 – 13 years age) was 4.26 ± 0.82 ng/ml, however these dogs were in different stages of the estrus cycle (anestrus (n = 3), proestrus (n = 1), estrus (n = 1), and diestrus (n = 5)) which would have increased the average circulating AMH (Turna Yilmaz et al., 2015). Extensive studies on ovarian aging such as those performed in women, rodents and food production animals are limited in dogs. As has been seen humans (Fleming et al., 2015) and in one study of dogs (Hollinshead et al., 2017), we found a progressive decline in circulating AMH concentration with advancing age in dogs. In women and in heifers, there is evidence to support that AMH concentrations vary minimally over the course of the menstrual/estrus cycle (Cook et al., 2000; Hehenkamp et al., 2006; Ireland et al., 2011) however a multivariate analysis in women (n = 17) under 38 years showed significantly larger intra-individual variation in AMH levels (Overbeek et al., 2012), with higher cyclic variations seen in the late follicular phase (Hadlow et al., 2013) and higher mean AMH levels (Kissell et al., 2014; Randolph et al., 2014) compared of women who were older. However it is generally accepted that these variations are not significant (Fleming et al., 2015). Our study established a significant variation in dogs that were baseline and those who were in proestrus as has been previously seen in by Nagashima et al. (n = 5) (Nagashima et al., 2016). We limited collecting serum samples from animals who did not display overt signs of heat (estrus) such as swollen vulva, serosanguinous discharge, as Nagashima et al. have shown a 2-fold increase between anestrus (mean 0.30 ± 0.01 ng/ml) and estrus, reaching concentrations as high as 0.99 ng/ml (mean 0.64 ± 0.03 ng/ml, 1.5 – 4 years) (Nagashima et al., 2016). The positive correlation of circulating AMH of baseline dogs with age as opposed to AMH from dogs in proestrus, suggests 72 that baseline AMH might be a better indicator of fertility. We are however unable to confirm the exact status of dogs with progesterone levels higher than baseline and it is possible that there may be overlap in animals in proestrus and diestrus. The substantial variation of circulating AMH during the estrus cycle is possibly because dog is a litter bearing animal therefore having a proportionally larger group of reserve follicles that enter into the growth phase and proportionally larger numbers that ovulate. This study had a few limitations. It was not feasible to process entire ovaries in these dogs to count the total numbers of primordial and primary follicles and perhaps this is a reason why the relationship between AMH and follicle reserves could not be established. Another limitation in this study was the inability to perform a longitudinal analysis of the study cohort and therefore it was not possible to characterize AMH variation within the entire estrous cycle and changes seen with advancing age. CONCLUSION In conclusion, this is the first study to demonstrate the relationship between ovarian reserves and advancing age. This study has also affirmed that anestrous AMH is significantly lower than proestrus AMH concentrations, unlike in women. Therefore, before serum AMH concentration in breeding dogs is utilized as an indicator of ovarian numbers, the knowledge of the stage of estrus cycle will be critical. Similar to studies in women (Fleming et al., 2015; Gomez et al., 2016), this study demonstrated a strong negative correlation between dog ovarian follicle reserves and advancing age. 73 APPENDICES 74 Reagent d-Limonene Ethanol 100% Ethanol 90% Ethanol 80% Ethanol 70% Hematoxylin Tap water Acid 0.3% alcohol Tap water 2x 2x 1x 1x 1x 1x 1x 1x 1x Time 10 min each 5 min 1 min 1 min 1 min 2 min 2 min 2 dips 3 dips Notes We use a 12-slide coplin staining jar for most of these reagents made fresh every day made fresh every day made fresh every day Will need to be changed occasionally Standing tap water Add commercial grade ethanol (350 ml) + distilled water (150 ml) + concentrated hydrochloric acid (1.5 ml) Standing tap water 0.2% Ammonia water solution: Add concentrated Ammonium hydroxide (1ml) to distiller water (500 ml) Appendix 4.1. Hematoxylin and eosin staining protocol Place the slides in a hot air oven for 25 minutes at 55-60 °C to drip out the excess paraffin. Proceed with the following steps Blueing agent 1x 2 min tap Running water Ethanol 70% Eosin Ethanol 80% Ethanol 90% Ethanol 100% d-Limonene Mounting 1x 1x 1x 1x 1x 2x 2x Place the slide holder in a tray filled with gently running water. Place at an angle, so that the excess water runs off. Eosin stain loses its efficiency after 4-5 uses. made fresh every day made fresh every day May use xylene. We have not used it. DPX mountant and place a cover slip avoiding air bubbles. 8 min 3 min 2 min 5 min 10 sec 10 each 10 each sec sec 75 Appendix 4.2. VDL Endocrinology laboratory Progesterone assay ranges and their implication for breeding Estimate for Ovulation (no. of days) Estimated Time for Breeding (no. of days) Not applicable Earliest estimated window for breeding is from 4-6 days, but could be longer Estimated window for breeding is from 3-5 days, but could be longer Estimated window for breeding is 2-4 days Estimated window for breeding is 1-3 days Estimated window for breeding is 0-2 days Breed at once (0-1 day) • Concentration of Progesterone 0-2 nmol/L 3-6 nmol/L 7-12 nmol/L 13-18 nmol/L 19-31 nmol/L 32-64 nmol/L 65-90 nmol/L >90 nmol/L Baseline concentration, too early to estimate ovulation Minimum of 2 days before ovulation is expected. Results of 3-4 nmol/L may persist for a week or longer before increasing Minimum of 1 day before ovulation is expected Ovulation is impending or has just occurred Ovulation recently occurred Ova have matured, optimal potential for fertility Ova have matured by aging, decreased potential for fertility Too late or very reduced potential for fertility 76 REFERENCES 77 2016a. Corning® BioCoat™ Fibronectin Cell Culture Inserts. REFERENCES https://www.thomassci.com/Laboratory-Supplies/Tissue-Culture-Cellware 2016b. OMIA - Online Mendelian Inheritance in Animals. http://omia.angis.org.au/home/ 2016c. U S Food and Drug Administration Home Page. http://www.fda.gov/ 2016d. U.S. Pet Ownership Statistics. https://www.avma.org/KB/Resources/Statistics/Pages/Market-research-statistics-US-pet- ownership.aspx. (Accessed December 2016). Abdel-Ghani, M.A., Shimizu, T., Suzuki, H., 2014. 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An agarose-gel based method for transporting cell lines. 88 Curr Chem Genomics 3, 50-53. CHAPTER 5. SUMMARY AND FUTURE DIRECTIONS 89 SUMMARY The project had three specific aims. To first aim was to establish a dog ovarian cortical explant tissue culture model. The second aim was to model the effects of TAM on the dog ovarian cortical explants. And finally, the third aim was to characterize AMH in dogs. The study showed superiority of using 1.5% agarose gel system for promoting primordial and primary follicle survival. This method had the smallest SEM, was economical and uncomplicated. This study also showed that high dose CTXH caused marked follicular loss (P < 0.05) whereas treatment with TAM decreased follicular loss (P < 0.05) from CTX in-vitro, thus demonstrating that TAM had a similar protective action on dog ovaries as has been previously demonstrated in rodents. TAM alone did not have an effect on morphologically normal reserve follicle counts (P > 0.05). Finally, the current study demonstrated that spayed and intact female dogs can be successfully identified using a readily available human AMH ELISA. The results also demonstrated a significant negative relationship between AMH and advancing age and that there was a significant difference in serum AMH concentrations at anestrus and proestrus. FUTURE DIRECTIONS Optimize the ovarian cortical explant culture system Although, the agarose gel system was superior over the fibronectin coated inserts, there was a significant loss of viable follicles after 72-hours of culture suggesting that we are yet to harness the correct media and microenvironment to optimize and sustain viable follicles for longer periods. There have been significant strides made in recent years with respect to dog embryo in-vitro 90 fertilization by Nagashima et. al. in recent years where they produced the first ever successful live births from in-vitro fertilized embryos (Nagashima et al., 2015). They however still were dependent upon in-vivo matured oocytes. The lack of understanding has restricted the ability to develop the dog as the preferred model for translation reproductive research. Determine the mechanism of action of TAM This study was limited by the number of sections that could be obtained from a pair of ovaries since there was a vast difference in sizes of ovaries depending on the dog breed. Although there was a subjective decrease in apoptotic primordial and primary follicles in ovarian sections co treated with TAM and CTX, we could not demonstrate a significant association between CTX and apoptosis in the dog ovarian follicle sections as has been demonstrated in rodents (Piasecka-Srader et al., 2015). This was perhaps because of the smaller sample sizes that were used for this part of the study. Another limitation of our study was our inability to demonstrate the mechanism by which TAM annulled the effect of CTX on dog ovaries. Piesecka-Srader et al. demonstrated that TAM decreased the expression of multiple inflammation related genes, such as mediators of lipoxygenase and prostaglandin production, cytokine binding ,second messenger signaling, tissue modelling and vasodilation (Piasecka-Srader et al., 2015). Another study looking at radiation induced follicular loss implicated enhancement of insulin like- growth factor -1 by tamoxifen (Mahran et al., 2013). Because this study was limited by number of sections that could be obtained per ovary set, increasing the sample size, culturing and using a subset of cultured sections for mechanistic should be undertaken. 91 Better characterization of circulating AMH in dogs We demonstrated that circulating AMH varied significantly between dogs in anestrus and proestrus. We were unable to differentiate dogs that were in proestrus or diestrus based on serum progesterone. One of the long-term objectives is to use serum AMH in breeding dogs and service dogs as a marker for fertility to make sound breeding decisions. More comprehensive data and serum sampling is needed to determine absolute variations seen over the course of an estrus cycle and trends with advancing age. Mossa et. al. have demonstrated a significant variation between beef and dairy heifers (Mossa et al., 2017). The current study’s cohort was predominantly made up of mixed breeds, predominantly Chihuahua mixes and Pitt mixes and we could not establish a relationship based on breed sizes, in part perhaps because of smaller sample size. Hollinshead et. al. has shown significant effect of bitch size on serum AMH (Hollinshead et al., 2017). However, all their samples were collected during estrus and may have exaggerated the results. To conclude, to our knowledge this is the first study that examines the effects of coadministration of TAM with CTX in dogs and demonstrates that TAM has protective effects on ovarian follicles when it is co-administered with cyclophosphamide. This is the first study to demonstrate the relationship between ovarian reserves and advancing age. This study has also affirmed that anestrous AMH is significantly lower than proestrus AMH concentrations, unlike in women. Therefore, before serum AMH concentration is utilized as an indicator of ovarian numbers, the knowledge of the stage of estrus cycle will be critical. Similar to studies in women (Fleming et al., 2015; Gomez et al., 2016), this study demonstrated a strong negative correlation between dog ovarian follicle reserves and advancing age. 92 REFERENCES 93 2016a. Corning® BioCoat™ Fibronectin Cell Culture Inserts. REFERENCES http://www.thomassci.com/laboratory-Supplies/tissue-culture-cellware/ 2016b. 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