THE EFFECT OF EXCESSIVE DOSES OF FOLLICLE-STIMULATING HORMONE DURING OVARIAN STIMULATION ON OVULATORY-FOLLICLE FUNCTION AND OOCYTE QUALITY IN THE SMALL OVARIAN RESERVE HEIFER MODEL By Kaitlin Rose Karl A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Animal Science – Doctor of Philosophy 2023 ABSTRACT High doses of follicle stimulating hormone (FSH) during assisted reproductive technologies (ART) are correlated with decreased live birth rates in women and decreased ovarian function in cattle. Thus, the overarching hypothesis for my thesis research is that high FSH doses during ovarian stimulation are detrimental to ovulatory follicle function and oocyte quality. The small ovarian reserve (number of morphologically healthy oocytes) heifer (SORH) model was chosen to test this hypothesis because it mimics traits in infertile women with a small ovarian reserve who require ART to achieve pregnancy. The excessive FSH (Ex-FSH) doses not further increasing number of ovulatory-size follicles during ovarian stimulation were detrimental to ovulatory follicle function because they decreased circulating concentrations of estradiol and ovulation rate and caused severe abnormalities in cell-signaling pathways in granulosa and cumulus cells and oocytes resulting in aberrant luteinization, cumulus expansion, and resumption of meiosis. The prematurely expanded cumulus-oocyte complexes (COCs) contained poor quality oocytes while capacity of COCs to respond to an ovulatory hCG stimulus and resume meiosis was impaired. Circulating concentrations of anti-Mullerian hormone (AMH) and antral follicle count (AFC) in the SORH model were correlated positively with number of functional and dysfunctional ovulatory-size follicles developing during ovarian stimulation with various FSH doses (including Ex-FSH). These results support the conclusions that: i) ovarian stimulation with Ex-FSH impairs ovulatory follicle function and oocyte quality, thereby increasing oocyte wastage and decreasing ART outcomes, and ii) AMH and AFC are not predictive of the impact of Ex-FSH doses during ovarian stimulation on number of high-quality oocytes available for ART. This thesis is dedicated to my family. Thank you for always believing in me. iii ACKNOWLEDGMENTS I would first like to thank my major professor Dr. James Ireland for taking a chance on hiring a tenacious undergraduate student that was eager to learn about assisted reproduction. The unwavering dedication you have provided me over the years is invaluable. I would also like to thank my various lab mates throughout the years: Mrs. Janet Ireland, Dr. Fermin Jimenez- Krassel, Dr. Zaramasina Clark and Dr. Lilian Martins; without your knowledge and expertise, this research would not have been possible. Thank you to my committee members: Dr. Richard Pursley, Dr. Keith Latham, Dr. Robert Tempelman and Dr. Karl Olson, for your guidance as I completed my program. I would also like to acknowledge the Reproductive and Developmental Sciences Program at Michigan State University for its support. Thank you to Green Meadow Farms, Inc. for allowing me to select heifers from their herd for these projects. An additional thank you to the Michigan State University Beef Cattle and Research Center and all of the employees for allowing us to conduct our experiments using their facilities, and to West Michigan Beef Co LLC for allowing us access to their slaughter facility to collect abattoir ovaries for crucial experimental development. Finally, a special thank you to my family and friends. Your unwavering confidence in my abilities was an immense support during this undertaking. I am eternally grateful for your patience as I worked through my degree. This study was made possible and supported by the Agriculture and Food Research Initiative Competitive USDA-NIH Dual Purpose Program Grant no. 2017-67015-26084 from the USDA National Institute of Food and Agriculture, the Eunice Kennedy Shriver National Institute of Child Health & Human Development of the National Institutes of Health under Award Number T32HD087166, MSU AgBioResearch, and Michigan State University. iv TABLE OF CONTENTS LIST OF ABBREVIATIONS .............................................................................................vii CHAPTER 1. Introduction .................................................................................................1 REFERENCES ................................................................................................................12 APPENDIX ......................................................................................................................20 CHAPTER 2. Negative Impact of High Doses of Follicle-Stimulating Hormone During Superovulation on the Ovulatory Follicle Function in Small Ovarian Reserve Dairy Heifers ...................................................................................................................................22 Abstract ...........................................................................................................................24 Introduction ......................................................................................................................25 Materials and Methods .....................................................................................................28 Results............................................................................................................................. 33 Discussion ........................................................................................................................37 REFERENCES ................................................................................................................43 APPENDIX ......................................................................................................................49 CHAPTER 3. Excessive Follicle-Stimulating Hormone During Ovarian Stimulation of Cattle May Induce Premature Luteinization of Most Ovulatory-Size Follicles ............55 Abstract ............................................................................................................................57 Introduction ......................................................................................................................58 Materials and Methods .....................................................................................................60 Results ..............................................................................................................................65 Discussion ........................................................................................................................67 REFERENCES ................................................................................................................75 APPENDIX ......................................................................................................................81 CHAPTER 4. Follicular Hyperstimulation Dysgenesis: New Explanation for Adverse Effects of Excessive FSH in Ovarian Stimulation .............................................................87 Abstract ............................................................................................................................89 Introduction ......................................................................................................................90 Materials and Methods .....................................................................................................93 Results ..............................................................................................................................103 Discussion ........................................................................................................................114 REFERENCES ................................................................................................................123 APPENDIX ......................................................................................................................131 CHAPTER 5. Ovarian Stimulation with Excessive FSH Doses Causes Cumulus Cell and Oocyte Dysfunction in Small Ovarian Reserve Heifers ............................................140 Abstract ............................................................................................................................142 Introduction ......................................................................................................................143 Materials and Methods .....................................................................................................145 Results ..............................................................................................................................154 Discussion ........................................................................................................................157 v REFERENCES ................................................................................................................163 APPENDIX ......................................................................................................................168 CHAPTER 6. Limitations in Use of Ovarian Reserve Biomarkers to Predict the Superovulation Response in Small Ovarian Reserve Heifers.......................................... 187 Abstract ............................................................................................................................189 Introduction ......................................................................................................................190 Materials and Methods .....................................................................................................193 Results ..............................................................................................................................198 Discussion ........................................................................................................................201 Conclusion .......................................................................................................................206 REFERENCES ................................................................................................................207 APPENDIX ......................................................................................................................213 CHAPTER 7. Synopsis, Conclusion, and Practical Applications ....................................219 Synopsis ...........................................................................................................................220 Overall Conclusion ..........................................................................................................237 Practical Applications ......................................................................................................238 REFERENCES ................................................................................................................242 APPENDIX ......................................................................................................................247 vi AFC AMH ART BSA CC CL LIST OF ABBREVIATIONS antral follicle count anti-Müllerian hormone assisted reproductive technology bovine serum albumin cumulus cells corpus luteum/corpora lutea COC cumulus-oocyte complex cCOC and/or comCOC compact cumulus-oocyte complex cpFSH d D DEGs DPBS E EA EI EEI eCG commercial FSH-enriched porcine pituitary preparation of follicle- stimulating hormone day(s) day of treatment period differentially expressed genes Dulbecco’s Phosphate Buffered Saline estrogen estrogen-active estrogen-inactive extreme-estrogen-inactive equine chorionic gonadotropin eCOC and/or expCOC expanded cumulus-oocyte complex E:P Ex-cpFSH estrogen:progesterone excessive commercial FSH-enriched porcine pituitary preparation of follicle-stimulating hormone vii FDR FET FF FHD FPKM FSH GC GIFT GV GVBD h hCG hFSH false discovery rate frozen embryo transfer follicular fluid follicle hyperstimulation dysgenesis fragments per kilobase of exon per million mapped fragments follicle-stimulating hormone granulosa cells gamete intrafallopian transfer germinal vesicle germinal vesicle breakdown hours(s) human chorionic gonadotropin recombinant human follicle-stimulating hormone HH medium HEPES buffered medium HMG IBMX ICSI im IU IVF IVM kg lbs human menopausal gonadotropin 3-isobutyl-1-methylxanthine intracytoplasmic sperm injection intramuscular international units in vitro fertilization in vitro maturation kilograms pounds viii LH mg min mL m mm mmHG MI MII luteinizing hormone milligram minute(s) milliliter meter millimeter mm of mercury metaphase I metaphase II n and/or N number NPPC O OPU PG P PMSG SEM SORH ZIFT μg μl C-type natriuretic peptide oxytocin oocyte pick-up prostaglandin F2α progesterone pregnant mare serum gonadotropin standard error of the mean small ovarian reserve heifer zygote intrafallopian transfer microgram microliter ix CHAPTER 1. Introduction 1 Infertility and ART: Every person has the right to experience the enjoyment of having a family. However, the World Health Organization reports that approximately 1 of every 6 individuals of reproductive age (e.g., 1.9 billion women are 15-49 years old [1]) worldwide experiences infertility, which is defined as a failure to achieve a pregnancy after 12 months of unprotected sexual intercourse [2] in their lifetime [2, 3]. Moreover, infertility is increasing 5 – 10% per year globally [4]. Assisted reproductive technologies (ART) are medical procedures used to treat infertility, which usually results from a small ovarian reserve (total number of morphologically healthy oocytes), blocked fallopian tubes, endometriosis, low sperm count, polycystic ovarian disease, uterine problems, or unexplained problems [4]. Because of high costs of ART ($5,000 to $73,000, with 85% of these costs being paid out of pocket [2]) and poor patient access to ART (e.g., geographic disparities [5]), only 5% of all infertile couples use ART even though it has resulted in birth of over 9 million babies worldwide [4, 6] since the first successful embryo transfer in women in 1978 [7]. ART is not only costly but inefficient: The medical procedures used during ART usually require mechanical recovery of sperm and oocytes which are then combined during in vitro fertilization (IVF) or intracytoplasmic sperm injection (ICSI). The fertilized oocyte (embryo) is then transferred mechanically back into the female to produce a pregnancy. ART also includes gamete intrafallopian transfer (GIFT), zygote intrafallopian transfer (ZIFT) and frozen embryo transfer (FET). A critical and commonly used technique during ART is ovarian stimulation with pharmacological doses of follicle-stimulating hormone (FSH), which is a pituitary glycoprotein hormone required for ovarian follicular growth and development. FSH is typically injected daily into women for one to two weeks during an ART cycle to stimulate maximal growth of ovulatory-size follicles. A relatively large number of ovulatory-size follicles developing in 2 response to the FSH (e.g., a dozen) injections enhances the likelihood that the clinician will recover enough high-quality oocytes for successful IVF. Approximately 2 to 5 days after IVF of the oocytes, the highest quality embryo(s) are transferred back into the donor or a recipient resulting in an approximate 24% chance of a live birth per ART cycle [4, 8]. However, ovarian stimulation with FSH results in extremely variable responses typically ranging from 0 to 52 ovulatory-size follicles [9]. Moreover, this high variability in number of ovulatory-size follicles often results in the need for repetitive ovarian stimulation cycles during ART (range = 1 – 9, [10]) before a patient achieves a successful pregnancy. The reason multiple ART cycles are required for a successful pregnancy is primarily because most oocytes recovered from ovulatory- size follicles do not fertilize [11], or do not develop normally after IVF [12, 13]. Indeed, 90-95% of all oocytes retrieved from ovulatory-size follicles for IVF do not result in a live birth [14]! High FSH doses during ART are linked to low live birth rates: The high variability in responsiveness of women to ovarian stimulation, coupled with the absence of reliable biomarkers to accurately predict responsiveness of patients to ovarian stimulation, very likely explains why the FSH doses typically used during ovarian stimulation procedures range up to 20-fold among different ART clinics [15]. These high FSH doses during ART, however, are associated with poor oocyte quality and reduced embryo survival [16, 17], as well as high oocyte and embryo wastage [14-16, 18-22]. For example, a recent study with over 500,000 women reported a negative relationship between FSH dose, number of oocytes retrieved [23] and live birth rate independent of age, weight or health of the donor [15]. In this large study, the highest FSH doses were associated with nearly a 50% reduction in live birth rates while relatively low FSH doses resulted in the highest live birth rates. Despite the inverse relationship between FSH dose and live birth rate, nearly 70% of women in this study received high FSH doses during their ovarian 3 stimulation protocols [15]. Taken together, these studies support the overarching hypothesis for my thesis research that high doses of FSH during ovarian stimulation are detrimental to ovulatory follicle function and oocyte quality. The bovine is a relevant biomedical model to evaluate pharmacological FSH action during ART: Most of the data supporting my overarching hypothesis are correlative. Thus, the direct effect of high FSH doses and the associated mechanism of action of FSH on ovarian function, oocyte quality and embryo survival have heretofore not been examined. I chose the bovine as a relevant biomedical model to test my hypothesis for several reasons: Firstly, cattle share many reproductive attributes similar to those in women, including multiple follicular waves during each reproductive cycle [24, 25], a single ovulation during each reproductive cycle [24, 25], a relatively long reproductive cycle and similar gestation length [26- 28], and similar ovarian stimulation protocols during embryo transfer [28, 29]. Secondly, studies in cattle also imply that high FSH doses may be detrimental to ART outcomes. For example, high doses of FSH or FSH-like factors (e.g., pregnant mare serum gonadotropin, human menopausal gonadotropin) used to induce superovulation decrease fertilization rate [30-32], embryo yield [32, 33] or quality [30, 32] and number of transferable embryos [31-34]. In addition, high FSH doses are linked to high oocyte and embryo wastage [30- 36]. Some studies reported negative effects of high FSH doses on number of corpora lutea (CL) [35, 37], circulating progesterone [35] and estradiol [37] concentrations, and ovulation rate [35]. Still other studies show that high FSH doses increase the number of antral follicles [38, 39], estradiol [35] and progesterone concentrations [37], while others report no effects of high FSH doses on CL number [30-32, 38-40], circulating progesterone [38] or estradiol [30] concentrations, or ovulation rate [30, 39]. Such variability among studies may reflect use of 4 diverse sources and doses of FSH or FSH-like products to superovulate cattle of various breeds, ages, or parities and with unknown numbers of follicles in the ovarian reserve. Thirdly, most women seek ART because they have a small ovarian reserve [41], and women with small ovarian reserves share many similarities with small ovarian reserve cattle [42- 45]. These similarities include a low antral follicle count (AFC, [42]), hypersecretion of FSH [42-44] but low circulating anti-Müllerian hormone (AMH) [45], progesterone [46] and testosterone [47] concentrations during the reproductive cycle, and poor responsiveness to ovarian stimulation [42]. Because the reproductive traits, ART procedures, and the negative effects of high FSH doses during ovarian stimulation on ART outcomes are similar between cattle and women, the small ovarian reserve heifer (SORH) was chosen as a highly relevant biomedical model to test my overarching hypothesis that high doses of FSH during ovarian stimulation are detrimental to ovulatory follicle function and oocyte quality. Numerous precautions were taken in my studies to improve interpretation of the effects of high FSH during ovarian stimulation on ovulatory follicle function and oocyte quality in the SORH model: Like women, responsiveness of cattle to ovarian stimulation is also typically highly variable ranging from 0 to 100 ovulatory-size follicles developing in response to FSH treatments [48]. Many reviews [49-54] have identified a variety of factors that cause or contribute to the high variability in response of cattle to ovarian stimulation, including for example breed [49, 52, 54, 55], age [49, 52, 54], parity [52, 54], nutrition [49, 52, 54], follicular waves [50-52, 54], presence of dominant follicle [49-51], antral follicle count and the ovarian reserve [56-58], season [49, 52, 54], and source of hormone [51, 52]. To prevent or minimize the 5 potential impact of variables that may confound interpretation of the effects of high FSH doses on ovulatory follicle function and oocyte quality, I standardized my studies as follows: Firstly, only reproductively mature 11- to 12-month-old Holstein heifers weighing 800 to 900 lbs and purchased from a single commercial dairy farm (Green Meadow Farms Inc, Ovid- Elsie, MI) were used to complete my planned studies. In addition, once heifers were transported to campus, they were fed the same balanced diet and housed in the same location (Michigan State University Beef Cattle Teaching and Research Center, East Lansing, MI) throughout my studies. Secondly, the size of the ovarian reserve is highly variable amongst cattle (e.g. at birth, 1,000 to 400,000 oocytes [59]) with 15% to 20% of the herd usually having a relatively small ovarian reserve compared with age-matched counterparts [42, 44, 45, 60-62]). Thus, only heifers with a relatively low AFC (≤15 follicles, determined via serial ovarian ultrasonography [56, 63]), and correspondingly a small ovarian reserve (80% fewer oocytes compared with heifers with a high AFC, [64]) were purchased from Green Meadow Farms Inc and used in my studies. This rigid heifer selection criteria based on AFC was expected to further reduce variability in response to ovarian stimulation and to improve the biomedical relevance of the SORH model, especially because a small ovarian reserve is the primary reason women seek ART [4, 41]. Thirdly, ovarian stimulation in cattle typically requires twice daily injections of FSH (12 hours between injections) for 4 days. However, another source of variation in response to FSH during ovarian stimulation is the timing of the first FSH injection during the 21-day estrous cycle of cattle. Heifers typically have two or three independent follicular waves approximately 7 to 10 days in length depending on the number of waves occurring during the estrous cycle [65]. Each follicular wave is characterized by the emergence and atresia of dozens of antral follicles 6 culminating in selection and development of a single ovulatory-size dominant follicle. However, only the dominant follicle developing during the follicular phase of the estrous cycle ovulates while the others developing during the luteal phase undergo atresia [65]. Response to FSH injections during ovarian stimulation is greater if the injection is initiated early in the follicular wave (day of emergence or the day after) before a dominant follicle is selected [66-68]. Moreover, responsiveness to FSH is greater during the first compared with subsequent follicular waves [66]. Thus, the FSH injections during ovarian stimulation in my studies were standardized to begin prior to selection of the dominant follicle and initiation of the first follicular wave, which is ±1 day from ovulation (Day 1 of estrous cycle, Figure 1). Fourthly, the timing of the first FSH injection to coincide with initiation of the first follicular wave in my studies, instead of the second or third wave as commonly observed in heifers undergoing ovarian stimulation and embryo transfer [50, 66, 69], ensured that circulating concentration of progesterone was relatively low [70]. A series of prostaglandin F2α (PG) injections coinciding with subsequent FSH injections during ovarian stimulation (Figure 1) was then used in the SORH model to ensure that the circulating progesterone concentrations remained low (< 0.5 ng/ml, [56]) throughout FSH treatments. This protocol was administered to: i) reduce the potential impact of high progesterone on FSH action and dominant follicle function, and ii) to better mimic the low progesterone concentrations during the follicular phase of the menstrual cycle in women when ovarian stimulation protocols are typically administered during ART [71-73]. Fifthly, a single commercial source of FSH was used during ovarian stimulation of the SORH model. FSH is the key driver of folliculogenesis in all mammals [74, 75]. Thus, FSH is the primary hormone used for ovarian stimulation during ART in women and cattle. Historically, 7 attempts to purify hormones, such as human chorionic gonadotropin (hCG) and equine chorionic gonadotropin (eCG, PMSG), for ovarian stimulation began in the late 1920’s and early 1930’s [76]. Efforts progressed to the 1960’s when human menopausal gonadotropin (HMG) was extracted. However, it wasn’t until the 1990’s that recombinant human FSH was successfully used during ovarian stimulation of women to produce a pregnancy through IVF [76]. Similarly, in bovine, eCG, PMSG and HMG were the standard stimulatory drugs used during ART procedures. However, in the late 1970’s, a crude porcine pituitary extract containing FSH with low LH contamination was used to improve responsiveness of cattle to ovarian stimulation [50]. Today, this pituitary extract is called Folltropin-V and sold commercially by Vetoquinol Inc (Princeville, Quebec) and is the only FDA approved drug for superovulation in cattle. Folltropin-V (NIH-FSH-P1) is an extract of porcine pituitary glands that contains 700 IU (equivalent to 400 mg NIH-FSH-P1 with 0.25% LH contamination or <1 mg NIH-LH-S19) of FSH per 20 ml vial [77]. Since FSH has a relatively short half-life in bovine (approximately 5 h), twice daily intramuscular injections of Folltropin-V (87.5 IU/injection) at 12-hour intervals for four consecutive days (a total of 8 injections) using a decreasing dose regimen (half doses for last 4 injections) is the typical industry recommended injection protocol during ovarian stimulation of cattle [77]. Folltropin-V continues to be widely and successfully used for ovarian stimulation in the bovine embryo transfer industry. Although recombinant bovine FSH products have been developed [78-80], they are not available for commercial use in cattle. Thus, Folltropin-V was used for ovarian stimulation of the SORH model in my studies. Hereafter, this commercial porcine pituitary extract of FSH is referred to as cpFSH. To minimize potential batch to batch variation in cpFSH bioactivity during production, I used the same batch of cpFSH during ovarian stimulation for each study. 8 Sixthly, dose response studies were conducted to evaluate responsiveness of the SORH model to ovarian stimulation with cpFSH. The doses of cpFSH used in my studies were 35 IU (20 mg /1 ml), 70 IU (40 mg/2 ml), 140 IU (80 mg /4 ml) and 210 IU (120 mg/6 ml) per injection. The cpFSH dose range is 60% lower and 240% higher than the Vetoquinol industry standard recommended dose. Note, this four-fold range in cpFSH doses is much narrower than the 20-fold range in doses of human recombinant FSH injected into women during ART [81]. Each of the 8 doses of cpFSH during ovarian stimulation of the SORH model were equal to eliminate any potential confounding effects of a decreasing dose regimen. During all ovarian stimulation protocols, the SORH model was injected with PG twice to synchronize estrous cycles. As depicted in Figure 1, the first injection of cpFSH was given 36 hours after the last PG injection (± 1 day of ovulation and emergence of the first follicular wave) followed by 7 additional cpFSH injections at 12-hour intervals. To regress the newly formed CL after ovulation and maintain low progesterone concentrations throughout the treatment period, a further three PG injections were given at 12-hour intervals starting at the time of the 7th cpFSH injection (Day 4-5 of estrous cycle, Figure 1). In some studies, a single 2500 IU injection of hCG, which is sufficient to ovulate 40 to 60 follicles in heifers [82-84], was given coincident with the third PG injection (12 hours after the last (or 8th) cpFSH injection) to induce ovulation (Figure 1). This cpFSH and PG injection protocol was used in all my studies to evaluate FSH action during ovarian stimulation on ovarian function (Figure 1). Seventhly, the crossover experimental designs used in my FSH dose response studies (Chapters II, V, VI) using the SORH model were novel because they further minimized the sources of variation that could confound interpretation of the direct effects of cpFSH on ovulatory follicle function and oocyte quality. For example, when FSH dose response studies 9 were conducted using the Williams Latin Square Design (as explained in Chapters II and VI), or when an excessive (210 IU cpFSH) was compared with the industry-standard cpFSH (70 IU) dose using a Crossover Design (as explained in Chapter V), each animal received each of the FSH doses being tested and thus served as its own control. In addition, the sequence of FSH doses during each ovarian stimulation protocol was unique for every heifer and each FSH dosage followed every other FSH dosage an even number of times, thereby being balanced for potential carryover effects (e.g., the effects of one treatment altering the effect of a subsequent condition). Another advantage of the this design, is it minimizes the potential impact of nuisance variables including differences in seasonal temperature, rations or animal genotypes, thereby facilitating greater precision on estimating the effects of the different FSH doses on the endpoints measured in my studies [85, 86]. To test the hypothesis that high FSH doses during ovarian stimulation are detrimental to ovulatory follicle function and oocyte quality, my objectives were to determine if different FSH doses during ovarian stimulation of the SORH model: i) impacted ovarian function (Chapter II [56]), ii) ovulatory follicle function (Chapter III [87]), iii) expression of genes critical for ovulatory follicle function and oocyte quality (Chapter IV [88]), iv) cumulus function and oocyte quality (Chapter V [89]), and v) the reliability of the ovarian reserve biomarkers, AFC and AMH, as predictors of responsiveness to ovarian stimulation (Chapter VI [58]). The results of my studies illustrate that the 210 IU cpFSH dose during ovarian stimulation is excessive because it does not further increase the number of ovulatory-size follicles developing during ovarian stimulation. This excessive FSH dose is also detrimental to ovulatory follicle function because it is linked to the disruption of numerous pathways in granulosa and cumulus cells and oocytes vital to ovulatory follicle function and oocyte quality. 10 These findings support my hypothesis and the conclusion that high FSH doses during ovarian stimulation are detrimental to ovulatory follicle function and oocyte quality. In addition, AMH and AFC were not predictive of the impact of excessive FSH doses during ovarian stimulation on the number of high-quality oocytes available for ART. 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Excessive follicle-stimulating hormone during ovarian stimulation of cattle may induce premature luteinization of most ovulatory-size follicles. Biol Reprod 2022; 106:968-978. Clark ZL, Ruebel ML, Schall PZ, Karl KR, Ireland JJ, Latham KE. Follicular hyperstimulation dysgenesis: new explanation for adverse effects of excessive FSH in ovarian stimulation. Endocrinology 2022; 163. Kaitlin R. Karl PZS, Zaramasina L. Clark, Meghan L. Ruebel, Jose Cibelli, Robert J. Tempelman, Keith E. Latham, James J. Ireland. Ovarian stimulation with excessive FSH doses causes sumulus cell and oocyte dysfunction in small ovarian reserve heifers. Molecular Human Reproduction 2023. 19 APPENDIX Synchronization Stimulation Folltropin + PG Ovulation Rest Period PG PG FSH FSH FSH FSH PG hCG PG PG Half-way Check-In End Day 1 PG 11 d FSH FSH FSH PG FSH 4 d of stimulation 12 h post final FSH PG 28 d 21 d rest Figure 1. Experimental design. During all ovarian stimulation protocols, the SORH model was injected with PG twice to synchronize estrous cycles. The first injection of cpFSH was given 36 hours after the last PG injection (± 1 day of ovulation and emergence of the first follicular wave) followed by 7 additional cpFSH injections at 12-hour intervals. To regress the newly formed CL after ovulation and maintain low progesterone concentrations throughout the treatment period, a further three PG injections were given at 12-hour intervals starting at the time of the 7th cpFSH injection (Day 4-5 of estrous cycle). In some studies, a single 2500 IU injection of hCG was given coincident with the third PG injection (12 hours after the last (or 8th) cpFSH injection) to induce ovulation. Nine days after hCG-induced ovulation when development of CL was monitored, two PG injections were administered at 12-hour intervals to regress the CLs and allow a spontaneous, non-stimulated estrous cycle to occur to allow circulating hormone concentrations and AFC to return to a basal level before the subsequent ovarian stimulation cycle would begin. This timeframe was referred to as the “rest period” and 20 Figure 1 (cont’d) was a minimum of 21 days, with a half-way check-in (11 days) ultrasonography scan to monitor that ovary size and AFC were returning to normal, that CL were regressing, and no cystic follicles were present. This cpFSH and PG injection protocol was used in all my studies to evaluate FSH action during ovarian stimulation on ovarian function. 21 CHAPTER 2. Negative Impact of High Doses of Follicle-Stimulating Hormone During Superovulation on the Ovulatory Follicle Function in Small Ovarian Reserve Dairy Heifers1 1 Karl, Kaitlin R., Fermin Jimenez-Krassel, Emily Gibbings, Janet LH Ireland, Zaramasina L. Clark, Robert J. Tempelman, Keith E. Latham, and James J. Ireland. "Negative impact of high doses of follicle- stimulating hormone during superovulation on the ovulatory follicle function in small ovarian reserve dairy heifers." Biology of Reproduction, 104(3), (2021): 695-705. By permission of Oxford University Press. 22 Title: Negative Impact of Excessive Doses of Follicle-Stimulating Hormone During Superovulation on Ovulatory Follicle Function in Small Ovarian Reserve Dairy Heifers Running Title: Excessive FSH Doses During Superovulation are Detrimental to Ovulatory Follicle Function Summary Sentence: Excessive doses of Folltropin-V in cattle with a low antral follicle count and small ovarian reserve are detrimental to ovulatory follicle function. Keywords: ovarian stimulation, estradiol, anti-Müllerian hormone, reduced ovulation rate, decreased ovulatory follicle function, decreased number of ovulatory follicles, bovine Authors and affiliations: Kaitlin R. Karl1, Fermin Jimenez-Krassel1, Emily Gibbings1, Janet L.H. Ireland1, Zaramasina L. Clark1, Robert J. Tempelman1, Keith E. Latham1, James J. Ireland1,2 Molecular Reproductive Endocrinology Laboratory, Department of Animal Science1, Michigan State University, East Lansing, MI 48824, Grant support: 1 1 This study was supported by the Agriculture and Food Research Initiative Competitive USDA-NIH Dual Purpose Program Grant no. 2017-67015-26084 from the USDA National Institute of Food and Agriculture and AgBioResearch at Michigan State University. 2 Correspondence: James J. Ireland, Molecular Reproductive Endocrinology Laboratory, Department of Animal Science, Michigan State University, East Lansing, Michigan, 48824, USA. Tel.+1 517 432 1384. Email: ireland@msu.edu 23 Abstract When women with small ovarian reserves are subjected to assisted reproductive technologies (ART), high doses of gonadotropins are linked to high oocyte and embryo wastage and low live birth rates. We hypothesized that excessive FSH doses during superovulation are detrimental to ovulatory follicle function in individuals with a small ovarian reserve. To test this hypothesis, heifers with small ovarian reserves were injected twice daily for four days beginning on Day 1 of the estrous cycle with 35 IU, 70 IU, 140 IU, or 210 IU doses of Folltropin-V (FSH). Each heifer (n = 8) was superovulated four different times (21-24 d between superovulations) with each of the four FSH doses using a Williams Latin Square Design. During each superovulation regimen, three prostaglandin F2α injections were given at 12 h intervals starting at the 7th FSH injection to regress the newly formed corpus luteum (CL). Human chorionic gonadotropin (hCG) was injected 12 h after the last (8th) FSH injection to induce ovulation. Daily ultrasonography and blood sampling were used to determine number and size of follicles and CL, uterine thickness, and circulating concentrations of estradiol, progesterone, and anti- Müllerian hormone (AMH). The highest doses of FSH did not increase AMH, progesterone, ovulatory follicle number or uterine thickness, but decreased estradiol production, number of CL, and ovulation rate compared with lower FSH doses, indicating detrimental effects on ovulatory follicle function. 24 Introduction Assisted reproductive technologies (ART) include intravaginal ultrasound-guided needle aspiration of ovarian follicles to recover eggs, also referred to as oocyte pick-up (OPU) in cattle, or egg retrieval in women, for in vitro fertilization (IVF). ART also includes traditional embryo transfer primarily used in cattle to propagate offspring from genetically superior donors at a greater than normal rate [1]. Both ART techniques use exogenous follicle stimulating hormone (FSH) to stimulate growth of large numbers of ovulatory follicles to obtain oocytes for fertilization [2]. Response to superovulation, however, is highly variable among women [3] and cattle [2, 4] often resulting in costly, unpredictable outcomes [5-7]. Many reasons may exist for this variability, such as age [8, 9] and inherent differences in size of the ovarian reserve [10]. Several studies indicate that high FSH doses may contribute to the variability in superovulation response and outcomes. For example, dose response studies in cattle indicate that high doses of FSH or FSH-like factors (e.g., pregnant mares serum gonadotropin, human menopausal gonadotropin) during superovulation decrease fertilization rate [11-13], embryo yield [13, 14] or quality [11, 13] and number of transferable embryos [12-15]. High FSH doses are also linked to high egg and embryo wastage in cattle [11-17] and women [3, 5, 6, 18-22]. A recent study with over 500,000 women reports a negative relationship between FSH dose and live birth rate independent of age, weight or health of the donor in women [3]. These data support the hypothesis that high FSH doses during superovulation may be detrimental to ovulatory follicle function, which in turn may impair oocyte quality and embryo survival. The large study supporting detrimental effects of high FSH doses on live birth rates during ART in women is correlative [3]. However, determination of the potential direct negative effects of excessive FSH doses on embryo survival in women is unlikely to occur in ART clinics. 25 Consequently, we chose the bovine as an experimental model to examine FSH action during ovarian stimulation because the bovine is a single-ovulating species with a relatively long reproductive cycle like women; and, during embryo transfer, the bovine undergoes ovarian stimulation (superovulation) protocols similar to those used during ART in women. In addition, a major contributing factor compelling women to seek ART is a small ovarian reserve [23]. Thus, another significant advantage of the bovine model is that we have established procedures to identify cattle with small ovarian reserves to evaluate FSH action during superovulation [24-27]. Variable results have emerged using the bovine model with respect to FSH dose on ART outcomes. Some studies reported no effect of high FSH doses on fertilization rate [28-31], embryo yield [28-32] or number of transferable embryos [11, 28-30, 32], but other studies reported negative effects of high FSH doses on embryo transfer outcomes [11-15]. Interestingly, some studies reported negative effects of high FSH doses on number of corpora lutea (CL) [16, 29], circulating progesterone [16] and estradiol [29] concentrations, and ovulation rate [16]. Still other studies report that high FSH doses increased the number of antral follicles [28, 30], estradiol [16] and progesterone concentrations [29] while others reported no effects of high FSH doses on CL number [11-13, 28, 30, 32], circulating progesterone [28] or estradiol [11] concentrations, or ovulation rate [11, 30]. Such variability between studies may reflect use of diverse sources and doses of FSH or FSH-like products to superovulate cattle of various breeds, ages, or parities and with unknown numbers of follicles in the ovarian reserve. Moreover, the FSH dose response studies typically used cross-sectional experimental designs with individual cattle assigned at random to one of several FSH doses with limited numbers of animals per dose (usually 4 or 5), single point in time measurements of most endpoints, different superovulation 26 regimens starting at unknown stages of a follicular wave or CL development, and on different days of the estrous cycle. To minimize the sources of variation and achieve a direct test of effects of high FSH doses on ovarian function in cattle, we employed a powerful experimental design that included small ovarian reserve Holstein heifers and the use of a Williams Latin Square Design for statistical analyses. The small ovarian reserve model minimizes variability in the response to superovulation attributable to age-related differences in the ovarian reserve and mimics several characteristics of women with small ovarian reserves including a low antral follicle count (AFC, [24]), hypersecretion of FSH [24-26], low circulating AMH concentrations [27] during the reproductive cycle, and poor response to superovulation [24]. To test our hypothesis, the same eight 11- to 12-month old Holstein heifers were subjected to superovulation beginning on Day 1 of the estrous cycle (near beginning of first follicular wave) with four different FSH doses using a Williams Latin Square Design [33]. The Williams Latin Square Design was chosen because each animal serves as its own control thus reducing the total number of animals needed to complete the dose response study. Furthermore, because the same heifers were superovulated multiple times, the design balances, and tests for potential carryover effects of one superovulation on response to the next superovulation. Ovulatory follicle function was monitored daily during each superovulation regimen by measuring alterations in circulating ovarian hormones, as well as development of ovulatory follicles. This rigorous study design revealed significant negative effects of high FSH doses on ovarian function. 27 Materials and Methods Identification of small ovarian reserve heifers Our previous studies have established that 11- to 12-month-old heifers with ≤15 follicles of ≥3 mm in diameter during ovarian follicular waves (15% to 20% of a herd) have an 80% smaller ovarian reserve compared with age-matched counterparts with a high AFC (≥ 25 follicles) [24, 26, 27, 34-36]. To identify small ovarian reserve heifers for the study, two groups of 39 and 40 Holstein heifers (11-12 months of age, weighing 318 to 408 kg) located at Green Meadow Farms Inc, Ovid-Elsie, MI were subjected to ovarian ultrasonography to determine AFC and follicle sizes [26]. Approximately half of the heifers from each group had an AFC of ≤10 follicles. These low AFC heifers received two intramuscular injections (25mg/2mL) of prostaglandin F2α (PG, Lutalyse HighCon, Zoetis) 10 d apart to induce luteolysis, which triggers ovulation and development of the first follicular wave of the estrous cycle. Four days after the last PG injection, (about Day 1 to 2 of the estrous cycle and beginning of the first follicular wave), heifers were subjected to ultrasonography to determine AFC. The result was eight heifers identified with an AFC ranging from 5 to 9, subsequently housed at the Michigan State University Beef Cattle Teaching and Research Center for the duration of the project. These heifers were fed maintenance diets that met NRC requirements [37]. The Institutional Animal Care and Use Committee at Michigan State University sanctioned all procedures involving cattle. Superovulation of small ovarian reserve heifers Heifers received three PG injections to synchronize estrous cycles. The first two injections were 10 d apart and the third PG injection was given 12 h after the second PG injection. Each heifer was subjected to daily ultrasonography beginning 36 h after the last PG 28 injection to detect ovulation and the emergence of the first follicular wave. Heifers were then treated with Folltropin-V (Vetoquinol USA Inc), an extract of porcine pituitary glands that contains 700 IU (equivalent to 400 mg NIH-FSH-P1 with 0.25% LH contamination or <1 mg NIH-LH-S19) of FSH per 20 ml vial. Hereafter, because of the minimal amount of LH contamination, Folltropin-V is referred to as FSH. To minimize confounding variables and improve our understanding of the FSH dose effects on ovulatory follicle function, we selected a Williams Latin Square Design. Each of the eight heifers were superovulated a total of four times, but a different FSH dose was administered during the 1st, 2nd, 3rd and 4th superovulation regimen. Furthermore, the sequence of FSH doses during each superovulation differed for every animal. For example, the actual FSH dose sequence for Heifer #1 in our study was twice daily injections for 4 d of 35 IU, 70 IU, 140 IU, and 210 IU FSH during their 1st, 2nd, 3rd, and 4th superovulation, respectively. In contrast, the actual FSH dose sequence for Heifer #2 in our study was twice daily injections for 4 d of 140 IU, 210 IU, 35 IU, and 70 IU FSH for the 1st, 2nd 3rd, and 4th superovulations, respectively. Note that each dosage follows every other dosage treatment exactly twice, thereby being balanced for carryover effects. For example, the dosage of 70 IU follows the dosage of 35 IU twice, i.e., once in Heifer #1 and again in Heifer #8. Another advantage of this Latin Square Design, is that it minimizes the potential impact of other nuisance variables including differences in seasonal temperature, rations or animal genotypes, thereby facilitating greater precision on estimating the effects of the different FSH doses on the endpoints measured in our study [38, 39]. Each heifer was injected intramuscularly twice daily (12 h apart) for four days (8 injections total) with one of four different FSH doses per injection: 1) 35 IU or 20 mg NIH-FSH-P1/1ml (x 8 injections = 280 IU or 160 mg total), 2) 70 IU or 40 mg/2ml (560 IU or 320 mg total), 3) 140 IU or 80 mg /4ml 29 (1120 IU or 640 mg total), 4) 210 IU or 120 mg/6ml (1680 IU or 960 mg total). Hereafter, dose of FSH is referred to as IU per injection. The FSH dose range per injection was 60% lower and 240% higher than the Vetoquinol recommended dose per injection of 87.5 IU. FSH injections began 36 h after the last PG injection which was ±1 d from ovulation and initiation of the first follicular wave in all heifers in our study. To regress the newly formed CL, a further three PG injections were given 12 h apart starting at the time of the 7th FSH injection (about Day 4-5 of estrous cycle). A single 2,500 IU injection of human chorionic gonadotropin (hCG, Chorulon HCG 10,000 IU, Merck Animal Health, USA), which is sufficient to ovulate up to 40 to 60 follicles in heifers [40], was given coincident with the third PG injection (which is 12 h after the last (or 8th) FSH injection) to induce ovulation (see Fig. 2). The FSH treatment period is hereafter defined as Days 1 to 4 when heifers received twice daily FSH injections for 4 days and Day 5 when hCG was injected to stimulate ovulation. Likewise, the post-hCG period encompasses the nine days post-hCG administration (Day 1 to 9) during which CL development was monitored. Blood sampling and ultrasonography To monitor hormone concentrations, coccygeal vein blood samples (10 ml) were collected twice daily (12 h apart) coincident with FSH injections, then once daily for 2 d beginning at the time of the hCG injection (i.e., 12 h after the last FSH injection), and then once every other day over the course of 9 d. To determine follicle number and size, number of CL, and uterine and endometrium thickness, cattle were subjected daily to serial ovarian and uterine ultrasonography beginning on the morning of the day of the last PG injection and continuing until the experiment ended, which was 9 d after the hCG injection. Hereafter, follicle sizes are defined as follows: AFC = all follicles ≥3 mm to <10 mm in diameter and ovulatory follicles ≥10 30 mm in diameter. Uterine and endometrium thickness were measured from cross sections of both the left and right uterine horns approximately ~2 cm from the uterine body bifurcation as previously explained [34, 41]. Immunoassays Serum concentrations of AMH were determined for blood samples collected daily beginning on the morning coincident with the first FSH injection and ending the evening of the day for the last FSH injections (n = 4 samples per heifer). A commercially available AMH ELISA kit for bovine (MOFA Global, Verona, WI) was used to measure AMH concentrations in duplicate 20 µl serum samples in cattle per kit instructions. The two-site AMH assay was previously validated [27] for use in cattle and does not cross react with other members of the transforming growth factor beta (TGFβ) superfamily including TGFβ, bone morphogenic factor- 4 (BMP4), inhibin or activin [42]. The inter- and intra-assay coefficients of variation for the AMH assay were 5.4% and 5.6%, respectively. Serum estradiol-17β concentrations were determined for blood samples collected daily beginning on the morning coincident with the first FSH injection and ending on the morning of the hCG injection (12 h after the last FSH injection) and every other day for 9 d beginning 24 h after hCG (n = 14 samples per heifer) by RIA as previously published [43, 44]. Inter- and intra- assay coefficients of variation for estradiol-17β assays were 7.6% and 3.7%, respectively. Serum concentrations of progesterone were determined for blood samples collected as explained for estradiol (n = 14 samples per heifer) per instructions using a previously validated commercial kit (BC-1113 MP Progesterone Enzyme Immunoassay Test Kit, Catalog, MP Biomedicals Diagnostics Division Orangeburg, NY). The inter- and intra-assay coefficients of variation were 7.8% and 6.8%, respectively. 31 Statistical Analysis All statistical analyses were performed using Statistical Analysis System (SAS 9.4 Institute, Cary, NC) PROC UNIVARIATE and PROC GLIMMIX [45]. Response outcomes analyzed included AFC, ovulatory follicle number, CL number, ovulation rate, uterine and endometrium thickness and circulating concentrations of AMH, estradiol and progesterone. In order to render responses that were more normally distributed, AFC, ovulatory follicle number, and CL number were analyzed using a square root transformation, whereas uterine and endometrium thickness, as well as circulating concentrations of AMH, estradiol and progesterone were analyzed using a logarithmic transformation. Each response variable was analyzed using a repeated measures mixed model accounting for the main effects of period, dose, and, where applicable, day of response within period and all possible interactions. The repeated measures specification allowed for a stronger correlation between repeated measures across days within periods, as opposed to across periods within heifers. Main effects of factors and/or their interactions were considered significant for the corresponding response variable analysis conducted if P≤0.05. The statistical model also accounted for first order carryover effects [46], i.e., effects of the previous superovulation regimen affecting responses within the current superovulation period on each variable. Any interaction involving the FSH dosage factor that was determined to be statistically significant was analyzed using the slice statement in SAS [45] to determine mean differences between dosages separately within levels of the other factor involved in the interaction. When the ANOVA test for FSH was significant, Fisher’s LSD [45] was used to determine if the highest FSH dose differed significantly (P≤0.05) from other FSH doses. All estimated means and standard errors were back-transformed to the scale of observation. 32 Results A key strength of the present study’s experimental design was that each animal received each of the four FSH doses being tested. We observed no carryover effects (P>0.05) for any response variable, implying no evidence that a superovulation period applied within one period impacted the response within the subsequent superovulation regimen. In addition, since this study required 8 months to complete, the use of the Williams Latin Square Design minimized potential confounding effects due to aging and weight gain, changes in season and temperature, or other unknown nuisance variables on the response of Holstein heifers to different FSH doses during superovulation. Consequently, study outcomes informed us about effects of FSH dosage without confounding effects of these other variables. Effect of FSH dose during superovulation on circulating concentrations of AMH Circulating concentrations of AMH were measured during Days 1 to 5 of the FSH treatment. However, AMH concentrations were unaltered during this time period or by the different doses of FSH used to superovulate heifers (Fig. 2A). Effect of FSH dose during superovulation on AFC Although AMH concentrations were unaltered during superovulation, AFC increased (P<0.0001) in heifers during Days 1 to 5 of FSH treatment (Fig. 2B). During Days 1 to 3 of FSH treatment, AFC was similar for heifers treated with all different FSH doses. On Days 4 to 5 of FSH treatment, AFC was similar for heifers treated with 70 IU, 140 IU, and 210 IU FSH doses but greater (P≤0.05) for heifers treated with the 210 IU compared with the 35 IU FSH dose (Fig. 2B). During Days 1 to 9 after hCG administration, AFC decreased (P<0.0001) in heifers treated with the different FSH doses (Fig. 2C). Heifers treated with the 70 IU, 140 IU and 210 IU 33 FSH dose had a similar AFC on Days 1 to 9 post-hCG. However, heifers treated with the 210 IU FSH dose had a lower (P≤0.05) AFC on most days post- hCG compared with heifers treated with the 35 IU FSH dose (Fig. 2C). Effect of FSH dose during superovulation on number of ovulatory follicles The number of ovulatory follicles (≥10 mm in diameter) increased (P<0.0001) during Days 1 to 5 of the FSH treatment for heifers treated with all the different FSH doses (Fig. 3A). Number of ovulatory follicles was similar for all heifers treated with different FSH doses on Days 1 to 3 of FSH treatment. During Days 4 and 5 of FSH treatment, heifers treated with the 210 IU FSH dose had a greater (P≤0.05) number of ovulatory follicles compared with heifers treated with 35 IU FSH dose (Fig. 2A). During these same days, however, heifers treated with the 70 IU, 140 IU and 210 IU FSH doses had a similar number of ovulatory follicles (Fig. 3A). In contrast to the increase in number of ovulatory follicles during Days 1 to 5 of FSH treatment, the number of ovulatory follicles decreased (P<0.0001) during Days 1 to 9 post-hCG for heifers treated with all the different FSH doses (Fig. 3B). The number of ovulatory follicles was similar each day post-hCG for heifers treated with the 70 IU, 140 IU and 210 IU FSH doses. However, heifers treated with the 210 IU FSH dose had a greater (P≤0.05) number of ovulatory follicles on most days post-hCG compared with heifers treated with the 35 IU FSH dose (Fig. 3B). The higher number of ovulatory follicles detected post-hCG for the heifers treated with the 210 IU FSH dose implies these heifers also had fewer follicles ovulate and thus a potential lower ovulation rate compared with heifers treated with the 35 IU FSH dose. Effect of FSH dose during superovulation on circulating concentrations of estradiol Coincident with the increase in AFC (Fig. 2B) and ovulatory follicle number (Fig. 3A), the circulating estradiol concentrations in heifers also increased (P<0.0001) during Days 1 to 5 34 of FSH treatment for heifers treated with all the different FSH doses (Fig. 3C). During Days 1 to 5 of FSH treatment, estradiol concentrations were similar on Days 1 to 3 for heifers treated with all the different FSH doses. Estradiol concentrations were also similar on Days 4 and 5 of FSH treatment for heifers treated with the 35 IU and 210 IU FSH doses. In contrast, heifers treated with the 210 IU FSH dose had lower (P≤0.05) estradiol concentrations on Days 4 and 5 of the FSH treatment compared with heifers treated with the 70 IU and 140 IU FSH doses (Fig. 3C). Estradiol concentrations decreased (P<0.0001) in heifers treated with all the different FSH doses after Day 1 post-hCG (Fig. 3D). However, on Day 1 post-hCG, heifers treated with the 210 IU FSH dose had higher (P≤0.05) estradiol concentrations compared with heifers treated with the 35 IU FSH dose. In contrast, heifers treated on Day 1 post-hCG with the 210 FSH dose had lower (P≤0.05) estradiol concentrations compared with heifers treated with the 140 IU FSH dose (Fig. 3D). Effect of FSH dose during superovulation on number of corpora lutea The number of CL detected in heifers by ovarian ultrasonography decreased (P<0.02) for heifers treated with all the different FSH doses during Days 1 to 5 of FSH treatment (Fig. 4A). However, number of CL during this period, was similar among the heifers treated with the different FSH doses (Fig. 4A). During Days 1 to 9 post-hCG, the number of CL increased (P<0.0001) for all heifers treated with the different FSH doses (Fig. 4D). However, the number of CL during this period was unaltered in heifers treated with the different FSH doses (Fig. 4D). Effect of FSH dose during superovulation on circulating concentrations of progesterone Concomitant with the decrease in number of CL during Days 1 to 5 of FSH treatment (Fig. 4A), circulating progesterone concentrations also decreased (P<0.009) for heifers treated 35 with the different FSH doses during this same period (Fig. 4C). This decrease in progesterone concentrations during Days 1 to 5 of FSH treatment was likely in response to the multiple PG injections on Days 4 and 5 of FSH treatment (Fig. 4A). The decrease in progesterone concentrations during Days 1 to 5 of FSH treatment, however, was unaltered by treatment of heifers with the different FSH doses (Fig. 4C). Concomitant with the increase in number of CL during Days 1 to 9 post-hCG (Fig. 4B), the circulating concentrations of progesterone also increased (P<0.0001) during the same time period. However, the increase in progesterone concentrations post-hCG was unaltered by treatment of heifers with the different FSH doses (Fig. 4D), which is likely explained by the absence of an effect of FSH doses on number of CL (Fig. 4B). Effect of FSH dose during superovulation on uterine and endometrial thickness Uterine and endometrial (data not shown) thickness was unaltered during Days 1 to 5 of FSH treatment or by the different FSH doses used to superovulate heifers (Fig. 4E). In contrast, during Days 1 to 9 post-hCG, uterine and endometrial (data not shown) thickness decreased (P<0.0001) ~35% from its peak thickness on Day 2 (Fig. 4F). During this period, however, treatment of heifers with the different FSH doses did not alter uterine and endometrial thickness (Fig. 4F). Effect of FSH dose on ovulation rate Ovulation rate was calculated by dividing the number of CL on Day 7 after the hCG (Fig. 4B) injection by number of ovulatory follicles (≥10 mm) present at time of the hCG injection (Fig. 3A). Ovulation was not different between the four groups examined individually. However, ovulation rates for heifers treated with the two highest FSH doses combined were lower (P≤0.001) compared with heifers treated with the two lowest FSH doses combined (Fig. 5). 36 Discussion This is the first study to address the consequences of different FSH doses on ovarian function within individual animals with comparable ovarian reserves in any mammalian species. The most compelling findings of this study are that higher doses of FSH during superovulation of healthy, nulliparous, young adult Holstein heifers with small ovarian reserves: 1) do not result in a dose response effect for any endpoints of ovulatory follicle function measured, 2) are excessive (lower doses were just as effective) and do not improve follicular growth and response to superovulation, 3) potentially detrimental to ovulatory follicle function, and 4) do not alter uterine or endometrial thickness. FSH has a well-established role in regulating follicular function. FSH action during superovulation is mediated by FSH receptors located exclusively on granulosa and cumulus cells [47]. In addition, the interaction of FSH with its receptors has a critical role in regulation of steroidogenesis, especially estradiol and progesterone production, cumulus cell mass expansion and, cell metabolism, all of which contribute to oocyte competence in ovulatory follicles [47]. Despite the well-established role of FSH in regulating follicular function, results of the present study did not demonstrate a positive FSH dose-response effect on any of the endpoints measured. In fact, the highest FSH doses (140 and 210 IU FSH per injection) did not enhance circulating AMH concentrations (marker for growth of healthy preantral and small antral follicles, [10, 48]), AFC ( ≥3 to <10 mm), ovulatory follicle ( ≥10 mm) numbers, circulating estradiol or progesterone concentrations, CL number or ovulation rate compared with the 70 IU FSH dose. This demonstrates that the highest FSH doses used to superovulate heifers with a low AFC were both excessive and economically wasteful. Interestingly, these high FSH doses did not reduce the number of growing follicles, which may explain why there were no carryover effects, including 37 the impact of the different FSH dose sequences, or of superovulation regimen on the subsequent regimen, for any endpoint measured in the present study. The reason for the absence of a positive FSH dose-response effect on ovulatory follicle number and function is unknown. However, heifers with a low AFC and small ovarian reserve, as used in the present study, do not inherently respond as well to superovulation as heifers with a higher AFC [24, 26]. It is well-established that women with a small ovarian reserve respond poorly to ovarian stimulation protocols during ART [3, 49]. In addition, our previous FSH dose response study used granulosa cells isolated from antral follicles of cows with low or a high AFC [50]. The size of antral follicles in this in vitro study were similar to sizes of the follicles at initiation of superovulation of heifers here. Results of the in vitro study demonstrate that FSH action on granulosa cell function is biphasic resulting in both a positive and negative window for responsiveness to FSH action. For example, the lower FSH doses showed a positive relationship with estradiol secretion, while the highest doses decreased estradiol secretion. Furthermore, the physiological window of positive responsiveness of granulosa cells to FSH action (that is, before onset of premature luteinization and loss of estradiol producing capacity) was positively associated with AFC and size of the ovarian reserve [50]. Thus, peak response of granulosa cells to FSH action occurs at much lower FSH doses for cattle with a low compared to high AFC [50]. These in vitro observations imply that a much narrower FSH dose range than the one used in the present study will likely be necessary to achieve a positive FSH dose-response on the endpoints measured. The potential relationship between the AFC and the window for the positive effects of FSH action on granulosa cells during superovulation may also explain the conflicting results among previous FSH dose response studies [11-14, 28-32] in cattle with an unknown AFC. For 38 example, if the different FSH doses used to superovulate cattle did not span the putative physiological windows for both positive and negative effects of FSH action on granulosa cells [50], results would be expected to produce either positive, negative or no effects on endpoints measured. We also show here that high FSH doses not only fail to improve response to superovulation, they are also potentially detrimental to ovulatory follicle function. Estradiol is a well-established marker for ovulatory follicle function in cattle [51] and primarily produced by dominant ovulatory follicles [52, 53]. Thus, the results of the present study implied that the highest dose of FSH used to superovulate heifers impaired ovulatory follicle function. In further support of the potential detrimental impact of the high FSH doses, the capacity of the low estradiol-producing ovulatory follicles to ovulate in response to an hCG injection was also reduced in the present study. For example, even though the number of ovulatory follicles was similar, there was a tendency for number of CL to be lower for heifers treated with the highest compared with the two intermediate FSH doses. In addition, ovulation rate was lower for heifers treated with the two highest compared with the two lowest FSH doses in the present study. These combined observations illustrate that superovulation of small ovarian reserve heifers with the highest FSH doses reduced the capacity of ovulatory follicles to produce estradiol, ovulate and form CL in response to hCG. This finding is unlikely restricted to the Holstein heifers with a low AFC and small ovarian reserve since other FSH dose response studies using different breeds and ages of cattle with unknown AFC also show similar results [16, 29]. These observations raise the question of whether the quality of oocytes recovered from ovulatory follicles subjected to high FSH doses is also compromised, which could explain at least part of the high oocyte wastage during ART in 39 cattle [11-17] and women [3, 5, 6, 18-22]. Taken together, these observations indicate that high FSH doses are not only excessive and economically wasteful, but also detrimental to ovulatory follicle function. Many studies have used doses of Folltropin-V higher than the industry standard of 70 IU per injection to improve the superovulation response or reduce number of times cattle are injected during superovulation to defray costs associated with embryo transfer [40, 54-56]. However, the two highest FSH doses (140 IU, 210 IU) per injection in the present study are unlikely to be used commercially to superovulate cattle. In contrast, very high FSH doses, similar to those in the present study, are likely used during ART cycles in some women. For example, although there was a 6-fold range in total FSH doses used to superovulate heifers (280 IU to 1680 IU) in the present study, the total FSH doses during ART cycles in women have a 20- fold range (<1,000 IU to 20,000 IU) [3]. However, the same FSH standard was not used to determine IU for the FSH preparations used in cattle and women. Thus, a direct comparison between the FSH doses during superovulation of heifers in this study and ART cycles in women cannot be established. Nevertheless, relatively high total FSH doses decrease oocyte and embryo quality and yield during embryo transfer in cattle [11, 13, 14] albeit this finding is controversial [28-32]. In addition, high FSH doses are positively correlated with oocyte and embryo wastage in cattle [11-17] and women [3, 5, 6, 18-22] and with the lowest live birth rates during ART cycles in women [3]. Whether decreased ovulatory follicle function following high FSH dose ovarian stimulation protocols, as observed in the present study for the small ovarian reserve heifers, also impairs oocyte quality remains to be established. High FSH and LH doses uncouple gonadotropin receptors from their respective signaling system in granulosa, theca and luteal cells in animal models and humans, altering ovarian 40 function [57-59]. Whether the loss of the capacity of ovulatory follicles to produce estradiol and ovulate is directly attributable to the highest FSH doses used to superovulate heifers could not be unequivocally established here. However, LH stimulates granulosa and luteal cells to produce progesterone [60, 61] and ovulatory follicles to undergo luteinization and produce progesterone during a preovulatory LH surge [60, 61]. Because Folltropin-V is a pituitary source of porcine FSH (pFSH) it contains minor (0.25%) amounts of porcine pituitary LH (pLH). Thus, it is possible that the LH contamination in the highest doses of Folltropin-V caused or contributed to the diminished estradiol-producing capacity of granulosa cells in ovulatory follicles in the present study. Arguing against such an effect, the circulating progesterone concentrations remained low and decreased, and were unaltered by different doses of Folltropin-V during the FSH treatment period. Consequently, the minor LH contamination even in the highest dose of Folltropin-V appears insufficient to modify granulosa or luteal cell function. A more plausible explanation for why the highest Folltropin-V doses caused the loss of capacity of ovulatory follicles to produce estradiol and ovulate during superovulation in the present study is that the high Folltropin-V doses triggered premature luteinization and diminished the capacity of granulosa cells in ovulatory follicles to produce estradiol, as observed in our previous in vitro study [50]. Endometrial thickness increases during the follicular phase and declines during the luteal phase of reproductive cycles despite increasing circulating progesterone concentrations in cattle [34] and women [62]. This is similar to the patterns shown in the present study. However, FSH dose and the corresponding changes in ovarian steroid hormone production did not alter uterine or endometrial thickness. Nevertheless, the pattern of change in endometrial thickness in the superovulated low AFC heifers was remarkably similar to changes we previously observed in 41 unstimulated heifers with a high rather than low AFC [34]. The reason for this difference in patterns between studies is unknown, but may be explained by the greater circulating progesterone concentrations in superovulated low AFC heifers in the present study compared untreated low AFC heifers in our previous study [34]. Although controversial [63, 64], greater endometrial thickness is linked positively to embryo survival in women [65, 66]. Consequently, progesterone therapy is often used during ART in women to enhance luteal progesterone concentrations [67-69] and to improve outcomes but these results are also mixed [70, 71]. Results of the present study in small ovarian reserve heifers, however, imply that high FSH doses during ART in women are unlikely to diminish endometrial thickness and in turn potentially contribute to poor embryo survival. The results of the present study combined with results of others in cattle [11, 12, 14-16, 28-32] and women [3, 5, 6, 18-22] support the conclusion that superovulation of small ovarian reserve cattle with excessive FSH doses is costly and potentially detrimental to ovulatory follicle function. While the detrimental effects of high FSH doses on ovulatory function was established in small ovarian reserve in cattle the present study, it remains to be determined if excessive FSH doses also impair oocyte quality and reproductive outcome. Such an effect could help explain poor embryo yield and quality in cattle [11-15] and low live birth rates during ART in women [3]. 42 1. 2. 3. 4. 5. 6. 7. 8. 9. REFERENCES Mapletoft RJ, Hasler JF. 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The role of progesterone therapy in early pregnancy: from physiological role to therapeutic utility. Gynecol Endocrinol 2017; 33:421-424. 48 APPENDIX Figure 2. Effect of different doses of FSH on circulating concentrations of anti-Mϋllerian hormone (AMH) and antral follicle count (AFC) for heifers with a small ovarian reserve. Beginning on Day 1 of the estrous cycle near the beginning of the first follicular wave, heifers were superovulated with twice daily injections of each of four different doses (35 IU ● , 70 IU □ , 140 IU ○ , 210 IU ♦) of FSH (Folltropin-V) for four days (depicted by arrows) using a Williams Latin Square Design as explained in Methods. Thus, each heifer was superovulated a total of four times with 21 to 24 d between superovulation regimens. Symbols represent means (±SEM) for the same 8 heifers. As depicted by arrows, three prostaglandin F2α (PG) injections were given 12-h apart starting on Day 4 of FSH treatment to regress the corpus luteum. A single human chorionic gonadotropin (hCG) injection (arrow) was given 12 h after the last FSH injection on Day 5 of treatment to induce ovulation. AMH concentrations were determined every 24 h throughout Folltropin-V treatment but not post hCG. Number of antral follicles ≥ 3 mm to <10 mm in diameter (Antral Follicle Count, AFC) was determined daily by ovarian ultrasonography until 3 d after the hCG injection when AFC was determined every other day until the end of the study. As depicted within each figure, results of the Type III ANOVA indicate whether significant (P≤0.05) differences existed in overall AMH concentrations and AFC in heifers during the different days of treatment (Day) and between the FSH doses (Dose), and between heifers treated with the different 49 Figure 2 (cont’d) FSH doses for any day during the FSH treatment period (Day*Dose). Asterisks indicate that means differ (* = P<0.05, ** = P<0.01) when compared with the 210 IU FSH dose. 50 Figure 3. Effect of different doses of FSH on ovulatory follicle number and estradiol concentrations for heifers with a small ovarian reserve. Heifers were superovulated beginning on Day 1 of the estrous cycle with four different doses (35 IU ● , 70 IU □ , 140 IU ○ , 210 IU ♦) of FSH (Folltropin-V, arrows depict injections) and treated with PG and hCG (depicted by arrows) as explained in the legend for Fig. 1. Symbols depict means (±SEM) for the same 8 heifers. The number of ovulatory size follicles (≥10mm) were determined by daily ultrasonography measurements and then every other day starting 3 d after the hCG injection. Estradiol concentrations were determined at 24-h intervals during FSH treatments and at 48-h intervals post hCG. As depicted within each figure, results of the Type III ANOVA indicate whether significant (P≤0.05) differences exist in overall number of ovulatory follicles and circulating estradiol concentrations in heifers during the different days of treatment (Day) and between the FSH doses (Dose), and between heifers treated with the different FSH doses for any day during the treatment period (Day*Dose). Asterisks indicate that means differ (* = P<0.05, ** = P<0.01) when compared with the 210 IU FSH dose. 51 Figure 4. Effect of different doses of FSH on corpora lutea number, progesterone concentrations and uterine thickness for heifers with a small ovarian reserve. Heifers were superovulated beginning on Day 1 of the estrous cycle with four different doses (35 IU ● , 70 IU □ , 140 IU ○ , 210 IU ♦) of FSH (Folltropin-V, arrows depict injections) and treated with PG and hCG (depicted by arrows) as explained in the legend for Fig. 1. Symbols depict means (±SEM) for the same 8 heifers. Number of corpora lutea and uterine thickness were determined by daily ultrasonography measurements and then every other day starting 3 d after the hCG injection. Progesterone concentrations were determined every 24 h during FSH treatment and every 48 h after the hCG injection. As shown within each figure, results of the Type III ANOVA indicate whether significant (P≤0.05) differences exist in overall number of corpora lutea, progesterone 52 Figure 4 (cont’d) concentrations and uterine thickness for heifers during the different days of treatment (Day) and between the FSH doses (Dose), and between heifers treated with the different FSH doses for any day during the treatment period (Day*Dose). Asterisks indicate that means differ (* = P<0.05, ** = P<0.01) when compared with the 210 IU FSH dose. 53 Figure 5. Effect of different doses of FSH on ovulation rate for heifers with a small ovarian reserve. Heifers were superovulated beginning on Day 1 of the estrous cycle with four different doses (35 IU, 70 IU, 140 IU, 210 IU) of FSH (Folltropin-V, arrows depict injections) and treated with PG and hCG (depicted by arrows) as explained in the legend for Fig. 1. Bars depict means (±SEM) for the same 8 heifers. Ovulation rate was calculated by dividing the number of corpora lutea on Day 7 post-hCG (Fig. 3B) by number of ovulatory follicles present at time of hCG (Fig. 2A). As depicted within the figure, results of the Type III ANOVA indicated that a significant (Dose = P=0.03) difference existed in ovulation rate for heifers treated with the different doses of FSH. Asterisks indicate that the pooled mean (±SEM) for ovulation rates for the 140 IU and 210 IU doses was lower (P<0.001) compared with the pooled mean (±SEM) for the 35 IU and 70 IU doses. 54 Excessive Follicle-Stimulating Hormone During Ovarian Stimulation of Cattle May Induce Premature Luteinization of Most Ovulatory-Size Follicles1 CHAPTER 3. 1 Clark, Zaramasina L., Kaitlin R. Karl, Meghan L. Ruebel, Keith E. Latham, and James J. Ireland. "Excessive follicle-stimulating hormone during ovarian stimulation of cattle may induce premature luteinization of most ovulatory-size follicles." Biology of Reproduction, 106(5), (2022): 968-978. 55 Title: Excessive FSH during ovarian stimulation of cattle induces premature luteinization of most ovulatory-size follicles Running Title: Excessive FSH induces premature luteinization Summary sentence: Excessive doses of Folltropin-V (FSH) during ovarian stimulation of heifers with a small ovarian reserve reduce intrafollicular estradiol while inducing premature luteinization of the majority of ovulatory-size follicles. Keywords: ovarian stimulation, small ovarian reserve, ovulatory-size follicles, estradiol, progesterone, oxytocin, cumulus cell expansion, premature luteinization Authors and affiliations: Zaramasina L. Clark1, Kaitlin R. Karl1, Meghan L. Ruebel1, Keith E. Latham1, James J. Ireland1 1Reproductive and Developmental Sciences Program, Department of Animal Science, Michigan State University, East Lansing, MI 48824. Grant support1 Correspondence2 Present address 3 1 This study was supported by the Agriculture and Food Research Initiative Competitive USDA-NIH Dual Purpose Program Grant no. 2017-67015-26084 from the USDA National Institute of Food and Agriculture, the Eunice Kennedy Shriver National Institute of Child Health & Human Development of the National Institutes of Health under Award Number T32HD087166, and AgBioResearch at Michigan State University. 2 James J. Ireland, Molecular Reproductive Endocrinology Laboratory, Department of Animal Science, Michigan State University, East Lansing, Michigan, 48824, USA. Tel.+1 517 432 1384. Email: ireland@msu.edu 3 Zaramasina L. Clark. Present address: School of Biological Sciences, Victoria University of Wellington, Wellington, New Zealand. Meghan L. Ruebel. Present address: Anschutz Medical Campus, University of Colorado, Denver, CO, USA. 56 Abstract High FSH doses during ovarian stimulation are detrimental to ovulatory follicle function and decrease live birth rate. However, the mechanism whereby excessive FSH causes ovarian dysfunction is unknown. This study tested the hypothesis that excessive FSH doses during ovarian stimulation induce premature luteinization of ovulatory-size follicles. Small ovarian reserve bovine heifers were injected twice daily for four days with 70 IU (industry standard dose; N = 7 heifers) or 210 IU (excessive dose, N = 6 heifers) Folltropin-V (FSH), at the emergence of a new follicular wave. Ovulatory-size (≥10 mm) follicles were excised from ovaries 12 h after the last FSH injection (no hCG injected) and hormone concentrations in follicular fluid (FF) were determined using ELISA. Luteinization was monitored by assessing cumulus cell-oocyte complex (COC) morphology and measuring concentrations of estradiol (E), progesterone (P) and oxytocin (O) in FF. For a subset of follicles, COC were classified as having compact (cCOC) or expanded (eCOC) cumulus cell layers, and as estrogen-active (EA, E:P in FF ≥1), estrogen- inactive (EI, E:P in FF ≤1>0.1) or extreme-estrogen-inactive (EEI, E:P in FF ≤0.1). A high proportion (72%) of ovulatory-size follicles in 210 IU, but not 70 IU, dose heifers displayed eCOC. The high dose also produced higher proportions of EI or EEI follicles which had lower E:P ratio and/or E but higher P and/or O concentrations compared with the 70 IU dose heifers. Excessive FSH doses during ovarian stimulation are concluded to induce premature luteinization of most ovulatory-size follicles in heifers with small ovarian reserves. 57 Introduction The use of high gonadotropin doses in women undergoing assisted reproductive technology (ART) causes high oocyte and embryo wastage and decreases live birth rate [1, 2]. The mechanisms whereby high follicle-stimulating hormone (FSH) doses impair ovulatory follicle function are poorly understood. Ovarian follicle development becomes progressively reliant and ultimately dependent on gonadotropin (FSH, and luteinizing hormone, LH) signaling to enhance growth and function. In cattle, the transition of follicles to gonadotropin dependence occurs in 4 mm antral follicles and continued follicle growth and maturation are dependent on adequate support from circulating FSH [3]. FSH has a range of functions within the follicle and interacts with many different signaling pathways that contribute to ovulatory follicle function, and oocyte development and quality. These include steroid hormone synthesis, cell proliferation and survival, induction of LH receptor expression and FSH receptor downregulation [4]. Given the broad, complex intrafollicular effects of FSH of follicular function, the effects of exogenous FSH on ovulatory follicle function likely vary with dosage. For example, estradiol secretion by bovine granulosa cells in response to different FSH doses is curvilinear; estradiol production initially increases in a dose-dependent manner but the highest FSH doses decrease estradiol production [5]. This supports our recent observations of decreased estradiol production by ovulatory-size follicles during ovarian stimulation of cattle with high FSH doses [6]. Taken together, these observations imply that high FSH doses during ovarian stimulation may be detrimental to granulosa cell function in ovulatory follicles, although the mechanism and potential impact of such an effect on ovulatory follicle function and oocyte quality are unclear. 58 Ovarian stimulation is a method of ART designed to provide continued FSH support by exogenous administration of FSH to promote development of the gonadotropin-dependent follicles to ovulatory-size. Subsequent development and completion of final maturation of ovulatory-size follicles in preparation for ovulation is dependent on the ability of these follicles to respond to LH [7, 8]. Luteinization is the process, regulated by LH, by which the ovulatory follicle is remodeled following ovulation to form the corpus luteum and concomitantly switch from primarily estradiol to progesterone production [9]. Premature luteinization is a term applied to clinical observations of high circulating progesterone concentrations or reduced estradiol:progesterone (E:P) ratio on the day of the ovulatory stimulus in women [10-20]. Premature luteinization impacts 12.3 to 46.7% of ART cycles in women contributing to poor ART outcomes [21]. There is evidence that premature luteinization can be induced by high doses of FSH administered during ovarian stimulation in patients undergoing ART [18, 22] and ovarian stimulation in animal models [23]. However, the impact of the altered circulating endocrine profile on ART outcomes is contentious and very few, if any, of the studies in women have directly evaluated the ovulatory-size follicles developing in response to ovarian stimulation, particularly with excessive FSH doses (i.e., doses beyond what is needed to achieve a maximum ovulatory response). Importantly, we observed that high FSH doses decrease estradiol production in vitro [5] and during superovulation of cattle with a small ovarian reserve [6]. It is unknown to what extent excessive FSH doses during ART cause premature luteinization of ovulatory follicles, thereby compromising oocyte quality and ART outcomes. The present study, therefore, tests the hypothesis that excessive FSH doses during ovarian stimulation can induce premature luteinization of ovulatory-size follicles. We employ the small 59 ovarian reserve heifer model validated in our laboratory because it shares numerous phenotypic characteristics of women with small ovarian reserves including, a diminished ovarian reserve and low antral follicle count (AFC) [24], low circulating concentrations of AMH [25], heightened FSH secretion [24] and reduced responsiveness to ovarian stimulation [26]. The present study measured alterations in well-established intrafollicular markers of luteinization of ovulatory follicles in cattle including cumulus cell expansion [27] and intrafollicular concentrations of estradiol [28-30], progesterone [28-30] and oxytocin [31] in individual ovulatory-size follicles developing in response to ovarian stimulation with either the industry standard (70 IU) or excessive (210 IU) FSH doses [6]. We provide evidence that in the absence of an LH or LH-like stimulus (e.g., hCG), excessive FSH doses induce premature luteinization of a high proportion of the ovulatory-size follicles developing in response to ovarian stimulation. Premature luteinization may also impair oocyte quality, enhance oocyte wastage, and contribute to poor ART outcomes in cattle with small ovarian reserves. These results also indicate a possible need for caution in selecting high gonadotropin doses for human ART. Materials and Methods Selection of heifers with a low AFC and small ovarian reserve, ovarian stimulation protocol, excision of ovulatory-size follicles from ovaries, and recovery of follicular fluid (FF) and cumulus cell-oocyte complexes (COC) from ovulatory-size follicles Selection of 14 heifers aged 11 to 12 months with a small ovarian reserve and ovarian stimulation procedures were performed as described [6]. Briefly, the number of antral follicles ≥3 mm in diameter (AFC) in two groups of 50 Holstein heifers (Green Meadow Farms Inc., Ovid-Elsie, MI) were determined using ovarian ultrasonography. Animals were ranked according to AFC and luteolysis was induced in the 25 heifers with the lowest AFC in each group using 60 two prostaglandin F2α (PG; 12.5 mg/mL, Lutylase HighCon, Zoetis) injections administered 10 days apart. AFC for each heifer was determined prior to each PG injection and 4 days after the final PG injection. Individuals determined to consistently have an AFC ≤15 (N = 14 heifers; mean ±SEM AFC = 6.8±0.4) on these days were selected for this study. We have previously shown that age-matched heifers with an AFC ≤15 also have an ovarian reserve (total number of morphologically healthy follicles/oocytes in ovaries) 80% smaller than heifers with an AFC ≥ 25 [25]. Heifers in the present study were housed at the Michigan State University Beef Cattle Teaching and Research Center. All procedures described herein were sanctioned by the Institutional Animal Care and Use Committee at Michigan State University. Heifers were injected with 70 IU (40 mg, N = 7 heifers) or 210 IU (120 mg, N = 7) doses of Folltropin-V (Vetoquinol USA Inc). Folltropin-V is an extract of porcine pituitary glands that contains 700 IU (equivalent to 400 mg NIH-FSH-P1 with 0.25% LH contamination or <1 mg NIH-LH-S19) of FSH per 20 mL vial. However, one heifer treated with the 210 IU dose was excluded from the final analysis due to the absence of a response to ovarian stimulation. Hereafter, because of the minimal amount of LH contamination, Folltropin-V is referred to as FSH. The 70 IU and 210 IU FSH doses were chosen as the industry standard and excessive doses, respectively, based on our previous study [6]. The 210 IU FSH dose compared with the 70 IU dose is excessive because it does not increase the number of ovulatory-sized follicles but decreased circulating estradiol concentration and ovulation rate following ovarian stimulation [6]. To synchronize estrous cycles and the day ovarian stimulation began, heifers were administered PG injections 10 days apart followed by a third PG injection 12 h later. On Day 1-2 of the estrous cycle, which is near the time of ovulation and emergence of the first follicular 61 wave (~36 h after last PG injection), animals were randomly assigned to treatment with either 70 IU or 210 IU FSH twice daily for 4 days. Antral follicle size and number were monitored by daily ovarian ultrasonography as previously explained [6]. As depicted in Table 1, AFC prior to ovarian stimulation did not differ between FSH doses. Ovaries were collected from a single heifer per day alternating between FSH dose groups until the study was completed. Heifers were euthanized 12 h after the final FSH injection and both ovaries recovered. Note that heifers were not injected with LH or an LH-like stimulus (for example, human chorionic gonadotropin; hCG) during the ovarian stimulation regimen. Time from euthanasia of heifers in our study to collection of ovaries was 33±3 minutes. In agreement with our previous study [6], the number and diameter of ovulatory-size follicles did not differ between FSH doses (Table 1). Ovaries were rinsed with 70% ethanol, washed in Dulbecco’s Phosphate Buffered Saline (DPBS), and placed in fresh DPBS for excision of ovulatory-size follicles. FF was aspirated from all ovulatory-size follicles (≥10 mm). For a subset of follicles, the FF was immediately transferred to a separate dish and searched using a dissecting microscope to find the COC. Between 5 and 15 COCs were recovered from FF of ovulatory-size follicles per heifer. Each COC was classified as having compact or expanded layers of cumulus cells as depicted in the representative images in Fig. 6. Hereafter, COC with compact layers of cumulus cells are referred to as cCOC whereas COC with expanded layers of cumulus cells are referred to as eCOC. After the COC was removed from FF, the residual FF was recovered, and the volume measured. A protease inhibitor cocktail (cOmplete™, EDTA-free Protease Inhibitor Cocktail, Roche, USA) was added to each FF sample at a 1:10 dilution to minimize protein degradation. The FF samples were centrifuged to pellet cellular debris and 200 µL aliquots of the supernatant stored at -80 °C. Thereafter, each remaining ovulatory-size follicle on both ovaries was aspirated and the 62 FF processed as described above but without recovery of the COC. The time elapsed from removal of ovaries, excision of ovulatory-size follicles from ovaries, aspiration of FF, and recovery and classification of COCs for each heifer ranged from 2 to 5 hours. Estradiol (E), progesterone (P) and oxytocin (O) assays Concentrations of E and P were measured in FF of all ovulatory-size follicles regardless of whether a COC was recovered, whereas O was measured in a subset of ovulatory-size follicles based on COC morphology and ratio of E:P in FF as explained in Results. Estradiol and P concentrations were measured in the same aliquot of FF from each ovulatory-size follicle using commercially available ELISA kits (Cayman Chemical Company, MI, USA). The inter- and intra-assay coefficients of variation for the E assays were 9.3% and 7.0%, respectively, and 11.0% and 9.9%, respectively for the P assays. Oxytocin concentrations in FF were determined using the Oxytocin ELISA kit (ENZO Life Sciences, Inc). Serial dilutions of bovine serum and FF from small (3-5 mm), medium (8-10 mm) and large (12-15 mm) bovine follicles were evaluated against the standard curve to confirm parallelism. The inter- and intra-assay coefficients of variation were 7.5% and 7.9%, respectively. Classification of ovulatory-size follicles based on ratio of E:P in FF Cattle typically have one or two non-ovulatory follicular waves and one ovulatory follicular wave during a 21-day estrous cycle [32]. We previously established that cattle have both estrogen-active (EA) and estrogen-inactive (EI) follicles during a follicular wave [28-30, 33]. EA follicles are healthy, growing, dominant, non-ovulatory or ovulatory follicles during follicular waves, whereas EI follicles are either dominant ovulatory follicles undergoing luteinization in response to the preovulatory LH surge or dominant non-ovulatory follicles destined for atresia at the end of a follicular wave [28-30, 33]. In the present study, ovulatory- 63 size follicles were not only classified based on whether the COC had compact or expanded layers of cumulus cells, but also based on the ratio of E:P concentrations in FF as follows: estrogen- active (EA, E:P in FF ≥1), estrogen-inactive (EI, E:P in FF ≤1>0.1) or extreme-estrogen-inactive (EEI, E:P in FF ≤0.1). Statistical analyses All statistical analyses were performed using the Statistical Analysis System (SAS 9.4) software [34]. Data were transformed before analyses if not normally distributed. Specifically, data for E, P and O concentrations were log transformed, whilst data for E:P ratios were arcsine transformed. When overall treatment effects were significant (P≤0.05), Fisher LSD [34] was used to determine if means differed (P≤0.05). Note that degrees of freedom during statistical analyses were based on the number of heifers (N = 7 heifers treated with 70 IU FSH dose or 6 heifers treated with 210 IU FSH dose), not the number of ovulatory-size follicles. To assess whether the excessive, 210 IU, vs industry-standard, 70 IU, FSH dose during ovarian stimulation induced premature luteinization of the ovulatory-size follicles, five separate analyses were performed on the present data set. Firstly, Fisher’s exact test was used to determine if the proportion of ovulatory-size follicles with compact- or eCOC differed (P≤0.05) between heifers undergoing ovarian stimulation with the 70 IU vs 210 IU FSH dose. Secondly, given that the industry standard dose did not yield any eCOC, two t-tests were performed. The first t-test evaluated the effect of dose by comparing the concentrations of E, P, O and E:P in compact follicles from the 70 IU vs 210 IU FSH dose. The second t-test evaluated the impact of COC expansion, unique to the 210 IU dose group, by comparing the concentrations of E, P, O and E:P in the cCOC 210 IU follicles to those in the eCOC 210 IU follicles. Thirdly, ANOVA was used to determine if concentrations of E, P, O and ratio of E:P in FF of the ovulatory-size 64 follicles differed (P≤0.05) between heifers treated with the 70 IU vs 210 IU FSH dose. Fourthly, chi-square analysis was used to determine if proportion of ovulatory-size follicles classified as EA, EI and EEI differed (P≤0.05) between heifers undergoing ovarian stimulation with the 70 IU vs 210 IU FSH dose. Fifthly, ANOVA was used to determine if concentrations of E, P, O and E:P in FF of ovulatory-size follicles with cCOC or eCOC differed (P≤0.05) between follicles classified as EA, EI or EEI for the heifers undergoing ovarian stimulation with 70 IU vs 210 IU FSH dose. Results Analysis 1: Effect of FSH dose during ovarian stimulation on proportion of ovulatory-size follicles in heifers with cCOC or eCOC Only cCOC were observed in ovulatory-size follicles in heifers undergoing ovarian stimulation with the 70 IU FSH dose whereas ovulatory-size follicles in heifers treated with the 210 IU FSH dose had both cCOC and eCOC. As shown in Table 2, the heifers undergoing ovarian stimulation with 210 IU vs 70 IU FSH dose had a lower (P<0.001) proportion of ovulatory-size follicles with cCOC (28±3% vs 100±0%, mean ± SEM) but a higher (P<0.001) proportion of ovulatory-size follicles with eCOC (72±3% vs 0%). Analysis 2: Effect of COC morphology (cCOC vs eCOC) on concentrations of E, P, O and E:P in FF of ovulatory-size follicles for heifers undergoing ovarian stimulation with the 70 IU or 210 IU FSH dose This analysis was designed to determine if concentrations of E, P, or O or E:P in FF differed between the ovulatory-size follicles with cCOC vs eCOC for the heifers undergoing ovarian stimulation with the 70 IU or 210 IU FSH dose. Ovulatory-size follicles with cCOC had similar E, P, and O concentrations and ratio of E:P, regardless of FSH dose (Fig. 7). However, 65 for heifers undergoing ovarian stimulation with the 210 IU FSH dose, the ovulatory-size follicles with eCOC not only had a decreased E and E:P ratio but also increased P and O concentrations in FF compared with the ovulatory-size follicles with cCOC (Fig. 7). Analysis 3: Overall effect of FSH dose during ovarian stimulation of heifers on concentrations of E, P, and O and E:P in FF Estradiol concentrations and E:P in FF did not differ between heifers treated with the 70 or 210 IU FSH doses (Fig. 8). In contrast, P and O concentrations were 7- and 6-fold higher, respectively, in heifers undergoing ovarian stimulation with the 210 IU vs 70 IU FSH dose (Fig. 8). Analysis 4: Effect of FSH dose during ovarian stimulation on diameter and proportion of ovulatory-size follicles in heifers classified as EA, EI and EEI We examined whether FSH dose impacted the diameter and proportion of ovulatory-size follicles classified as EA vs EI or EEI. As depicted in Table 2, the ovulatory-size follicles for heifers stimulated with the 70 IU FSH dose were classified as EA (79±6%) or EI (21±6%) whereas the ovulatory-size follicles for heifers stimulated with the 210 IU FSH dose were classified as EA (54±9 %), EI (24±8%) or EEI (22±3%). Diameters were similar among the EA, EI and EEI ovulatory-size follicles regardless of FSH dose or COC morphology (Table 2). However, the heifers stimulated with 210 IU vs 70 IU FSH dose had a lower (P<0.001) proportion of ovulatory-size follicles classified as EA (54±7% vs 79±6%), a similar proportion classified as EI (24±8% vs 21±6%) and a higher (P<0.001) proportion classified as EEI (22±3% vs 0%). 66 Analysis 5: Effect of COC morphology (cCOC vs eCOC) on concentrations of E, P, O and E:P in FF of ovulatory-size follicles classified as EA, EI or EEI in the heifers undergoing ovarian stimulation with different FSH doses Dominant follicles classified as EA [28, 30] and containing cCOC [35] are representative of ovulatory follicles in cattle. Thus, the E, P and O concentrations and E:P in FF in the EA ovulatory-size follicles with cCOC in the heifers stimulated with the industry-standard, 70 IU, FSH dose were considered the control values to use to determine if FSH dose, COC morphology or the E:P follicle classification scheme altered the E, P, or O concentrations or E:P in FF. In comparison with the control (70-C-EA), the E:P ratio and E concentrations were lower in the EI ovulatory-size follicles with cCOC for heifers stimulated with the 70 IU FSH (70-C-EI) dose and in the EI (210-E-EI) and EEI (210-E-EEI) ovulatory-size follicles which contained eCOC in the heifers stimulated with the 210 IU FSH dose (Table 2). In addition, compared with the control (70-C-EA), P in FF was higher in the 210-E-EI and 210-E-EEI ovulatory-size follicles, while O was only higher in the 210-E-EEI ovulatory-size follicles, both of which were from heifers stimulated with the 210 IU FSH dose and contained eCOC (Table 2). Discussion The present results provide direct evidence in the small ovarian reserve heifer model that in the absence of a preovulatory LH or LH-like (e.g., hCG) stimulus, most ovulatory-size follicles developing in response to ovarian stimulation with an excessive FSH dose (3-fold greater than industry-standard) have: i) expanded layers of cumulus cells, ii) decreased capacity to produce E but enhanced capacity to produce P resulting in a decreased ratio of E:P in FF, and/or iii) enhanced capacity to produce oxytocin. Because these phenotypic changes occur in ovulatory follicles in response to the preovulatory gonadotropin surge and in dominant non- 67 ovulatory follicles destined for atresia at the end of a follicular wave in cattle [27-31, 36], they are hereafter collectively referred to as premature luteinization of ovulatory-size follicles. The most compelling data in the present study supporting premature luteinization of ovulatory-size follicles was the overt observation that most (72±3%) ovulatory-size follicles in heifers undergoing ovarian stimulation with the excessive FSH dose contained eCOC. In contrast, all the ovulatory-size follicles in heifers undergoing ovarian stimulation with the industry-standard FSH dose contained a cCOC. This was an unexpected finding because cumulus cell expansion is one of the key events initiated by the preovulatory gonadotropin surge in cattle [27, 36] and other species including women [37, 38]. However, the ovaries in the present study were recovered without the administration of an ovulatory stimulus (hCG). This observation of prematurely expanded COC may be explained for example by availability of sufficient LH during ovarian stimulation, either due to endogenous LH secretion or the presence of LH-contamination in the Folltropin-V preparation, to stimulate cumulus cell expansion [38]. However, several studies show that ovarian stimulation protocols utilizing FSH (including Folltropin-V) reduce LH pulse frequency and mean peripheral LH concentrations [39-41]. Additionally, we observed no instances of ovulation that would be expected with an endogenous LH surge or even enhanced endogenous LH secretion. Moreover, whilst Folltropin-V contains very minor LH contamination, we previously demonstrated in the same small ovarian reserve heifer model, that there were no increases in circulating estradiol or progesterone concentrations during ovarian stimulation [6]. This was true regardless of the fact that four different Folltropin- V doses, including the industry-standard and excessive FSH doses utilized herein, were evaluated [6]. This suggests that the minimal LH contamination in Folltropin-V was unlikely sufficient to modify granulosa, cumulus or theca cell function in the heifers stimulated with the 68 excessive FSH dose in the present study [6]. These considerations indicate that premature cumulus cell expansion is unlikely to be a response to endogenous or exogenous LH signalling. The ovarian stimulation protocol used here, coupled with hormonal measurements and classification of individual ovulatory-size follicles based on COC morphology and ratio of E:P in FF, provides new insights into the impact of excessive FSH doses on the development of ovulatory-size follicles. For instance, the FSH injections in heifers were initiated at the emergence of the first follicular wave (Day 1 of the estrous cycle) when about a dozen 3 to 4 mm antral follicles were present. These FSH injections continued for 3 additional days resulting in development of ~13 to 19 ovulatory-size (≥10 mm) during the ovarian stimulation period. However, the excessive, compared with industry-standard, FSH dose did not increase the number of ovulatory-size follicles, as observed previously using the same heifer model [6]. Instead, it increased the heterogeneity of the ovulatory-size follicles, which were separated into four distinct phenotypes. The ovulatory-size follicles from heifers undergoing ovarian stimulation with the industry-standard FSH dose had mostly (79±6%) EA ovulatory-size follicles with cCOC. These are the hormonal and morphological characteristics of normal ovulatory follicles in cattle [28, 30, 33]. In contrast, the heifers undergoing ovarian stimulation with excessive FSH doses had ovulatory-size follicles with relatively equal proportions of four distinct phenotypes. One follicular phenotype (EA with cCOC; 210-C-EA) mimicked the control phenotype while the remaining three follicular phenotypes exhibited one (expanded cumulus; EA with eCOC; 210-E- EA), two (expanded cumulus and high P in FF; EI with eCOC; 210-E-EI) or three (expanded cumulus and high P and O in FF; EEI with eCOC; 210-E-EEI) phenotypic characteristics of prematurely luteinized ovulatory-size follicles. Whether these four types of ovulatory-size follicles represented sequential changes (for example, from an EA ovulatory-size follicle with 69 cCOC into the EEI ovulatory-size follicle with an eCOC) or distinct trajectories that ovulatory- size follicles follow in response to ovarian stimulation with excessive FSH doses, is yet to be determined. Nevertheless, this heterogeneity very likely contributes to the inverse relationship we have reported between FSH dose during ovarian stimulation with oocyte recovery and live birth rate during ART in women [1, 2] and reduced estradiol production and decreased ovulation rate in cattle [6]. However, the excessive FSH dose did not cause premature luteinization of all the ovulatory-size follicles developing in response to ovarian stimulation in the present study. In fact, the control phenotype for ovulatory-size follicles was also observed in 28±3% of ovulatory- size follicles in the heifers that were stimulated with the excessive FSH dose. The reason for this is unclear but perhaps may be explained by differences in the capacity of the granulosa and cumulus cells in individual ovulatory-size follicles to respond to the excessive FSH dose or differences in exposure to serum FSH. In sheep, granulosa cells from different follicles from the same individual have varying capacities to respond to stimulation with FSH and hCG as measured by in vitro cAMP production [42]. Furthermore, follicular development is hierarchical during follicular waves in cattle [43]. Differences in ovarian vascular supply to individual follicles [44, 45] may cause different distributions of exogenous FSH administered during ovarian stimulation to individual follicles. This unequal distribution of FSH to individual follicles could explain why some ovulatory-size follicles in heifers undergoing ovarian stimulation with the excessive FSH dose in the present study did not undergo premature luteinization and the observed heterogeneity of follicles. Expansion of the cumulus cell layers of the COC is not only the predominate symptom of premature luteinization, but it also can occur independent of concomitant changes in 70 intrafollicular hormone concentrations in the ovulatory-size follicles in heifers undergoing ovarian stimulation with the excessive FSH dose in the present study. Nearly 30% of the ovulatory-size follicles (210-E-EA) in heifers undergoing ovarian stimulation with excessive FSH doses had eCOC but no alterations in intrafollicular hormone concentrations compared not only with controls (70-C-EA), but also with the EA, ovulatory-size follicles with cCOC (210-C- EA) in heifers undergoing ovarian stimulation with the excessive FSH dose. Although the implications of the premature eCOC phenotype are unknown, this suggests that other maturational processes linked to cumulus cell expansion, including for example increased glucose utilization [46], ovulation [38], oocyte maturation [47], and the developmental potential of oocytes in cattle [36, 48] and other species [38, 49], may also be altered during ovarian stimulation of heifers with excessive FSH doses. Clinical methods to distinguish the EA, ovulatory-size follicles with cCOC from prematurely luteinized ones, have not been developed, but may be necessary to improve ART outcomes. There is considerable in vivo and in vitro evidence that the oocyte plays an integral role in preventing premature luteinization. The oocyte prevents luteinization and downregulation of FSHR and modulates FSH-induced steroidogenesis, thus enhancing E production while concomitantly inhibiting P production in granulosa cells [50-52]. The evidence of premature cumulus cell expansion coupled with the shift from E to P production by ovulatory-size follicles in heifers undergoing ovarian stimulation with the excessive FSH dose observed in the present study implies that excessive FSH doses may also impair the capacity of the oocyte to regulate luteinization. Our previous study using the same small ovarian reserve heifer model demonstrated that excessive FSH doses decreased capacity of ovulatory-size follicles to ovulate in response to an ovulatory stimulus [6]. However, it is unknown if premature expansion of the 71 COC in ovulatory-size follicles developing in response to the excessive FSH dose observed in the present study also impairs responsiveness of these follicles to an ovulatory stimulus like hCG or GnRH. If so, this could explain why we observed a reduced ovulation rate in heifers undergoing ovarian stimulation with excessive FSH doses in our previous study [6]. Ovarian stimulation of heifers with the excessive FSH dose in the present study altered intrafollicular production of hormonal markers of luteinization. These include established markers seen in cattle and other species including humans, such as changes in E [28, 30, 33], P [28, 30, 33] and O [5, 53-55]. This observation provides direct endocrine evidence, linked to the differences in COC morphology, that excessive FSH doses induce premature luteinization in a large proportion of the ovulatory-size follicles developing in response to ovarian stimulation. For example, a higher proportion (46%) of the ovulatory-size follicles in heifers undergoing ovarian stimulation with the excessive, compared with the industry-standard, FSH dose in the present study were classified as EI and EEI with eCOC. These same ovulatory-size follicles also had E:P that were lower while P concentrations in FF were higher compared with the EA ovulatory-size follicles in heifers with cCOC undergoing ovarian stimulation with the industry-standard FSH dose (control). However, whether this intrafollicular shift from E to P production by ovulatory- size follicles developing during ovarian stimulation of heifers with excessive FSH doses also resulted in enhanced circulating concentrations of progesterone, similar to that observed during ART in some women [16, 18, 20], was not determined in the present study. Although beyond the scope of the present study, it is critical to understand the mechanism whereby excessive FSH doses during ovarian stimulation induce premature luteinization of the developing ovulatory-size follicles. High concentrations of purified FSH trigger ovulation in rodents and primates [37]. In addition, in vitro maturation (IVM) 72 experiments demonstrate that high FSH alone induces cumulus cell expansion [37], and this is particularly important, given the absence of evidence for functional LH receptor expression in the cumulus cells [56-58]. Even in vivo, cumulus cells rely on a paracrine signalling loop to respond to the preovulatory LH surge. Specifically, LH stimulates production of EGF-like factors in the granulosa cells which then signal via the EGF receptor, expressed on the cumulus cells, mediating the effects of LH on the cumulus cells [59]. However, FSH can also induce the expression of EGF-like factors including AREG and EREG in the granulosa and cumulus cells, and supplementation of IVM media with AREG and EREG resulted in cumulus cell expansion and oocyte maturation [60-62]. It is worth noting in these and other in vitro embryo production experiments, the proportions of COCs that underwent cumulus cell expansion following IVM varied considerably, a further indication of the heterogeneity in cumulus cell responsiveness to FSH [57, 60]. Based on similarities in cumulus cell gene expression profiles, others have suggested that in vitro, FSH signalling mimics some of the downstream pathways induced by LH in vivo [27]. Thus, the variable capacities of COCs to respond to supraphysiological concentrations of FSH may explain the dose-dependent differences in the proportion of ovulatory-size follicles with eCOC and why most, but not all, of the ovulatory-size follicles in the heifers undergoing ovarian stimulation with the excessive FSH dose had eCOC in the present study. The eCOC phenotype observed herein provides compelling evidence that excessive FSH doses during ovarian stimulation of heifers with a low AFC and small ovarian reserve induces premature luteinization of a large proportion of the ovulatory-sized follicles. In summary, the heifers undergoing ovarian stimulation with the excessive, 210 IU, compared with the industry-standard, 70 IU, FSH dose had ovulatory-size follicles containing primarily eCOC and classified as EI or EEI. The EI or EEI ovulatory-size follicles with eCOC in 73 the excessive dose heifers had a lower E or E:P ratio but higher P or O concentrations compared with ovulatory-size follicles in the 70 IU FSH dose heifers. The excessive FSH doses used during ovarian stimulation are concluded to induce premature luteinization of the majority of ovulatory-size follicles in heifers with small ovarian reserves. Whether premature luteinization of ovulatory-size follicles also impairs their capacity to ovulate in response to an LH stimulus, initiates atresia or alters oocyte maturation and quality thereby contributing to oocyte wastage, is unknown. Moreover, it is also unknown whether premature luteinization of ovulatory-size follicles during ovarian stimulation with excessive FSH doses is unique to heifers with a small ovarian reserve. The answers to these questions are, however, critical to improving the efficiency of ovarian stimulation protocols and ART outcomes not only in cattle, but perhaps in other mono-ovulatory mammals, like women. 74 REFERENCES 1. 2. 3. 4. 5. 6. 7. 8. Baker VL, Brown MB, Luke B, Smith GW, Ireland JJ. Gonadotropin dose is negatively correlated with live birth rate: analysis of more than 650,000 assisted reproductive technology cycles. Fertil Steril 2015; 104:1145-1152.e1145. Clark ZL, Thakur M, Leach RE, Ireland JJ. 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Antral follicle count (AFC, follicles ≥3 mm in diameter) prior to ovarian stimulation and number, and diameter of ovulatory-size (≥10 mm) follicles developing in response to ovarian stimulation of heifers with the industry standard, 70 IU, or excessive, 210 IU, FSH dose. Heifers with a low AFC (≤15 follicles) and small ovarian reserve were subjected to PG injections to synchronize estrous cycles and injected with 70 IU (n = 7 heifers) or 210 IU (n = 6 heifers) FSH during ovarian stimulation as explained in Methods. AFC was determined prior to the first FSH injection whereas number and diameter of ovulatory-size follicles was determined 12 h after the last FSH injection. ANS = P>0.05 81 Figure 6. Representative images of cumulus cell-oocyte complexes (COC) with compact (cCOC) or expanded (eCOC) layers of cumulus cells from heifers treated with the industry- standard, 70 IU, or excessive, 210 IU, FSH dose. Holstein heifers with a low AFC and small ovarian reserve were injected with either the 70 IU or 210 IU FSH dose during ovarian stimulation and ovulatory-size follicles (≥10 mm) excised from ovaries 12 h after the last FSH injection as explained in Methods. COCs recovered from FF of each follicle were classified under the dissecting microscope as having a compact-COC (two images on left) or expanded-COC (two images on right). The white scale bar in the bottom right corner of each image represents 130 µm, the diameter of the oocyte for an ovulatory-size follicle [63]. 82 Figure 7. Effect of FSH dose (IU) and COC morphology (cCOC, eCOC) on concentrations of estradiol (E), progesterone (P), oxytocin (O) and ratio of E:P in follicular fluid (FF). Holstein heifers with a low AFC and small ovarian reserve were injected with either the industry- standard, 70 IU, or the excessive, 210 IU, FSH dose during ovarian stimulation and ovulatory-size follicles were excised from ovaries 12 h after the last FSH injection as explained in Methods. COC morphology was assessed in 5 to 15 ovulatory-size follicles per heifer and the concentrations of E, P and O were measured in the FF of these follicles. Total number of ovulatory-size follicles in each FSH dose-COC morphology group of heifers for the E, P and E:P measurements follows: 70 IU-cCOC = 59; 210 IU-cCOC = 20; 210 IU-eCOC = 51. For O, a random subset of ovulatory-size follicles was selected for analysis in each FSH dose-COC morphology group of heifers as follows: 70 IU-cCOC = 22; 210 IU-cCOC = 12; 210 IU-eCOC = 36. Bars represent the mean ± SEM for heifers (not follicles). Lines above bars indicate which means were compared whereas asterisks (* P<0.05, *** P<0.001) denote differences between means for heifers. N = number of heifers. 83 Figure 8. Effect of FSH dose (IU) on concentrations of estradiol (E), progesterone (P), oxytocin (O) and ratio of E:P in follicular fluid (FF). Holstein heifers with a low AFC and small ovarian reserve were injected with either the industry-standard, 70 IU, or the excessive, 210 IU, FSH dose during ovarian stimulation and ovulatory-size follicles were excised from ovaries and FF aspirated 12 h after the last FSH injection as explained in Methods. Concentrations of E and P in FF were measured in 5 to 24 follicles per heifer (n = a total of 93 ovulatory-size follicles from 7 heifers) treated with 70 IU FSH and in 15 to 23 follicles per heifer (n = 114 ovulatory-size follicles from 6 heifers) treated with 210 IU FSH dose. Oxytocin was measured in 2 to 5 follicles per heifer (n = 22 of the 93 ovulatory-size follicles from the 7 heifers) treated with 70 IU FSH and in 7 to 9 follicles per heifer (n = 48 of the 114 ovulatory-size follicles from the 6 heifers) treated with 210 IU FSH. N within bars indicates number of heifers treated with each FSH dose and the bars represent the mean (± SEM) concentration of E, P, E:P ratio and O for heifers (not follicles). Asterisks indicate differences between means were significant (*** P<0.001). 84 Group FSH COC E:P ratio (IU) Morphology in FF 70-C-EA 70 Compact 70-C-EI 70 Compact 210-C-EA 210 Compact 210-E-EA 210 Expanded 210-E-EI 210 Expanded 210-E-EEI 210 Expanded EA [4.11±0.83] EI** [0.68±0.04] EA [2.61±0.45] EA [2.17±0.41] EI*** [0.59±0.12] EEI*** [0.03±0.01] Proportion Follicle Concentration (ng/mL) (%) per diameter heifer (mm) 79±6 12.0±0.8 21±6 11.4±1.5 28±3 11.5±0.5 26±5 12.3±0.7 24±8 11.7±0.5 Estradiol Progesterone Oxytocin 191±28 49±10 0.28±0.03 (N=7, 48)A (N=7, 48) (N=7, 12) 59±10** 114±41 0.25±0.04 (N=5, 11) (N=5, 11) (N=5, 10) 207±33 104±23 0.40±0.04 (N=6, 20) (N=6, 20) (N=6, 12) 203±31 110±10 0.34±0.05 (N=6, 21) (N=6, 21) (N=6, 12) 116±27 268±75* 0.76±0.15 (N=6, 16) (N=6, 16) (N=6, 12) 22±3 11.6±0.5 33±10*** 2643±1027*** 6.80±2.43*** (N=6, 15) (N=6, 15) (N=6, 12) Table 2. Effect of COC morphology (Compact, Expanded) and classification of follicles as estrogen-active (EA), estrogen-inactive (EI) or extreme-estrogen-inactive (EEI) during ovarian stimulation of heifers with 70 IU (industry standard) or 210 IU (excessive) FSH doses on ratio of E:P in follicular fluid (FF), proportion of follicles in each COC and follicle classification category, and on diameter and concentrations of estradiol (E), progesterone (P), and oxytocin (O) in ovulatory-size follicles. Holstein heifers with a low AFC and small ovarian reserve were injected with either the 70 IU or 210 IU FSH dose and ovulatory-size follicles were excised from ovaries as explained in Methods. The COC recovered from the FF of 5 to 15 ovulatory-size follicles per 85 Table 2 (cont’d) heifer were classified as either having compact or expanded layers of cumulus cells and each ovulatory-size follicles was also classified based on E:P in FF as EA (E:P in FF ≥1), EI (E:P in FF <1>0.1) or EEI in FF= ≤0.1) as explained in Methods. This table summarizes the follicular characteristics in this subset of follicles in which we had information on COC morphology as well as intrafollicular ratio of E:P. The hormone concentrations in the EA ovulatory-size follicles with a compact COC in heifers undergoing ovarian stimulation with the 70 IU FSH dose (depicted in the first row of this table; 70-C-EA) were considered representative of preovulatory follicles in cattle [28, 30, 35] and thus used as control values. Asterisks (* P<0.05, ** P<0.01, *** P<0.001) denote differences between the means (±SEM) follicles). vs control AN = number of heifers, or number of ovulatory-size follicles groups heifers heifer other (not for 86 Follicular Hyperstimulation Dysgenesis: New Explanation for Adverse Effects of Excessive FSH in Ovarian Stimulation1 CHAPTER 4. 1 Zaramasina L Clark, Meghan L Ruebel, Peter Z Schall, Kaitlin R Karl, James J Ireland, Keith E Latham, “Follicular Hyperstimulation Dysgenesis: New Explanation for Adverse Effects of Excessive FSH in Ovarian Stimulation.” Endocrinology. 2022. Volume 163, Issue 9. By permission of Oxford University Press. 87 Title: Follicular hyperstimulation dysgenesis: new explanation for adverse effects of excessive FSH in ovarian stimulation Running Title: Follicular hyperstimulation dysgenesis Keywords: excessive FSH doses, ovarian stimulation, transcriptome analysis, cumulus cells, granulosa cells, oocytes, ovulatory-size follicles Authors and affiliations: Zaramasina L. Clark1,2,3 Meghan L. Ruebel1,2,4, Peter Z. Schall1,2,5, Kaitlin R. Karl2, James J. Ireland2, Keith E. Latham2* 1 Equal contributions. 2Reproductive and Developmental Sciences Program, Department of Animal Science, Michigan State University, East Lansing, MI, USA; 3Present address: School of Biological Sciences, Victoria University of Wellington, Wellington, New Zealand; 4Present address: University of Colorado-Anschutz Medical Campus, Aurora, CO, USA; 5Present address: University of Michigan, Ann Arbor, MI, USA *Correspondence: Reproductive and Developmental Sciences Program, Department of Animal Science, Michigan State University, East Lansing, MI 48824, USA. Tel: +1517 353-7750; E- mail: latham1@msu.edu :-0003-1206-9059 Grant support: This study was supported by the Agriculture and Food Research Initiative Competitive USDA-NIH Dual Purpose Program (Grant no. 2017-67015-26084), the USDA National Institute of Food and Agriculture, the Eunice Kennedy Shriver National Institute of Child Health and Human Development of the National Institutes of Health under Award Number T32HD087166, and AgBioResearch at Michigan State University. 88 Abstract High FSH doses during ovarian stimulation protocols for assisted reproductive technologies (ART) are detrimental to ovulatory follicle function and oocyte quality. However, the mechanisms are unclear. In a small ovarian reserve heifer model, excessive FSH doses lead to phenotypic heterogeneity of ovulatory-size follicles, with most follicles displaying signs of premature luteinization, and a range in severity of abnormalities. By performing whole transcriptome analyses of granulosa cells, cumulus cells and oocytes from individual follicles of animals given standard or excessive FSH doses, we identified progressive changes in the transcriptomes of the three cell types, with increasing severity of follicular abnormality with the excessive doses. The granulosa and cumulus cells each diverged progressively from their normal phenotypes and became highly similar to each other in the more severely affected follicles. Pathway analysis indicates a possible dysregulation of the final stages of folliculogenesis, with processes characteristic of ovulation and luteinization occuring concurrently rather than sequentially in the most severely affected follicles. These changes were associated with disruptions in key pathways in granulosa and cumulus cells, which may account for previously reported reduced estradiol production, enhanced progesterone and oxytocin production and diminished ovulation rates. Predicted deficiencies in oocyte survival, stress response and fertilization suggest likely reductions in oocyte health, which could further compromise oocyte quality and ART outcomes. 89 Introduction High follicle-stimulating hormone (FSH) doses are incorporated into assisted reproductive technologies (ART) in women and cattle to increase the number of ovulatory-size follicles and oocytes available for in vitro or in vivo fertilization and embryo transfer. However, high FSH doses during ovarian stimulation are inversely related to oocyte recovery and live birth rates in women [1,2], and decreased ovarian function and embryo transfer success in cattle [3-7]. These observations support the hypothesis that high FSH doses during ovarian stimulation protocols are detrimental to ovulatory follicle function and oocyte quality, but the mechanisms of these effects are unclear. During natural reproductive cycles, ovulatory follicles undergo luteinization in response to the preovulatory luteinizing hormone (LH) surge, characterized by an intrafollicular shift from primarily estradiol to progesterone production. However, high doses of recombinant human FSH (hFSH) administered to women [8,9] and animals [10] induces premature luteinization, characterized by a reduced circulating estradiol:progesterone (E:P) ratio on the day of the ovulatory stimulus [9,11-17]. Premature luteinization occurs in 12.3 to 46.7% of ART cycles in women but its impact on ART outcomes is unclear, due in part to inconsistent assignment of a premature luteinization phenotype [18]. We are not aware of any FSH dose-response studies in women or other relevant animal models that have directly examined the ovulatory-size follicles developing in response to ovarian stimulation with excessive FSH doses (i.e., doses greater than needed to achieve a maximum ovulatory response). Consequently, it is unknown if excessive FSH doses during ovarian stimulation cause differentiation (e.g., luteinization) of its target sites, the granulosa (GC) and cumulus cells (CC), resulting in premature luteinization [9,11-17], impaired oocyte quality and ART outcomes. 90 The small ovarian reserve heifer (11 to 12 months old, SORH) is a unique and ideal model for identifying excessive FSH doses and their impacts on ovulatory follicle function. Aside from being a mono-ovulatory model that like women are subjected to ART, it mimics the well-established characteristics of women with small ovarian reserves. These include a diminished number of morphologically healthy follicles (and thus oocytes) in the ovarian reserve, FSH hypersecretion, low circulating anti-Müllerian hormone, progesterone and androgen concentrations during reproductive cycles, and reduced responsiveness to ovarian stimulation compared with age-matched individuals with a larger ovarian reserve [19-21]. Our previous work demonstrated that increasing the FSH dose to elicit a greater response to ovarian stimulation during ART is detrimental to number of oocytes retrieved and live birth rate [1,2]. Although, the primary reason women seek ART is due to having a small ovarian reserve, it is unclear why these individuals respond poorly to high FSH doses during ovarian stimulation [22,23]. Using the SORH model in a dose-response study, we found that doses 3-fold higher than industry-standard doses of Folltropin-V, a commercial FSH-enriched porcine pituitary preparation (cpFSH), during ovarian stimulation were excessive because they did not increase the number of ovulatory-size follicles but reduced hCG-induced ovulation rate and decreased circulating estradiol concentrations [3]. In response to the industry-standard cpFSH dose, ovulatory-size follicles contained compact cumulus-oocyte complexes (cCOCs), and most (80%) had higher intrafollicular E:P [24]. These follicles were considered morphologically healthy, estrogen-active (EA) dominant follicles, mimicking the well-established phenotype of dominant preovulatory follicles developing during the follicular phase of estrous cycles prior to the preovulatory gonadotropin surge in cattle [25-27]. In stark contrast, most (72%) of the ovulatory- 91 size follicles developing in response to the excessive cpFSH doses contained expanded COCs (eCOCs) and >50% of these eCOC-containing follicles had lower E:P (estrogen-inactive, EI) and/or high oxytocin concentrations [24]. These EI ovulatory-size follicles with eCOCs mimicked the phenotypes of dominant ovulatory follicles developing during the follicular phase of estrous cycles following the preovulatory gonadotropin surge (but prior to ovulation), or the dominant non-ovulatory follicles developing during the luteal phase of estrous cycles as they lose dominance and undergo atresia during a non-ovulatory follicular wave in cattle [25,26,28]. Thus, most ovulatory-size follicles developing in response to ovarian stimulation with excessive cpFSH doses undergo premature luteinization, characterized by CC expansion, reduced estradiol but enhanced intrafollicular progesterone and/or oxytocin production, and a diminished sensitivity to an LH-like (human chorionic gonadotropin; hCG) stimulus resulting in a reduced ovulation rate. FSH is the predominant gonadotropin in Folltropin-V, however LH is a minor (<1%) contaminant [29]. Regardless, high Folltropin-V doses during ovarian stimulation do not enhance circulating progesterone or estradiol production, or induce early ovulation, as would be expected if the preparation contained biologically significant amounts of LH [3]. Additionally, Folltropin- V injections do not enhance overall circulating LH concentrations in cattle [30]. These combined observations indicate that the LH contamination even in high doses of Folltropin-V is likely insufficient to impact ovarian function. Other reports support the conclusion that premature luteinization is likely attributable to excessive FSH concentrations [10,18]. Overall, this supports the hypothesis that excessive FSH doses during ovarian stimulation disrupt pathways in GC and CC directly and indirectly regulated by FSH, resulting in premature luteinization of ovulatory- size follicles, leading to impaired oocyte quality. 92 To test this hypothesis, and to discover the molecular and cellular mechanisms responsible for adverse effects of excessive FSH doses, we used the SORH model and applied RNA sequencing (RNA-seq) and a bioinformatics analysis pipeline incorporating Ingenuity Pathway Analysis (IPA; QIAGEN Inc., https://www.qiagenbioinformatics.com/products/ingenuity-pathway-analysis, [31]) to identify transcriptome differences between ovulatory-size follicles developing during ovarian stimulation with industry-standard vs excessive cpFSH doses. We acquired transcriptome data for each cell type (GC, CC and oocyte) from individual ovulatory-size follicles either without (industry- standard dose) or with (excessive dose) varying degrees (based on COC morphology and intrafollicular estradiol, progesterone, and oxytocin concentrations) of the premature luteinization phenotype. The results provide new mechanistic insight into the impact of excessive cpFSH doses on cellular phenotypes within individual ovulatory-size follicles, including remarkable changes in molecular and cellular pathways, and functions predictive of diminished oocyte quality that could not fully be explained by premature luteinization alone. These results point to potential risks of using excessive FSH doses during ART. Materials and Methods Supplemental data files are available at Figshare (RRID:SCR_004328), https://scicrunch.org/resolver/RRID:SCR_004328, doi: 10.6084/m9.figshare.19208604. https://doi.org/10.6084/m9.figshare.19208604.v1 [32]. Small ovarian reserve heifer (SORH) selection, ovarian stimulation protocol and excision of ovulatory-size follicles Heifers with a small ovarian reserve used in this study (n=14) were selected as previously described [3,24]. Briefly, ultrasonography was used to determine the antral follicle count (AFC; 93 number of antral follicles ≥3 mm in diameter) of two groups of 50 Holstein heifers (11-12 months old; Green Meadow Farms Inc., Ovid-Elsie, MI). Using the AFC, animals were ranked and the twenty-five heifers with the lowest AFC in each group were administered two prostaglandin F2α (PG; 12.5 mg/mL, Lutylase HighCon, Zoetis) injections ten days apart to induce ovulation and the initiation of the first follicular wave. The AFC was measured prior to each PG injection and four days after the final PG injection. Animals with an AFC ≤15 (14 heifers) on these days were selected for inclusion in this study and housed at the Michigan State University Beef Cattle Teaching and Research Centre, for the duration of the study. Our previous study has shown that age-matched individuals with a AFC ≤15 also have an 80% smaller ovarian reserve (total number of morphologically healthy follicles/oocytes in ovaries) than heifers with a high (≥25) AFC [20]. All procedures described herein were approved by the Institutional Animal Care and Use Committee at Michigan State University. The two cpFSH doses investigated herein were chosen as the industry standard (70 IU) and excessive (210 IU) doses based on previous data [3, 24]. For ovarian stimulation, heifers were treated with either 70 IU (40 mg, n = 7 heifers) or 210 IU (120 mg, n = 7) cpFSH (Folltropin-V, Lot# 499213, Vetoquinol USA Inc) per injection. The cpFSH contains 700 IU (equivalent to 400 mg NIH-FSH-P1 with <1 mg NIH-LH-S19) of FSH per 20 mL vial [29]. One heifer treated with the 210 IU dose was excluded from the final analysis due to an absence of response to ovarian stimulation. Heifers were synchronized by administering PG injections ten days apart followed by a third PG injection 12 h later. At the time of ovulation or emergence of the first follicular wave (~36 h after the last PG injection), animals were randomly assigned to treatment with either 70 or 210 IU cpFSH twice daily for four days (8 injections total). Daily ultrasonography was used to monitor the AFC and diameter of these follicles. The AFC prior to 94 superovulation did not differ between cpFSH dose [24]. Heifers were euthanized 12 h after the final cpFSH injection and the pairs of ovaries were recovered in lieu of administration of an ovulatory dose of hCG that is standard for this superovulation protocol [3]. The number and size of ovulatory-sized follicles produced in response to ovarian stimulation did not differ between cpFSH doses [24]. Ovaries were washed briefly in 70% ethanol, then Dulbecco’s Phosphate Buffered Saline (DPBS) and then placed in fresh DPBS for the duration of tissue collection. For a subset of follicles (5-15/heifer), ovulatory-size (≥ 10 mm) follicles were individually dissected from the ovary. Follicular fluid was aspirated to recover the COC. The aspirated follicular fluid was immediately transferred to a separate dish and processed as described below. The follicular fluid was recovered and stored for measurement of progesterone, estradiol, and oxytocin. Measurement of progesterone, estradiol and oxytocin and determination of estradiol:progesterone ratio in follicular fluid of ovulatory-size follicles As described [24], estradiol, progesterone and oxytocin concentrations were measured in follicular fluid samples from all ovulatory-size follicles where a COC was recovered (n = 5- 15/heifer). Estradiol and progesterone concentrations were measured using commercially available ELISA kits (Estradiol, RRID:AB_328054; Progesterone, RRID:AB_2811273; Cayman Chemical Company, MI), enabling the calculation of the estradiol:progesterone (E:P) ratio. The E:P ratios were used to classify follicles as EA (E:P ratio ≥1) or EI (E:P ratio <1). Follicular fluid oxytocin concentrations, another established marker of premature luteinization [33-36], were also measured using the Oxytocin ELISA kit (RRID:AB_2815012; ENZO Life Sciences, Inc, NY, USA). 95 Follicle types and nomenclature, and statistical analyses of follicle characteristics Type 1 follicles were those from cattle that receive the industry-standard dose (70 IU) of cpFSH. Three of the high-dose (210 IU) follicular phenotypes were chosen for analysis by RNA- seq, designated as Types 2-4, respectively [32]. The statistical analyses performed on data in Table 1 were undertaken using the Statistical Analysis System (SAS 9.4) software program [37]. Data were arcsine- (E:P ratio) or log- (estradiol, progesterone and oxytocin) transformed before overall treatment (i.e. follicle type) effects were evaluated, and Fisher LSD was used to determine if means differed (P<0.05). Type 1 and Type 2 follicles were highly similar to each other and presumably represent the most healthy, normal follicles having cCOC and EA characteristics. Based on these parameters, it is important to note that the only difference between Type 1 and 2 follicles was the cpFSH dose used during ovarian stimulation, 70 and 210 IU, respectively. With progressively more severe abnormalities, Types 3 and 4 follicles contained eCOC, and Type 4 follicles additionally displayed an EI endocrine profile, making them phenotypically the most divergent from Type 1 follicles. Samples from seven heifers receiving the standard dose and six heifers receiving the excessive dose were obtained. One sample per relevant follicle type per heifer was included where possible and were randomly selected and excised from the ovaries, and processed to yield a GC sample, a CC sample and an oocyte sample for RNA-seq. Only high-quality RNA-seq libraries were included in the analysis of transcriptome effects for GC (Type 1 n=7, Type 2 n=6, Type 3 n=5, Type 4 n=4), CC (Type 1 n=6, Type 2 n = 6, Type 3 n=6, Type 4 n=4), and oocytes (Type 1 n=5, Type 2 n=3, Type 3 n=6, Type 4 n=4). Transcriptome profiles were compared between doses within cell type, and between cell types to assess the molecular changes that 96 accompany and may underlie the observed morphological and endocrine features of these four follicle types. Oocyte and cumulus cell isolation The COC were identified within the follicular fluid that was aspirated from each individual follicle. Dishes were examined under a heated (38.5 °C) stereomicroscope and then were moved to a 35 mm dish filled with HH medium (HEPES buffered medium with 3mg/mL BSA) using a Drummond Micropipette with a glass tip. Morphology of the COC, either compact (cCOC) or expanded (eCOC) were recorded [24]. COCs were incubated in 0.1 % hyaluronidase (Sigma-Aldrich, St. Louis, MO) in HH medium at 37 °C for 5 min to separate the CC from the oocyte. The CC mass and oocyte were then transferred to a dish with HH medium to remove the hyaluronidase. The CC mass was divided into 2-3 aliquots and then washed through three drops of ~200 µL of HH medium. Cumulus cell aliquots were transferred to 200 µL drops of DPBS and moved to a 1.5 mL tube. Tubes were spun down until cells pelleted. The DPBS was removed and 100 µL of PicoPure Extraction Buffer™ was used to resuspend the cell pellet. Tubes were transferred to a 42 °C incubation for 30 min and stored at -80° C. After separation from the CC, the oocyte was incubated in 0.1% pronase in HH medium and observed for removal of the zona pellucida. Denuded oocytes were transferred into 3, ~150 µL drops of HH medium to wash off the pronase solution. The oocytes were then moved into a tube with 50 µL PicoPure Extraction buffer™, which was incubated at 42 °C for 30 min and stored at -80° C. Granulosa cell isolation The follicle wall was bisected, and the GCs were gently scraped into DPBS. The follicle wall was removed from the dish and small sections of GC were isolated and transferred into a 97 tube where 100 µL of PicoPure Extraction Buffer™ was added and processed as described above for the CC. All samples were stored at -80° C until RNA extraction. RNA Extraction and RNA-seq library preparations Following the manufacturer’s protocol, total RNA was isolated from oocytes, CC, and GC using the PicoPure RNA Isolation Kit™ (ThermoFischer Scientific, Waltham, MA) including a DNase digestion (RNase-Free DNase; Qiagen, Hiden, Germany) to remove contaminating DNA. A randomly selected subset of samples was assessed for RNA quality and all samples fell in the range of 9.2 – 10.0 RIN values (mean ± SEM = 9.9 ± 0.1), indicating that the RNA isolation method was robust and yielded high quality RNA. For RNA-seq library preparation of oocytes, the Ovation® SoLo RNA-Seq System kit (TECAN, Redwood, CA) was used, including a bead purification, end repair, adaptor ligation and first round library amplification and purification steps. Then 20–30 ng of each library was used for the remainder of library preparation, which included the use of AnyDeplete bovine primers for rRNA depletion, and a second round of library amplification and purification. With the SoLo kit, enzymatic shearing was applied rendering RNA-seq libraries between 300 and 350 bp in length. For CC and GC, library construction was conducted using the TECAN Universal RNA- Seq with NuQuant kit which included Bovine AnyDeplete primers for rRNA depletion. Library construction steps included cDNA synthesis, fragmentation using a Covaris-2 sonicator that mechanically fragments the cDNA to an average length of 300 bp, end repair, adaptor ligation, strand selection, AnyDeplete, and final library amplification steps following the manufacturer’s protocol. All RNA-seq libraries were assessed for quantity using the Qubit DNA High Sensitivity Assay Kit (ThermoFisher Scientific) and quality using the Agilent High Sensitivity DNA kit 98 (Agilent Technologies) on a Bioanalyzer, following product instructions. A total of 23 oocytes (1 oocyte was excluded due to poor library quality), 24 CC, and 24 GC barcoded libraries were pooled and sequenced on an Illumina HiSeq 4000 (Illumina, San Diego, CA, USA) at the Michigan State University- Genomics Core. During the demultiplexing of the oocyte sequencing results, two samples were flagged for potential inversion of barcode identification and were subsequently removed from downstream analysis. An additional two CC, two GC, and two oocyte samples were excluded due to low quality output relating to low generation of reads, low complexity, etc. This resulted in a total of high-quality samples numbering 22 for CC and GC, and 18 for oocytes. RNA sequencing data are available at the Gene Expression Omnibus (RRID:SCR_005012) https://scicrunch.org/resolver/SCR_005012 (accession number GSE197116) [38]. Sequencing data processing Sample information included follicle type membership, dose, morphology, and exonic reads [32]. Raw sequencing data in fastq format underwent initial sequencing quality metrics using FastQC (https://www.bioinformatics.babraham.ac.uk/projects/fastqc). Resultant FastQC data identified aberrant nucleotide distribution in the first 5 basepairs. Trimming was conducted using TrimGalore [39] with the following parameters: minimum quality threshold of 20, minimum length of 20 basepairs, removal of Ns from ends of reads, and a hard trim of the first 5 basepairs. The bovine cDNA genome (ARS-UCD1.20, build 102) was downloaded from Ensembl and indexed with Kallisto (v0.44.0) [40] and mRNA abundance quantitation were calculated using standard settings. 99 Differential expression calculation Resultant Kallisto outputs were processed with DESeq2 (v1.30.1) [41] within R (v4.1.0). For normalization and differential expression calculations, the somatic cells (CC and GC) were processed jointly while the oocyte samples were processed independently. Transcript abundance was collapsed to gene abundance using biomaRt (v2.45.8) [42], using Ensembl gene identifiers. Pre DEG (differentially expressed genes) filtering of lowly expressed genes was conducted, using a threshold of at least 1 FPKM in at least one sample. For CC and GC, all pairwise comparisons were conducted: comparing follicle types within cell type and comparing follicle types across cell type. Within the oocyte samples, pairwise comparisons were calculated comparing follicle types. Level of significance for DEG was set at false discovery rate (FDR) <0.01. We considered limiting the analysis to a set of highly homologous genes identified using the MetaPhOrs database to address possible concerns related to gene annotation differences between bovine and the other species from which much of the data in the IPA database have been derived [43]. However, because the application of sequence-based cross-species consistency scores can exclude some well-studied genes, even apparent gene homologs that share a gene symbol annotation, we elected not to apply this additional filter in our analysis. IPA analysis Identification of Canonical Pathways (CPs), Disease and Functions (DFs), and Upstream Regulator (URs) was conducted by submitting the resultant DEG lists to Ingenuity Pathway Analysis (QIAGEN Inc., https://www.qiagenbioinformatics.com/products/ingenuity-pathway-analysis) (IPA database content as of 01/2021) [31]. IPA was selected due to the robustness of its manually curated Knowledge Base, which contains >7M observations (Qiagen.com, March 100 2021) including molecular interactions organized into >700 pathways, >800,000 expression datasets and reported associations of molecules with diseases and biological functions, and >30 integrated third-party databases (Qiagen IPA in-program description), and its ability to compare multiple datasets. Like standard gene set enrichment methodology, submitted gene lists were compared to the genes associated with each CP/DF/UR to calculate a level of significant overlap (p-value; significance set at 0.05). With the known impact of up- or down-regulating genes on a given IPA CP or BF entry, the software can also calculate a direction of CP or BF modulation (activation or inhibition), denoted as positive and negative z-scores, respectively (significance set at z>|1.96|). For the UR analysis, the activity of an UR is predicted based on the direction of change for the downstream DEG targets. It should be noted that the magnitudes of gene expression changes do not factor into the calculations, only the direction of change. To remove potential spurious results, IPA data were limited to those entries with greater than one DEG present. DF entries were filtered to remove disease/cancer related entries, and the term Biological Functions (BF) applied. To prioritize the consideration and presentation of IPA results, we employed a hierarchical approach. IPA natively identifies entities that satisfy two statistical measures. P- value: level of significant overlap between the number of DEGs and the number of molecules in an IPA entry; significance set at p<0.05. Z-score: level of directionality based on the known interactions; significance set at |z|>1.96. All entries are limited to significant p-values. We then noted those IPA entries with significant z-scores. The remaining IPA entries, lacking a significant z-score, were examined with respect to p-value and other parameters, such as mRNA expression level and regulation. 101 The combined approach of whole-transcriptome RNAseq plus IPA allows a robust assessment of gene expression changes coupled with the identification of potentially affected pathways and functions, which can further implicate affected upstream regulators, providing novel direction for hypothesis-driven research. While other methods such as RT-qPCR can be useful for quantifying the expression of a selected set of genes, such methods are limited in scope as they can only assay known sequences and can be subject to artifacts related to hybridization. The combination of RNAseq and IPA can provide an evaluation of whether a pathway or network is affected in a given situation. An important value of the whole- transcriptome approach is that it avoids bias and assays a large number of interacting or target molecules, the expression of which provides a statistically powerful assessment of whether a pathway or function is affected. Luteinization Marker Identification Because the IPA database did not include luteinization as a BF, we generated a manually curated list of 71 markers of luteinization, selected from previously published works evaluating changes in DEGs that were characteristic of the transition to a luteinized state [44-52]. The expected changes during luteinization were determined based on the observed increase or decrease during this transition to luteinization. We acknowledge that this is not an exhaustive list of markers characteristic of luteinization (that is, occurring in response to the preovulatory LH surge). Moreover, they are not considered exclusive markers of premature luteinization, that is, luteinization occurring before or in the absence of an ovulatory stimulus, such as LH or hCG. DEG and IPA Figures The R package ComplexHeatmap (v2.6.2) [53] was utilized to generate heatmaps of DEGs and IPA results arranged with cowplot (v1.1.1) [54]. Correlation data were generated with 102 the R package corrplot (v0.90) via the Pearson method and plotted using the ComplexHeatmap package. Results Study design and data set The overall goal of the study was to compare the transcriptomes of oocytes, CC and GC isolated from ovulatory-size follicles in ovaries of SORH that received either a standard (70 IU) or excessive (210 IU) dose of cpFSH, to ascertain the effects of cpFSH dose and to assess potential cell-cell interactions at the transcriptome level. In an earlier report [24] we determined the different ovulatory-size follicle phenotypes based on morphological and endocrine features (i.e. cCOC vs. eCOC, and EA vs. EI). Ovulatory-size follicles from ovaries of animals receiving the industry standard dose were predominantly of a single phenotype (Type 1), whereas multiple follicular phenotypes were identified in animals receiving the excessive dose. Samples from Type 1 follicles were classified as the control follicle type for subsequent major comparisons (i.e., 2 v 1, 3 v 1 and 4 v 1), based on the published characteristics (for example, EA and containing a cCOC [25-27]) of the ovulatory follicle in cattle and the fact that these samples were collected from heifers treated with the industry-standard, 70 IU, cpFSH dose. RNAseq library preparation yielded high quality libraries [32]. We calculated an average of 13 million exonic reads for CC, 17.9 million for GC, and 16.9 million for oocytes. This resulted in detection and quantitation of mRNAs from an average of 14,865 genes for CC, 15,255 for GC and 14,658 for oocytes. Principal component analysis (PCA) indicated a high degree of reproducibility within cell type and follicle type [32]. 103 Progressively more severe follicular abnormality is accompanied by progressively more severe transcriptome alterations in all three cell types DEGs were identified between follicle Types 2-4 and control, Type 1, follicles [32]. Among the excessive dose follicles analyzed, transcriptome divergence from Type 1 follicles increased progressively from Type 2 to Type 3 and then to Type 4 [32]. The PCA plot [32] indicated that all three cell types from Type 4 follicles were most divergent from those from the control, Type 1 follicles. Volcano plots (Figure 9A) reveal not only an increasing number of DEGs in the progression from Type 2 to Type 4, but an additional shift to more genes being overexpressed (indicated in red) in Types 3 and 4 relative to Type 1 follicles for CC and GC. Comparisons of DEGs between Types 2-4 and Type 1 revealed significant overlap in DEGs for follicle Types 2-4 for CC and GC, but also reveal unique DEGs for each follicle type (Figure 9B). Oocytes displayed the fewest DEGs, with just a single DEG for Types 2 and 3 follicles, and 283 DEGs for Type 4 follicles (FDR<0.01), with no overlaps between follicle types (Figure 1B). DEGs with reduced expression include oogenesis factors: Folliculogenesis-specific BHLH Transcription Factor (FIGLA), Serine-threonine kinase 11-interacting protein (STK11IP, a.k.a. LIP1), and ADP ribosylation factor GTPase activating protein 3 (ARFGAP3). CC displayed, 24, 133, and 3,131 DEGs (FDR<0.01) from follicle Types 2-4, respectively, and thus there were many more affected DEGs for Type 4 CC than GC (below). Most of the Type 2 CC DEGs overlapped with Types 3 and/or 4, and most of the Type 3 CC DEGs overlapped with Type 4. Again, CCs from each follicle type displayed a small number of unique DEGs. Of the 24 transcripts in the 2 v 1 follicle type with FDR<0.01, eight were identified as down-regulated in Type 2 relative to Type 1 follicles. The DEGs with the largest 104 magnitude differences (±15 log2(fold-change), relative to Type 1) included both up- (e.g. FGG, MEPE, NGFR, LIF, MDFI, NCS1, RGS2, VGF, AREG and TGFA) and down- (KIAA1324 and AQP11) regulated transcripts [32]. All of the Type 2 upregulated DEGs were also identified as the DEGs with the largest magnitude changes in the Types 3 and 4 follicles. GC displayed 15, 148 and 1,368 DEGs for Types 2-4 follicles, respectively. Fewer than half the Type 2 GC DEGs were shared with Types 3 and 4 follicles, whereas a majority of Type 3 GC DEGs were shared with Type 4 follicles. A small number of DEGs were unique to GC of each follicle type. Of the 15 transcripts in the 2 v 1 follicle types with FDR<0.01, 8 were annotated (SERPINA14, NPS and MASP1 were up-regulated and TMEM255A, FGG, LRRC7, TNFSF4 and CRM1 were down-regulated) [32]. SERPINA14 and MASP1 were also identified as DEGs (FDR<0.01) in the GC of the Types 3 and 4 follicles. In Type 3 follicles, only 5 of 148 DEG were down-regulated. The DEGs with the largest differences were all up-regulated (≥ 10 log2(fold-change)), and included SCG3, SERPINA11, EDN2, SERPINA14, GRO1, CSRP3, XCL1 and ADAMTS20 [32]. In Type 4 follicles, <20% of the 1368 DEGs (FDR<0.01) were down- regulated in GC. The genes with the largest differences were again all up-regulated (≥ 10 log2(fold-change), relative to Type 1 GC), and included SERPINA11, SCG3, SERPINA14, GRO1, NPS, EDN2, ADAMTS20, TNN, XCL1, CSRP3 and AMKRD55 [32]. More severe follicular phenotypes are associated with enhanced expression of a subset luteinization markers Our previous study reported that the effects of excessive cpFSH dose on COC morphology and intrafollicular endocrine profile resembled a premature luteinization effect [24]. To explore this idea further, we compared Types 2-4 to Type 1 follicles for effects in GC and CC on a manually curated panel of 71 luteinization markers (Fig. 10A, Column 1) identified from 105 the literature [44-52]. In comparison with Type 1 follicles, progressively more markers were altered in a manner consistent with luteinization in in Type 2, 3 and 4 ovulatory-size follicles in GCs (26/71 in Type 4) and CCs (33/71 in Type 4), even so, less than half of this list of luteinization markers were affected in any of the follicle types. Many of the affected markers were progressively upregulated in both cell types in Types 3 and 4 (with eCOC morphology) follicles. Many of the luteinization markers were more highly expressed in the CC than GC (Figure 10A, Column 4, denoted by yellow coloring). Three luteinization markers (KIAA1324, VNN1 and AQP11) were affected opposite to the predicted directions in Type 2 or 3 CCs. An additional four markers (IGFBP4, OXTR, ANGPT2, HSD17B7) displayed changes opposite to the predicted directions in Type 4 CC. Analysis of the correlation between CC and GC of the different follicle types with respect to the expression of these luteinization markers revealed that Types 1, 2 and 3 CC were highly similar; Type 1 and Type 2 GCs were highly similar; Type 4 CCs diverged markedly from Types 1-3 CCs; and Type 3 and 4 GCs diverged noticeably from Types 1 and 2 GCs, indicating a stronger deviation with respect to luteinization marker gene expression with more severe follicular abnormality (Figure 10B). CC and GC progressively become more alike as follicular phenotype becomes more abnormal An interesting relationship between CC and GC was seen wherein CC and GC became more like each other across this progression from less to more severe follicular abnormality. This relationship emerged in multiple analyses. The volcano plots (Figure 11A) revealed similar numbers of GC-CC DEGs for Type 1 and 2 follicles (1,511 and 1,463, respectively) but fewer for Types 3 and 4 follicles (559 and 277, respectively). We performed a correlation analysis of all expressed genes for CC and GC normalized against CC and GC in Type 1 follicles (Figure 11B) and found increased similarities between CC4 and both GC3 and GC4. This was also 106 evident for the luteinization markers, progressively fewer differences between CC and GC were seen moving across Types 2, 3 and 4 follicles (Figure 10A, fourth column). We also performed a hierarchical clustering of correlation between the luteinization markers for CC and GC. Although CC and GC clustered separately, increased correlations were seen comparing CC4 with GC4, and comparing CC3 with GC1 and GC3 (Figure 10B). We also noted that out of a group of 15 luteinization markers reported for GCs at 22 h post-hCG in cattle [52], 13 were detected in CCs and in GCs in our study. Of these, 4 were upregulated in Type 4 GCs, and, importantly, 9 were upregulated in Type 4 CCs (Figure 10). This further illustrates that the Type 4 CCs acquire increased resemblance to GCs. More severe follicular phenotypes are associated with progressively increased effects on hormone-responsive genes Our analysis of luteinization markers revealed that many of the markers that were affected in CC and GC are regulated by either estrogen-, FSH- or LH-signaling, (14, 14, and 13, luteinization markers, respectively) or all three of these signaling pathways (6 luteinization markers), 13 by progesterone, some by testosterone (6 luteinization markers) or oxytocin (3 luteinization markers), and one (PTGS2) that is regulated by all of these hormones (Figure 10A, Column 5). To explore potential endocrine regulation of these and other DEGs across cell and follicle types, we examined expression levels of mRNAs encoding the relevant hormone receptors (Figure 12A). Estrogen receptor, ESR1 mRNA trended toward reduced expression and ESR2 mRNA was significantly reduced in both CC (p=0.0036) and GC (p=0.0184) of Type 4 follicles. FSH receptor mRNA expression was significantly downregulated in Type 4 CC (p=0.028) and appeared to be variably diminished in Type 4 GC but failing to reach significance. 107 LH receptor mRNA was expressed more highly in GC than CC as expected, with a trend toward downregulation in Type 4 GC, although not statistically significant. Analysis of the number of DEGs potentially regulated by the different hormones (Figure 12B) revealed progressively more target DEGs across the series of Type 2, 3 and 4 follicles, consistent with the increasing number of total DEGs. The largest number of target DEGs in CC and GC was seen for estrogen, followed in order of magnitude, with fewer effects elicited by FSH, LH, oxytocin, progesterone, and testosterone. The majority of estrogen, FSH, LH and oxytocin effects were increases in DEG expression. Progesterone yielded an enhanced number of decreases in DEG expression for CC. Ingenuity Pathway Analysis (IPA) reveals affected pathways and processes and implicates relevant upstream regulators To identify the major canonical pathways (CPs) and biological functions (BFs) impacted by the transcriptome alterations in follicle Types 2-4 compared to Type 1 within each cell type, and to identify potential causative upstream regulators (URs) directing major components of the transcriptome changes, we subjected DEG lists to Ingenuity Pathway Analysis (IPA). Oocyte IPA analysis for oocytes was limited to the Type 4v1 comparison, as only this comparison yielded a sufficient number (283, FDR<0.01) of DEGs for analysis. IPA identified 13 affected CPs (Figure 13) [32] one of which (Role of PKR in Interferon Induction and Antiviral Response) was predicted to be activated (z = 2.45), and another (EIF2 Signaling) predicted to be inhibited (z = -2.33). Other prominent affected CPs included the unfolded protein response, endoplasmic reticulum (ER) stress pathway, mTOR signalling, protein ubiquitination, clathrin-mediated endocytosis, and the sumoylation pathway. IPA also identified 115 affected 108 BFs (Figure 13) [32], including one predicted as activated (regeneration of axons) and five identified as inhibited (phagocytosis, cell survival, cell viability of leukemia cells, engulfment of cells, proliferation of kidney cells). A total of 108 URs showed significant effects with four identified as activated (Figure 13) [32], including RICTOR, UDP-D-glucose, CGA and SGPP2. A further 11 URs were identified as being inhibited (z ≤ -1.96) including MYCN, MLXIPL, MYC, TCR, CD3, XBP1, ATF4, ELL2, FBXO32, β-estradiol, and IL3. Additional URs were identified as affected, four of which were reduced in expression (HSP90B1, HSPA5, FIGLA and TMBIM6). Cumulus Cells IPA Analysis for CC revealed 14 affected CPs for Type 2 CC, none of which were predicted as activated or inhibited (Figure 14A) [32]. For Type 3 CC, there were 15 affected CP (P<0.05) [32], of which three were activated (z ≥ 1.96) [32] and none were inhibited. For Type 4 CCs, 240 CP were significantly affected, with 152 activated (P<0.05; z ≥ 1.96) and five inhibited (z ≤ -1.96) [32]. There was considerable overlap in affected CP for Type 4 CC and Type 4 GC (Figure 14A) [32], echoing the other similarities described above for these two cell populations. Prominent overlaps included 17 activated CP (e.g., STAT3, ILK, integrin, fibrosis, RHOA, among others) and one prominent inhibited CP (RHOGDI signaling) shown in Figure 6A. Overall, there were 18 shared activated CP and 46 shared inhibited CP [32]. Despite the limited number of affected CP for Type 2 CC, there were 171 BF that showed significant effects relative to Type 1 CC (P<0.05). One BF, sensory system development, was activated (z = 1.97) [32] and none were inhibited. For Type 3 CC, there were 310 BF that showed significant effects (P<0.05) [32], with 52 identified as being activated (z ≥ 1.96) and 8 identified as being inhibited (z ≤ -1.96). For Type 4 CC there were 261 affected BF (P<0.05) 109 [32], of which 138 were activated (z ≥ 1.96) and nine were inhibited (z ≤ -1.96) [32]. There was considerable overlap between Types 3 and 4 CC and Types 3 and 4 GC for affected BF, with 54 shared across all four cell types (e.g., angiogenesis, migration of cells, organization of cytoskeleton, several entries related to cell morphology, and several entries related to survival), and 98 shared across Type 4 CC and Types 3 and 4 GC, which include many entries related to cell movement (Figure 14C) [32]. Additional BF were shared between Type 3 CC and Type 4 GC, further echoing the similarities between these cell populations. Apoptosis was predicted to be an inhibited BF in Types 3 and 4 CC and Type 4 GC (Figure 14C) [32]. For Type 2 CC, 100 URs showed significant effects (P<0.05) with two activated (z ≥ 1.96) [32] and none inhibited. For Type 3 CC, 522 URs were affected (P<0.05) with 79 identified as activated (z ≥ 1.96) [32] and four identified as inhibited (z ≤ -1.96). For Type 4 CC, 1,044 URs were affected (P<0.05) with 240 activated (z ≥ 1.96) [32] and 76 inhibited (z ≤ -1.96). Many of the 64 URs were significantly activated across Types 2-4 CC and Types 3 and 4 GC, and many of these were predicted to be activated. Two URs (STAT3, RAF1) were predicted to be activated across all CC and GC populations, and 40 were predicted to be activated across Types 3 and 4 CC and GC (e.g., CTNNB1, EGF, TGFB1, TNF, IGF1, VEGF, SMAD4) (Figure 14B) [32]. Beta-estradiol was affected across all five cell populations and predicted to be activated in Type 3 CC and Types 3 and 4 GC, with positive z-scores (but z < 1.96) for the other cell populations. Progesterone was classified as activated for Types 3 and 4 CC. Additional affected URs were shared between two luteinization markers (PTGS2 and HIF1A). PTGS2 was activated in Type 4 CC and GC and upregulated in Types 2 and 3 CC and Type 3 GC. HIF1A was activated in Types 3 and 4 CCs and GCs. Type 4 CCs and GCs, again echoing the similarities between these two cell populations described above. Two URs (EPAS1, TGFB1) 110 were themselves upregulated in Type 3 CC and Types 3 and 4 GC (Figure 14B) [32]. Others were upregulated (STAT3, KITLG, SMAD7, IGF2, TCF4) or downregulated (F2RL1, FANCC) in Type 3 CC and Type 4 GC (Figure 14B) [32]. Alpha-catenin was predicted to be inhibited across all Type 2-4 CC and GC populations. Granulosa Cells IPA analysis for GC yielded significant results for comparisons of Types 2-4 follicles to Type 1 follicles. Due to the limited number of mRNAs identified, IPA analysis was not performed for Type 2 GCs. For Type 3 GCs, there were 45 affected CPs (P<0.05), with 4 CPs identified as being activated (z ≥ 1.96; Figure 14A) [32] including ILK signaling, hepatic fibrosis signaling, TGF-β signaling and synaptogenesis signaling, and none were inhibited. For Type 4 follicles, 69 CPs were affected (P<0.05), with 23 activated (including STAT3, ILK, hepatic fibrosis, integrin, TGF-β, actin cytoskeleton, NF-kB, RHOA, PDGF, paxillin, spingosine-1- phosphate, and other signaling pathways) and one (RHOGDI signaling) inhibited (Figure 14A) [32]. There were 351 significantly affected BF (P<0.05) for Type 3 GC, with 102 identified as activated (z ≥ 1.96; Figure 14C) [32] and eight inhibited (z ≤ -1.96). For Type 4 GC, there were 263 BF that showed significant effects (P<0.05) [32], with 153 identified as activated (z ≥ 1.96) and 12 identified as inhibited (z ≤ -1.96). There was considerable overlap between Type 3 and Type 4 affected BFs (Figure 14C) [32], including activation of angiogenesis, cell migration, vasculogenesis, and various entries related to cell movement and cell/organismal survival, as well as inhibition of several entries related to inflammation. Type 4 GC additionally displayed inhibition of apoptosis, which was also affected in Type 3 GC but without a significant z-score. 111 Type 3 GC yielded 1,235 affected UR (P<0.05). Of these, 201 URs were identified as activated (z ≥ 1.96) [32] and a further 26 were identified as being inhibited (z ≤ -1.96). For Type 4 GC, 1,549 URs showed significant effects (P<0.05), 391 URs were activated (z ≥ 1.96) and 118 were inhibited (z ≤ 1.96) [32]. Activated URs included many prominent reproductive or ovarian factors such as β-estradiol, TGFβ1, STAT3, YAP1, and many others. There was once again considerable overlap between Types 3 and 4 GC for affected URs (Figure 14B) [32]. These included predicted activation of STAT3, RAF1, CTNNB1, TGFβ1, among many others, and inhibition of alpha catenin. A number of affected URs were themselves among the DEG lists, including STAT3, KITLG, SMAD7, IGF2, and TCF4 which were upregulated, and F2RL1, FANCC and MECP2 which were downregulated in Type 4 GC, and EPAS1 and TGFβ1 which were upregulated in Types 3 and 4 GC. SMAD7 downregulation contrasted with its predicted activation in Type 4 GC. FSH impacts major CPs and BFs downstream of β-estradiol and KIT ligand The above IPA analysis revealed significant effects on many CP, BF and UR. To connect these effects to actions of FSH, we used the IPA Path Explorer tool to connect identified URs from Type 3 and 4 CC and to identify additional URs that connect FSH stimulation to downstream effects on CP and BF in CC. This analysis identified a system of extended networks shared in both CC and GC along with an affected extended network in oocytes (Figure 15). In CC and GC, the predominant effect was initiated through FSH and the FSHR. Although IPA indicated predicted activation of LH signaling in GC, this could reflect instead the extensive overlaps in downstream actions of the two hormones. FSHR was downregulated in Type 4 cells, but its activity was predicted to be activated in both CC and GC (Figure 12 and 15) [32]. FSHR activation was connected to increased β-estradiol signalling and to increased expression and 112 activity of TFGβ1, EPAS1 and STAT3 in both Types 3 and 4 cells, and to inhibited activity of estrogen receptor in Type 4 cells. FSHR was also connected to increased KITL expression and activation in Type 4 cells. Additional actions were predicted to be mediated via significant changes (no z-score) in arrestins: ARRB1 (Type 4 GC and CC) and ARRB2 (Type 3 and 4 GCs) with increased expression of ARRB1 in Type 4 CCs. These effects on URs then worked via effects on increased and decreased DEG expression ultimately to elicit predicted increases in CPs and BFs (integrin signaling, cytoskeleton organization, STAT3 signaling, and IL1 signaling), along with reduced RHOGDI signalling and apoptosis in Type 4 CC and GC. The effects on cytoskeleton organization and apoptosis were also seen for Type 3 CC and GC. In Type 4 oocytes, the IPA extended network analysis highlighted effects initiated by a predicted decrease in β-estradiol signaling with reduced expression of HSPA5 and increased activation of MYC signaling. These URs impacted 62 DEGs that were associated with predicted downstream effects including reduced cell survival, phagocytosis and EIF2 signaling (Figure 15). Assessment of possible atretic state in Type 4 follicles One possible impact of excessive FSH could be to induce follicular atresia. To assess whether this is occurring, a previously defined set of bovine ovarian follicular atresia markers totaling 197 were downloaded from a meta-analysis on bovine granulosa cells [55]. The reported 197 marker genes were filtered here to include only those with an average p-value less than 0.05 (n=183), and of these, 149 displayed detected expression in our dataset. These 149 genes were intersected with the DEG lists for both CCs and GCs, comparing follicle Types 2-4 to Type 1 [32]. Only 54 of the 149 were identified as DEGs in at least one of the six comparisons. Twenty were affected in both GC and CC for Type 4v1 and in the expected direction. Of the top 30 113 DEGs reported by [55], nine were not detected (NMB, OSAP, CRISPLD2, CLCA2,DEFB4, CD5L, SWAP70, DEFBS, FABP4), five displayed expected increased expression in both CCs and GCs, (TRIB1, SH3RF1, CKB, ETS2, PLAT), four displayed expected decreased expression (CALB2, SUSD4, TRC1D8, TOX2), three were affected only in GCs (FOLR2, GSDMB, CTSS), six were unaffected (INHBA, GRB14, NAPIL5, CITED1, LCRRC17, FDFT1, IL1A) and one displayed the opposite to expected effect (GSTA5). Effects of cpFSH dose on genes related to progesterone synthesis and oxytocin signaling A prominent feature of the cpFSH effect is an increase in follicular levels of progesterone and oxytocin, particularly in Type 3 and 4 follicles. Relative to Type 1 follicles, we observed increased expression of mRNAs encoding STAR in CC from Type 4 follicles and GC from Type 3 and 4 follicles, and HSD3B1 in CC from Type 4 follicles [32]. We also noted significant positive IPA results for ‘Steroidogenesis of hormone’ and ‘Synthesis of steroid’ in Type 3 CCs and GC, with a significant positive z-score for Type 3 CCs [32]. Likewise, relative to Type 1 follicles expression of the mRNA encoding oxytocin, OXT, was increased in CC from Type 4 follicles and GC from Type 3 and 4 follicles [32]. We did not observe an increase in oxytocin receptor mRNA expression in CC or GC. Discussion The main discovery from this study is that excessive cpFSH doses (3-fold above industry standard), associated with a premature luteinization phenotype (based on intrafollicular steroid hormone concentrations and COC morphology [24]) in most ovulatory size follicles, lead to progressively more severe changes in the transcriptomes of CC and GC as follicular abnormalities increased. These two cell types become more alike with the severity of follicular abnormality, while also diverging substantially from their normal phenotypes seen in follicles 114 from animals receiving the industry standard cpFSH dose. These changes are accompanied by less severe changes in the oocyte transcriptome, however the oocyte transcriptome changes suggest significant alterations in cellular physiology that may compromise oocyte quality. We recognize that oocytes execute major changes in their proteomes via temporally regulated translation and degradation of stored mRNAs as well as post-translational processes. However, observed changes in the relative abundances in abnormal follicles, particularly deficiencies in essential mRNAs or mRNAs related to essential oocyte processes can reasonably be interpreted as indicating potential compromise in oocyte quality, either through deficient mRNA content or through abnormal regulation of stabilization/destabilization impacting the capacity to produce key proteins when needed. Thus, although oocytes from Type 2 and 3 follicles display very minor transcriptome changes, the more extensive changes exhibited in oocytes from Type 4 follicles indicate a potential compromise in oocyte quality. Further studies to assess oocyte quality would be valuable, but given complexity and cost are beyond the scope of this report. Overall, about half (Types 3 and 4) of the ovulatory-size follicles in animals receiving the excessive cpFSH dose display substantial transcriptome changes, which may reflect loss of FSH sensitivity during folliculogenesis, for example through downregulation of FSH receptor signalling pathways. Such a loss of sensitivity would disrupt gene regulation, leading to the previously reported reduction in ovarian function during ovarian stimulation with excessive cpFSH doses in the SORH model [3, 24]. Moreover, our analysis indicates that the orderly progression of cellular and molecular changes typical of the final stages of folliculogenesis are severely disrupted, with aspects of ovulation, inhibited apoptosis and stress, and partial luteinization occurring together, and are associated with compromised oocyte health in the most severely affected follicles. Rather than simply reflecting a shift in the intrafollicular endocrine 115 milieu characteristic of premature luteinization, the transcriptome data indicate even more drastic alterations in ovulatory follicle function that do not compare readily to the normal follicular developmental progression. Collectively, these observations indicate that excessive cpFSH doses lead to follicular hyperstimulation dysgenesis negatively impacting ovulatory follicle function and oocyte viability. Previous studies in cattle found that prolonged FSH doses led to altered GC transcriptomes [56, 57]. The results here greatly extend our understanding of adverse effects of excess cpFSH on ovarian cells, and further indicate a significant risk of using FSH doses greater than is necessary to achieve a maximum response during ovarian stimulation protocols (i.e., excessive doses), with the potential to reduce desired ART outcomes by diminishing ovarian function and compromising oocyte quality. The mechanisms by which excessive FSH doses cause ovarian follicle dysfunction resulting in poor oocyte quality and ART outcomes have been unknown. Results from an earlier meta-analysis revealed a novel set of follicular cell biomarker genes that could be predictive of oocyte quality, and suggested that correct synchrony of the developmental transition in gene expression amongst the CC, GC and oocytes may be disrupted by high FSH doses, which could negatively impact oocyte quality [58]. Another meta-analysis of array studies examining developmental GC or CC transcriptome changes and oocyte transcriptome changes associated with FSH coasting also indicate effects on oocyte quality [59]. Using the SORH model, key changes in the different cell types that comprise an ovulatory-size follicle (CC, GC, and oocytes) following ovarian stimulation with an excessive 210 IU cpFSH dose were evaluated in detail using RNA-seq and provide new mechanistic insights into the effects of excessive FSH doses during ovarian stimulation. 116 Building on our previous work characterizing ovulatory-sized follicles according to follicular fluid E:P ratios and COC morphology [24], we show herein, that the increasing severity in follicle phenotypes with excessive cpFSH doses in the SORH model (i.e., progressing from Type 2 to 4) was accompanied by a similarly intensifying disruption in transcriptomes. This was particularly evident in the somatic (CC and GC) cells of the follicle. We also observed a significant downregulation of FSHR mRNA in CC and a non-significant decrease in GC of Type 4 follicles, indicating a potential desensitization of these follicles to FSH. This suggests either an intrinsic difference in the capacity to respond to cpFSH stimulation [60] or that individual follicles were exposed to different amounts of cpFSH as a result of differential vascular supply [61,62]. Either possibility may explain the observed variation in EA and EI states in follicles from the same heifer, given that β-estradiol and progesterone production by GC are induced by FSH [63]. Additionally, high cpFSH doses were associated with predicted disruptions in cell-cell communication and cellular physiology that may reduce oocyte quality. Overall, these data indicate that excessive FSH doses during ovarian stimulation result in varying degrees of abnormality in most ovulatory-size follicles, with significant negative impacts on key molecular and cellular processes. Antrum formation is typically associated with divergence of CC and GC phenotypes and function, as well as spatial separation within the follicle, with CC, but not GC remaining in direct contact with the oocyte [64-66]. The AFC for experimental animals in our study was monitored by ultrasonography prior to and throughout the ovarian stimulation period. We observed that the majority of follicles responding to stimulation with either cpFSH dose were antral follicles (≥3mm) present on the ovary prior to initiation of the ovarian stimulation protocol. Thus, the divergence of CC and GC prior to ovarian stimulation is inferred. With the high cpFSH dose, CC 117 and GC transcriptomes became progressively more different from normal transcriptome profiles, and became more alike with greater follicular abnormality, a relationship reflected in the markers of luteinization and in the IPA results, where there were a large proportion of shared URs and BFs. For example, in the luteinization markers analysis, the transcriptomes of CC from Type 3, and GC from Type 3 and 4 follicles, were similar to each other but distinct from the control Type 1 CC or GC. This change signifies an increasingly severe and profound disruption in follicular biology with high cpFSH doses. Several possibilities could explain such a change including, but not limited to, atresia, premature luteinization or differentiation into an unidentified cell type. The features of EI (e.g., Type 4) follicles are generally thought to reflect atresia [25, 26, 67]. The limited effects observed on the expression of bovine atresia marker genes could be interpreted as indicating a mild early atretic phenotype. However, the IPA analyses indicated that apoptosis and necrosis, both hallmarks of atresia, were inhibited in CC and GC of Type 3 and 4 follicles, and thus do not support this interpretation. Instead, IPA revealed a novel and complex phenotype, wherein increasing severity of follicle abnormality was accompanied by activation of URs (e.g. EGF, TGFB1, TNF) known to influence GC differentiation and proliferation [68-70] and numerous changes indicative of tissue remodelling (for example, angiogenesis, cell movement/migration, proliferation and differentiation of cells). Many of these are hallmarks of the complex and tightly regulated changes that occur in ovarian follicles in preparation for ovulation and resultant luteinization [71]. We presented evidence that the phenotypic differences (eCOC and follicular fluid E:P ratio) were indicative of premature luteinization, particularly in the Type 4 follicles [24]. However, premature luteinization refers to mostly clinical observations following ovarian stimulation with high FSH doses, where peripheral E:P concentrations were shifted favoring 118 progesterone production and suggesting that follicles were undergoing luteinization without an ovulatory (LH- or hCG) stimulus [11-13, 72]. Key steroidogenic enzymes involved in progesterone synthesis were also upregulated including STAR (increased in Type 4 CCs and GCs) and HSD3B1 (increased in Type 4 GCs). Examining a manually curated list of luteinization markers [44-51] revealed that a majority of luteinization markers were not affected in any cell type from any of the follicle types. These observations indicate that premature luteinization in this system is not due to a simple conversion of GC or CC to a corpus luteum- like state. Rather, both GC and CC progressively deviate toward a cell state characterised by a loss of cell-cell communication within the follicle, as evidenced by the morphological expansion of the COC and shared upregulation of BF relating to changes in cell morphology (including branching of cells and organization of cytoskeleton) and cell migration (including migration of cells and cell movement). Moreover, many of these changes predicted by the IPA analysis, including key pro-differentiation URs shared by GC and CC in Type 3 and 4 follicles (depicted in Figure 14 & 15; e.g. TGFB1, EGF, TNF, EPAS1) and BF (including interleukin signaling, integrin signaling, re-organization of the cytoskeleton, angiogenesis and immune or inflammatory reponses), support the idea that molecular changes within these cells are associated with reorganization of the follicles characteristic of ovulation [68, 71, 73]. This suggests a temporal dysregulation of the final stages of folliculogenesis, whereby a chaotic combination of processes characteristic of ovulation and luteinization occur concurrently, rather than sequentially. This may explain the increased incidence of anovulation observed in our previous study with the SORH model in response to ovarian stimulation with the excessive cpFSH dose [3]. 119 An unaddressed concern in the field of bovine ART has been the impact of LH- contamination present in the cpFSH preparations used for ovarian stimulation. Gonadotropin preparations with significant LH-activity or -contamination were utilized in many of the foundational studies leading to the development of ovarian stimulation in cattle (e.g. pregnant mare serum gonadotropin; PMSG [74]) and women (e.g. human pituitary gonadotropin; hPG [75]). Despite widespread use of recombinant hFSH in human ART, commercially available recombinant bovine equivalents are not currently available. Using the SORH model and Folltropin-V, a commonly used cpFSH preparation, we’ve shown the LH-contamination contained in this preparation was not sufficient to increase circulating progesterone concentrations or induce premature ovulation, even in excessive dose treated animals [3]. The only differences in gonadotropin receptor expression were observed in the CC where FSHR expression in Type 4 follicles was reduced, a pattern also observed in the GC, implicating FSH signalling as the major driver for differences in follicle types. Moreover, the convergence of CC and GC transcriptomes suggests excessive cpFSH, mimics some (but not all) aspects of LH signalling in a subset (Type 3 and 4) of follicles. As reflected in the expanded network analysis, the downstream impacts of FSH and LH on CC and GC are inextricably linked. For instance, EGF-like factors, key intermediaries of LH signalling in the follicle and with key roles inducing CC expansion and oocyte maturation, are also produced in response to FSH [76-78]. In in vitro maturation experiments with recombinant FSH, LH was still identified as a key UR supporting the argument that in vitro FSH may induce signalling events in COC similar to those observed in response to LH on in vivo follicle development [79, 80]. Based on the data herein, it could also be argued that excessive FSH doses in vivo also induce changes in a proportion of follicles that previously have been attributable to LH. 120 Despite the oocyte yielding the fewest DEGs, IPA analysis of the Type 4v1 oocyte data indicated potentially deleterious changes affecting key processes relating to oocyte function during maturation. In contrast to the CC and GC, the activity of the key UR, β-estradiol (reduced in follicular fluid in Type 4 follicles) appeared to be inhibited in the oocyte. This was accompanied by predicted inhibition of cell survival, phagocytosis and EIF2 signaling. Although estrogen signalling is not essential for oocyte maturation in vivo [81], estradiol and oocyte secreted factors have indispensable roles in maintaining the CC phenotype and function [66, 82]. Whether loss of CC communication with the oocyte preceded and/or induced the reported changes in oocyte function could not be determined. The EIF2 signalling pathway has a role during oocyte nuclear maturation, particularly at the germinal vesicle to metaphase II transition during meiosis [83, 84]. Thus, reduced EIF2 signaling may be indicative of compromised ER stress and unfolded protein response pathways, increasing oocyte susceptibility to stressors such as oxidative stress [85, 86]. Likewise, both engulfment of cells and phagocytosis were down regulated in Type 4 oocytes, potentially signifying a reduced capacity for fertilization [87]. However, in combination with the reduced cell survival, the decreased phagocytosis may instead reflect a shift in the balance between apoptosis and autophagy, known to be characteristic of oocyte cell death, reflecting an emerging atretic state [88-90]. This study has demonstrated that ovarian stimulation with an excessive cpFSH dose resulted in a convergence of CC and GC transcriptomes as phenotypic follicle abnormalities increased. We cannot state whether CCs became more GC-like or vice versa, or whether both cell types acquired a different abnormal phenotype as follicular abnormalities increased. Moreover, it does not appear that either GCs or CCs simply convert to a corpus luteum-like state, as would be expected for simple premature luteinization. Instead, it appears that the changes in 121 GCs and CCs reflect an overall loss of cell-cell communication within the follicle that resulted in the dysregulation of maturational processes in both CC and GC, associated with altered oocyte functions that could compromise oocyte quality. Together with our previous observations of reduced ovulation rates in SORH heifers treated with an excessive cpFSH dose during ovarian stimulation, and the fact that Type 3 and 4 follicles represented ~50% of ovulatory-sized follicles produced, the dysfunction observed in the CC, GC and the oocyte likely impair ovulation and would likely result in poor ART outcomes due to reduced oocyte quality. 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Panel A) Each plot displays DESeq2 results [-log10(FDR)], DEGs were called at a significance threshold greater than 2.0 (FDR<0.01), denoted by horizontal dashed line. Point color key: Black = non-DEG, Red = up- regulated DEGs, Blue = down-regulated DEGs. Panel B) Summary of DEG overlaps for each cell type comparing follicle types 2-4 versus type 1: oocyte, granulosa cell, and cumulus cell. 131 Figure 10. Overview of gene expression changes for manually curated luteinization markers identified from the literature. Panel A consists of five columns exploring the changes in luteinization marker expression in cumulus and granulosa cell samples. Panel A column 1 depicts the expected change in expression during luteinization, wherein red fill denotes an increase in expression, blue a decrease. Panel A columns 2 and 3 depict the changes in expression of markers in cumulus and granulosa cells, respectively, from different follicle types, fill of any color denotes 132 Figure 10 (cont’d) log2(fold-change), red equating to an increase and blue a decrease in expression. Panel A column 4 depicts the difference in marker expression when comparing cumulus and granulosa cells within follicle type; yellow fill denotes higher expression in cumulus cells and green higher expression in granulosa cells. Panel A column 5 identifies which of the markers are targets of hormones per the IPA database. For Panel A columns 2-4, DEG status is denoted by ** = FDR<0.01 and * = FDR<0.05. Panel B consists of a correlation heatmap with hierarchical clustering based on the average FPKM of the luteinization markers for both cumulus and granulosa cells among the four follicle types. Red fill indicates a higher correlation of expression values, while blue denotes less correlation. CC = cumulus cell and GC = granulosa cell. 133 Figure 11. Volcano plots comparing DEGs in cumulus and granulosa cells for each follicle type (1-4) and correlation heatmap with applied hierarchical clustering of expression values for cumulus and granulosa cells. For Panel A, each plot displays DESeq2 results [-log10(FDR)) and represents the DEGs in the cumulus and granulosa cells within a follicle type. DEGs were called at a significance threshold greater than 2.0 (FDR<0.01), denoted by horizontal dashed line. Point color key: Black = non-DEG, Red = up-regulated DEGs, Blue = down-regulated DEGs. CC = cumulus cells and GC = granulosa cells. Panel B) Within cumulus and granulosa cell types, FPKM values for follicle Types 2-4 were normalized against those for Type 1 as the denominator. The correlation between the types were calculated via the Pearson method. Color denotes correlation coefficient: red equating to more similar, and blue less similar. 134 Figure 12. Expression of hormone receptor mRNAs and the number of DEGs potentially affected by hormones. Panel A) Boxplots portray the FPKM values of ESR1, ESR2, FSHR, and LHCGR for each cell and follicle type. The x-axis denotes follicle Types 1-4 and the y-axis denotes the FPKM values for each sample. Significant differences between Types, within cell type were overlayed from the DESeq2 analysis. Horizontal lines denote those comparisons with an adjusted p-value below 0.05. Panel B) Barplots show the number of DEGs targeted by each hormone indicated, as predicted by the IPA database, for each cell type and for each comparison indicated on the x-axis. The y-axis quantifies the number of DEGs, with the color denoting the direction of regulation: red = up-regulated, blue = down-regulated, bifurcated by horizontal line at y=0. 135 Figure 13. Oocyte IPA pathway, regulator, and function heatmap. Heatmaps depict significantly affected canonical pathways, biological functions, and upstream regulators for the follicle Type 4v1 comparison in oocytes. Colored fill for cells in heat maps denote z-score: red = activated, blue = inhibited, black = no significant z-score. Within the Upstream Regulator panel, the circles within colored fills denote regulators that are also differentially expressed: blue = down- regulated. 136 Figure 14. Cumulus and granulosa cell IPA pathway, regulator, and function heatmap. Heatmap depicts IPA canonical pathways (Panel A), upstream regulators (Panel B), and biological functions (Panel C) for cumulus and granulosa cells. All panels consist of two columns (wrapped in two portions for panels B and C): left column denotes dataset of origin with the cell type and follicle type comparison indicated, right column (“LM”) denotes the number of luteinization markers (Figure 2) present in the pathway. Along the y-axis are the identified canonical pathways. 137 Figure 14 (cont’d) Colored fill for cells in heat maps denote z-score: red = activated, blue = inhibited, black = no significant z-score. Level of significant overlap was set at P<0.05, and cells with white fill denote those not meeting significance. Panel A) Entries were limited to those with significance in at least three datasets and/or significant z-scores in at least two. Panel B) Entries were limited to those with significance in at least four datasets and/or significant z-scores in at least three datasets. Panel C) Entries were limited to those with significance in at least five datasets and/or significant z- scores in at least four datasets. 138 Figure 15. Expanded network analysis of IPA results. Custom expanded network figure linking significant upstream regulators, their downstream DEG targets, and enriched pathways and functions, derived from the IPA software. Figure is split into two panels: left panel contains joint information for granulosa and cumulus cell (3v1 and 4v1, comparisons), right panel contains oocyte (4v1 comparison). Each panel is further divided into three sections: 1) Upstream regulators, 2) Downstream DEGs, and 3) Functions. Section one contains upstream regulators, connected in a network, which were identified with significant z-scores, p-values of overlap, and/or were differentially expressed. The Downstream DEG section tabulates the number of DEGs for each cell type, bifurcated by direction of regulation, and source of comparison. The Function section identifies the pathways and functions significantly enriched by the above DEGs and upstream regulators. For the granulosa and cumulus cell section, superscripts denote which comparison of origin: 3=3v1, 4=4v1. For both vertical sections, outer border colors for molecules denote significant predicted z-score (|z| > 1.96): blue=inhibited, red=activated. Interior solid coloring denotes molecules for which mRNAs are differentially expressed (FDR<0.05): blue=down- regulated, red=up-regulated. Outer borders in solid black indicate a significant effect without z score, while dashed outer borders denote instances of non-significant p-values. Left and right portions of borders and inner fills denote results for granulosa and cumulus cells, respectively. The “?” symbol by LH indicates that, although IPA suggests LH as a possible affected UR, there is substantial overlap between FSH and LH signaling (about 2/3 of pathway members). 139 Ovarian Stimulation with Excessive FSH Doses Causes Cumulus Cell and Oocyte Dysfunction in Small Ovarian Reserve Heifers1 CHAPTER 5. 1 Kaitlin R. Karl, Peter Z. Schall, Zaramasina L. Clark, Meghan L. Ruebel, Jose Cibelli, Robert J. Tempelman, Keith E. Latham, James J. Ireland. “Ovarian stimulation of the small ovarian reserve heifer with excessive FSH doses causes cumulus cell and oocyte dysfunction.” Molecular Human Reproduction, 29(10), (2023), gaad033. By permission of Oxford University Press 140 Title: Ovarian stimulation with excessive FSH doses causes cumulus cell and oocyte dysfunction in small ovarian reserve heifers Running Title: Excessive FSH dysregulates cumulus function and oocyte maturation Keywords: small ovarian reserve, excessive FSH, superovulation, oocyte pick-up, cumulus- oocyte complexes, dysregulation of cumulus function, FSH target genes in cumulus cells, premature cumulus expansion, resumption of meiosis Authors and affiliations: Kaitlin R. Karl1, Peter Z. Schall1,2, Zaramasina L. Clark1,3, Meghan L. Ruebel1,4, Jose Cibelli1,5, Robert J. Tempelman1, Keith E. Latham1,6 and James J. Ireland1* 1Reproductive and Developmental Sciences Program, Department of Animal Science, Michigan State University, East Lansing, MI, USA; 2Present affiliation: University of Michigan, Ann Arbor, MI, USA; 3Present affiliation: School of Biological Sciences, Victoria University of Wellington, Wellington, New Zealand; 4Present affiliation: USDA-ARS, Southeast Area, Arkansas Children’s Nutrition Center, Little Rock, AR, USA, 5Department of Large Animal Clinical Sciences, Michigan State University, East Lansing, MI, USA, 6Department of Obstetrics, Gynecology and Reproductive Science, Michigan State University, East Lansing, MI, USA *Correspondence: James J. Ireland, Molecular Reproductive Endocrinology Laboratory, Department of Animal Science, Michigan State University, East Lansing, Michigan, 48824, USA. Tel.+1 517 290 7074. Email: ireland@msu.edu, https://orcid.org/0000-0002-5923-8000 Funding: This study was supported by the Agriculture and Food Research Initiative Competitive USDA-NIH Dual Purpose Program Grant no. 2017-67015-26084 from the USDA National Institute of Food and Agriculture, the Eunice Kennedy Shriver National Institute of Child Health & Human Development of the National Institutes of Health under Award Number T32HD087166, MSU AgBioResearch, and Michigan State University. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. 141 Abstract Excessive FSH doses during ovarian stimulation in the small ovarian reserve heifer (SORH) cause premature cumulus expansion and follicular hyperstimulation dysgenesis (FHD) in nearly all ovulatory-size follicles with predicted disruptions in cell-signaling pathways in cumulus cells and oocytes (before ovulatory hCG stimulation). These observations support the hypothesis that excessive FSH dysregulates cumulus cell function and oocyte maturation. To test this hypothesis, we determined whether excessive FSH-induced differentially expressed genes (DEGs) in cumulus cells identified in our previously published transcriptome analysis were altered independent of extreme phenotypic differences observed amongst ovulatory-size follicles and assessed predicted roles of these DEGs in cumulus and oocyte biology. We also determined if excessive FSH alters cumulus cell morphology, and oocyte nuclear maturation before (premature) or after an ovulatory hCG stimulus or during in vitro maturation (IVM). Excessive FSH doses increased expression of 17 cumulus DEGs with known roles in cumulus cell and oocyte functions (responsiveness to gonadotropins, survival, expansion, and oocyte maturation). Excessive FSH also induced premature cumulus expansion and oocyte maturation but inhibited cumulus expansion and oocyte maturation post-hCG and diminished the ability of oocytes with prematurely expanded cumulus cells to undergo in vitro fertilization (IVF) or nuclear maturation during IVM. Ovarian stimulation with excessive FSH is concluded to disrupt cumulus cell and oocyte functions by inducing premature cumulus expansion and dysregulating oocyte maturation without an ovulatory hCG stimulus yielding poor-quality cumulus-oocyte complexes (COCs) that may be incorrectly judged morphologically as suitable for IVF during ART. 142 Introduction Diagnosis of a small ovarian reserve (total number of morphologically healthy follicles/oocytes in ovaries) is the primary reason women seek assisted reproductive technologies (ART) [1]. However, many women with small ovarian reserves respond poorly to treatments with follicle stimulating hormone (FSH) during ovarian stimulation protocols [2-4]. Moreover, total FSH doses during ovarian stimulation protocols vary from <1,000 IU to 20,000 IU [2, 4], and high FSH doses during ovarian stimulation are inversely linked to ovarian function, oocyte recovery, and live birth rate during ART in women [2, 4-11] and ovarian function and embryo transfer outcomes in cattle [12-18]. Whether excessive FSH doses during ovarian stimulation cause or contribute to poor ART outcomes, and the mechanisms responsible for such an effect are unknown. To better understand the impact of high FSH doses during ovarian stimulation on ovulatory follicle function, oocyte quality, and associated mechanisms, we developed the small ovarian reserve heifer (SORH) model. This unique model has biomedical relevance because it exhibits not only a relatively low number of morphologically healthy oocytes and a low antral follicle count (AFC) [19] but diminished circulating concentrations of anti-Műllerian hormone [20], progesterone and estradiol, hypersecretion of FSH [19, 21, 22] during reproductive cycles, and poor responsiveness to superovulation [19] like women with a small ovarian reserve. Although recombinant FSH is not available for use in cattle, ovarian stimulation of the SORH model with different doses of Folltropin-V, a commercial FSH-enriched porcine pituitary preparation (cpFSH) with minimal LH contamination (<1%), has provided new insights into the potential detrimental impact of high FSH doses on ovulatory follicle function. For example, we established that cpFSH doses only 3-fold higher than the industry standard are excessive and thus 143 economically wasteful because they do not increase number or size of ovulatory-size follicles compared with lower doses [23]. In addition, excessive cpFSH (Ex-cpFSH) doses are detrimental to ovulatory follicle function because they decrease both circulating estradiol concentrations and responsiveness of ovulatory-size follicles to hCG, thereby reducing ovulation rate [23]. Using the SORH model, we also established that ovarian stimulation with the Ex-cpFSH doses results in premature cumulus cell expansion and luteinization (as measured by low intrafollicular estradiol but high progesterone and oxytocin concentrations) prior to an ovulatory hCG stimulus in a large proportion of the ovulatory-size follicles [24]. Furthermore, based on whole transcriptome analysis data, we concluded that Ex-cpFSH doses during ovarian stimulation cause follicular hyperstimulation dysgenesis (FHD) in all ovulatory-size follicles [25]. Severe abnormalities in multiple cell-signaling pathways in granulosa and cumulus cells and oocytes critical for folliculogenesis, steroidogenesis, luteinization, cell survival, ovulation, and oocyte maturation and quality characterize this disorder. Consequently, although the precise mechanisms remain unclear, FHD likely causes or contributes to the inhibition of ovulatory-size follicle growth, reduction in estradiol production, promotion of premature cumulus expansion and luteinization, and diminution of ovulation rate in response to hCG, as we observed during ovarian stimulation of the SORH model with Ex-cpFSH [23-25]. Our observations support the hypothesis that Ex-cpFSH doses during ovarian stimulation dysregulate cumulus function and oocyte maturation. We tested our hypothesis here using the SORH model in two ways. First, we determined if Ex-cpFSH doses during ovarian stimulation resulted in dysregulation of cumulus cell genes critical for function and regulation of oocyte maturation. This was accomplished by manually interrogating our published whole transcriptome data set to determine whether any of the 144 previously identified Ex-cpFSH-induced cumulus cell DEGs [25] are altered in all ovulatory-size follicles independent of individual follicular phenotypic differences, and whether these DEGs have known roles in cumulus cell function and oocyte maturation. We reasoned that cpFSH- induced overexpression of cumulus genes in all ovulatory-size follicles could explain the premature cumulus expansion observed in nearly all ovulatory-size follicles developing in response to Ex-cpFSH doses in our studies [24, 25]. Second, we tested whether the Ex-cpFSH- induced premature cumulus expansion observed in our studies [24, 25] is accompanied by altered oocyte nuclear maturation before (premature) or after an ovulatory hCG stimulus, or during in vitro maturation (IVM). Materials and Methods Analysis of Ex-cpFSH-induced differentially expressed cumulus genes and their predicted roles in regulating cumulus cell function and oocyte maturation In our previously published study [25], we analysed by RNA sequencing four ovulatory- size follicle Types that had been identified based on FSH dose, COC morphology (expanded, compact), and differences in intrafollicular concentrations of estradiol, progesterone and oxytocin [25]. Type 1 ovulatory-size follicles were those from animals receiving the industry standard dose and mimic healthy ovulatory follicles during estrous cycles in cattle [26, 27], having compact cumulus-oocyte complexes (comCOCs), higher intrafollicular estradiol than progesterone concentrations, and relatively low oxytocin concentrations [24, 25]. Types 2, 3, and 4 follicles were identified in Ex-cpFSH treated SORH [24, 25]. Type 2 phenotypically resembles Type 1. Type 3 is like Type 1 and 2 but has an expanded layer of cumulus cells (expCOCs). Type 4 also has expCOCs but has much higher progesterone than estradiol and the highest oxytocin concentrations. Through RNA sequencing, we identified 4576 DEGs (characteristic of 145 the FHD phenotype) in granulosa and cumulus cells and oocytes of the Ex-cpFSH-induced Type 2, 3 or 4 compared with the industry-standard cpFSH-induced Type 1 (control) ovulatory-size follicles. Of these 4576 DEGs, 3288 were observed in cumulus cells of the Type 2, 3 or 4 ovulatory-size follicles [25]. In the present study (Fig. 16A), we manually interrogated each of these cumulus cell DEGs to identify those that were common to all three Ex-cpFSH ovulatory- size follicle types (Type 2, 3 and 4) and determined based on a literature search whether these DEGs have known roles in regulation of cumulus function and oocyte maturation (Fig. 16A). Effect of Ex-cpFSH on cumulus expansion and oocyte maturation We conducted three experiments using the SORH model (Fig. 16B). The first experiment (Exp 1, Fig. 16B) determined whether Ex-cpFSH doses during ovarian stimulation altered cumulus morphology and stage or timing of oocyte nuclear maturation. Each ovulatory-size (≥10 mm in diameter) follicle for each heifer (n = 10) was subjected to OPU 12 h after the last cpFSH dose (no hCG stimulus given) and each COC was classified morphologically as comCOCs or expCOCs, and stage of nuclear maturation attained for oocytes within each COC was determined. Exp 2 (Fig. 16B) determined whether Ex-cpFSH treatment during ovarian stimulation altered the capacity of cumulus cells and oocytes in ovulatory-size follicles to respond to an ovulatory hCG stimulus. Heifers were injected with an ovulatory dose of hCG 12 h after the last cpFSH injection. Each ovulatory-size follicle for each heifer (n = 16) was subjected to OPU 24 h post-hCG but prior to ovulation. The responsiveness of cumulus cells and oocytes to the ovulatory hCG stimulus was then evaluated by morphological classification of each COC as comCOCs or expCOCs, and determination of nuclear maturation stage attained for the oocyte within each COC. Exp 3 (Fig. 16B) determined if the Ex-cpFSH doses during ovarian stimulation altered the capacity of expCOCs to resume meiosis during in vitro maturation (IVM) 146 and their ability to be fertilized by IVF. Each ovulatory-size follicle for each heifer (n = 18) was subjected to OPU 12 h after the last cpFSH (no ovulatory hCG stimulus given) and each recovered COC was classified morphologically as comCOCs or expCOCs. The expCOCs from the Ex-cpFSH treated heifers (n = 8) were subjected to IVF and compared to comCOCs from abattoir ovaries that were also subjected to IVF. Additionally, a subset of Ex-cpFSH treated heifers (n = 10) was used to assess nuclear maturation stages of expCOCs vs comCOCs after IVM (Fig. 16B). Reagents Unless otherwise mentioned, chemicals and reagents were purchased from Merck KGaA (Darmstadt, Germany). Identification of heifers with a small ovarian reserve We previously established that 11- to 12-month-old heifers with a low antral follicle count (AFC, ≤15 follicles of ≥3 mm in diameter) during ovarian follicular waves (15% to 20% of a herd) also have 80% smaller ovarian reserves (total number morphologically healthy follicles and oocytes in ovaries) compared with age-matched counterparts with a high AFC (≥ 25 follicles) [19, 20, 22, 28-30]. In the present study, serial ovarian ultrasonography was used to identify 11- to 12-month-old Holstein heifers of similar weights with a low AFC and small ovarian reserve [23] for ovarian stimulation. Ovarian stimulation protocol To synchronize estrous cycles for ovarian stimulation, heifers received an initial 2 ml intramuscular injection of prostaglandin-F2α (PG, 12.5 mg dinoprost/ml, Lutalyse HighCon, Zoetis, Parsipanny, NJ, USA) followed by two additional PG injections 12 h apart 10 d later. Each heifer underwent daily ovarian ultrasonography to detect ovulation and emergence of the 147 first follicular wave. The 1st injection of Folltropin-V (porcine pituitary extract containing primarily FSH with 0.25% LH contamination (cpFSH), Vetoquinol USA Inc., Fort Worth, TX, USA) began 36 h after the last PG injection which was ±1 d from ovulation and initiation of the first follicular wave in all heifers. Heifers received 8 intramuscular injections of cpFSH (either 70 IU or 210 IU) at 12-h intervals. The cpFSH dose range per injection was 20% lower and 240% higher than the Vetoquinol recommended dose per injection of 87.5 IU. Hereafter, the 70 IU industry-standard cpFSH dose is referred to as the control while the 210 IU cpFSH is referred to as the Ex-cpFSH dose. Following superovulation, either two or three intramuscular PG injections were given 12 h apart starting at the time of the 7th cpFSH injection (about Day 4 or 5 of the estrous cycle) to regress the newly formed CL during each ovarian stimulation regimen. Oocyte pick-up (OPU) was then conducted 12 h after the last cpFSH and PG injections or 24 h after a single 2.5 ml (2,500 IU) intramuscular injection of human chorionic gonadotropin (hCG, Chorulon HCG 10,000 IU, Merck Animal Health USA, Rahway, NJ, USA) to induce oocyte maturation but prior to ovulation. Oocyte retrieval from ovulatory-size follicles For oocyte retrieval (or oocyte pick-up, OPU), heifers received caudal epidural anesthesia with lidocaine hydrochloride 2% (0.22 mg/kg; Lidocaine 2%, VetOne, Boise, ID, USA) mixed with xylazine hydrochloride 10% (0.025 mg/kg; AnaSed; VetOne, Boise, ID, USA) to minimize the stress of rectally manipulating ovaries for OPU. Follicles ≥ 10 mm were aspirated using a real-time B-mode ultrasound scanner (Ibex EVO; E.I. Medical Imaging, Loveland, CO, USA) equipped with an 8.0 MHz microconvex transducer housed in a plastic vaginal probe with a stainless-steel needle guide connected to the aspiration equipment. COCs were aspirated from each follicle using an 18-gauge x 3-inch disposable follicular aspiration needle (Partnar Animal 148 Health, Port Huron, MI, USA) connected to a Brazilian-style IVF tubing (Partnar Animal Health, Port Huron, MI, USA) and inserted into the stainless-steel needle guide. The contents of each follicle were aspirated into a 50 ml conical collection tube using an electric suction pump (K- MAR-5200, Cook Medical, Brisbane, Australia) at a variable negative pressure of 200 ± 1 mm Hg. Each 50 ml conical collection tube contained ~3 ml of medium, which consisted of protein- free chemically defined hamster embryo culture medium-6 (HECM [31]) and TALP-4-(2- hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES). The HECM-HEPES (HH) medium [32] was supplemented with 0.3% bovine serum albumin (BSA), 500 µM 3-isobutyl-1- methylxanthine (IBMX), and 100 nM C-type natriuretic peptide (NPPC). IBMX and NPPC were used to block oocyte maturation [33]. Before each OPU, the needle, IVF tubing, and 50 ml conical collection tubes were pre-coated with HH medium supplemented with 0.3% BSA and 5% polyvinyl propylene. Morphological classification of COCs To classify COCs, the contents of each 50 ml conical collection tube containing COCs were poured over an embryo filter (Hy-flow filter, SPI™, Canton, TX, USA) to isolate the COCs from each heifer. The filter was sprayed with HH medium to transfer COCs into a Petri dish. A stereomicroscope was used to classify each COC based on the number of cumulus cell layers [34], as reported by us for cattle subjected to ovarian stimulation [24]. Compact COCs (comCOCs) had one or more layers of cumulus cells surrounding the zona pellucida and oocyte whereas expanded COCs (expCOCs) had partially or totally expanded cumulus cells surrounding the zona pellucida and oocyte. Denuded oocytes devoid of cumulus cells were also recorded. 149 In vitro maturation When comCOCs or expCOCs were subjected to in vitro maturation (IVM), they were followed by four washes in drops of HH medium without IBMX and NPPC followed by four washes in the IVM medium. The IVM medium consisted of Medium 199 supplemented with 22 µg/ml sodium pyruvate, 4 IU/ml hCG (Chorulon; Merck & Co., Inc., Rahway, NJ, USA), 50 µg/ml gentamicin, and 100 µl/ml fetal bovine serum [32]. COCs were cultured in 10 µl drops of IVM medium in groups of 3 to 5 for 22 h at 38.5 °C in a 5% CO2 atmosphere with 100% humidity. To assess the nuclear maturation of COCs after IVM, our statistical power analysis limited us to groups of 3 to 5 COCs per drop from each heifer unless otherwise specified. However, a preliminary study was conducted to aspirate COCs from ovaries collected from cattle at a local abattoir (West Michigan Beef Co LLC, Hudsonville, MI, USA) to determine if 3 or 5 COCs per drop was representative of the proportion of oocytes reaching MII during IVM. Chi square analysis indicated that groups of 3 (n = 55) or 5 (n = 31) COCs per drop did indeed produce similar proportions of MII oocytes during IVM (mean ± SEM, 71 ± 2% vs 80 ± 6%, respectively P≥0.85). Data for heifers that had < 3 COCs recovered by OPU were not subjected to IVM and were omitted from statistical analysis. In vitro fertilization of expCOCs Unless specified, all in vitro fertilization (IVF) media were purchased from IVF Bioscience, Cornwall, UK. The expCOCs were incubated at 5% CO2 in air at 38.5° C in high humidity. All expCOCs were rinsed in HEPES-buffered HECM containing 1 mg/ml of bovine serum albumin (HH-BSA) [35] and placed in the incubator in a 44 µl drop of pre-equilibrated BO-IVF medium under mineral oil. Frozen sperm from a single high fertility bull (Lolo, 150 STgenetics, Navasota, TX, USA) was thawed at 38° C for 1 min and rinsed twice in BO- SEMENPREP medium using two consecutive 4 min centrifugations at 350 g at 22° C. After the final rinse, the sperm pellet was resuspended in BO-IVF at a final concentration of 2.0 x 106 sperm/ml in a 6 µl volume of the suspension, which was then added to 44 µl drops containing the expCOCs for 18 h. Subsequently, presumptive zygotes had their cumulus cells removed in 1 mg/ml of hyaluronidase in HH-BSA using a 140 µm internal diameter glass pipette. After three rinses in HH-BSA, embryos were incubated in BO-IVC medium for 7.5 d at 38.5° C. Classification of stages of nuclear maturation To determine nuclear maturation stages, oocytes were processed as reported [36]. The comCOCs and expCOCs were kept separate and washed three times in HH medium containing IBMX and NPPC and denuded by placing them into 2 ml microcentrifuge tubes containing 200 µl of 1% hyaluronidase solution at 37°C for 5 min followed by vortexing for 5 min. Denuded oocytes were then washed three times in HH medium containing IBMX and NPPC, transferred to four-well plates, and incubated for 15 min at room temperature in a permeabilization and fixation medium (6% Triton X-100 (v:v) in 3.7% paraformaldehyde solution (wt/vol)). The oocytes were washed three times again in Dulbecco’s Phosphate Buffered Saline (DPBS) without calcium and magnesium for 15 min at room temperature. Groups of ≤ 5 oocytes from comCOCs or expCOCs were transferred to 5 µl drops of VectaShield Plus Antifade Mounting Medium with 4',6-diamidine-2'-phenylindole dihydrochloride (DAPI; Vector Laboratories, Inc., Burlingame, CA, USA, H-2000-10) and mounted onto a glass slide under a coverslip. Oocytes were examined under an epifluorescence microscope to determine the stage of nuclear maturation [36, 37] (Fig. 17). Oocytes with intact germinal vesicles were classified as G0, G1, G2, and G3 as depicted (Fig. 17, Panels A, B, C, and D, respectively; described in Fig. 151 17 caption). The GV classifications were confirmed with anti-lamin A/C labeling as reported [36] (data not shown). The proportions of ovulatory-size follicles at the G0, G1, G2, or G3 nuclear stage were similar (P>0.05) independent of cpFSH dose (data not shown), and thus combined into a single nuclear stage, hereafter called germinal vesicle (GV). All other oocytes not in the GV stage were classified as germinal vesicle breakdown = GVBD, metaphase I = MI, or metaphase II = MII, as depicted (Fig. 17, Panels E, F and G, respectively). In addition, oocytes with no visible nuclear material or fragmented chromosomes were classified as degenerated. Statistical Analysis The R package DSeq2 [38] was used in our previous transcriptome study [25] to determine that 3288 genes in cumulus cells were significantly (FDR <0.01, except where noted as FDR<0.05 in Table 3) differentially expressed (DEGs) when Type 2, 3 and 4 ovulatory-size follicles were compared with Type 1 ovulatory-size follicles. In the present study, we manually interrogated each of these Ex-cpFSH-induced 3288 cumulus cell DEGs [24] to determine whether any of the cumulus DEGs were common to all ovulatory-size follicles independent of their individual phenotypes. For statistical comparisons where zero expression values are obtained for genes in some samples (see Fig. 18), DESeq2 uses a maximum likelihood estimate to assign very small values less than one to those entries. The maximum likelihood estimate reflects the maximum likelihood of detection with a small number of additional reads counted. We conducted three experiments (Exp) in the present study to determine if ovarian stimulation of the SORH model with Ex-cpFSH dysregulates cumulus function and oocyte nuclear maturation. Prior to data collection, a power analysis was conducted to determine the minimum number of animals and COCs per animal required to detect effects at a significance level of P≤0.05 at a power level of 0.80 using a balanced crossover design [39]. Analysis 152 determined that a minimum of 5 animals per dose (10 animals total) and a minimum of 3 to 5 COCs per animal would be sufficient to meet these criteria. To minimize animal numbers and confounding variables, a balanced crossover design was used. Each heifer acted as its own control, and each heifer was subjected to ovarian stimulation twice with a different cpFSH dose [70 IU (40mg/2ml) or 210 IU (120mg/6ml)] and with a different cpFSH dose sequence (e.g., 70 IU, 210 IU vs 210 IU, 70 IU) at the 1st and 2nd ovarian stimulation regimen [23]. One heifer in Exp 2 was hyper-responsive to both doses of cpFSH (70 IU and 210 IU; 60 and 70 ovulatory-size follicles, respectively). When normality and distribution were evaluated statistically, that heifer was a statistical outlier and thus removed from further analysis [40]. In Exp 3, several heifers from each cpFSH dose group were removed from statistical analysis because OPU resulted in < 3 COCs (control, 1 heifer removed; Ex- cpFSH, 4 heifers removed) as explained in Methods. The responses for Exp 1-3 were expressed as binomial proportions using total number of COCs analyzed as the denominator. The model statement for logistic regression analysis included dose of cpFSH, COC morphology, oocyte nuclear stage of maturation, and the associated interaction between these variables. Estimates were reported as the mean (±SEM) proportion of ovulatory follicles per heifer with comCOCs, expCOCs or denuded oocytes, or proportion of ovulatory-size follicles per heifer with comCOCs or expCOCs at GV, GVBD, MI, MII, or with degenerated oocytes. Unless specified otherwise, all statistical analyses were performed using Statistical Analysis System (SAS 9.4 Institute, Cary, NC, USA) PROC GLIMMIX and PROC LOGISTIC [40, 41]. When using PROC LOGISTIC, Firth’s Penalized Likelihood was also used for analyzing the stages of oocyte nuclear maturation data from each 153 Exp to mitigate non-convergence due to the presence of all-zero responses for a particular treatment within the dataset [42]. The cpFSH dose effects were considered significant if P≤0.05. Results Ex-cpFSH induces across all follicle phenotypes 17 DEGs that have predicted roles in regulation of cumulus cell function and oocyte maturation This analysis of our previously published data yielded 17 shared cumulus DEGs common to Type 2, 3 and 4 ovulatory-size follicles as depicted in Volcano (Fig. 18) and Box Whisker Plots (Fig. 19). These 17 Ex-cpFSH-induced cumulus cell DEGs all have well-established roles in modulation of FSH action and cumulus cell function including induction of cumulus cell expansion and regulation of oocyte maturation as indicated in Table 3. Ex-cpFSH did not increase number of ovulatory-size follicles or COC recovery The numbers of ovulatory-size follicles for each heifer were counted, and COCs were recovered 12 h after the last cpFSH injection in Exp 1 and 3 and 24 h after the ovulatory hCG stimulus in Exp 2. The number of ovulatory-size follicles, number of aspirated ovulatory-size follicles, number of COCs recovered, and COC recovery rate were similar (P>0.05) between heifers treated with the different cpFSH doses (Table 4). Ex-cpFSH induced premature cumulus expansion and resumption of meiosis prior to an ovulatory hCG stimulus The COCs recovered 12 h after the last cpFSH injection (Exps 1, 3) by OPU were classified as comCOCs, expCOCs, or denuded. Only comCOCs or denuded oocytes were recovered by OPU in the controls. In contrast, comCOCs, expCOCs, and denuded oocytes were recovered from heifers treated with Ex-cpFSH doses (Table 5, Exps 1, 3). The proportion of ovulatory-size follicles with expCOCs per heifer was higher (P<0.0001), while the proportion 154 with comCOCs was lower (P<0.01) in the Ex-cpFSH treated heifers compared with controls for Exp 1 and 3 (Table 6). The impact of the Ex-cpFSH treatment on nuclear maturation prior to the ovulatory hCG stimulus was only examined in Exp 1. Results show that 93% of ovulatory-size follicles in controls and 77% in the Ex-cpFSH treated heifers had comCOCs (Table 5, Exp 1). Of the comCOCs, 77% in controls and 97% in the Ex-cpFSH treated heifers were at the GV stage (Table 6). Although none of the comCOCs had resumed meiosis in controls, a very low proportion (2%) of the ovulatory-size follicles for the heifers treated with the Ex-cpFSH dose had comCOCs that had resumed meiosis (MI, MII, Table 6). However, the proportions of ovulatory-size follicles with comCOCs at MI or MII for the Ex-cpFSH treated heifers did not differ (P>0.05) from controls (Table 6). In contrast to controls, 22% of the ovulatory-size follicles had expCOCs (Table 5, Exp 1), and 49% of the expCOCs had resumed meiosis (GVBD+MI+MII) in the Ex-cpFSH treated heifers (Table 6). The remainder of the expCOCs in the Ex-cpFSH treated heifers were at the GV stage, had oocytes classified as degenerated or fragmented, or were lost during processing. Ex-cpFSH blocked cumulus expansion and resumption of meiosis in response to an ovulatory hCG stimulus In response to the ovulatory hCG stimulus, the proportion of ovulatory-size follicles per heifer with comCOCs was higher (P<0.01) while proportion with expCOCs was lower (P<0.05) for the heifers treated with Ex-cpFSH doses compared with controls (Table 5, Exp 2). Surprisingly, although prematurely expanded COCs (expCOCs, as observed in Exps 1, 3; see [24, 25] for pictures) could not be distinguished morphologically from expCOCs after the ovulatory hCG stimulus, the proportion of ovulatory-size follicles with expCOCs was nearly 155 identical to the results for the Ex-cpFSH treated heifers in Exps 1, 3 prior to an ovulatory hCG stimulus (Table 5). We observed that 71% of the expCOCs in controls and 65% of the expCOCs in the Ex- cpFSH treated heifers had resumed meiosis (GVBD+MI+MII) after the ovulatory hCG stimulus (Table 7). The Ex-cpFSH treated heifers had a higher (P<0.01) proportion of ovulatory-size follicles with expCOCs at MI but a lower (P<0.001) proportion at MII compared with controls. In addition, the Ex-cpFSH treated heifers had a lower (P<0.0001) proportion of ovulatory-size follicles with expCOCs at GV but a higher proportion at MI (P<0.01) and MII (P<0.001) compared with the comCOCs from the Ex-cpFSH treated heifers (Table 7). After the ovulatory hCG stimulus, we unexpectedly observed that 46% of the ovulatory- size follicles from controls and 72% of the ovulatory-size follicles from heifers treated with the Ex-cpFSH doses had comCOCs (Table 5). Moreover, only 1% of the ovulatory-size follicles with the comCOCs had resumed meiosis in the control compared with ~6% in the Ex-cpFSH treated heifers (Table 7). Like Exp 1, however, the proportion of ovulatory-size follicles with comCOCs at GVBD, MI, or MII were similar between treatment groups (Table 7). Ex-cpFSH-induced prematurely expanded COCs (expCOCs) responded poorly to IVF The proportion of expCOCs that cleaved and formed blastocysts or hatched blastocysts after IVF was much lower (P<0.05 to P<0.0001) compared with IVF results for IVM-matured comCOCs (Table 8). The very poor cleavage results for expCOCs are unlikely caused by the small number of COCs per IVF group, especially since 12 of 20 (60%) singlets and 38 of 52 (73%) pairs of comCOCs subjected to IVF cleaved by Day 2 of culture. 156 Ex-cpFSH reduced capacity of comCOCs and expCOCs recovered prior to an ovulatory hCG stimulus to resume meiosis during IVM The vast majority (91%) of the ovulatory-size follicle with comCOCs resumed meiosis during IVM (Table 9). Nevertheless, the proportion of ovulatory-size follicles with comCOCs that progressed to MII after IVM was lower (P<0.01) for the Ex-cpFSH treated heifers compared with controls (Table 9). None of the controls had expCOCs as observed in Exp 1. In contrast, 32% of the ovulatory-size follicles recovered 12 h after the last cpFSH injection (prior to the ovulatory hCG stimulus) had expCOCs (Table 5, Exp 3). However, only 38% of the premature expCOCs resumed meiosis and reached MII during IVM while 55% were classified as degenerated (Table 8). For the Ex-cpFSH treated heifers, the proportion of ovulatory-size follicles with expCOCs that reached MII during IVM was lower (P<0.0001) while the proportion classified as degenerated was higher (P<0.05) compared with comCOCs subjected to IVM (Table 9). Discussion The most significant findings in the present study are that Ex-cpFSH doses during the ovarian stimulation of the SORH model: i) increase expression of 17 genes in cumulus cells with well-established roles in cumulus expansion, function, and regulation of resumption of meiosis in all ovulatory-size follicles prior to an ovulatory hCG stimulus, ii) induce premature (prior to ovulatory hCG stimulus) expansion of the COC and resumption of meiosis in a moderate proportion of ovulatory-size follicles, iii) impair the capacity of prematurely expanded COCs to undergo IVF and resume meiosis during IVM, and iv) reduce responsiveness of comCOCs to an ovulatory hCG stimulus. Taken together, these observations in the SORH biomedical model support the conclusion that excessive FSH doses during ovarian stimulation dysregulate cumulus 157 cell function thereby impairing oocyte quality, contributing to oocyte wastage, and diminishing IVF success and ART outcomes. The present study extends our previous transcriptome analysis [24, 25] by showing that the Ex-cpFSH treatment induces differential expression of a small number of genes (17 of the 3288) in cumulus cells of all the ovulatory-size follicles independent of their extreme phenotypic differences. Many of these cumulus cell DEGs are regulated directly by FSH (AREG, IGFBP-3, IGFBP-5, PTX3, RGS2, TGFα, PLAT, FGG) in a variety of species. Moreover, most of these DEGs have well-established critical roles in regulation of gonadotropin action (GPR50, IGFBP- 3, IGFBP-5, RGS2) or secretion (VGF) and apoptosis (IGFBP-3, IGFBP-5, RGS-2), which could directly dysregulate cumulus cell function. Although IGFBP-3, IGFBP-5, and RGS-2 are pro- apoptotic, results of Ingenuity Pathway Analysis in our previous transcriptome study [24, 25] using a robust, high-quality data set did not indicate activation or inhibition of apoptosis. In addition, several of the cumulus cell DEGs identified in the present study are well-established regulators of cumulus expansion (AREG, LIF, MGAT5, MEPE, PTX3, TGFα, PLAT) and resumption of meiosis (AREG, IGFBP-3, TGFα, PLAT). However, other DEGs impact the extracellular matrix (MEPE, MGAT5, PTX3) and tight junctions and cell communication (CLDN11, NGFR), including calcium movement (NCS1, RGS2, VGF) in non-cumulus cell types that could also have an undiscovered yet critical role in cumulus function and capacity to regulate resumption of meiosis. Two DEGs serve other roles in cumulus cells (FGG, MDFI). FGG has a role in regulation of blood clotting [43] and MDFI inhibits myogenesis [44] and may have a role in ovulation [45, 46]. Thus, dysregulation of FGG and MDFI could contribute to the reduced ovulation rate observed for the heifers subjected to Ex-cpFSH during ovarian stimulation in our previous study [23]. 158 Taken together, the discovery of DEGs overexpressed in cumulus cells in response to Ex-cpFSH in all of the ovulatory-size follicles of Ex-cpFSH dosed animals provides new insight into potential mechanisms whereby excessive FSH action prior to an ovulatory hCG stimulus induces premature cumulus expansion, as observed in our previous [24, 25] and present studies, and premature resumption of meiosis. The 17 cumulus cell DEGs reported here for Ex-cpFSH treated heifers are induced prior to an ovulatory hCG stimulus and many of these genes are well-established to be involved in cumulus expansion. However, even though these DEGs are in COCs of all ovulatory-size follicles, we observed here and in our previous studies [24, 25] that premature cumulus expansion did not occur in all ovulatory-size follicles. This disconnect very likely represents a potential lag between the Ex-cpFSH induced alterations in gene expression with initiation of the morphological changes causing cumulus expansion (e.g., extracellular matrix). In addition, differential responsiveness of ovulatory-size follicles to cpFSH, which may explain the high within animal variability observed in gene expression (see Box Whisker Plots, Fig. 17), could also contribute to the disconnect between expression of genes involved in cumulus expansion and cumulus expansion. In the Ex-cpFSH treated heifers, a high proportion (75%) of ovulatory-size follicle had expCOCs in our previous study [24, 25] while only a moderate proportion (22% for Exp 1, 32% for Exp 3) of ovulatory-size follicles had expCOCs in the present study. We cannot rule out the possibility that animal variation or differences in cpFSH potency explain the differences in proportion of ovulatory-size follicles with expCOCs between our present and previous [24, 25] studies. However, it is highly likely that expCOCs, often described as very sticky in a variety of species [47, 48], are more fragile and difficult to remove from ovulatory-size follicles during 159 OPU than comCOCs. For example, OPU damages COCs if the aspiration pressure is not optimal [49-51], which could explain why denuded oocytes were observed after OPU in the present study. Consistent with this possible explanation, denuded oocytes were not observed when COCs were removed from excised ovulatory-size follicles using gentle pressure on a syringe and needle as in our previous study [24, 25]. We recognize a larger number of the Ex-cpFSH induced prematurely expanded COCs (expCOCs) from additional cattle will be necessary to confirm the inferior IVF results compared with not only controls in the present study but also with typical IVF results following IVM or artificial insemination of cattle reported by others [52]. Nevertheless, our findings indicate that the prematurely expanded COCs observed here are defective and have poor-quality oocytes. The precise reason for diminished oocyte competence is unknown but very likely arises due to dysregulated progression through meiosis (Table 6). However, we also observed in the present study, that nearly 38% of the expCOCs recovered by OPU before an ovulatory hCG stimulus reached the MII stage during IVM. This finding implies that IVM of the prematurely expanded COCs could have improved the poor IVF results observed in the present study (Table 8), although this was not examined. Nevertheless, we observed that the moderate proportion of ovulatory-size follicles with prematurely expanded COCs developing to MII during IVM was highly variable amongst heifers, while a higher proportion had degenerated during IVM (Table 7). Thus, we expect that IVM of prematurely expanded COCs would only marginally improve IVF success for a limited number of heifers. We established that Ex-cpFSH doses during ovarian stimulation of the SORH model reduced estradiol production [23-25] and decreased the hCG-induced ovulation rate [23]. These findings indicated that the Ex-cpFSH doses during ovarian stimulation impair responsiveness of 160 ovulatory-size follicles to an ovulatory hCG stimulus. If so, this could explain why both estradiol production and ovulation rate were reduced in our previous studies [23]. Results here further confirm the likelihood that Ex-cpFSH doses diminish the responsiveness of ovulatory-size follicles to an ovulatory hCG stimulus based on three comparisons. First, after the ovulatory hCG stimulus, the proportion of ovulatory-size follicles per heifer with comCOCs (Table 5, Exp 2) and proportion of comCOCs that remained at the GV stage (Table 7) were both higher in the Ex- cpFSH treated heifers compared with controls. Second, the proportions of ovulatory-size follicles per heifer treated with Ex-cpFSH doses that had comCOCs and expCOCs before (Table 5, Exp 1) and after (Table 5, Exp 2) the ovulatory hCG stimulus were nearly identical. Third, a higher proportion of the expCOCs remained at GV while a lower proportion reached MII in the Exp- cpFSH treated heifers compared with controls (Table 7) implying that the Exp-cpFSH doses impeded resumption and progression of nuclear maturation to MII. These combined findings, coupled with our previous results [23-25], provide compelling evidence that the Ex-cpFSH treatment during ovarian stimulation (prior to the ovulatory hCG stimulus) blocks responsiveness of the ovulatory-size follicles to hCG thereby reducing availability of high-quality hCG-matured COCs for ART. Previous studies in sheep show that typical ovarian stimulation regimens do not enhance responsiveness of all ovulatory-size follicles to FSH or LH [53]. This observation may explain why, although lower than the Ex-cpFSH treated heifers, a relatively high proportion (46%) of ovulatory-size follicles per heifer in controls had comCOCs after the ovulatory hCG stimulus. While the precise reason is unclear, the results imply that ovarian stimulation even with the industry-standard cpFSH doses (control) also hinders responsiveness to an ovulatory hCG stimulus. In support of this possibility, results of our previous study [23] show that ~20% of the 161 ovulatory-size follicles per heifer developing in response to the control doses do not ovulate in response to an ovulatory hCG stimulus. In summary, the results identified key Ex-cpFSH-induced DEGs in cumulus cells that may dysregulate cumulus function, enhance premature cumulus expansion, and impair oocyte quality in ovulatory-size follicles developing prior to an ovulatory hCG stimulus. In addition, the excessive FSH also induced inhibition of cumulus expansion and oocyte maturation post-hCG, and a reduced capacity of oocytes with prematurely expanded cumulus cells to undergo IVF or nuclear maturation during IVM. Although cumulus expansion is the hallmark for oocyte maturation in response to a preovulatory LH surge [54-57], our observations emphasize the risks of recovery of predominantly dysregulated cumulus-oocyte complexes (COCs). 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Fertil Steril 1999; 72:412-417. 167 A) Interrogate Prior RNA-seq Study that Identified 3288 Ex-cpFSH-induced Cumulus DEGs in Ovulatory -Size Follicle Types 2, 3 and 4 vs Type 1 (Clark et al., 2022b) Determine if any of the 3288 Ex-cpFSH-induced Cumulus DEGs are Common to All Ovulatory -size Follicle Types Literature Review to Determine Function of 17 DEGs APPENDIX B) Small Ovarian Reserve Heifers Ovarian Stimulation with Control or Ex -cpFSH Doses Exp 1-3 Ovulatory -size Follicle Number and COC Recovery Exp 1 COC Morphology and Meiotic Progression before hCG Exp 2 COC Morphology and Meiotic Progression after hCG Exp 3 Meiotic Progression of comCOCs after IVM IVF of Prematurely Expanded COCs Figure 16. Experimental approaches. To test the hypothesis that excessive FSH action during ovarian stimulation dysregulates cumulus cell function and oocyte maturation, we determined: A) if any of the 3288 Ex-cpFSH-induced cumulus DEGs in Types 2, 3 and 4 vs 1 ovulatory-size follicles of the SORH model, identified in our previously published transcriptome data set [25], were common to all ovulatory-size follicles independent of their Type and if these DEGs were critical for cumulus function and oocyte 168 Figure 16 (cont’d) maturation. We also determined: B) if Ex-cpFSH doses during ovarian stimulation of the SORH model altered number of ovulatory- size follicles and COC recovery rate (Exp 1-3), cumulus cell morphology (Exp 1-2), nuclear maturation of oocytes before (premature, Exp 1) or after an ovulatory hCG stimulus (Exp 2) or after in vitro maturation of compact COCs (comCOCs, Exp 3), and IVF of prematurely expanded COCs (Exp 3). Abbreviations used: Ex-cpFSH, excessive commercial FSH-enriched porcine pituitary preparation; DEGs, differentially expressed genes; SORH, small ovarian reserve heifer; COC, cumulus-oocyte complex; Exp, experiment; comCOCs, compact cumulus-oocyte complexes; expCOCs, expanded cumulus-oocyte complexes. 169 A B C D E F G H Figure 17. Representative images of the different stages of nuclear maturation of bovine oocytes. COCs were denuded, stained with DAPI and oocyte nuclear maturation stage determined as explained in Methods. The germinal vesicle (GV) stages of nuclear maturation of 170 Figure 17 (cont’d) oocytes are as follows: A. GV0 = diffuse filamentous pattern of chromatin in the whole nuclear area, occasionally surrounding the nonfluorescent nucleolus; B. GV1 = similar to GV0 except only a few condensed chromatin foci were detected in the nucleus; C. GV2 = chromatin was further condensed into clumps or a strand distributed throughout the nucleoplasm; or D. GV3 = chromatin condensed into a single clump within the nuclear envelope. Oocytes not classified at a GV stage were classified as: E. germinal vesicle breakdown = GVBD, F. metaphase I = MI, G. metaphase II = MII, or H. degenerated = oocytes with fragmented chromosomes or without visible nuclear material. Abbreviations used: COCs, cumulus-oocyte complexes; GV, germinal vesicle. 171 Figure 18. Volcano plots comparing cumulus cell DEGs for Type 2, 3 and 4 vs 1 ovulatory-size follicles. Volcano plots of cumulus cell transcriptome data noting locations of the 17 DEGs described in this analysis. Data are from the previously published transcriptome study [25]. The panels show data for the comparisons noted above each panel (left to right, Type 2, 3, and 4 vs Type 1). Each panel plots the Log2 (foldchange) along the x axis, and the -Log10 (FDR) from the DESeq2 output from our previous study [25]. Horizontal dashed lines denote two FDR thresholds: 0.01 and 0.05. Point color key: black = non-DEG, Red = upregulated DEGs, blue = downregulated DEGs. The 17 DEGs common to all follicle types are indicated by their gene symbols. Abbreviations used: DEGs, differentially expressed genes; FDR, false discovery rate. 172 Gene Symbol AREG CLDN11 Claudin 11 Gene Name Amphiregulin FGG Fibrinogen G Gamma G protein-coupled receptor 50 Insulin-like growth factor binding protein 3 Insulin-like growth factor binding protein 5 Leukemia-inhibitory factor MyoD family inhibitor Matrix extracellular phosphoglycoprotein Alpha-1,6-manosyl-glycoprotein beta- 1,6-acetylglucosaminyltransferase Functions ORM [58, 59], CE [58, 59] JXN [60-62] Cl (Simurda et al., 2020) G [58, 63], ORM [64] G [65], ORM [65], A[66] Ovulatory-size follicle types Control Type 1 (n=7) Type 2 (n=6) Ex-cpFSH Type 3 (n=6) Type 4 (n=5) 0 0.5 0 7 3 0.1 100 12 2474 189 25 226 28 2133 102 77 225 98 1564 243* G [67], A [68-71] 15* 479 429 21367 ORM [72, 73], CE [74] O [45] (Martin et al., 1983) ECM [75-77] 0 0 0 2 0.8 7 55 5 1 270 77 63 CE [78], ECM [78] 56 285 558 1111 Neuronal calcium sensor 1 Nerve growth factor receptor Plasminogen Activator, Tissue Pentraxin 3 Ca [79, 80] JXN [81], A [82] ECM [83], CE [84], ORM [84] CE [85], ECM [86-88] Regulator of G protein signaling 2 Transforming growth factor alpha G [89], Ca [90], A[91] G [92], ORM [93, 94], CE [58, 95] VGF nerve growth factor inducible G [96-98], Ca [99, 100] 0 0 39* 2 0 0 0 0.6 3 673 122* 6 0.1 7 7 74 727 4232 71 9 7 60 97 4460 5030 102 73 193 GPR50 IGFBP3 IGFBP5 LIF MDFI MEPE MGAT5 NCS1 NGFR PLAT PTX3 RGS2 TGFα VGF Table 3. Average FPKM values for differential expression of cumulus genes critical for regulation of cumulus function and oocyte maturation in Type 1 compared with Ex-cpFSH-induced Types 2, 3 and 4 ovulatory-size follicles. Heifers were subjected to ovarian stimulation with IS-cpFSH (control) or Ex-cpFSH doses, ovulatory-size (≥10 mm in diameter) follicles were excised from ovaries 12 h after the last cpFSH injection (no hCG given), and cumulus cells were removed from COCs and subjected to RNAseq and bioinformatic analyses. The ovulatory-size follicles developing in response to the control cpFSH doses were classified as Type 1 whereas those developing in response to Ex-cpFSH doses were classified as Type 2, 3 or 4 based on differences in estradiol, 173 Table 3 (cont’d) progesterone and oxytocin concentrations in follicular fluid and cumulus morphology [25]. The number of libraries (each library was from an individual ovulatory-size follicle of a single animal) analyzed is in parentheses under each follicle type. The values represent averages for fragments per kilobase per million mapped reads (FPKM). Zero = undetectable values. Box Whisker Plots of these results are depicted in Fig. 17. Type 1 ovulatory-size follicles differed (P<0.01, *P<0.05) from Type 2, 3, or 4 follicles for each gene, as explained in Fig. 17’s legend. Gene functions are abbreviated as follows: A = apoptosis, Ca = calcium movement, CE = cumulus expansion, Cl = clotting regulation, ECM = extracellular matrix, G = Gonadotropin action/secretion, JXN = tight junction and cell communication, O = ovulation, ORM = oocyte resumption of meiosis. Abbreviations used: FPKM, fragments per kilobase per million mapped reads; DEGs, differentially expressed genes; IS-cpFSH, industry-standard commercial FSH-enriched porcine pituitary preparation; Ex-cpFSH, excessive commercial FSH-enriched porcine pituitary preparation. 174 Figure 19. Expression of cumulus cell genes critical for regulation of cumulus function and oocyte maturation in Type 1 compared with Ex-cpFSH-induced Types 2, 3 and 4 ovulatory- size follicles. Heifers in our previous study [25] were subjected to ovarian stimulation with IS- cpFSH (control) or Ex-cpFSH doses and cumulus cells removed from the different types ovulatory-size follicles and subjected to RNAseq and bioinformatic analyses as explained in Table 3’s legend. Types 1, 2, 3 and 4 ovulatory-size follicles are depicted along the x axis. Data points within each panel reflect mean expression values for replicates of each gene based on fragments per kilobase of exon per million reads (FPKM, y axis) determined in our previous study [25]. Analysis of our previously published data identified 17 shared cumulus DEGs (FDR <0.05 to FDR <0.01) common to all the Type 2, 3 and 4 ovulatory-size follicles. These 17 Ex-cpFSH-induced cumulus cell DEGs all have well-established roles in modulation of FSH action and cumulus cell function including induction of cumulus cell expansion and regulation of oocyte maturation (Table 3). Gene symbols are depicted at the top of each panel and corresponding gene names and functions are in Table 3. Within each panel, the large black circles denote mean FPKM values, small gray circles denote each sample, and vertical lines denote standard errors of means. Based on statistical analysis results from our previous study [25], the horizontal lines at the top of each panel represent comparisons between Type 1 vs 2, 3, or 4 ovulatory-size follicles. Asterisks above each horizontal line indicate if the FDR values for each mean comparison differed significantly (* = P≤0.05, ** = P≤0.01, *** = P≤0.001, **** = P≤0.0001). Abbreviations used: IS-cpFSH, industry-standard commercial FSH-enriched porcine 175 Figure 19 (cont’d) pituitary preparation; Ex-cpFSH, excessive commercial FSH-enriched porcine pituitary preparation; FPKM, fragments per kilobase per million mapped reads; DEG, differentially expressed gene; FDR, false discovery rate. 176 Exp 1 Exp 2 Exp 3 Control Ex-cpFSH Control Ex-cpFSH Control Ex-cpFSH 19 ± 5 (6 – 56) 31 ± 6 (9 – 59) 21 ± 4 (6 – 60) 26 ± 4 (9 – 70) 17 ± 3 (5 – 46) 19 ± 2 (3 – 31) 16 ± 4 (5 – 41) 24 ± 4 (8 – 44) 17 ± 2 (5 – 32) 20 ± 3 (8 – 50) 18 ± 3 (13 – 56) 18 ± 2 (19 – 45) 10 ± 3 (2 – 28) 13 ± 2 (5 – 26) 12 ± 2 (2 – 27) 13 ± 2 (2 – 31) 10 ± 2 (5 – 41) 11 ± 1 (9 – 28) Number of ovulatory-size follicles (≥10 mm) Number of aspirated follicles Number of COCs recovered by OPU COC recovery rate 58% ± 5% (29 – 90%) 53% ± 6% (22 – 85%) 58% ± 3% (33 – 100%) 63% ± 5% (25 – 87%) 55% ± 5% (13 – 100%) 56% ± 5% (10 – 82%) Table 4. Impact of ovarian stimulation of small ovarian reserve heifers with control or Ex-cpFSH doses on number of ovulatory- size follicles and COC recovery. Small ovarian reserve Holstein heifers (Exp 1, n = 10/dose; Exp 2, n = 16/dose; Exp 3, n = 18/dose) were subjected to ovarian stimulation with IS-cpFSH (control) or Ex-cpFSH doses. The heifers in Exp 1 and 3 were subjected to OPU to recover COCs 12 h after the last cpFSH injection while the heifers in Exp 2 were subjected to an ovulatory hCG stimulus 12 h after the last cpFSH injection, and COCs were recovered by OPU 24 h after the hCG injection but before ovulation. The number of ovulatory- size follicles was determined at the time of OPU. Recovery rate = number of COCs recovered by OPU divided by the number of 177 Table 4 (cont’d) aspirated follicles. Data are expressed as means ± SEM per heifer with the range of the data collected in parentheses. There were no statistical differences between means. Abbreviations used: Exp, experiment; IS-cpFSH, industry-standard commercial FSH-enriched porcine pituitary preparation; Ex-cpFSH, excessive commercial FSH-enriched porcine pituitary preparation; OPU, oocyte pick-up; COCs, cumulus-oocyte complexes. 178 Morphological classification of COCs comCOC Exp 1 Exp 2 Exp 3 Control Ex-cpFSH Control Ex-cpFSH Control Ex-cpFSH 93% ± 3% (75 –100%) 77% ± 4%** (58 – 100%) 46% ± 9% (0 – 100%) 72% ± 5%** (27 – 100%) 91% ± 3% (0 – 94%) 65% ±5%**** (40 – 100%) expCOC 0% ± 0.4% (0 – 4%) 22% ± 5%**** (0 – 42%) 45% ± 8% (0 – 100%) 24% ± 4%* (0 – 39%) 3% ± 2% (0 – 29%) 32% ±5%**** (0 – 82%) Denuded 3% ± 3% (0 – 25%) 0.7% ± 1% (0 – 7%) 7% ± 2% (0 – 25%) 4% ± 1% (0 – 17%) 6% ± 3% (0 – 44%) 3% ± 1% (0 – 18%) Table 5. Impact of ovarian stimulation of small ovarian reserve heifers with control or Ex-cpFSH doses on morphology of COCs. Small ovarian reserve Holstein heifers (Exp 1, n = 10/dose, Exp 2, n = 16/dose, Exp 3, n = 18/dose) were subjected to ovarian stimulation with IS-cpFSH (control) or Ex-cpFSH doses, subjected to OPU to recover COCs, and COCs classified morphologically as compact (comCOC) or expanded (expCOC) and denuded oocytes recorded as explained in Table 4’s legend. Proportions per column may not total 100% because some COCs or oocytes were unidentifiable or lost during histological processing to assess nuclear maturation. These results were not impacted by treatment and thus not presented. Data are expressed as means ± SEM for proportions of ovulatory-size follicles per heifer with COCs classified as comCOC, expCOC, or denuded with the range of the data collected in parentheses. Asterisks (* = P≤0.05, ** = P≤0.01, **** = P≤0.0001) denote statistical differences between means within each experiment. Abbreviations used: Exp, experiment; IS-cpFSH, industry-standard commercial FSH-enriched porcine pituitary preparation; Ex-cpFSH, excessive commercial FSH-enriched porcine pituitary preparation; OPU, oocyte pick-up; COCs, cumulus-oocyte complexes; comCOC, compact cumulus-oocyte complex; expCOC, expanded cumulus-oocyte complex. 179 Control comCOC 9 ± 3 (1 – 27) Exp 1 Ex-cpFSH comCOC expCOC 7 ± 1 (1 - 14) 3 ± 1 (0 - 9) Number of oocytes evaluated per heifer Stage of nuclear maturation GV 77% ± 8% (30 – 100%) 97% ± 3% (75 – 100%) 19%±10%**** (0 – 100%) GVBD 0% ± 0% 0% ± 0% 1% ± 1% (0 – 11%) MI MII 0% ± 0% 0% ± 0% 1% ± 3% (0 – 13%) 1% ± 1% (0 – 13%) 40% ± 13%** (0 – 100%) 8% ± 5% (0 – 7%) Degenerated 2% ± 2% (0 – 14%) 0.8% ± 1% (0 – 33%) 8% ± 4% (0 – 33%) Table 6. Impact of ovarian stimulation of small ovarian reserve heifers with control or Ex- cpFSH doses on stage of nuclear maturation for comCOCs or expCOCs. Small ovarian reserve Holstein heifers (n = 10/dose) were subjected to ovarian stimulation with IS-cpFSH (control) or Ex-cpFSH doses then subjected to OPU to recover COC 12 h after the last cpFSH injection. COCs were classified morphologically as having a compact (comCOC) or expanded (expCOC) layer of cumulus cells. The stage of oocyte nuclear maturation was assessed in COCs as GV, GVBD, MI, MII, or Degenerated. Proportions per column may not total 100% because some COCs or oocytes were unidentifiable or lost during histological processing to assess nuclear maturation. These results were not impacted by treatment and thus not presented. Data are expressed as means ± SEM for proportion of ovulatory-size follicles with comCOCs or expCOCs at each nuclear maturation stage per heifer with the range of the data collected in parentheses. Asterisks (** = P≤0.01, **** = P≤0.0001) denote statistical differences between means within cpFSH dose. Abbreviations used: IS-cpFSH, industry-standard commercial FSH-enriched porcine pituitary preparation; Ex-cpFSH, excessive commercial FSH-enriched porcine pituitary preparation; OPU, oocyte pick-up; COCs, cumulus-oocyte complexes; comCOC, 180 Table 6 (cont’d) compact cumulus-oocyte complex; expCOC, expanded cumulus-oocyte complex; GV, germinal vesicle, GVBD, germinal vesicle breakdown; MI, meiosis I; MII, meiosis II. 181 Exp 2 Control Ex-cpFSH comCOC expCOC comCOC expCOC Number of oocytes evaluated per heifer 2 ± 0.5 (0 – 6) 4 ± 1 (0 – 14) 4 ± 0.8 (1 – 13) 3 ± 0.7 (0 – 9) Stage of nuclear maturation GV GVBD MI MII 68% ± 12% (0 – 100%) 4% ± 3%*** (0 – 40%) 86%± 3%++ (50 – 100%) 11% ± 5%**** (0 – 50%) 1% ± 1% (0 – 17%) 2% ± 2% (0 – 25%) 0% ± 0% 12% ± 6%* (0 – 75%) 4% ± 3% (0 – 50%) 1% ± 1% (0 – 17%) 0.7% ± 1% (0 – 11%) 30% ± 10%** (0 – 100%) 0% ± 0% 57%± 12%**** (0 – 100%) 0.7% ± 1% (0 – 11%) 34%±8%++++*** (0 – 100%) Degenerated 0% ± 0% 0% ± 0% 7% ± 6% (0 – 11%) 7% ± 3% (0 – 50%) Table 7. Impact of an ovulatory hCG stimulus following ovarian stimulation of small ovarian reserve heifers with control or Ex- cpFSH doses on stage of nuclear maturation for comCOC or expCOC. Small ovarian reserve Holstein heifers were subjected to ovarian stimulation with IS-cpFSH (control, n = 15) or Ex-cpFSH (n = 16) doses, subjected to an ovulatory hCG stimulus 12 h after 182 Table 7 (cont’d) the last cpFSH injection, and COCs were recovered by OPU 24 h after the hCG injection but before ovulation. COCs were classified morphologically as having a compact (comCOC) or expanded (expCOC) layer of cumulus cells. The oocyte nuclear maturation stage was assessed in COCs as GV, GVBD, MI, MII, or Degenerated. Proportions per column may not total 100% because some COCs or oocytes were unidentifiable or lost during histological processing to assess nuclear maturation. These results were not impacted by treatment and thus not presented. Data are expressed as means ± SEM for proportion of ovulatory-size follicles with COCs at each nuclear classification per heifer with the range of the data collected in parentheses. Asterisks (* = P≤0.05, ** = P≤0.01, *** = P≤0.001, **** = P≤0.0001) denote statistical differences between means within dose whereas plus (++++ = P≤0.0001) denote statistical differences between means but within COC classification across cpFSH doses. Abbreviations used: IS-cpFSH, industry-standard commercial FSH- enriched porcine pituitary preparation; Ex-cpFSH, excessive commercial FSH-enriched porcine pituitary preparation; OPU, oocyte pick- up; COCs, cumulus-oocyte complexes; comCOC, compact cumulus-oocyte complex; expCOC, expanded cumulus-oocyte complex; GV, germinal vesicle, GVBD, germinal vesicle breakdown; MI, meiosis I; MII, meiosis II. 183 COC morphology N Cleaved Blastocyst Hatched Blastocyst Exp 3 comCOC 182 124 (68%) 51 (28%) expCOC 37 2 (5.4%)**** 0 (0%)*** 22 (12%) 0 (0%)* Table 8. IVF results for bovine oocytes with prematurely expanded (expCOC) or compact layers (comCOC) of cumulus cells. The compact COCs (comCOC) were recovered from follicles of ovaries obtained at a local abattoir and subjected to IVM and IVF. The prematurely expanded COCs (expCOC) were obtained from Ex-cpFSH treated heifers (n = 8) 12 h after the last cpFSH injection but prior to an ovulatory hCG stimulus and subjected to IVF without IVM. IVF was performed using sperm from the same bull. Data are represented as the number of fertilized oocytes from comCOCs or expCOCs subjected to IVF that cleaved, formed blastocysts, or hatched blastocysts divided by total number of comCOCs or expCOCs subjected to IVF with the ratio in parentheses. Asterisks (* = P≤0.05, *** = P≤0.001, ****= P≤0.0001) = significant difference between ratios when comCOCs were compared with expCOCs. Abbreviations used: comCOC, compact cumulus-oocyte complex; expCOC, expanded cumulus-oocyte complex; Ex-cpFSH, excessive commercial FSH-enriched porcine pituitary preparation; N, number of comCOC or expCOC. 184 Exp 3 Control Ex-cpFSH comCOC comCOC expCOC 9 ± 2 (2 – 33) 7 ± 1 (2 – 20) 4 ± 0.8 (0 – 11) Number of oocytes evaluated per heifer Stage of nuclear maturation after IVM GV 8% ± 3% (0 – 43%) 9% ± 2% (0 – 25%) 8% ± 6% (0–50%) GVBD 6% ± 2% (0 – 25%) 5% ± 2% (0 – 25%) 0% ± 0% MI MII 3% ± 2% (0 – 33%) 6% ± 2% (0 – 25%) 0% ± 0% 82% ± 4% (50 – 100%) 70%±6%++ (17–100%) 38%±16%**** (0–100%) Degenerated 1% ± 1% (0 – 9%) 10% ± 4% (0 – 50%) 55% ± 14%* (0–100%) Table 9. Impact of ovarian stimulation of small ovarian reserve heifers with control or Ex- cpFSH doses on stage of nuclear maturation of comCOCs or expCOCs after IVM. Small ovarian reserve Holstein heifers (n = 17/control, n = 14/Ex-cpFSH; because some heifers had < 3 COCs and eliminated from study) were subjected to ovarian stimulation with IS-cpFSH (control) or Ex-cpFSH doses then subjected to OPU to recover COCs 12 h after the last cpFSH injection. COCs recovered by OPU were classified morphologically as having a compact (comCOC) or expanded (expCOC) layer of cumulus cells and subjected to IVM for 22 h. After IVM, oocyte nuclear maturation stages for COCs were assessed as GV, GVBD, MI, MII, or Degenerative. Proportions per column may not total 100% because some COCs or oocytes were unidentifiable or lost during histological processing to assess nuclear maturation. These results were not 185 Table 9 (cont’d) impacted by treatment and thus not presented. Data are expressed as means ± SEM for proportion of ovulatory-size follicles with comCOCs or expCOCs at each nuclear maturation stage per heifer with the range of the data collected in parentheses. Asterisks (* = P≤0.05, **** = P≤0.0001) denote statistical differences between means within cpFSH dose whereas pluses (++ = P≤0.01) denote statistical difference between means across dose but within COC classification across cpFSH doses. Abbreviations used: COCs, cumulus-oocyte complexes; IS-cpFSH, industry-standard commercial FSH-enriched porcine pituitary preparation; Ex-cpFSH, excessive commercial FSH- enriched porcine pituitary preparation; OPU, oocyte pick-up; comCOC, compact cumulus-oocyte complex; expCOC, expanded cumulus-oocyte complex; GV, germinal vesicle, GVBD, germinal vesicle breakdown; MI, meiosis I; MII, meiosis II. 186 Limitations in Use of Ovarian Reserve Biomarkers to Predict the Superovulation Response in Small Ovarian Reserve Heifers1 CHAPTER 6. 1 Kaitlin R. Karl, Janet LH Ireland, Zaramasina L. Clark, Robert J. Tempelman, Keith E. Latham, and James J. Ireland. "Limitations in use of ovarian reserve biomarkers to predict the superovulation response in small ovarian reserve heifers." Theriogenology 182 (2022): 53-62. 187 Title: The ovarian reserve biomarkers, anti-Mullerian hormone concentrations and antral follicle count, are positively linked to number of dysfunctional ovulatory-size follicles developing in response to superovulation of heifers with a small ovarian reserve Running Title: AMH and AFC are positively linked to number of dysfunctional ovulatory-size follicles developing in response to superovulation Summary Sentence: The ovarian reserve biomarkers, circulating AMH concentration and AFC, are positively associated with number of functional and dysfunctional ovulatory-size follicles independent of the FSH dose used during superovulation of heifers with a small ovarian reserve. Keywords: anti-Mϋllerian hormone; AMH; low antral follicle count; AFC; estradiol; small ovarian reserve; superovulation; dysfunctional ovulatory-size follicles; bovine; biomarkers Authors and affiliations: Kaitlin R. Karl1, Janet L.H. Ireland1, Zaramasina L. Clark1, Robert J. Tempelman1, Keith E. Latham1, James J. Ireland1,2 1Molecular Reproductive Endocrinology Laboratory, Department of Animal Science, Michigan State University, East Lansing, MI 48824. 2 Correspondence: James J. Ireland, Molecular Reproductive Endocrinology Laboratory, Department of Animal Science, Michigan State University, East Lansing, Michigan, 48824, USA. Tel.+1 517 432 1384. Email: ireland@msu.edu This study was supported by the Agriculture and Food Research Initiative Competitive USDA- NIH Dual Purpose Program Grant no. 2017-67015-26084 from the USDA National Institute of Food and Agriculture, the Eunice Kennedy Shriver National Institute of Child Health & Human Development of the National Institutes of Health under Award Number T32HD087166, MSU AgBio Research, and Michigan State University. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. 188 Abstract High FSH doses during superovulation of heifers with a small ovarian reserve increase the number of dysfunctional ovulatory-size follicles that do not ovulate in response to human chorionic gonadotropin (hCG). Thus, anti-Müllerian hormone (AMH) and antral follicle count (AFC), two well-established biomarkers of responsiveness of individuals to superovulation, are hypothesized to be positively linked to number of dysfunctional ovulatory-size follicles developing in response to superovulation with high FSH doses. To test this hypothesis, heifers with a small ovarian reserve were stimulated beginning on Day 1 of the estrous cycle with twice daily treatments for 4 days with each of four Folltropin-V (FSH) doses (35 IU, 70 IU (industry standard), 140 IU, or 210 IU) followed by prostaglandin F2α to regress corpora lutea (CL) from the previous estrous cycle and hCG to induce ovulation. Ovulatory-size follicles were classified as functional or dysfunctional based on whether they ovulated and formed CL in response to hCG. FSH dose did not impact the relationship between AMH, AFC and the number of functional or dysfunctional ovulatory-size follicles developing in response to superovulation. Thus, data from the four superovulations were averaged for each heifer. AMH and AFC were positively associated with the subsequent number of functional and dysfunctional ovulatory-size follicles and the proportion of ovulatory-size follicles that are dysfunctional after superovulation. Because measurements of AMH concentration and AFC predict the number but not functionality of ovulatory-size follicles, which may also impact oocyte quality, these ovarian reserve biomarkers are concluded to be unlikely useful to improve IVF or embryo transfer outcomes in heifers with a small ovarian reserve. 189 Introduction A major pitfall of FSH stimulation protocols during assisted reproductive technologies (ART) is the unpredictability [1-3] and inherent high variability in response of both women [4] and cattle [5, 6] to superovulation. This variability in the superovulation response results in a high rate of oocyte and embryo wastage resulting in costly unsuccessful ART outcomes [1-3]. Thus, discovery of biomarkers predictive of the donor’s response to superovulation should enable practitioners to better predict poor-, good-, and hyper-responders, and thereby improve current superovulation and embryo transfer protocols to optimize ART outcomes [7]. Such biomarkers could include circulating concentrations of anti-Müllerian hormone (AMH) or antral follicle count (AFC). Anti-Müllerian hormone concentrations are widely used to predict responsiveness of donors to superovulation protocols. Anti-Müllerian hormone concentration is static during reproductive cycles in cattle [8-11] and women [12, 13] and positively correlated with size of the ovarian reserve (total number of morphologically healthy follicles in ovaries) in cattle [11, 14], women [15] and mice [16], and with number of antral follicles growing during ovarian follicular waves in cattle [8]. Cattle with greater circulating AMH concentrations have higher AFCs and correspondingly larger ovarian reserves, greater numbers of healthy growing large preantral and small antral follicles [17], and display better responses to superovulation compared with their age-matched counterparts with low circulating AMH concentrations [18]. These observations explain why numerous studies in women [19, 20] and cattle [14, 21-24] also report a positive correlation between circulating AMH concentrations, AFC, and subsequent individual response to superovulation. However, few if any studies have evaluated the association of AMH and AFC prior to superovulation with subsequent number and function (e.g., capacity to ovulate in response to LH or LH-like stimulus) of the ovulatory-size follicles developing in response to 190 superovulation with FSH. Moreover, whether the positive association between the ovarian reserve biomarkers, AMH and AFC, with response to superovulation also exists among individuals with a relatively small ovarian reserve has not been examined. Consequently, the reliability of AMH and AFC measurements to predict number and function of ovulatory-size follicles developing in response to superovulation of individuals with a small ovarian reserve is unknown. Anti-Müllerian hormone is secreted primarily by the granulosa cells of morphologically healthy growing large preantral and small antral follicles [18, 23, 25], which undergo atresia during reproductive cycles unless they are stimulated with FSH to develop into ovulatory-size antral follicles during ART. Typically, a wide range of FSH doses are used by ART practitioners to enhance the superovulatory response in women with a small ovarian reserve [4]. However, FSH doses used during ART are inversely associated with oocyte recovery [26] and live birth rate independent of age, body mass index or health of donors [4]. These correlative findings in women are supported by dose response studies in cattle showing that high FSH doses decrease estradiol production and ovulation rate by ovulatory-size follicles developing response to superovulation [7], and reduce fertilization rate [27-29], embryo yield [29, 30] or quality [27, 29], and number of transferable embryos [28-31]. Taken together, these results support the overall working hypothesis for our studies that high FSH doses during superovulation are detrimental to ovulatory follicle function, oocyte quality and embryo survival in cattle. To examine the impact of high FSH doses during superovulation on ovulatory follicle function, we used the small ovarian reserve heifer model. This model mimics key characteristics of women with small ovarian reserves including a low AFC [32], hypersecretion of FSH [32-34], low circulating AMH concentrations [8] during the reproductive cycle, and poor response to 191 superovulation [32]. In our recent study [7], a Williams Latin Square Design was used to rigorously evaluate the effects of different FSH doses during superovulation of the same group of heifers with a small ovarian reserve on ovarian function. Results showed that the use of high FSH doses to improve the superovulatory response did not alter the circulating AMH concentrations or number of ovulatory-size follicles but did have a detrimental impact on the capacity of ovulatory-size follicles to produce estradiol and ovulate. Taken together, these observations support the hypothesis that AMH and AFC, which are well-established predictors of responsiveness of individuals to superovulation, are positively associated with number of dysfunctional ovulatory-size follicles developing in response to superovulation with high FSH doses. To test this hypothesis, we examined the relationship between circulating AMH concentrations and AFC with number of dysfunctional and functional ovulatory-size follicles developing in response to superovulation of heifers with a small ovarian reserve with four different FSH doses. Ovulatory-size follicles were defined as dysfunctional or functional based on whether they ovulated and formed a CL in response to hCG. We find that the ovarian reserve biomarkers, circulating AMH concentrations and AFC, prior to superovulation are positively linked to number of both dysfunctional and functional ovulatory-size follicles developing in response to superovulation independent of the FSH dose. Because AMH and AFC measurements prior to superovulation predict total number but cannot distinguish between functional and dysfunctional ovulatory-size follicles with potentially poor-quality oocytes developing in response to superovulation, these ovarian reserve biomarkers may not be consistently useful to improve IVF or embryo transfer outcomes in cattle with a small ovarian reserve. 192 Materials and Methods The experimental design for the present study, which was previously described in detail , will be briefly explained here. The Institutional Animal Care and Use Committee at Michigan State University sanctioned all procedures involving cattle. Selection of heifers with a small ovarian reserve We established that 11- to 12-month-old heifers with a low AFC (≤15 follicles of ≥3 mm in diameter) during ovarian follicular waves (15% to 20% of a herd) also have 80% smaller ovarian reserves and a reduced response to superovulation compared with age-matched counterparts with a high AFC (≥ 25 follicles) [7, 8, 32, 34-37]. In our previous study [7], serial ovarian ultrasonography was used to identify the nine, 11- to 12-month-old Holstein heifers of similar weights with a low AFC and small ovarian reserve used in the present study. Superovulation protocol, blood sampling and ovarian ultrasonography To synchronize estrous cycles for superovulation prior to FSH administration, these heifers were treated twice with prostaglandin-F2α (PG, 12.5mg PG/mL im, Lutalyse HighCon, Zoetis) 10 d apart followed by a 3rd PG 12 h after the 2nd PG. Each heifer was subjected to daily ovarian ultrasonography to detect ovulation and emergence of the first follicular wave. The 1st treatment of Folltropin-V (porcine pituitary extract containing primarily FSH with <1% LH contamination, Vetoquinol USA Inc) began 36 h after the last PG, which was ±1 d from ovulation and initiation of the first follicular wave in all heifers. Folltropin-V is hereafter referred to as FSH. The superovulation regimen in our previous study consisted of a total of eight im treatments of FSH at 12-h intervals. To regress the CL of the previous estrous cycle during superovulation, three additional PG treatments were given 12 h apart starting at the time of the 7th FSH treatment (about Day 4 to 5 of estrous cycle). To induce ovulation of the ovulatory-size 193 follicles developing in response to superovulation with FSH, heifers were treated with a 2,500 IU of human chorionic gonadotropin (hCG, Chorulon HCG 10,000 IU im, Merck Animal Health, USA), which is sufficient to ovulate up to 40 to 60 follicles in heifers [38], coincident with the 3rd PG (which is 12 h after the last (or 8th) FSH treatment). As explained in our previous study [7], serial ovarian ultrasonography was used daily to monitor alterations in AFC (follicles ≥3 mm in diameter), number of ovulatory-size follicles (follicles ≥10 mm in diameter) and CL beginning at the time of the 1st FSH treatment and ending nine days after the hCG for each superovulation regimen. In addition, blood samples (10 ml) were taken from the tail vein of each heifer once or twice daily beginning and ending concurrently with ultrasonography. A previously validated [8] commercially available AMH ELISA kit for cattle (MOFA Global, Verona, WI), which does not cross-react with other members of the transforming growth factor beta (TGFβ) superfamily including TGFβ, bone morphogenic factor-4 (BMP4), inhibin or activin [16], was used to measure circulating AMH concentrations in duplicate 20 µl serum samples per kit instructions [7]. The inter- and intra-assay coefficients of variation for the AMH assay (n = 6 plates) were 5.4% and 5.6%, respectively [7]. To measure circulating estradiol concentrations, serum estradiol-17β concentrations were determined by RIA as previously published [39, 40]. Inter- and intra-assay coefficients of variation for estradiol-17β assays (n = 15 plates) were 7.6% and 3.7%, respectively [7]. Experimental design The Williams Latin Square Design [41, 42] was used in our previous study [7] to minimize animal numbers, potential carry-over effects, and confounding variables such as environment and aging that could contribute to variability in responsiveness to superovulation [41, 42]. Each of the nine heifers used in the present study were superovulated as explained 194 above a total of four times with an interval of approximately 4 weeks between each superovulation regimen [7]. Each superovulation regimen used a total of eight equal FSH dose treatments of each different FSH dose: 35 IU (20mg NIH-FSH-P1/ml), 70 IU (40mg/2ml), 140 IU (80mg/4ml), or 210 IU (120mg/6ml). These FSH doses ranged from 60% lower and 240% higher than the Vetoquinol recommended dose per treatment of 87.5 IU. Hereafter, dose of FSH is referred to as IU per treatment. The data used in the present study to evaluate the association of AMH and AFC with number of dysfunctional and functional ovulatory-size follicles developing in response to superovulation with the different FSH doses was generated in our previous study [7] as follows. Anti-Müllerian hormone concentrations were determined from blood samples taken immediately prior to the 1st FSH treatment for all nine heifers subjected to each of the four FSH doses used during the superovulation regimens [7]. The AFCs for the heifers were determined by use of ovarian ultrasonography to count follicles ≥3 mm in diameter 1 to 2 h before the 1st FSH treatment for each of the four different FSH doses used during the superovulation regimens [7]. To determine number of ovulatory-size follicles that developed in response to superovulation with the different FSH doses and the corresponding circulating estradiol concentrations, the total number of follicles ≥10 mm in diameter for each heifer were determined at the time of the hCG treatment for each different superovulation regimen [7]. Time of hCG was chosen to determine number of ovulatory-size follicles because it coincides with the maximum number of ovulatory-size follicles that developed during each superovulation regimen in our previous study [7]. Capacity to ovulate in response to a preovulatory LH surge is a well-established characteristic of normal functioning ovulatory follicles in cattle [43-47]. Thus, the total number 195 of ovulatory-size follicles determined at time of the hCG treatment was separated into two different categories (functional, dysfunctional) based on whether the ovulatory-size follicles formed CL in response to the hCG treatment after each superovulation regimen. Number of functional ovulatory-size follicles was therefore defined as the total number of CL counted on Day 7 post-hCG [7] whereas the number of dysfunctional ovulatory-size follicles not ovulating in response to the hCG treatment was determined by subtracting the number of CL (or number of functional ovulatory-size follicles) from the total number of ovulatory-size follicles determined at time of the hCG treatment. Statistical analysis Box and whisker plots were used to show the distribution of all variables (AMH concentrations, AFC, total number of ovulatory-size follicles, number of dysfunctional and functional ovulatory-size follicles, proportion of total number of ovulatory-size follicles that were dysfunctional) before these data were transformed for statistical analyses. A Shapiro-Wilk test for normality of data distribution indicated all data for each FSH dose were normally distributed (P>0.05) after transformations. Anti-Müllerian hormone measurements were subjected to Log10 transformation whereas all other variables were subjected to square root transformation before correlation and linear regression analysis. To further assess variability in the biomarkers used in this study, repeatability of measurements of AMH and AFC within individuals was determined for the nine heifers used in the present study. AMH and AFC measurements were made in the individual heifers four different times 1.5 to 2 months apart, which is the interval between each superovulation regimen with the four different FSH doses. Repeatability of each AMH or AFC measurement within individuals was determined using transformed data as previously reported [34]. Repeatability 196 (range 0-1, where 1 = perfect) is defined as the proportion of the total variance attributable to animal variance, which is [48] calculated as σ2 animal/(σ2 animal + σ2 error) [49]. Variance components and repeatability estimates were determined using the PROC MIXED model approach of SAS [50], where heifers were accounted for as a source of variation and a common residual variance was used for all response variables across FSH doses. Linear regression analysis was performed using Statistical Analysis System (SAS 9.4 Institute, Cary, NC) with PROC GLIMMIX [50] to determine if a statistically significant (P≤0.05) linear relationship existed between AMH and AFC measurements before each superovulation with corresponding alterations in total and number of dysfunctional and functional ovulatory-size follicles in response to the different FSH doses during each superovulation regimen. PROC GLIMMIX [50] was then used to determine if the means and slopes generated during linear regression analysis differed (P≤0.05) among the four FSH doses used to superovulate heifers. Statistical Analysis System (SAS 9.4 Institute, Cary, NC) was also used to conduct a Pearson Correlation with PROC CORR [50] to determine if a statistically significant (P≤0.05) correlation existed between AMH concentrations and AFC measurements before each superovulation with corresponding total number of ovulatory-size follicles, number of functional and dysfunctional ovulatory-size follicles, and proportion of total number of ovulatory-size follicles that were dysfunctional. 197 Results Individual variability in circulating AMH concentration and AFC prior to superovulation with FSH and the corresponding variability in the endpoints used to measure response of each heifer to the different superovulation regimens The nine heifers in this study were of the same age, breed, weight, and all had a low AFC (≤15 follicles ≥3 mm in diameter). These selection criteria could have resulted in a relatively narrow degree of variability in the data collected from the nine heifers, thereby hindering interpretation of the linear regression and correlation results in our present study [51]. To examine this possibility, box and whisker plots [52] are provided to depict the distribution of the values examined in the present study, which included AMH (Fig. 20A), AFC (Fig. 20B), total number of ovulatory-size follicles (Fig. 22A), number of functional ovulatory-size follicles (Fig. 22B), number of dysfunctional ovulatory-size follicles (Fig. 24A), and proportion of total number of ovulatory-size follicles that were dysfunctional (Fig. 24B) for the nine heifers treated with each of the four FSH doses (35 IU, 70 IU, 140 IU, 210 IU) used during each superovulation regimen. Extensive variability existed in circulating AMH concentrations and AFC prior to superovulation and in all variables measured as endpoints of the response to the different superovulation regimens among the nine heifers. However, some variables at the different FSH doses (e.g., AFC at the 30 IU and 70 IU doses in Fig. 20B) were skewed (horizontal line within box representing the median value does not divide box into two equal halves) as depicted by the data points (solid dark circles) within boxes and by the whiskers (capped line running through the center of each box showing lowest and highest data point values) for each box in Figs. 20, 22 and 24. Thus, all data shown in Fig. 20, 22 and 24 were transformed (e.g., as depicted in Figs. 198 21, 23, 25) to ensure normality before statistical analyses were conducted. As shown in the Box and Whisker Plots (Figs. 20, 22, 24), the variability in both AMH concentrations and AFC observed among the nine heifers with a small ovarian reserve was sufficiently widespread to reliably evaluate not only if AMH concentration and AFC were repeatable within individual heifers, but also to determine if size of the ovarian reserve as measured by circulating AMH concentrations and AFC prior to superovulation with FSH was linked to the subsequent number and functionality (capacity to ovulate in response to hCG) of the ovulatory-size follicles developing in response to superovulation with different FSH doses. Repeatability of circulating AMH concentration and AFC within individual heifers prior to superovulation with the different FSH doses Circulating AMH concentrations and AFC were highly variable among the individual heifers (Fig. 20). In addition, a total of 8 months was required to complete the four superovulation regimens with the four different FSH doses (9 heifers per dose [7]). Thus, the high variability in AMH and AFC prior to superovulation with FSH among heifers, coupled with potential decline in the ovarian reserve during aging in cattle [53], could contribute to the high variability of responsiveness of heifers to superovulation depicted in Fig. 20. However, we have shown that circulating AMH concentrations and AFC are reliable biomarkers for size of the ovarian reserve in heifers [8]. Thus, to examine the potential age-related changes in size of the ovarian reserve, the four different measurements of AMH concentrations and AFC at approximately 2-month intervals prior to each superovulation regimen for each heifer were used to determine repeatability of circulating AMH concentrations and AFC within individuals. Although circulating AMH concentrations varied 41-fold between individuals ranging from 0.42 to 701 pg/ml (Fig. 20A), AMH concentrations were highly repeatable within 199 individuals (0.94, 1 = perfect). In addition, AFC varied 4.7-fold between individuals ranging from 3 to 14 (Fig. 20B) and repeatability of AFC within individuals was moderate (AFC = 0.39). The linear relationship and correlation of circulating AMH concentrations with AFC prior to superovulation of heifers with the different FSH doses The linear regression analysis indicated that the overall means and linear relationships (slopes) between AMH concentrations with AFC prior to superovulation with the four different FSH doses were similar (P>0.10) among heifers (data not shown). Thus, the average for circulating AMH concentrations (n = 4 measurements per heifer) or AFC (n = 4 measurements per heifer) for the nine heifers was determined. These data were then re-analyzed by linear regression and correlation analyses to further evaluate the association of AMH concentrations with AFC. A positive linear relationship and positive correlation existed between circulating AMH concentrations with AFC prior to superovulation of heifers with FSH (Fig. 21). The linear relationship and correlation between AMH concentration or AFC prior to superovulation with FSH with the endpoints used to measure the response of heifers to the different superovulation regimen The linear regression analysis indicated that the overall means and linear relationships (slopes) between AMH or AFC prior to superovulation with FSH with the endpoints used to measure the response of heifers to the four different superovulation regimens were similar (P>0.10, data not shown). Because there was no evidence that FSH dose during superovulation impacted the relationship between AMH and AFC with the response to superovulation, data collected prior to and during the four superovulation regimens were pooled and averages for each variable (AMH, AFC, total and number of functional and dysfunctional ovulatory-size follicles, proportion of total number of ovulatory-size follicles that were dysfunctional) were determined 200 for each heifer. These data for the nine heifers were then re-analyzed by linear regression and correlation analyses to further evaluate the association between the ovarian reserve biomarkers, AMH and AFC, prior to superovulation with FSH, with the subsequent number and functionality of the ovulatory-size follicles developing in response to superovulation. A positive linear relationship and positive correlation existed in heifers between circulating AMH concentrations and AFC with the subsequent total number of ovulatory-size follicles (Fig. 23) and with number of functional (Fig. 23) and dysfunctional (Fig. 25) ovulatory-size follicles and proportion of total number of ovulatory-size follicles that were dysfunctional (Fig. 25) developing in response to superovulation with FSH during the superovulation regimen. Discussion The goal of the present study was to determine if circulating AMH concentrations and AFC, which are well-established ovarian reserve biomarkers predictive of the responsiveness of individuals to superovulation, were positively associated with number of dysfunctional ovulatory-size follicles developing in response to superovulation with high FSH doses. The most significant finding of this study is that measurements of circulating AMH concentrations and AFC prior to superovulation with FSH were highly predictive of total number of ovulatory-size follicles developing in response to superovulation. However, the differences in AMH concentrations and AFC prior to superovulation with FSH were positively linked to an increased number of both dysfunctional and functional ovulatory-size follicles as well as an increased proportion of ovulatory-size follicles that were dysfunctional independent of the FSH dose used during superovulation. Because measurements of AMH concentration and AFC predict total number but not functionality of the ovulatory-size follicles developing in response to superovulation, which may also impact oocyte quality, these biomarkers are unlikely to be 201 consistently useful to improve IVF or embryo transfer outcomes in heifers with a small ovarian reserve. Another key finding of the present study showed that, despite use of 11- to 12-month-old heifers with a low AFC and small ovarian reserve, circulating AMH concentrations and AFC and correspondingly size of the ovarian reserve, varied extensively among animals but were highly or moderately repeatable, respectively, within individuals and positively associated with each other during the 8 months required to complete the different superovulation regimens. While use of the ovarian reserve biomarkers, AMH and AFC, may be useful to select a more uniform group of heifers (with similar AMH concentrations, AFC, and ovarian reserves) that also respond similarly to superovulation, small-ovarian-reserve-specific superovulation protocols designed to optimize development of functional ovulatory-size follicles will also likely be necessary to enhance ART outcomes. The highest FSH doses used to superovulate heifers in the present study did not impact the relationship between AMH concentrations and AFC with number and functionality of the ovulatory-size follicles developing in response to superovulation. This finding was surprising because the highest FSH doses during superovulation of the same heifers in the present study negatively impacted estradiol production and ovulation rate in our previous study [7]. However, we recognize that only nine animals were used per FSH dose in the present study, thus larger animal numbers may be necessary to further assess effects of high FSH doses on the biomarker- superovulation response relationship. Nevertheless, differences in size of the ovarian reserve, as measured by AMH concentrations and AFC in the heifers in the present study, were positively linked, independent of the FSH dose, with the total number of ovulatory-size follicles and with number of functional (or number of CL post hCG) ovulatory-size follicles developing in 202 response to superovulation. These positive relationships have also been reported by others using cattle [21, 54-56] and women [57, 58] with mixed ages and unknown ovarian reserves. Taken together, these observations support the value of both biomarkers, AMH and AFC, to not only predict the number of ovulatory-size follicles developing in response to superovulation but also the number of functional ovulatory-size follicles capable of ovulation in response to hCG (number of CL). Using reliable biomarkers such as AMH or AFC to predict responsiveness of individuals to superovulation enables ART practitioners to improve superovulation protocols for example to avoid excessive use of FSH, estimate number of oocytes potentially available for IVF or number of transferable embryos, and better inform patients/clients of expected ART outcomes. Nevertheless, the ovarian reserve biomarkers, AMH and AFC, were also positively associated with number of dysfunctional ovulatory-size follicles that did not ovulate in response to an hCG treatment and the proportion of total number of ovulatory-size follicles that were dysfunctional. The capacity to ovulate in response to a preovulatory LH surge is well-recognized by reproductive biologists as a physiologically indispensable criteria for a normal functioning ovulatory follicle with high quality oocytes during reproductive cycles of most mammals, including cattle. Thus, the strong positive correlation between both ovarian reserve biomarkers, AMH and AFC, with number of dysfunctional ovulatory-size follicles, as observed in the present study, implies that size of the ovarian reserve is positively linked not only with number of functional (capable of ovulation and forming CL or number of CL) but also with number of dysfunctional (incapable of ovulation) ovulatory-size follicles independent of the FSH doses used to superovulate heifers with a small ovarian reserve. The reason circulating AMH concentrations and AFC are positively linked with number of dysfunctional ovulatory-size follicles is unknown. Nevertheless, previous studies show that 203 superovulation diminishes capacity of bovine oocytes to develop into blastocysts [59, 60] and an inverse relationship exists between responsiveness to superovulation and proportion of ovulatory-size follicles with high quality oocytes in cattle [27, 29, 30] and women [4, 26, 61]. For example, superovulation of heifers with a high compared with a low AFC result in a greater number of CL, embryos/oocytes recovered and transferable embryos per animal but a lower proportion of transferable embryos and higher proportion of fertilized non-transferable embryos [8, 32]. In addition, studies in cattle with unknown AFC report that the proportion of high-quality embryos transferred is inversely linked to the responsiveness of superovulation [32, 62, 63], and that the number of transferable embryos declines as the number of oocytes recovered increases in women [61]. Taken together, these results indicate that responsiveness of individuals to superovulation is positively linked to increased number of both functional and dysfunctional ovulatory-size follicles with corresponding differences in oocyte quality. Whether oocyte quality is also impaired in the putative dysfunctional ovulatory-size follicles was beyond the scope of the present study. These questions, however, are important to answer to establish new methods to minimize formation of dysfunctional ovulatory-size follicles, reduce oocyte wastage, and improve efficiency of superovulation protocols to improve ART outcomes. Circulating AMH concentrations and AFC in the heifers with small ovarian reserves were positively correlated with each other prior to superovulation with FSH thus confirming results of our previous study that used the mixed group of heifers with low, intermediate or a high AFC [11, 36] as well as results of others using cattle [14, 22, 23, 64, 65] and women [66-68] with a mixed AFC and different ages. In addition, during the 1.5- to 2-month intervals between superovulation regimens in the present study, AMH concentration at emergence of the first follicular wave for each different estrous cycle was highly repeatable (0.91) within individuals. 204 This observation indirectly implies that, despite having a low AFC and small ovarian reserve, growth of the large preantral and small antral follicles potentially responsive to the FSH treatment during each superovulation regimen was also relatively constant at the beginning of each follicular wave within individuals in the present study. Thus, the high variation in response to superovulation observed here is more likely attributable to differences in size of the ovarian reserve among individuals rather than erratic growth of follicles during follicular waves within individuals. Unlike circulating AMH concentrations, AFC reflects the total number of antral follicles ≥3 mm in diameter during a follicular wave and thus is a direct measurement of follicles potentially responsive to superovulation. However, in contrast to AMH concentrations, repeatability of AFC during the first follicular waves within the same individual heifers in the present study was moderate (0.39) and lower than repeatability (0.85 to 0.95) of peak AFC during follicular waves within individuals determined in our previous studies which used heifers with a low, intermediate, or high peak AFC [34]. The precise reason for the disparity in repeatability of AFC within individuals between our studies is unknown. However, FSH secretion is inversely associated with AFC [25, 34] and factors such as heat stress [69] and photoperiod [70] alter FSH secretion and can negatively impact repeatability of AFC in cattle. Whether FSH secretion and, in turn, growth of the larger antral follicles (≥3 mm in diameter) during follicular waves is less repeatable for individual heifers with a small ovarian reserve, as used in our present study, compared with age-matched individuals with a higher AFC is unknown. Alternatively, it is also possible that determination of peak AFC as in our previous study [34] is more repeatable within individuals than determination of AFC at an unknown stage of a follicular wave as in the present study. Nevertheless, grouping individual heifers based on 205 both circulating AMH concentrations and AFC, although time consuming, may be necessary to produce more uniform responses of individuals to superovulation. Conclusion In summary, availability of multiple, developmentally normal ovulatory-size follicles following superovulation is the pivotal physiological event to ensure enough high-quality oocytes are available for successful in vivo or in vitro fertilization and embryo transfer in cattle. Results of the present study, however, showed that the ovarian reserve biomarkers, AMH concentration and AFC, were positively associated with total number of ovulatory-size follicles, number of functional and dysfunctional ovulatory-size follicles, and proportion of total number of ovulatory-size follicles that were dysfunctional developing during superovulation with FSH. Because AMH concentration and AFC predict subsequent number but not functionality of ovulatory-size follicles developing in response to superovulation, which may also impact oocyte quality, measurement of these ovarian reserve biomarkers prior to superovulation with FSH is concluded to unlikely be consistently useful to improve IVF or embryo transfer outcomes in heifers with a small ovarian reserve. However, ovarian-reserve-specific superovulation protocols that optimize number of functional ovulatory follicles capable of ovulation in response to an LH or LH-like stimulus may be necessary to improve efficiency of superovulation procedures and reliability of circulating AMH and AFC to predict ART outcomes in individual heifers with a small ovarian reserve. 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Biol Reprod, 1999. 60(2): p. 349-354. Jeppesen, J.V., et al., Which follicles make the most anti-Müllerian hormone in humans? Evidence for an abrupt decline in AMH production at the time of follicle selection. Mol Hum Reprod, 2013. 19(8): p. 519-527. Takahashi, C., et al., Anti-Müllerian hormone substance from follicular fluid is positively associated with success in oocyte fertilization during in vitro fertilization. Fertil Steril, 2008. 89(3): p. 586-591. Lonergan, P., et al., Effect of follicle size on bovine oocyte quality and developmental competence following maturation, fertilization, and culture in vitro. Mol Reprod Dev, 1994. 37(1): p. 48-53. 60. Blondin, P., et al., Superovulation can reduce the developmental competence of bovine embryos. Theriogenology, 1996. 46(7): p. 1191-1203. 61. Meniru, G.I. and I.L. Craft, Utilization of retrieved oocytes as an index of the efficiency of superovulation strategies for in-vitro fertilization treatment. Hum Reprod, 1997. 12(10): p. 2129-2132. 62. 63. 64. 65. Shaw, D., et al., Effect of retinol palmitate on ovulation rate and embryo quality in superovulated cattle. Theriogenology, 1995. 44(1): p. 51-58. De Roover, R., et al., Characterisation of low, medium and high responders following FSH stimulation prior to ultrasound-guided transvaginal oocyte retrieval in cows. Theriogenology, 2005. 63(7): p. 1902-1913. Cardoso, C.J.T., et al., Anti-Müllerian hormone (AMH) as a predictor of antral follicle population in heifers. Anim Reprod, 2018. 15(1): p. 12-16. Gobikrushanth, M., et al., Repeatability of antral follicle counts and anti-Müllerian hormone and their associations determined at an unknown stage of follicular growth and 211 an expected day of follicular wave emergence in dairy cows. Theriogenology, 2017. 92: p. 90-94. Visser, J.A., et al., Anti-Müllerian hormone: a new marker for ovarian function. Reprod, 2006. 131(1): p. 1-9. Barbakadze, L., et al., The correlations of anti-mullerian hormone, follicle-stimulating hormone and antral follicle count in different age groups of infertile women. Int J Fertil Steril, 2015. 8(4): p. 393-398. Nardo, L.G., et al., Anti-Müllerian hormone levels and antral follicle count in women enrolled in in vitro fertilization cycles: Relationship to lifestyle factors, chronological age and reproductive history. Gynecol Endocrinol, 2007. 23(8): p. 486-493. 66. 67. 68. 69. Wolfenson, D. and Z. Roth, Impact of heat stress on cow reproduction and fertility. Anim Front, 2018. 9(1): p. 32-38. 70. Critser, J.K., et al., Effect of photoperiod on LH, FSH and prolactin patterns in ovariectomized oestradiol-treated heifers. J Reprod Fertil, 1987. 79(2): p. 599-608. 212 APPENDIX Figure 20. Box and whisker plots showing the distribution of AMH concentrations and AFC prior to superovulation of nine heifers with small ovarian reserves with different doses of Folltropin-V (FSH). The same group of Holstein heifers (11- to 12-months old) were superovulated four times with four different FSH doses (n = 9 heifers per dose) followed by a single treatment of hCG 12 h after the last FSH treatment for each superovulation regimen as explained in Methods. AMH concentration and AFC prior to each superovulation among the nine heifers were determined for each different FSH dose. 213 4 3 2 ) t r q s ( t n u o c e l c i l l o f l a r t n A 1 1.0 Y = 0.9X + 1.0 ** r = 0.86 ** 1.5 2.0 AMH (Log10) 2.5 3.0 Figure 21. Regression analysis and correlation coefficients for the relationship between concentrations of anti-Mϋllerian hormone (AMH) and antral follicle count (AFC) before superovulation of heifers with a small ovarian reserve. The same group of Holstein heifers (11- to 12-months old) were superovulated four times with four different FSH doses (n = 9 heifers per dose) followed by a single treatment of hCG 12 h after the last FSH treatment for each superovulation regimen as explained in Methods. Because there was no FSH dose effect, data was pooled. Thus, each symbol represents the mean ±SEM for the four superovulations for each heifer. Concentrations of AMH and AFC (≥3 mm in diameter) were determined 1 to 2 h before the first treatment of FSH. Asterisks indicate level of statistical significance (** = P<0.01). 214 Figure 22. Box and whisker plots showing the distribution of the number of ovulatory-size follicles and the number of functional ovulatory-size follicles observed following superovulation of nine heifers with small ovarian reserves with different doses of Folltropin- V (FSH). The same group of Holstein heifers (11- to 12-months old) were superovulated four times with four different FSH doses (n = 9 heifers per dose) followed by a single treatment of hCG 12 h after the last FSH treatment for each superovulation regimen as explained in Methods. The number of ovulatory-size follicles was determined at time of hCG. The number of functional ovulatory-size follicles was determined by counting the number of ovulatory-size follicles that had formed CL 7 d after the hCG. 215 Figure 23. Regression analysis and correlation coefficients for the relationship between concentrations of anti-Mϋllerian hormone (AMH) and antral follicle count (AFC) before superovulation with total number of ovulatory-size follicles and with number of functional ovulatory-size follicles following superovulation of heifers with a small ovarian reserve. The same group of Holstein heifers (11- to 12-months old) were superovulated four times with four different FSH doses (n = 9 heifers per dose) followed by a single treatment of hCG 12 h after the last FSH treatment for each superovulation regimen as explained in Methods. Because there was no FSH dose effect, data were pooled. Thus, each symbol represents the mean ±SEM for the four superovulations for each heifer. Concentrations of AMH and AFC (≥3 mm in diameter) were determined 1 to 2 h before the first treatment of FSH. The total number of ovulatory-size follicles (≥10 mm) and number of functional ovulatory-size follicles was determined as explained in the legend for Fig. 3. Plus signs or asterisks indicate level of statistical significance (++ = P≤0.07, +++ = P≤0.06, * = P<0.05, ** = P<0.01, *** = P<0.001). 216 Figure 24. Box and whisker plots showing the distribution of the number of dysfunctional ovulatory-size follicles and proportion of total number of ovulatory-size follicles that were dysfunctional following superovulation of nine heifers with small ovarian reserves with different doses of Folltropin-V (FSH). The same group of Holstein heifers (11- to 12-months old) were superovulated four times with four different FSH doses (n = 9 heifers per dose) followed by a single treatment of hCG 12 h after the last FSH treatment for each superovulation regimen as explained in Methods. The number of dysfunctional ovulatory-size follicles that did not ovulate and form CL in response to hCG was determined by subtracting the number of CL formed 7 d after the hCG (number of functional ovulatory-size follicles) from the total number of ovulatory-size follicles observed at time of hCG. The proportion of the total number of ovulatory-size follicles that were dysfunctional was determined by dividing the number of dysfunctional ovulatory-size follicles by the total number of ovulatory-size follicles at time of hCG. 217 Figure 25. Regression analysis and correlation coefficients for the relationship between concentrations of anti-Mϋllerian hormone (AMH) and antral follicle count (AFC) before superovulation with number and proportion of ovulatory-size follicles that were dysfunctional following superovulation of heifers with a small ovarian reserve. The same group of Holstein heifers (11- to 12-months old) were superovulated four times with four different FSH doses (n = 9 heifers per dose) followed by a single treatment of hCG 12 h after the last FSH treatment for each superovulation regimen as explained in Methods. Because there was no FSH dose effect, data was pooled. Thus, each symbol represents the mean ±SEM for the four superovulations for each heifer. Concentrations of AMH and AFC (≥3 mm in diameter) were determined 1 to 2 h before the first treatment of FSH. The number and proportion of dysfunctional ovulatory-size follicles were determined as explained in the legend for Fig. 5. Asterisks indicate level of statistical significance (* = P<0.05, ** = P<0.01, *** = P<0.001). 218 CHAPTER 7. Synopsis, Conclusion, and Practical Applications 219 Synopsis High FSH doses are used during ovarian stimulation of infertile women to maximize the number of oocytes available for IVF even though high FSH doses are well-established to be inversely correlated with live birth rate during ART. These observations provided rationale to use the SORH model to test the hypothesis that high FSH doses during ovarian stimulation are detrimental to ovulatory follicle function and oocyte quality. The following is a synopsis of the findings from the five published studies (Chapters II, III, IV, V, VI) that comprise my thesis research: Chapter II: Negative impact of high doses of follicle-stimulating hormone during superovulation on ovulatory follicle function in small ovarian reserve dairy heifers [1]. To determine the effects of high FSH doses on ovarian function, the SORH model (n = 9 heifers per dose) was treated with four different doses (8 injections at 12-h intervals per dose) of cpFSH during the ovarian stimulation protocol and then given an ovulatory dose of hCG 12 h after the last cpFSH injection. Serial ovarian ultrasonography and frequent blood sampling were used to monitor follicular growth during ovarian stimulation, ovulation in response to hCG, and alterations in ovarian hormone production. Results demonstrated the following: a) The use of the 210 IU cpFSH dose (3x industry-standard) compared with lower cpFSH doses during ovarian stimulation did not further increase the number or the size of ovulatory-size follicles developing during ovarian stimulation. Thus, the 210 IU cpFSH dose was hereafter referred to as “excessive”. b) Use of the excessive cpFSH (Ex-cpFSH) during ovarian stimulation decreased both the circulating estradiol concentrations and ovulation rate, which are well-established hallmarks of ovulatory follicle function. 220 Conclusion: Use of the 210 IU cpFSH dose during ovarian stimulation of the SORH model is potentially detrimental to ovulatory follicle function. This seminal observation provided the rationale to investigate the impact of Ex-cpFSH action on ovulatory follicle function directly in the next study. Chapter III: Excessive FSH during ovarian stimulation of cattle may induce premature luteinization of most ovulatory-size follicles [2]. To examine the effects of Ex-cpFSH on ovulatory follicle function, the SORH model was treated with the Ex-cpFSH or the industry-standard cpFSH (IS-cpFSH) dose during ovarian stimulation. Ovaries were removed from the SORH model 12 h after the last cpFSH injection, and each ovulatory-size follicle was excised from both ovaries. Follicular fluid (FF) was collected from each excised follicle and analyzed for concentrations of estradiol, progesterone, and oxytocin. Cumulus-oocyte complexes (COCs) were also recovered from the FF of each ovulatory-size follicle and classified as having a compact (comCOC) or expanded (expCOC) layer of cumulus cells. Results demonstrated the following: a) The majority (79%) of ovulatory-size follicles developing in response to the IS-cpFSH dose were classified as estrogen-active (EA) because they had higher intrafollicular concentrations of estradiol than progesterone. The EA follicles also had relatively low intrafollicular oxytocin concentrations and comCOCs. Hereafter, these IS-cpFSH-induced ovulatory-size follicles were referred to as Type 1 (control). b) In contrast to the predominant phenotype (Type 1) of the ovulatory-size follicles developing in response to the IS-cpFSH dose, there were nearly equal proportions (22%-28%) of four different phenotypes for the ovulatory-size follicles developing in each heifer in response to treatment with the Ex-cpFSH dose. The Ex-cpFSH induced phenotypes were EA with comCOCs 221 (Type 2, like Type 1), EA with expCOCs (Type 3), EI (estrogen-inactive, higher intrafollicular concentrations of progesterone than estradiol) with expCOCs (no Type assigned because they were omitted from RNAseq analysis in Chapter III), or EEI (extremely estrogen-inactive with very high intrafollicular concentrations of progesterone and oxytocin) with expCOCs (Type 4). The diversity of phenotypes of the ovulatory-size follicles developing in response to Ex- cpFSH during ovarian stimulation may reflect differences in exposure of each follicle in the first follicular wave to different concentrations of cpFSH because of inherent differences of blood flow from the ovarian arteries into the theca cell capillary network [3, 4] of each growing follicle as depicted in Figure 26. The differences in the theca capillary network likely result from relatively minor differences in the age, stage of differentiation, and size [5] of the small antral follicles at the beginning of the first follicular wave of the estrous cycle. If this possibility is correct, the unequal distribution of cpFSH into the small antral follicles that comprise the follicular wave would therefore also be expected to occur in the IS-cpFSH treated animals. However, the concentrations of cpFSH in each follicle of the IS-cpFSH treated heifers are postulated to be below the threshold necessary to dysregulate follicular function. Thus, the IS- cpFSH treatment during ovarian stimulation is hypothesized to result in a “normal” physiological range of cpFSH concentration in the thecal vasculature of most of the growing follicles in the first wave, thereby resulting in the development of predominantly healthy, EA ovulatory-size follicles with low oxytocin levels and comCOC, as observed in this study and depicted in Figure 26. c) Overall, in response to the Ex-cpFSH doses, 46% of the ovulatory-size follicles were EI while 72% of the ovulatory-size follicles had expCOCs. These observations demonstrated that the use of Ex-cpFSH doses during ovarian stimulation induces premature (prior to an hCG 222 ovulatory stimulus) luteinization (simultaneous loss of intrafollicular capacity to produce estradiol but enhanced capacity to produce progesterone and oxytocin) and premature cumulus expansion (marker for resumption of meiosis) of most ovulatory-size follicles. d) The Ex-cpFSH-induced premature luteinization and associated reduction in intrafollicular estradiol production (and presumed loss of FSH responsiveness) of most ovulatory-size follicles likely explains why circulating estradiol concentrations and perhaps responsiveness to the hCG ovulatory stimulus (presumed loss of FSH responsiveness could also block upregulation of the LH receptor) were both decreased following ovarian stimulation with Ex-cpFSH doses in my first study [1]. However, whether the ovulatory-size follicles developing in response to Ex-cpFSH doses that failed to ovulate in response to the hCG ovulatory stimulus, as observed in my first study [1], were also EI and prematurely luteinized, as shown in the present study (Figure 26), remains a critical question to answer. Moreover, I should point out that even the IS-cpFSH treated heifers have a relatively small proportion of EI ovulatory-size follicles (21%, not depicted in Figure 26) that may also be prematurely luteinized, which could explain why ovulation rates in the controls was ~80% rather than 100%. e) Results of the present study also show that Ex-cpFSH doses during ovarian stimulation induce premature cumulus expansion in most ovulatory-size follicles. This finding implied that cumulus function and, in turn, oocyte quality in the ovulatory-size follicles developing in response to the Ex-cpFSH may also be impaired. This observation provided rationale to examine the impact of excessive FSH action on cumulus function and oocyte quality directly in Chapter IV. 223 Conclusion: Use of the Ex-cpFSH doses during ovarian stimulation of the SORH model induces premature luteinization and cumulus expansion, and thus is detrimental to ovulatory follicle function. The next study was designed to better understand the mechanisms whereby excessive FSH action during ovarian stimulation impairs ovulatory follicle function by determining if the Ex-cpFSH disrupts signaling pathways in granulosa and cumulus cells and oocytes critical for function and oocyte quality in the Type 2, 3 and 4 ovulatory-size follicles. Chapter IV: Follicular hyperstimulation dysgenesis: new explanation for adverse effects of excessive FSH in ovarian stimulation [6]. To examine the effect of Ex-cpFSH during ovarian stimulation on signaling pathways, RNAseq and bioinformatic analyses were used to determine if gene expression in the granulosa and cumulus cells and oocytes harvested from a subset of the excised ovulatory-size follicles described in Chapter III (also depicted in Figure 26) differed between the IS-cpFSH-treated Type 1 (control) compared with the Ex-cpFSH-treated Type 2, 3 and 4 ovulatory-size follicles (n = 4 to 6 ovulatory-size follicles or heifers per cell type per cpFSH dose). Highlights of the results of this study follow: a) mRNAs for ~15,000 of the bovine’s 22,000 genes were detected in each cell type during RNAseq analysis resulting in a robust coverage of the bovine’s transcriptome. b) A total of 3,108 unique (no overlap) differentially expressed genes (DEGs) were identified when the Type 1 (controls) were compared with the Type 2, 3 and 4 ovulatory-size follicles. c) There was usually a progressive increase in the total number of DEGs, the number of the DEGs up- or down-regulated by hormones well-established to control ovulatory follicle 224 function (e.g., estrogen-, FSH-, or LH-signaling, progesterone, testosterone, or oxytocin), and the degree of over-expression of the DEGs identified in granulosa and cumulus cells and oocytes when the Type 2 vs 3 vs 4 were compared with the Type 1 (control) ovulatory-size follicles. d) The majority of the DEGs (total for all cell types combined = 5,104) were in cumulus (3,131/3,288 = 95%) and granulosa (1,368 of 1,531 total = 89%) cells and oocytes (283/285 = 99%) of the Type 4 ovulatory-size follicles. Thus, the Type 4 phenotype, which had extremely high intrafollicular progesterone and oxytocin concentrations (prematurely luteinized) and expCOC and comprised ~25% of the ovulatory-size follicles developing in response to ovarian stimulation with Ex-cpFSH (Chapter III, Figure 26), also displayed the most abnormal transcriptome. e) As observed in Chapter III, ~50% of the ovulatory-size follicles developing in response to Ex-cpFSH were classified as EI (includes Type 4) and thus considered prematurely luteinized. However, only 35% and 45% of the list of 71 manually curated published luteinization markers in the present study were identified as DEGs in the cumulus or granulosa cells, respectively, primarily in the Type 4 ovulatory-size follicles. In addition, the direction (up or down) of gene expression of these DEGs was occasionally inconsistent with literature expectations. These observations imply that the premature luteinization of the ovulatory-size follicles induced by Ex-cpFSH during ovarian stimulation is very likely abnormal compared with the process of luteinization of ovulatory follicles induced by a preovulatory gonadotropin (LH and FSH) surge during the bovine estrous cycle. f) Expression of the FSH receptor (FSHR) mRNA was lower (tendency) in cumulus and granulosa cells of the Type 4 vs Type 1 (control) ovulatory-size follicles. FSH is well- established to up-regulate expression of FSHR, the LH receptor (LHCGR) and aromatase gene 225 (CYP19A1, critical for estradiol production) in granulosa [7, 8] and cumulus [9, 10] cells. Thus, the finding in the present study that FSHR expression is reduced in granulosa and cumulus cells implies that responsiveness of these cell types to FSH stimulation was diminished. If so, the loss of FSH responsiveness may also explain why both the LHCGR and CYP19A1 were not up- regulated in the present study and why intrafollicular estradiol concentrations (Chapter III) and expression of the estradiol receptor (ESR2) mRNA in granulosa and cumulus cells were both lower in the Type 4 vs Type 1 (control) ovulatory-size follicles in the present study. g) In contrast to the downregulation of CYP19A1 and estradiol production, the Type 4 ovulatory-size follicles had increased expression of mRNAs for enzymes critical for progesterone synthesis (STAR, HSD3B1) and for oxytocin production (OXT) in both granulosa and cumulus cells. These combined (f, g) observations potentially explain why intrafollicular progesterone and oxytocin production were enhanced, while estradiol production is abrogated in Type 4 ovulatory-size follicles (Chapter III). However, whether the excessive Ex-cpFSH action per se, or other indirect mechanisms such as those linked to premature luteinization (Chapter III) or over-expression of other regulatory factors that may regulate progesterone production, are sufficient to enhance production of the enzymes critical for progesterone and oxytocin production in Type 4 ovulatory-size follicles is unknown. h) The number of DEGs (Volcano plots) and expression values for DEGs (correlation heatmap) indicated that granulosa and cumulus cells were most alike in the extreme Type 4 phenotype, perhaps because both cell types were undergoing abnormal luteinization in response to Ex-cpFSH doses during ovarian stimulation. i) Whether the aberrant and prematurely luteinized Type 4 ovulatory-size follicles were possibly also atretic was examined by comparing the DEGs for the Type 4 vs Type 1 (control) 226 with a list of 183 published bovine ovarian atresia markers. However, only a few of these atretic markers were identified as DEGs in granulosa and cumulus cells implying that apoptosis was not predominant in the Type 4 follicles. j) Because of the enormous number of DEGs (5,104) identified in the different cell types for the Type 2, 3 and 4 compared with the Type 1 (control) ovulatory-size follicles, Ingenuity Pathway Analysis (IPA) and activation z-score analysis were used to identify which major canonical pathways, biological functions, and upstream regulators of transcription were potentially activated or inhibited during ovarian stimulation with the Ex-cpFSH. IPA uses proprietary software based on a manually structured and integrated Knowledge Base of an extraordinarily large number of published data sets. For example, 2,000,000 peer- reviewed publication and a total of 14.5 million findings, and >30 third-party databases are used to capture a wide range of data including metabolomics and proteomics from a variety of tissues and models. The information generated during IPA is used to determine which pathways, biological functions and upstream regulators may be impacted by the DEGs based on statistically significant over-representation of the DEGs specific to each pathway, biological function, or upstream regulator. k) The number of canonical pathways, biological functions, and upstream regulators with significant activation z-scores (implies an activated or inhibited state of action for the pathway, function, or upstream regulator) was progressively increased when the granulosa and cumulus cells for the Type 2 or 3 and 4 were compared with the Type 1 (control) ovulatory-size follicles. However, the cell types for the Type 4 ovulatory-size follicles had the majority of activated (A) or inhibited (I) canonical pathways (cumulus: A =152 of 155 or 98% of the total for Type 2, 3 and 4 combined or 152/155 (98%), I=5/5 (100%); granulosa: A=23/27 (85%), I=1/1 (100%), 227 oocyte: A=1/1 (100%), I=1/1 (100%)), biological functions (cumulus: A=138/191 (72%), I=9/17 (53%); granulosa: A=153/257 (60%), I=12/20, (60%) oocyte: A=1/1 (100%), I=5/5 (100%)), and upstream transcriptional regulators (cumulus: A=240/321 (75%), I=76/80 (95%); granulosa: A=391/592 (66%), I=118/144 (82%), oocyte: A=4/4 (100%), I=11/11 (100%)) with significant activation z-scores. This observation further illustrates the high degree of abnormality in the transcriptomes for the granulosa and cumulus cells and oocytes of Type 4 compared with the Type 2 and 3 ovulatory-size follicle phenotypes during ovarian stimulation with Ex-cpFSH. l) The Type 4 ovulatory-size follicles had the most extreme phenotype characterized by diminished intrafollicular estradiol production but very high intrafollicular production of progesterone and oxytocin and prematurely expanded COC (Chapter III). Type 4 also had the greatest number and degree of over-expression of DEGs, and the most activated or inhibited canonical pathways, biological functions, and upstream transcriptional regulators (Chapter IV). Thus, the IPA Causal Network Analysis tool used the upstream transcriptional regulators identified by IPA in this study to identify the upstream transcriptional regulators linked to FSH stimulation and in turn to the downstream effects on DEGs, canonical pathways and biological functions. This information was used to define a putative mechanism by which Ex-cpFSH alters the cellular phenotype of the granulosa and cumulus cells and oocytes in Types 3 and 4 ovulatory-size follicles. m) Results of the IPA Causal Network Analysis predicted activation of FSH and LH signaling in granulosa and cumulus cells (Chapter IV, Figure 15). Activation of LH signaling, however, likely reflects the well-established overlap (70% in this study) between FSH and LH signaling pathways. Activation of FSH signaling was linked to increased gene expression or 228 activation of downstream signaling pathways in both granulosa and cumulus cells for Type 3 and 4 ovulatory-size follicles. This includes predicted increases in signaling through FSH (FSHR) and LH (LHCGR) receptorsA [11, 12], arrestin (ARRß1 and ARRß2)B [13], ß-estradiol production, and increased expression of mRNAs encoding Kit Ligand (KITLG)C [14], transforming growth factor ß1 (TGFß1)D [15], endothelial PAS domain protein 1 (EPAS1)E [16], and signal transducer and activator of transcription 3 (STAT3)F [17], but inhibition of signaling through the estrogen receptor complex (consists of ERβ and ERα)G [18]. These up-stream transcriptional regulators are well-established to have a critical role in follicular differentiation, function, survival, and ovulation and were established to up- or down-regulate many of the DEGs identified in the granulosa and cumulus cells of Type 3 and 4 ovulatory-size follicles in the present study. These predicted upstream events and their connection to the affected DEGs predicted activation of integrin signalingH [19], STAT3 signaling, interleukin signaling (IL)I [19, A Critical for growth, differentiation and function of granulosa and cumulus cells. B Regulates desensitization/re-sensitization of G protein-coupled receptors and thus cAMP and phosphatidylinositol signaling. C Cytokine regulating signaling pathways such as STAT involved in cell survival, growth, and function. D Regulates SMAD transcription factors that regulate cell growth, differentiation, apoptosis, and modulates actions of other growth factors. E Hypoxia-inducible factor 2–alpha or HIFA, regulates blood vessel formation. F Regulates cell growth and movement, apoptosis, follicular function, meiotic resumption, and bovine cumulus expansion. G G-protein-coupled estrogen receptor, critical for growth, differentiation, steroidogenesis, and ovulation. H Transmembrane receptors facilitating extracellular matrix adhesion involved in cumulus expansion and luteinization. I Cytokine involved in inflammation, cumulus expansion and luteinization and resumption of meiosis. 229 20], and cytoskeleton organizationJ [21], along with inhibition of Rho GDP-dissociation inhibitor signaling (RHOGDI)K [22, 23]. These results of the IPA Causal Network Analysis thus define a potential pathway whereby Ex-cpFSH doses during ovarian stimulation may alter many upstream transcriptional regulators and downstream DEGs that in turn activate or inhibit several key signaling pathways and biological functions in granulosa and cumulus cells. These affected functions would, in turn, impact a range of cellular functions critical for follicular function, ovulation and fertility, such as oocyte meiosis, cumulus expansion, extracellular matrix adhesion, luteinization and apoptosis. These Ex-cpFSH-induced alterations in granulosa and cumulus function may have caused, contributed to, or were a consequence of the aberrant premature luteinization (and associated loss of intrafollicular capacity to produce estradiol but enhanced capacity to produce progesterone and oxytocin) and premature cumulus expansion observed in the Type 3 and 4 ovulatory-size follicles (Chapter III). n) Results of the IPA Causal Network Analysis also identified three interconnected upstream transcriptional regulators predicted to be affected in oocytes of Type 4 ovulatory-size follicles (Chapter IV, Figure 15). The predicted decrease in ß-estradiol signaling was linked to inhibition of HSPA5 expressionL [24]. HSPA5 is an oocyte stress protein [25] involved in protein folding in the endoplasmic reticulum (ER) and signal transduction at the cell surface. Downregulation of HSPA5 may promote ER stress and resumption of meiotic maturation [26] J Consist of filaments and fibres in cytoplasm critical for changes in cell shape such as luteinization and cumulus expansion. K Downregulates Rho GTPases to maintain GTPase activity in cytosol which in turn enhances actin production, which is likely necessary for luteinization, cell adhesion, membrane trafficking and bovine cumulus expansion and apoptosis. L Codes for binding immunoglobulin protein (BiP). 230 and increased activation of MYC signalingM [27]. In addition, overexpression of MYC induces apoptosis during meiosis of spermatocytes [28] and results in global transcription of active genes and dysregulation of chromatin interactions [29]. These upstream regulators were predicted to be linked to downstream inhibition of several processes important for oocyte biology and quality, including EIF2 signalingN [30], cell survivalO [31], and phagocytosisP [32]. o) Many additional upstream transcriptional regulators, canonical pathways and biological functions were not linked by our IPA Causal Network Analysis to Ex-cpFSH stimulation. The absence of such connections in IPA does not negate the possible existence of such connections, as this may reflect an absence of data in the IPA Knowledge Base. Because these other effects were induced by Ex-cpFSH stimulation, however, it is reasonable to infer that such effects constitute additional, indirect consequences of excessive FSH, even if not previously reported in the literature. Conclusion: Ex-cpFSH doses during ovarian stimulation simultaneously disrupt multiple cell- signaling pathways in granulosa and cumulus cells and oocytes critical for folliculogenesis, steroidogenesis, luteinization, cell survival, ovulation, and oocyte maturation and quality resulting in follicular hyperstimulation dysgenesis (FHD) in most ovulatory-size follicles (Figure 26). Thus, the FHD likely contributes to, or causes, the reduced intrafollicular estradiol but enhanced progesterone and oxytocin production, premature luteinization and cumulus expansion, reduced responsiveness to hCG and potentially impaired oocyte quality observed in most M Family of genes coding for phosphoproteins involved in cell replication. N Eukaryotic Initiation Factor 2, phosphorylation of EIF2 inhibits initiation of translation in a variety of stress response genes. O Apoptosis. P Important for fertilization. 231 ovulatory-size follicles developing in response to Ex-cpFSH doses during ovarian stimulation of the SORH model. These observations provided the rationale to examine cumulus function and oocyte quality more directly in the next study. Chapter V: Ovarian stimulation with excessive FSH doses causes cumulus cell and oocyte dysfunction in small ovarian reserve heifers [33]. To further examine the effects of Ex-cpFSH during ovarian stimulation on cumulus function and oocyte quality, I conducted the following two-part study. First, to determine if Ex- cpFSH altered expression of genes critical for cumulus function and oocyte maturation, the RNAseq data generated in Chapter IV was manually interrogated to determine if DEGs critical for cumulus function and oocyte quality were identified in Type 2, 3 and 4 compared with Type 1 (control) ovulatory-size follicles. Second, to determine the impact of Ex-cpFSH on cumulus cell morphology and nuclear maturation of oocytes, the SORH models (n = 19 to 20 animals per dose) was subjected to ovarian stimulation with IS-cpFSH or Ex-cpFSH doses as explained in Chapter II. Oocyte pick-up (OPU) was used to recover COCs either 12 h after the last cpFSH injection (before hCG injection) or 24 h after an ovulatory dose of hCG (before ovulation). COCs were classified as comCOCs or expCOCs as explained in Chapter III. The stage of nuclear maturation (germinal vesicle, GV; germinal vesicle breakdown, GVBD; meiosis I, MI; or meiosis II, MII) of oocytes was then determined. The stage of nuclear maturation was also determined after comCOCs collected prior to an hCG stimulus were subjected to in vitro maturation (IVM). Additionally, a subset of comCOCs and expCOCs were subjected to in vitro fertilization (IVF) to determine their ability to undergo successful fertilization. Results demonstrated the following: 232 a) Ex-cpFSH doses during ovarian stimulation prematurely increased expression of 17 genes in cumulus cells (AREG, CLDN11, FGG, GPR50, IGFBP3, IGFBP5, LIF, MDFI, MEPE, MGAT5, NCS1, NGFR, PLAT, PTX3, RGS2, TGFα, VGF) with well-established roles in regulation of cumulus function (Chapter V, Table 3) independent of their ovulatory-size follicle phenotype (Type 2, 3, or 4; Figure 27). Over-expression of these cumulus genes during ovarian stimulation with Ex-cpFSH may have caused or contributed to the premature cumulus expansion and resumption of meiosis observed in most ovulatory-size follicles prior to an ovulatory hCG stimulus in this study. b) In response to ovarian stimulation with Ex-cpFSH, nearly all (17 of 19) cumulus genes were over-expressed in the cumulus but not the granulosa cells in Type 2, 3 and 4 compared with the Type 1 (control) ovulatory-size follicles (Figure 27). However, the Type 2 ovulatory-size follicles had similar intrafollicular concentrations of estradiol, progesterone and oxytocin and contained comCOCs like Type 1 (control). This finding implies that Ex-cpFSH induced over- expression of the cumulus genes precedes the morphological changes resulting in cumulus expansion and precedes the disruption in intrafollicular hormone production leading to premature luteinization in the more extreme Type 3 and 4 phenotypes. This observation provides rationale to determine if the Ex-cpFSH-induced cumulus cell DEGs are potential secretory biomarkers not only for premature cumulus expansion but for ovulatory follicle health and oocyte quality during ovarian stimulation in future studies. c) Ex-cpFSH induced premature (prior to ovulatory hCG stimulus) cumulus expansion and premature resumption of meiosis in a moderate proportion of ovulatory-size follicles (Figure 27). 233 d) Ex-cpFSH reduced the capacity of prematurely expanded COCs to undergo IVF and resume meiosis during IVM (Figure 27). e) Ex-cpFSH reduced the capacity of the COCs to undergo cumulus expansion and resume meiosis in response to the ovulatory hCG stimulus (Figure 28). However, the effect of the ovulatory hCG stimulus on the prematurely over-expressed 17 cumulus genes observed in this study was not evaluated. f) The proportion of ovulatory-size follicles with expanded cumulus cells before and after an ovulatory hCG stimulus were nearly identical during ovarian stimulation with the Ex-cpFSH. This observation implies that Ex-cpFSH doses during ovarian stimulation blocked responsiveness of cumulus cells to an LH or LH-like stimulus. Thus, only the poor quality, prematurely expanded COCs (expCOCs) would be available for IVF following the ovulatory hCG stimulus (Figure 27). Nevertheless, the expCOCs collected post-hCG were not subjected to IVF in this study. Therefore, future studies will be necessary to evaluate whether the comCOCs recovered after Ex-cpFSH treatment and the ovulatory hCG stimulus are capable of IVF. The outcome of this future study would firmly establish in the SORH model if the Ex-cpFSH doses during ovarian stimulation are a major cause of the high oocyte and embryo wastage observed during ART [34-41]. g) A high proportion of the ovulatory-size follicles developing in response to ovarian stimulation with the IS-cpFSH doses had comCOCs and thus failed to respond to the ovulatory hCG stimulus. Consequently, even use of the IS-cpFSH doses during ovarian stimulation may be detrimental (e.g., down-regulation of the cumulus LH receptor and thus reduced sensitivity to hCG [42-44]) to ovulatory follicle function, especially when coupled with the observation that 234 ~20% of the ovulatory-size follicles in controls fail to ovulate in response to an ovulatory hCG stimulus as observed in Chapter II. Conclusion: Ovarian stimulation with Ex-cpFSH dysregulates cumulus function and prematurely induces cumulus expansion and oocyte maturation, and inhibits responsiveness to hCG thereby raising the risk of recovery of a high proportion of poor-quality expCOCs (incapable of being fertilized) that are indistinguishable from healthy expCOCs for IVF during ART. The next study determined if the well-established ovarian reserve biomarkers, antral follicle count (AFC) and anti-Müllerian hormone (AMH), could be used to reliably predict responsiveness of the SORH model to ovarian stimulation with different cpFSH doses. Chapter VI: Limitations in use of ovarian reserve biomarkers to predict the superovulation response in small ovarian reserve heifers [45]. To determine if the ovarian reserve biomarkers, AFC and AMH concentration, predicted responsiveness to ovarian stimulation, both biomarkers were measured prior to ovarian stimulation of the SORH model (n = 9 heifers per dose) with the four different cpFSH doses as explained in Chapter II. AFC and AMH prior to ovarian stimulation were then compared with the total number of ovulatory-size follicles, the number of functional ovulatory-size follicles (the number that ovulated in response to hCG based on the total number of corpora lutea), and the number of dysfunctional ovulatory-size follicles (the number that did not ovulate in response to hCG based on the total number of ovulatory-size follicles minus the number of functional ovulatory-size follicles or corpora lutea) determined 12 h after the last cpFSH injection. Results demonstrated the following: 235 a) The dose of cpFSH, including the Ex-cpFSH dose, used during ovarian stimulation did not impact the degree of positive correlation between AFC or AMH with the total number of ovulatory-size follicles or the number of functional or dysfunctional ovulatory-size follicles (Figure 29). b) Numerous precautions were taken to minimize variability in the responsiveness of individual heifers to ovarian stimulation in this study, including standardization of age, weight, breed, parity, diet, housing, size of the ovarian reserve, timing of FSH injections relative to follicular waves, and source of FSH. Nevertheless, the variability in responsiveness of each heifer to ovarian stimulation remained extremely high ranging from 3 to 70 ovulatory-size follicles. c) Despite the high variability in response to ovarian stimulation among individuals, measurements of AFC and circulating AMH concentrations prior to ovarian stimulation were highly positively correlated with the total number of ovulatory-size follicles, and the number of functional and dysfunctional ovulatory-size follicles, independent of cpFSH dose used during ovarian stimulation (Figure 29). d) AFC and circulating AMH concentrations were also positively correlated with the proportion of the total number of ovulatory-size follicles developing in response to ovarian stimulation that were dysfunctional. This observation implies that there is an inverse relationship between size of the ovarian reserve (as measured by AFC or AMH) and the number of functional ovulatory follicles capable of ovulation in response to an hCG ovulatory stimulus during ovarian stimulation, independent of the cpFSH dose used. This finding supports previous results of others (see Chapter VI for references) and may imply that ovarian stimulation per se is detrimental to ovulatory follicle function and perhaps oocyte quality. 236 Conclusion: Measurements of the ovarian reserve biomarkers, AFC and AMH, prior to ovarian stimulation are predictive of the number of both functional and dysfunction ovulatory-size follicles. Consequently, I conclude that these biomarkers are unlikely to be useful to consistently improve IVF or embryo transfer outcomes in the SORH model. Once new protocols are developed to maximize the number of functional estrogen-active ovulatory-size follicles developing during ovarian stimulation, however, AFC and AMH measurements would be useful biomarkers to identify groups of relatively uniform responders to ovarian stimulation, albeit time-consuming and expensive. This positive outcome would be expected to avoid excessive use of FSH, better estimate the number of oocytes potentially available for IVF or the number of transferable embryos, and better inform patients/clients of expected ART outcomes. Overall Conclusion The results of the five studies described in my thesis support the overall conclusions that: i) ovarian stimulation with Ex-cpFSH alters gene expression and dysregulates numerous signaling pathways in granulosa and cumulus cells and oocytes critical for ovulatory follicle function and oocyte quality, thereby increasing the likelihood of oocyte wastage and decreased ART outcomes, and ii) AMH and AFC are not predictive of the impact of Ex-cpFSH doses during ovarian stimulation on the number of high-quality oocytes available for ART. Based on the results of the studies described in my thesis, I accept my over-arching hypothesis that high FSH doses during ovarian stimulation are detrimental to ovulatory follicle function and oocyte quality. 237 Practical Applications ART in women: ART is a $4.9 billion industry contributing ~2% of the infants born annually in the U.S. [46]. However, despite the high costs of ART and high probability of unsuccessful ART outcomes potentially attributable to the inverse relationship between FSH doses and live birth rates, clinicians continue to use the high-FSH-dose approach to reverse infertility in couples that desire to start a family. The high-FSH-dose protocols embedded in many of the 489 fertility clinics performing ART in the U.S. [47] will likely continue until clinicians have a better mechanistic understanding of the consequences of ovarian stimulation with high FSH doses on ovulatory follicle function and oocyte quality. My studies, however, circumvented the ethical barrier of using women for such studies because the SORH model is biomedically relevant and thus produced results potentially significant to ART practices. The doses of cpFSH used in the SORH model in my studies cannot be directly compared with doses of recombinant hFSH (rhFSH) used during ART in women (total dose range = < 1000 to 20,000 IU [48]). Nevertheless, the studies in my thesis provide compelling evidence supporting the warnings of Nobel Prize winner R. G. Edwards [49, 50] that doses of FSH during ovarian stimulation are detrimental to oocyte quality and embryo survival. In my studies, doses of cpFSH only three-fold higher than the industry-standard cpFSH doses during ovarian stimulation were detrimental to ovulatory follicle function because they reduced intrafollicular capacity to produce estradiol, triggered premature luteinization, cumulus expansion and resumption of meiosis, and reduced oocyte quality prior to an hCG ovulatory stimulus to induce oocyte maturation for IVF. My studies also provided mechanistic insight into how the excessive FSH action during ovarian stimulation caused ovulatory follicle dysfunction by showing that numerous genes and multiple signaling pathways in granulosa and cumulus cells and oocytes, well-established to regulate 238 folliculogenesis, steroidogenesis, luteinization, cell survival, cumulus expansion, ovulation, and oocyte maturation and quality, were altered in most ovulatory-size follicles. Moreover, a high proportion of all ovulatory-size follicles developing in response to ovarian stimulation with Ex- cpFSH contained poor quality expCOCs incapable of fertilization that cannot be distinguished morphologically from high quality hCG-matured expCOCs. These findings raise a cautionary red flag to clinicians that after a patient receives excessive FSH doses during the ovarian stimulation protocol, there is a high risk of oocyte wastage and recovery of poor quality hCG- matured expCOCs for IVF. However, it is currently impossible to determine when during the ovarian stimulation protocol, the FSH doses are excessive and thus detrimental to oocyte quality. Moreover, whether an FSH dose is excessive very likely varies from patient to patient. However, the discovery during my thesis research that excessive FSH doses reduce circulating estradiol concentrations and cause a significant over-expression of 17 genes specific to the cumulus in the SORH model may have practical application towards improvement of ART outcomes in women. Nevertheless, future studies using the SORH model must demonstrate that the decrease in circulating estradiol concentrations or the enhanced secretion of some of the over-expressed cumulus gene products during ovarian stimulation with excessive FSH doses are reliable biomarkers for impending follicular hyperstimulation dysgenesis and cumulus dysregulation. I also need to establish that a reduction in FSH dosages based on these biomarkers will indeed increase the number of high-quality oocytes available for IVF, thereby improving ART outcomes. Art in cattle: The International Embryo Technology Society (IETS) 29th Annual Report [51] on embryo transfer in cattle indicates that North America (primarily the United States) is the leading country for in vivo and in vitro embryo production. Data for 2019 indicates that of the 239 total of 744,009 embryos transferred into recipients, 218,926 embryos (total = ~1.5 million globally) were in vivo derived (IVD) whereas 525, 078 (total globally = 1,010,680) were in vitro produced (IVP) embryos. The IETS report also indicates there is a global trend since 2001 of decreasing numbers of IVD but increasing numbers of IVP embryos including frozen transfers into recipients. For example, in the past 5 years, IVP in the U.S. has increased 145% [51]. Regardless of whether IVD or IVP procedures are used to generate embryos for cattle in the U.S., cattle undergo ovarian stimulation protocols with cpFSH to maximize the number of ovulatory-size follicles and thus high-quality oocytes available for IVD or IVP embryos [52]. The cpFSH dosages used during ovarian stimulation by the bovine embryo transfer industry are usually 70 IU (IS-cpFSH) for the first four injections followed by a decreasing dose regimen which is much lower than the excessive 210 IU dose of cpFSH used in my studies. My results indicate that AFC and AMH are potentially useful to the bovine embryo transfer industry to predict responsiveness to ovarian stimulation, as explained by the caveat articulated in Chapter VI. My results also caution against the use of high FSH doses, for example, to potentially reduce the number of cpFSH injections during ovarian stimulation as done by others [53-55]. Moreover, as explained in Chapter V, a high proportion of COCs in the ovulatory-size follicles developing in response to the IS-cpFSH dose failed to respond to an ovulatory hCG stimulus. Thus, the IS- cpFSH dose may also have detrimental effects on ovulatory-follicle function and oocyte quality. Therefore, conducting further studies utilizing the SORH model may not only have outstanding translational relevance to human ART practices, but generate valuable information for the bovine embryo transfer industry by advancing the use of even lower cpFSH doses than are typically used during ovarian stimulation protocols. The use of lower cpFSH doses during ovarian stimulation would not only reduce the exorbitant costs of cpFSH (700 IU per 20-ml vial of 240 Folltropin-V = $225; recommended dosage is 2.5 ml x 8 injections or 1 vial per animal), but enhance ART outcomes, potentially independent of the ovarian reserve size in cattle. 241 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 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Hayden CB, Sala RV, Pereira DC, Moreno JF, García-Guerra A. Effect of use and dosage of p-follicle-stimulating hormone for ovarian superstimulation before ovum pick up and in vitro embryo production in pregnant Holstein heifers. J Dairy Sci 2023. https://doi.org/10.3168/jds.2023-23576 Yamamoto M, Ooe M, Kawaguchi M, Suzuki T. Superovulation in the cow with a single intramuscular injection of FSH dissolved in polyvinylpyrrolidone. Theriogenology 1994; 41:747-755. 54. Mapletoft R, Bó G. Innovative strategies for superovulation in cattle. Anim Reprod 2018; 10:174-179. 55. Carvalho PD, Hackbart KS, Bender RW, Baez GM, Dresch AR, Guenther JN, Souza AH, Fricke PM. Use of a single injection of long-acting recombinant bovine FSH to superovulate Holstein heifers: a preliminary study. Theriogenology 2014; 82:481-489. 246 APPENDIX FSH FSH SORH O P E Type 1 O P E Type 2 Type 3 Type 4 P E P O E O P O E P O E Follicular Hyperstimulation Dysgenesis Figure 26. Model depicting the differences in amounts of cpFSH (FSH, green circles) in blood flowing from the ovarian arteries (horizontal red cylinders) into each follicle’s theca capillary network (vertical red cylinders) as the smaller antral follicles develop ( ) into Type 1, 2, 3 and 4 ovarian follicles during ovarian stimulation with the control (left panel) vs excessive (right panel) cpFSH doses. The arrows above the small ovarian reserve heifer (SORH) model depict timing (12-h intervals) of Folltropin-V (commercial FSH) or prostaglandin (PG) injections or ovariectomy. The differences in the theca capillary network for each follicle type is postulated to result from relatively minor differences in the age, stage of differentiation, and size of the small antral follicles at the beginning of the first follicular wave of the estrous cycle. Ovarian stimulation of the SORH model with 247 Figure 26 (cont’d) the control cpFSH doses results in development of Type 1 ovulatory-size follicles which are estrogen-active (EA) and have high intrafollicular concentrations of estradiol (E) but low progesterone (P) and oxytocin (O) and a compact layer of cumulus cells surrounding the oocyte. In contrast, the ovulatory-size follicles developing in response to excessive cpFSH doses exhibit a heterogenous phenotype defined as follows: Type 2 - similar to Type 1, Type 3 – EA but with an expanded layer of cumulus cells, Type 4 – estrogen- inactive with low E but high P and O and an expanded cumulus. The Type 2, 3 and 4 phenotyes produced by the excessive cpFSH treatment (no hCG ovulatory stimulus) exhibit Follicular Hyperstimulation Dysgenesis (FHD) characterized by a disruption of multiple cell-signaling pathways in granulosa and cumulus cells and oocytes critical for folliculogenesis, steroidogenesis, luteinization, cell survival, ovulation, and oocyte maturation and quality. The control cpFSH doses are hypothesized to be below the threshold of cpFSH concentration that cause FHD, resulting in the development of predominantly healthy Type 1 EA ovulatory-size follicles with a compact layer of cumulus cells surrounding the oocyte. 248 Figure 27. Impact of excessive cpFSH doses prior to an hCG ovulatory stimulus on expression of genes critical for cumulus cell function and oocyte maturation, resumption of meiosis and IVF. The arrows above the small ovarian reserve heifer (SORH) model depict timing (12-h intervals) of Folltropin-V (commercial FSH) or prostaglandin (PG) injections, ovariectomy or oocyte pick-up (OPU). Follicle types are explained in Figure 26’s legend. A total of ~15,000 genes was detected in cumulus and granulosa cells and oocytes during RNAseq analysis (left panel) and 3,288 cumulus genes were differentially expressed (DEGs) in cumulus cells when the Type 2, 3 and 4 ovulatory-size follicles developing in response to ovarian stimulation with the excessive cpFSH dose were compared with the Type 1 ovulatory-size follicles developing in response to the control cpFSH doses (right panel). However, only 17 (AREG, CLDN11, FGG, GPR50, IGFBP3, IGFBP5, LIF, MDFI, MEPE, MGAT5, NCS1, NGFR, PLAT, PTX3, RGS2, TGFα, VGF) of the 3,288 DEGs 249 Figure 27 (cont’d) (right panel) were identified independent of the follicle phenotype and with well-established roles in regulation of FSH action, cumulus function and oocyte maturation. Only compact cumulus-oocyte-complexes (COCs) and oocytes at the germinal vesicle stage (GV) of nuclear maturation were observed during ovarian stimulation with the control cpFSH doses (left panel). In contrast, prematurely expanded COCs and premature resumption of meiosis (MII) of oocytes were only observed during ovarian stimulation with the excessive cpFSH doses (right panel). In addition, rates of both in vitro maturation (IVM) and in vitro fertilization (IVF) of the prematurely expanded COCs (right panel) were greatly diminished compared with controls. Over-expression of the 17 cumulus genes during the excessive cpFSH-induced follicular hyperstimulation dysgenesis in the Type 2, 3 and 4 ovulatory-size follicles is hypothesized to have caused or contributed to the premature cumulus expansion and resumption of meiosis observed in most ovulatory-size follicles prior to an ovulatory hCG stimulus. 250 Figure 28. Impact of excessive cpFSH doses during ovarian stimulation on responsiveness of ovulatory-size follicles to an ovulatory hCG stimulus. The arrows above the small ovarian reserve heifer (SORH) model depict timing (12-h intervals) of Folltropin- V (commercial FSH), prostaglandin (PG), human chorionic gonadotropin (hCG) injections and oocyte pick-up (OPU). Follicle types are explained in Figure 26’s legend. Following ovarian stimulation with excessive cpFSH doses, the Type 2, 3 and/or 4 ovulatory-size follicles have a reduced responsiveness to hCG-induced ovulation and to hCG-induced oocyte maturation (MII) compared with the Type 1 ovulatory-size follicles developing in response to control cpFSH doses. 251 Figure 29. Usefulness of circulating concentration of anti-Müllerian hormone (AMH) to predict the responsiveness of the small ovarian reserve heifer (SORH) model to ovarian stimulation with different FSH doses. Prior to the first cpFSH injection, a single blood sample was taken and AMH concentration was determined. The SORH model was then subjected to ovarian stimulation with control (left panel) or excessive (right panel) cpFSH doses and number of ovulatory-size follicles 12 h after the last cpFSH injection and number that ovulated (functional) or did not ovulate (dysfunctional) in response to hCG was determined. Independent of cpFSH dose during ovarian stimulation, the AMH concentration prior to ovarian stimulation was highly positively correlated not only with number of ovulatory-size follicles prior to an hCG injection but with number of functional and dysfunctional ovulatory-size follicles after the hCG ovulatory dose. Although the Type 2, 3 and 4 ovulatory-size follicles developing in response to excessive cpFSH doses exhibit 252 Figure 29 (cont’d) different degrees of follicular hyperstimulation dysgenesis, which could negatively impact responsiveness to hCG, the phenotype (Type 2, 3, 4) of the dysfunctional ovulatory-size follicles that do not ovulate in response to hCG has yet to be established. Measurement of AMH prior to ovarian stimulation is predictive of the number of functional and dysfunction ovulatory-size follicles, thus unlikely useful to improve IVF or embryo transfer outcomes in the SORH model. 253