Variation in the Ovarian Reserve Is Linked to Alterations in ...

BIOLOGY OF REPRODUCTION 80, 954–964 (2009) Published online before print 21 January 2009. DOI 10.1095/biolreprod.108.073791

Variation in the Ovarian Reserve Is Linked to Alterations in Intrafollicular Estradiol Production and Ovarian Biomarkers of Follicular Differentiation and Oocyte Quality in Cattle1 J.J. Ireland,6 A.E. Zielak-Steciwko,3,4 F. Jimenez-Krassel,6 J. Folger,6 A. Bettegowda,7 D. Scheetz,6 S. Walsh,4 F. Mossa,4 P.G. Knight,8 G.W. Smith,7 P. Lonergan,4,5 and A.C.O. Evans2,4,5 School of Agriculture Food Science and Veterinary Medicine4 and Conway Institute,5 College of Life Sciences, University College Dublin, Dublin, Ireland Molecular Reproductive Endocrinology Laboratory,6 Laboratory of Mammalian Reproductive Biology and Genomics,7 Department of Animal Science, Michigan State University, East Lansing, Michigan School of Biological Sciences,8 University of Reading, Reading, United Kingdom (CYP19A1) production by granulosal cells, function and survival of thecal cells (TBC1D1), responsiveness of cumulus cells to estradiol (ESR1, ESR2), and cumulus cell determinants of oocyte quality (CTSB).

ABSTRACT The mechanisms whereby the high variation in numbers of morphologically healthy oocytes and follicles in ovaries (ovarian reserve) may have an impact onovarian function, oocyte quality, and fertility are poorly understood. The objective was to determine whether previously validated biomarkers for follicular differentiation and function, as well as oocyte quality differed between cattle with low versus a high antral follicle count (AFC). Ovaries were removed (n ¼ 5 per group) near the beginning of the nonovulatory follicular wave, before follicles could be identified via ultrasonography as being dominant, from heifers with high versus a low AFC. The F1, F2, and F3 follicles were dissected and diameters determined. Follicular fluid and thecal, granulosal, and cumulus cells and the oocyte were isolated and subjected to biomarker analyses. Although the size and numerous biomarkers of differentiation, such as mRNAs for the gonadotropin receptors, were similar, intrafollicular concentrations of estradiol and the abundance of mRNAs for CYP19A1 in granulosal cells and ESR1, ESR2, and CTSB in cumulus cells were greater, whereas mRNAs for AMH in granulosal cells and TBC1D1 in thecal cells were lower for animals with low versus a high AFC during follicle waves. Hence, variation in the ovarian reserve may have an impact on follicular function and oocyte quality via alterations in intrafollicular estradiol production and expression of key genes involved in follicle-stimulating hormone action (AMH) and estradiol

estradiol, follicle, follicular development, oocyte development, ovary

INTRODUCTION The nonreplenishable pool of healthy follicles and oocytes in ovaries (ovarian reserve) of women [1–3] and cattle [4, 5] is highly variable at birth, and the number of follicles and oocytes and oocyte quality decrease rapidly during aging. Consequently, adults have a highly variable ovarian reserve throughout their reproductive lifespan [1–5]. Many studies report that relatively high numbers of follicles are positively associated with enhanced ovarian function and a variety of indices of fertility in women [6–12] and cattle [4, 13–17]. These observations imply not only that individuals with low follicle numbers have diminished ovarian function, poor oocyte quality, and suboptimal fertility, but that the variation in nonovulatory follicle numbers has an important role in the reproductive success of single-ovulating species. However, the mechanisms whereby the inherently high variation in follicle numbers [1, 4] may have an impact on ovarian function, oocyte quality, and fertility in humans and cattle are poorly understood, especially because appropriate animal models to examine these potentially important linkages have not been developed. We have taken advantage of the bovine model to examine the physiological significance of the variation in follicle numbers, because cattle have relatively long reproductive cycles with two or three well-characterized follicle-stimulating hormone (FSH)-induced waves of growth and atresia of antral follicles [18, 19] similar to women [20, 21]. Most importantly, however, serial ovarian ultrasonography shows that the peak number of antral follicles growing during each different follicular wave is very highly repeatable within individuals (0.85 to 0.95; 1 ¼ perfect), despite the 7-fold variation in follicle numbers among young adult cattle [22, 23]. These observations clearly illustrate that cattle can be phenotyped reliably based on antral follicle count (AFC) during follicular waves [22, 23], and that cattle are an appropriate model to elucidate the physiological significance of the high variation in follicle numbers in ovaries. Recent studies using the bovine model also demonstrate that circulating FSH concentrations are inversely [17, 22–24] associated with number of morphologically healthy follicles

1

Supported by National Research Initiative Competitive grants 200401697 and 2007-01289 from the U.S. Department of Agriculture Cooperative State Research, Education, and Extension Service and by the Michigan State University Agriculture Experiment Station to J.J.I., and by Science Foundation Ireland (02/IN1/B78) and the Department of Agriculture Fisheries and Food under the National Development Plan through the Research Stimulus Fund (RSF 06 328) to A.C.O.E. and P.L. The opinions, findings and conclusions or recommendations expressed in this material are those of the authors and do not necessarily reflect the views of the funding agencies. 2 Correspondence: A.C.O. Evans, UCD Veterinary Sciences Centre, Belfield, Dublin 4, Ireland. FAX: 353 1 716 6253; e-mail: [email protected] 3 Current address: Institute of Animal Breeding, Faculty of Biology and Animal Science, Wroclaw University of Environmental and Life Sciences, 50-375 Wroclaw, Poland. Received: 29 September 2008. First decision: 21 October 2008. Accepted: 6 January 2009. Ó 2009 by the Society for the Study of Reproduction, Inc. eISSN: 1259-7268 http://www.biolreprod.org ISSN: 0006-3363

954

OVARIAN RESERVE, FOLLICLE FUNCTION, OOCYTE QUALITY

955

TABLE 1. List of ovarian biomarkers (hormones, growth factors, mRNAs) measured in each follicle cell type or follicular fluid for individual follicles that were used to evaluate whether follicular differentiation, function and atresia, and oocyte function and competence differed between cattle with low versus a high AFC during follicular waves. Parameter Follicles Differentiation Granulosal cells Thecal cells Cumulus cells Function Follicular fluid Atresia Granulosal cells Thecal cells Follicular fluid Oocytes Differentiation Oocytes Quality Oocytes Cumulus cells

Biomarker

Messenger RNAs for FSHR [50–52], LHCGR [50–52], ESR1 [53, 54], ESR2 [47, 53, 54], CYP19A1 [52, 55], AMH [56], BCAR1 [52], and MRPL41 [57] Messenger RNAs for LHCGR [50, 51], ESR1 [54, 58], ESR2 [54, 59], TBC1D1 [57], PRNP [60], CAMK1 [52], and SRF [61] Messenger RNAs for FSHR [62], ESR1 [63], ESR2 [63], and AMH [56] Estradiol [50, 51], progesterone [50, 51], FST [64], and AMH [65] Messenger RNAs for TGFBR3 [47, 52], FIBP [52], and CARTPT [66] Messenger RNAs for FOXO3 [61], TGFBR3 [47] INHA [43, 67, 68], ACT [43] Messenger RNAs for ESR2 [63], JY-1 [69], INHBA [70], and INHBB [70] Messenger RNA for FST [71] Messenger RNAs for CTSB, CTSS, CTSK, and CTSZ [72]

and oocytes during follicular waves in young adult cattle. In contrast, circulating anti-Mu¨llerian hormone (AMH) concentrations are positively associated with follicle numbers in cattle [25]. Moreover, healthy young adult cattle with a consistently low versus a high AFC during follicular waves also have a reduced number of morphologically healthy oocytes and follicles in ovaries [25], reduced responsiveness to superovulation and an accompanying reduced yield of high-quality transferable embryos, and diminished in vitro blastocyst development [22, 23]. These same phenotypic differences between young adult cattle with a low versus a high AFC are also reported for older versus younger women [3, 6, 8, 10, 12, 26–30] or cattle [4, 15–17, 31–34], young women with relatively high versus normal circulating FSH concentrations [35], and young women born small versus normal size for gestational age [36, 37]. However, it is unknown whether the inherently high variation in follicle numbers and the corresponding chronic alterations in circulating FSH and AMH concentrations during follicular waves of young adult cattle are also associated with alterations in ovarian function. It is well established that FSH has a fundamentally important positive role in folliculogenesis. For example, FSH receptors are exclusive to granulosal cells of follicles [38, 39], and FSH is required for growth and differentiation [40] of antral follicles during follicular waves [41–43]. The AMH receptors are also located in granulosal cells [44], but AMH is reported to inhibit FSH action in rodent models [45]. Taken together, the inherently high variation in follicle numbers during follicular waves of bovine estrous cycles and the corresponding alterations in circulating FSH and AMH concentrations are hypothesized to alter differentiation of follicles growing during follicular waves, which in turn has an impact on follicular function and oocyte quality. The present study, therefore, was designed to begin to test this hypothesis by determining whether ovarian biomarkers in follicular fluid (steroids and growth factors), somatic cells (mRNAs in thecal, granulosal, and cumulus cells), and oocytes (mRNAs), previously shown in the bovine to be potentially important for follicle development and oocyte quality, differed between healthy age-matched young adult cattle with a consistently low versus a high AFC during follicular waves.

MATERIALS AND METHODS Animals, Tissue Collection, and Follicle Classification Cross-bred beef heifers (Hereford 3 Angus 3 Charolais, n ¼ 32, 11–13 mo old) were synchronized with two injections of prostaglandin F2a (PG; Prosolvin; Intervet Ireland Ltd., Dublin, Ireland) given 11 days apart. Follicle development was monitored daily in each heifer by ovarian ultrasonography. Heifers were assigned to one of two groups based on the peak number of follicles (3 mm in diameter) in the first follicular wave of the estrous cycle: low (15 follicles, n ¼ 5 animals) or high (25 follicles, n ¼ 5 animals). Remaining animals in the intermediate category (n ¼ 22) were not used for subsequent studies. All experiments involving animals were licensed by the Department of Health and Children of Ireland in accordance with the cruelty to animals act (Ireland 1897) and the European Community Directive 86/609/EC. Animals in the low and high groups were resynchronized with PG as explained above and slaughtered at the emergence of the first follicle wave, which was 24–48 h after ovulation as first detected by ultrasonography (Days 2–3 of the estrous cycle) but before the wave’s dominant follicle could be identified based on its distinctly larger size. This early stage of the follicular wave was selected as the time for surgical removal of ovaries because the largest follicles are rapidly growing in response to the postovulatory FSH surge that initiates the first follicular wave [41]. Moreover, the largest growing follicles should have relatively similar diameters (;5–7 mm [46]), and thus be at potentially similar stages of growth and differentiation. Pairs of ovaries were collected, weighed, and measured. The three largest follicles per pair of ovaries were dissected, diameters measured, follicular fluid (FF) aspirated, and thecal and granulosal cells collected from each individual follicle as described previously [47]. Cumulus-oocyte complexes were also retrieved from the FF of each follicle, and cumulus cells and denuded oocytes were isolated as described previously [48] and stored at 808C in lysis buffer (Ambion, Huntingdon, U.K.) until RNA isolation. All dissections were carried out in ice-cold RNAlater (Ambion). Theca and granulosa cells were snap frozen in 1 ml of TRIZOL (Invitrogen, Carlsbad, CA) using liquid nitrogen and stored at 808C. Ovarian volume was calculated using the formula volume ¼ (4/3) p 3 (length 3 width 3 height) [49].

Selection of Ovarian Biomarkers to Quantify in Growing Follicles and Oocytes A variety of steroids and growth factors in FF and mRNAs in granulosal, thecal, and cumulus cells and oocytes previously shown primarily in the bovine to be potentially important for dominant follicle development and oocyte quality were selected as ovarian biomarkers for cell differentiation, follicular function, atresia, and differentiation and quality of oocytes (Table 1).

Analyses of FF Estradiol concentrations were measured in FF samples as described previously [73] using a Biodata Estradiol MAIA kit (Biochem Immunosystems,

956

IRELAND ET AL.

TABLE 2. Gene name, accession number, and forward (F) and reverse (R) primer sequences used for quantitative real-time PCR of bovine granulosal and thecal cells. Gene (symbol)

Primer sequencesa

GenBank accession no.

Luteinizing hormone/chorionic gonadotropin receptor (LHCGR) Follicle stimulating hormone receptor (FSHR)

NM174381 L22319

F: R: F: R: F: R: F:

5 0 -TGACCATGGCCCGTCTAAAA-3 0 5 0 -TACTACCCAAAGCAATTTATAGATTCAATG-3 0 5 0 -TGGTCCTGTTCTACCCCATCA-3 0 5 0 -GAAGAAATCCCTGCGGAAGTT-3 0 5 0 -CACCCATCTTTGCCAGGTAGTC-3 0 5 0 -ACCCACAGGAGGTAAGCCTATAAA-3 0 5 0 -GGCCCAGGGCACAAGAA-3 0 5 0 -GGGACCGCTGTGTCAGCTT-3 0 5 0 -GAAGGCAAAGTCCCCAAAGC-3 0 5 0 -CATCGTCTTCTTTGCCAACAATC-3 0 5 0 -CCACGAGCTGCTAGAGTTTGC-3 0 5 0 -CGTCTGGTACACGTCCTCCAT-3 0 5 0 -AGCTGGAAGAGCTTGCCTTCCT-3 0 5 0 -GCCAAGCAGCTCTTTCTGGAGA-3 0 5 0 -GCAGGGAGCGCGATATTG-3 0 5 0 -TGCGGCCCGTCAACA-3 0 5 0 -AGGGACGGCACCACTTATTTATT-3 0

Cytochrome P450, family 19, subfamily A, polypeptide 1 (CYP19A1)

NM174305

Transforming growth factor, beta receptor III, betaglycan or inhibin co-receptor (TGFBR3)

XM001924

Fibroblast growth factor (acidic) intracellular binding protein (FIBP)

AF010187

Breast cancer anti-estrogen resistance 1 (BCAR1)

NM014567

Mitochondrial ribosomal protein L41 (MRPL41)

BM251860

Forkhead box O3 (FOXO3)

AF032886

Homo sapiens serum response factor or c-fos serum response element-binding transcription factor (SRF)

XM011430

R: F: R: F: R: F: R: F: R: F:

Prion protein (p27–30) (present in Creutzfeldt-Jakob disease, Gerstmann-Strausler-Scheinker syndrome, and fatal familial insomnia (PRNP)

NC007311

R: 5 0 -TGCGGCCCGTCAACA-3 0 F: 5 0 -TCTCTGGTACTGGGTAATGCACAT-3 0

Calcium/calmodulin dependent protein kinase 1 (CAMK1)

XM516263

TBC1 (tre-2/USP6, BUB2, cdc 16) domain family, member 1 (TBC1D1)

BM252120

Cocaine and amphetamine regulated transcript (CARTPT) a

NM001007820

R: F: R: F:

5 0 -GGGAGAACTTCACCGAAACTGA-3 0 5 0 -CGGTGAGTGAGCAGATCAAGAA-3 0 5 0 -CCCCACGACTCGCTGTCT-3 0 5 0 -TCTACATTCCTGACCCTAAAATGCT-3 0

R: 5 0 -GTGAACTTTGTGGCTCTATTTTGGTT-3 0 F: 5 0 -CGCCATGGATGATGATATTGC-3 0 R: 5 0 -AAGCCGGCCTTGCACAT-3 0

All primers were used at 900 nM.

Bologna, Italy). All FF was diluted 1:100 in PBS prior to assay. The interassay coefficients of variation (CVs) were 6.7%, 6.6%, and 7.9% for low, medium, and high (17.9, 51.9, and 199.9 pg/ml) reference samples, respectively, and the intraassay CV values were 9.9%, 6.6%, and 11.7% for the same samples, respectively. Progesterone concentrations in FF were measured by fluoroimmunoassay (AutoDELFIA Progesterone; Wallac Oy, Turku, Finland). Prior to assay, FF was diluted 1:100 in PBS. The interassay CV values were 10.6%, 5.2%, and 3.2% for low, medium, and high (0.25, 2.37, and 6.09 ng/ml) reference samples, respectively, and the intraassay CV values were 2.1%, 6.6%, and 11.7% for low, medium, and high reference samples, respectively. Concentrations of inhibin-A in FF were determined using a two-site ELISA specifically developed for bovine/ovine samples [74]. Purified bovine 32-kDa inhibin [75] was used as standard, and the detection limit of the assay was 20 pg/ml. Within- and between-plate CVs were both ,12%. Concentrations of total ACT in FF were measured using the two-site ELISA [76]. Recombinant human ACT was used as standard, and the assay detection limit was 100 pg/ml. Within- and between-plate CVs were both ,10%. Concentrations of total follistatin in FF were measured using the two-site ELISA [77]. The assay detection limit was equivalent to 100 pg/ml recombinant human FST, and within- and between-plate CVs were both ,16%. The commercially available human MIS/AMH ELISA kit (DSL-10–14400; Beckman Coulter Inc., Fullerton, CA) previously validated in the bovine [25] was used to measure AMH concentrations in FF (diluted 1:200) in cattle per the kit instructions. The two-site AMH assay does not cross-react with other members of the transforming growth factor b (TGFb) superfamily, including TGFb1, BMP4, or ACT [78]. Intraassay CV for an overall average AMH concentration of 1.62 6 0.07 ng/ml was 10% (n ¼ 1 assay).

random hexamers. The quantities of both RNA and cDNA were measured using a NanoDrop ND-100 (NanoDrop Technologies, Wilmington, DE) spectrophotometer. Quantitative real-time PCR was carried out using an Mx3000P Q-RT-PCR machine (Stratagene, La Jolla, CA) in 96-well plates to determine differences in mRNA abundance for genes of interest in follicular cell types from heifers with low versus high AFCs. Primers for real-time PCR for genes of interest were designed using Primer Express, Version 2.0 (Applied Biosystems, Foster City, CA) and are shown in Table 2. In brief, each reaction (25 ll) contained 20 ng of cDNA, 900 nM each primer, 12.5 ll of Brilliant SYBR Green QPCR Master Mix (Stratagene), and remaining volumes of nuclease-free water, and each sample was assayed in duplicate. Two nontemplate control samples were included on each plate for each primer set. The thermal cycling program used to amplify target cDNAs sequences involved 508C for 2 min (one cycle), 958C for 10 min (one cycle), 958C for 15 sec, and 608C for 1 min (40 cycles). Specificity of amplification with each primer set in each assay was confirmed by melting-curve analysis. The mRNA abundance for each target gene was normalized against the levels of the constitutive housekeeping gene ACTB. The mean target gene threshold cycle (Ct) and mean exogenous control (ACTB) Ct for each sample were calculated from duplicate wells. The mean Ct of the control was then subtracted from the target gene Ct of samples to give the DCt. In each experiment, the mean of one experimental group served as a control (calibrator). Subsequently, the DCt of sample was then subtracted from the calibrator DCt to give the DDCt. The relative amounts of target gene expression (Table 2) for each sample were then calculated using the formula 2(DDCt) [79].

RNA Isolation and Quantitative Real-Time PCR for Granulosal and Thecal Cells of Individual Follicles

Total RNA was extracted from single oocytes and matching cumulus cells isolated from the three largest follicles collected from beef heifers with low or high AFCs using the RNAqueous micro kit (Ambion, Austin, TX) according to the manufacturer’s instructions. Before RNA extraction, each sample was spiked with 250 fg of green fluorescence protein (GFP) synthetic RNA as an exogenous control for RNA recovery and efficiency of cDNA synthesis [48]. The average GFP recovery from oocytes for all animals was 64.9%, and there was no difference in GFP recovery between animals with a high or a low AFC

Total RNA was extracted from thecal and granulosal cell samples of each follicle using the standard TRIZOL protocol followed by DNase treatment steps to minimize genomic DNA contamination as described previously [47]. For cDNA synthesis, 2 lg of total RNA was reverse transcribed using the SuperScript II RNase H-Reverse Transcriptase (Invitrogen) and primed by

RNA Isolation and Quantitative Real-Time PCR for Oocytes and Cumulus Cells of Individual Follicles

957


TABLE 3. Gene name, accession number, and forward (F) and reverse (R) primer sequences used for quantitative real-time PCR of cumulus cells and oocytes. Gene (symbol)

GenBank accession no.

Follistatin (FST)

BF774514

Inhibin bA subunit (INHBA)

AW658434

Inhibin bB subunit (INHBB)

AW669304

Estrogen receptor 1 or alpha (ESR1)

U64962

Estrogen receptor 2 or beta (ESR2)

Y18017

FSH Receptor (FSHR)

L22319

JY-1 Anti-Mu¨llerian hormone (AMH)

M13151 BF868324

Cathepsin S (CTSS)

BE482678

Cathepsin K (CTSK)

BF230198

Cathepsin Z (CTSZ)

BE752253

18S rRNA

BC102293

Green fluorescent protein (GFP) a

F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R: F: R:

EF642496

Cathepsin B (CTSB)

Primer sequencesa

—

5 0 -CAGAGCTGCAAGTCCAGTACCA-3 0 5 0 -CATGTAGAGCTGCCTGGAACAGA-3 0 5 0 -GAGCAGTCGCACAGACCTTTC-3 0 5 0 -CCGGTGAGGATGGTCTTCAG-3 0 5 0 -GCATCTCCAGAATGCCTTCAC-3 0 5 0 -GTGGAAATGACTCTTATGCAATGG-3 0 5 0 -GGCCAACTCCTCCTCATCCT-3 0 5 0 -CTTGCACTTCATGCTGTATAGATGCT-3 0 5 0 -AGGTCAGAGGCTGAAGCATGA-3 0 5 0 -CACATAGCAGAAAGCCAAGGAA-3 0 5 0 -TGGTCCTGTTCTACCCATCA-3 0 5 0 -GAAGAAATCCCTGCGGAAGTT-3 0 5 0 -TTGGAACTTCCATGGACGACC-3 0 5 0 -ATTTGCTGGTGATCCCAAGAG-3 0 5 0 -CAGGGAAGAAGTCTTCAGCA-3 0 5 0 -AAGGTGGTCAAGTCACTCAG-3 0 5 0 -CGATGCCCGGGAACAGT-3 0 5 0 -GAGCCAGGATCCCTGATC-3 0 5 0 -TCGTGGTTGGCTATGGTAACC-3 0 5 0 -TGCAGGCCCCAGCTGTT-3 0 5 0 -CATATGAACTGGCCATGAACCA-3 0 5 0 -TGAGTCCAGTCATCTTCTGAACCA-3 0 5 0 -GGGAGAAGATGATGGCAGAAAT-3 0 5 0 -TCTTTTCGGTTGCCATTATGC-3 0 5 0 -GTGGTGTTGAGGAAAGCAGACA-3 0 5 0 -TGATCACACGTTCCACCTCATC-3 0 5 0 -CAACAGCCACAACGTCTATATCATG-3 0 5 0 -ATGTTGTGGCGGATCTTGAAG-3 0

All primers were used at 300 nM.

(67.3% 6 4% versus 62.4% 6 3%; P ¼ 0.30). Residual genomic DNA was removed by DNAse I digestion (Ambion). RNA was eluted twice from the silica-based microfilter cartridge using 10-ll volumes of prewarmed (758C) elution solution according to the manufacturer’s instructions. The RNA from each oocyte or cumulus cell sample was divided into two 10-ll aliquots. Both aliquots of extracted total RNA were used for cDNA synthesis. Total RNA (10 ll in duplicate) from each sample for real-time PCR analyses was used for reverse transcription using oligo dT(15) primers as described previously [48]. After termination of cDNA synthesis, each RT reaction was then diluted with nuclease-free water (Ambion) to a final volume of 20 ll. The quantification of all gene transcripts was done by real-time quantitative RT-PCR using SYBR Green PCR Master Mix (Applied Biosystems). Primers were designed using the Prime Express program (Applied Biosystems) and derived from bovine gene sequences found in GenBank (Table 3), and the amplicon size for each of the genes studied ranged from 80 to 270 bp. Each reaction mixture consisted of 2 ll of cDNA, 300 nM each of forward and reverse primers, 7.5 ll of nuclease-free water, and 12.5 ll of SYBR Green PCR Master Mix in a total reaction volume of 25 ll (96-well plates). Reactions were performed in duplicate for each sample in an ABI Prism 7500 Sequence Detection System (Applied Biosystems). Two nontemplate control samples were included on each plate for each primer set. Specificity of amplification with each primer set in each assay was confirmed by melting-curve analysis. The thermal cycler program consisted of 40 cycles of 958C for 15 sec and 608C for 1 min. For real-time PCR experiments, amounts of mRNAs of interest were normalized relative to an exogenous control (GFP) to control for differences in RNA recovery and efficiency of RT [48], and also were separately normalized relative to abundance of an endogenous control (18S rRNA) to account for RNA concentrations between samples. Data for INHBA, INHBB, and FSHR (Table 3) were expressed relative to 18S rRNA and quantified using the relative quantification method [79] as described above. Copies of ESR1, ESR2, FST, AMH, CTSB, CTSS, CTSK, CTSZ, JY-1, 18S rRNA, and GFP (Table 3) were quantified using the standard curve method for absolute quantification [80]. Data for these genes are expressed as copies of mRNA per copy of 18S rRNA or GFP mRNA.

Culture of Granulosal Cells Isolated from Animals with a Low Versus a High AFC Bovine granulosal cells were isolated from small antral follicles (2–5 mm in diameter) present on the surface of ovaries collected at a local abattoir. Ovaries

were selected based on the number of antral follicles per pair of ovaries as follows: ovaries in the low group had 15 follicles that were 3 mm in diameter, whereas ovaries in the high group had .25 follicles. For each group, only five follicles were isolated per ovary, and the granulosal cells from five ovaries were pooled. Four different pools of granulosal cells were then cultured in serum-free media as described previously [81]. In brief, 100 000 live cells were plated in 96-well Falcon Primaria plates for 2 days without media change. Concentrations of estradiol in spent media were measured at the end of culture using a commercially available kit (Diagnostic Products Corp., Los Angeles, CA). Estradiol concentrations were then normalized to the number of live granulosal cells at the end of the culture. Number of live cells was determined following trituration to remove cells from wells, staining of cells with Trypan blue for 1 min, and use of a hemacytometer to count the number of live versus dead cells.

Statistical Analysis ANOVA was used to determine whether statistically significant differences (P , 0.05) existed for concentrations of hormones and growth factors in FF or media and abundance of mRNAs in granulosal, thecal, and cumulus cells and oocytes in F1, F2, and F3 follicles for animals in the low versus high groups [82]. When main effects were significant, individual means were compared using protected least significant difference [82].

RESULTS Similar to results previously reported for this animal model [22, 23, 25], the overall average and peak of number follicles during the first follicle wave, number of follicles counted after surgical removal of ovaries, and paired ovarian weight and volume were much greater (P , 0.01) in animals from the high versus the low group (Table 4). Diameters of F1, F2, and F3 follicles differed (P , 0.0001; Fig. 1), but concentrations of estradiol (Fig. 1) and other hormones and growth factors in FF and the abundance of mRNAs for the ovarian biomarkers measured in follicles and oocytes (Table 1) were similar (P . 0.10; data not shown) among the different follicle size classes. Thus, data for the

958

IRELAND ET AL.

TABLE 4. Number of follicles (3 mm diameter) and the volume and weight of both ovaries at emergence of the first nonovulatory follicle wave on Days 2 to 3 of bovine estrous cycle for cattle with consistently low versus a high AFC during follicular waves. AFC Parameter No. of Follicles Overalla Peakb Paired ovarian Volume (cm3) Weight (g)

Low

High

P value

10.9 6 1.1 13.2 6 0.8

27.3 6 1.2 33.6 6 3.3

,0.001 ,0.001

7.3 6 0.9 9.6 6 0.7

14.4 6 1.9 15.5 6 2.1

0.009 0.014

a Mean number of follicles per day counted (by ovarian ultrasonography) during the first seven days of the first follicular wave of the estrous cycle before ovariectomy. b Mean peak number of follicles counted (by ovarian ultrasonography) during the first follicular wave of the estrous cycle.

largest three follicles were pooled for each animal in the low or high groups for further statistical analysis. The ovarian biomarkers are listed in Table 1. Each biomarker was chosen because previous studies (see references by each marker in Table 1), primarily in the bovine, demonstrated that the specific hormone, growth factor, or mRNA of interest was potentially important for follicular development. Results showed that although diameters were similar between low- and high-AFC groups (Table 5), several potentially key ovarian biomarkers in FF and the different cell types differed (P , 0.05) for the three largest follicles combined between the two groups of animals as follows: TABLE 5. Diameter, concentrations of hormones and growth factors in follicular fluid, and abundance of mRNAs in granulosal and thecal cells of individual follicles at emergence of the first nonovulatory wave on Days 2 to 3 of the estrous cycle that did not differ statistically (P 0.05) between cattle with consistently low versus a high AFC during follicular waves. AFC Follicle analysis Diameter (mm) FF (ng/ml) Progesterone INHA ACT FST AMH Granulosal cell mRNAa LHCGR FSHR TGFBR3 FIBP MRPL41 ESR1 ESR2 CARTPT BCAR1 Thecal cell mRNAa LHCGR TGFBR3 FOXO3 PRNP CAMK1 ESR1 ESR2 SRF a

Low

High

P value

6.1 6 0.3

5.4 6 0.3

0.26

34 21 1.9 3.7 286

6 6 6 6 6

7 2 0.2 0.4 21

42 19 1.4 6.6 438

6 6 6 6 6

15 2 0.2 1 92

0.48 0.72 0.41 0.21 0.17

6 95 18 12 3 5 22 18 2

6 6 6 6 6 6 6 6 6

3 27 4 0.4 0.2 7 3 7 0.4

1 82 22 3 4 5 28 71 3

6 6 6 6 6 6 6 6 6

0.3 16 5 0.3 1 0.8 3 40 0.3

0.73 0.85 0.75 0.86 0.72 0.92 0.11 0.25 0.09

12.3 61.1 19.4 15.8 3.5 1.5 4.4 8

6 6 6 6 6 6 6 6

3 23 4 3 1 0.3 0.9 1

15 50.6 20.5 16.6 5.3 1.7 5.4 6

6 6 6 6 6 6 6 6

5 14 5 4 2 0.4 0.6 1

0.24 0.99 0.84 0.90 0.31 0.61 0.18 0.10

Data expressed relative to ACTB mRNA 31000.

FIG. 1. Diameters and estradiol concentrations (mean 6 SEM, n ¼ 5 animals per group with three follicle classes) in FF for F1, F2, and F3 follicles at emergence of the first follicle wave on Days 2–3 of the estrous cycle of animals with low versus high AFCs during follicular waves. ANOVA indicated that diameters differed (P , 0.0001) between F1, F2, and F3 follicles but were similar (P . 0.26) for animals with low versus high follicle numbers. In contrast, estradiol concentrations were similar (P . 0.32) between F1, F2, and F3 follicles but higher (P , 0.02) for animals with low versus high follicle numbers.

1. Follicular fluid: Concentrations of a variety of wellestablished biomarkers for follicular function in the bovine were measured in FF (Table 1). Although intrafollicular concentrations of progesterone, ACT, INHA, FST, and AMH were similar (Table 5), estradiol concentrations in FF were about 2-fold higher (P , 0.02) in animals with a low versus a high AFC (Figs. 1 and 2). In addition, basal capacity to produce estradiol was ;3-fold greater (P , 0.01) for granulosal cells isolated from small antral follicles of animals with a low versus a high AFC (Fig. 3). 2. Thecal cells: Expression of genes for LHCGR, ESR1, ESR2, TBC1D1, PRNP, CAMK1, and SRF in thecal cells has previously been shown to be altered during follicular development in cattle (Table 1). Results of the present study showed that abundance of nearly all of these mRNAs was similar, except for TBC1D1, which was ;50% lower (P , 0.02; Fig. 4) in thecal cells for animals with a low versus a high AFC. 3. Granulosal cells: Expression of genes for FSHR, LHCGR, ESR1, ESR2, MRPL41, CYP19A1, and AMH in granulosal cells


FIG. 2. Estradiol concentrations in FF and abundance of CYP19A1 (aromatase) mRNA in granulosal cells for F1–F3 follicles combined for cattle with a consistently low versus a high AFC during follicular waves. Results for aromatase mRNA are expressed as the abundance of CYP19A1 mRNA per ACTB mRNA. Each bar represents the mean 6 SEM for a total of 15 follicles (three follicles in each of five animals). Asterisks indicate statistical significance between means, ***P , 0.01.

have previously been shown to be altered in bovine or rodent granulosal cells during follicular development (Table 1). Although abundance of mRNAs for FSHR, LHCGR, ESR1, ESR2, and MRPL41 was similar, abundance of CYP19A1 mRNA was 4-fold greater (P , 0.01; Fig. 2), but AMH mRNA was ;20-fold lower (P , 0.01; Fig. 4) for animals with a low versus a high AFC. There was also a tendency for BCAR1 mRNA to be lower (P , 0.09) for animals with a low versus a high AFC (Table 5). 4. Cumulus cells: Bovine cumulus cells contain receptors for FSH and estradiol, and they produce AMH (Table 1). However, despite marked differences in FSH and AMH secretion between cattle with a low versus a high AFC [22, 23, 25], abundance of mRNAs in cumulus cells for FSHR and AMH (Table 1) were similar (Table 6), but mRNAs for ESR1 and ESR2 were about 2-fold greater (P , 0.01; Fig. 5) for animals with low versus high follicle numbers. 5. Follicular atresia: Alterations in abundance of mRNAs for TGFBR3, FIBP, and CARTPT in granulosal cells, FOXO3 and TGFBR3 in thecal cells, and INHA and ACT concentrations in FF have been established to be reliable indices of atresia in bovine follicles (Table 1). However, no differences in these biomarkers were detected between animal groups (Table 5).

959

FIG. 3. Basal estradiol production by bovine granulosal cells. Bovine granulosal cells isolated from small antral follicles of animals with a low versus a high AFC were cultured serum free for 2 days, and estradiol concentrations were measured at the end of culture. Bars represent the mean (6SEM) for four different pools of granulosal cells. Asterisks indicate statistical significance between means, **P , 0.01.

6. Oocytes: Expression of genes for JY-1, INHBA, and INHBB are significantly altered during maturation of bovine oocytes (Table 1). Intrafollicular estradiol concentrations differed markedly between animal groups in the present study, and high estradiol inhibits maturation of bovine oocytes [63]. Consequently, it was determined whether alterations in intrafollicular estradiol result in alterations in abundance of mRNAs for ESR2, JY-1, INHBA, and INHBB (Table 1) in individual oocytes isolated from the two different animal groups. Although JY-1, INHBA, and INHBB mRNAs were similar, there was a tendency (P , 0.09) for ESR2 mRNA to be more abundant in oocytes from animals with a low versus a high AFC (Table 6). In addition, mRNA abundance of FST in oocytes and CTSB, CTSS, CTSK, and CTSZ in cumulus cells has been shown to be a reliable marker for oocyte quality in the bovine (Table 1). Although alterations in mRNA abundance of FST in oocytes and CTSK and CTSZ in cumulus cells did not differ (Table 6), abundance of CTSB mRNA was 2-fold higher (P , 0.01; Fig. 6), whereas there was a tendency for CTSS to also be higher (P , 0.09; Table 6) for animals with a low versus a high AFC. DISCUSSION The most significant results of the present study demonstrated for the first time in a single-ovulating species that the inherently high variation in follicle numbers during follicular

960

IRELAND ET AL. TABLE 6. Abundance of mRNAs in cumulus cells and oocytes of individual follicles at emergence of the first nonovulatory wave on Days 2 to 3 of estrous cycle that did not differ statistically (P , 0.05) for cattle with consistently low versus a high AFC during follicular waves. AFC Follicle analysis Cumulus cell mRNA CTSSa CTSZa CTSKa FSHRb AMHb Oocyte mRNA ESR2c INHBAb INHBBb JY-1c FSTc a b c

FIG. 4. Abundance of AMH mRNA for granulosal cells and TBC1D1 mRNA for thecal cells for cattle with a consistently low versus a high AFC during follicular waves. Results for mRNA abundance are expressed relative to ACTB. Each bar represents the mean 6 SEM for a total of 15 follicles (three follicles in each of five animals). Asterisks indicate statistical significance between means, ***P , 0.01, **P , 0.02.

waves is also associated with significant alterations in intrafollicular estradiol production, which is the hallmark for follicular function, and expression of key genes important for differentiation, function, and survival of thecal (TBC1D1), granulosal (CYP19A1, AMH), and cumulus cells (ESR1, ESR2, CTSB) in the largest three follicles growing during follicular waves. In the present study, it is important to emphasize that there were not widespread differences in sizes or amounts of many well-characterized bovine ovarian biomarkers, including those for atresia, in the three largest follicles obtained from each pair of ovaries. This important observation clearly demonstrates that follicles were dissected from ovaries of both groups of cattle at very similar stages of development during the first follicular wave. Thus, the differences in estradiol production and gene expression observed in the present study were more likely attributable to the inherently high variation in follicle numbers rather than differences in the timing of surgical removal of follicles from ovaries during a follicular wave between the cattle with low or high follicle numbers. Taken together, these results provided important new evidence that potential physiologically significant differences exist in gene expression related to steroidogenic capacity, hormonal responsiveness, and differentiation of thecal, granulosal, and cumulus cells between cattle with a low versus a high AFC.

Low

High

P value

8.7 76.3 90.5 1013.5 3.1

6 6 6 6 6

7 20 40 101 0.7

1.4 57.9 58.4 946.6 2.1

6 6 6 6 6

0.4 6 11 134 0.6

0.09 0.61 0.86 0.31 0.15

1.7 1396.9 1317 15.6 249.3

6 6 6 6 6

0.4 445 445 9 52

1 1316.6 984 37.3 194.4

6 6 6 6 6

0.2 566 304 15 61

0.09 0.68 0.32 0.19 0.20

Data expressed as copy of mRNA per 1000 copies of 18S rRNA. Data expressed relative to 18S rRNA 31000. Data expressed as copy of mRNA per 1000 copies of GFP mRNA.

Direct determinations of whether these differences, however, also have a negative impact on oocyte competence and fertility in young adult cattle with inherently low versus high numbers of healthy follicles and oocytes will be important to determine, but this was beyond the scope of the present study. Nevertheless, the results of the present study, coupled with our previous results [22, 23, 25], provide important new insights and directions into potential mechanisms whereby the variation in follicle numbers and, correspondingly, in circulating FSH and AMH concentrations, may have an impact on ovarian function in single-ovulating species, like cattle and perhaps humans. Circulating FSH concentrations are well established to be higher in cattle with relatively low numbers of follicles growing during follicular waves [17, 22–24]. Moreover, FSH is a positive regulator of aromatase [83], estradiol receptors [84], and estradiol production [83] by granulosal cells. Consequently, the enhanced intrafollicular and granulosal cell estradiol production, coupled with much higher expression of the gene transcripts for aromatase (CYP19A1) in granulosal cells and the estrogen receptors (ESR1 and ESR2) in cumulus cells (and a tendency for ESR2 in oocytes), for animals with low follicle numbers in the present study are most likely explained by chronically higher circulating FSH concentrations in cattle with low versus high follicle numbers [17, 22–24]. The reason estrogen receptors were selectively increased in these cell types but not mural granulosal or thecal cells, despite the high intrafollicular estradiol concentrations, is unknown. Nevertheless, previous reports support differential regulation of the same genes in the different cell types of follicles [85, 86]. Surprisingly, however, high physiological concentrations of estradiol block maturation of bovine oocytes in vitro and cause chromosomal aberrations [63]. Moreover, relatively high FSH concentrations in the presence of insulin in rodents [87], heightened secretion of FSH in transgenic rodents [88], and superovulation of cattle [89, 90], which would be expected to greatly enhance ovarian estradiol production, also diminish developmental competence of oocytes and fertility. These intriguing findings imply that the much higher (2-fold) intrafollicular estradiol concentrations, coupled with the potentially enhanced responsiveness of cumulus cells and oocytes to estradiol observed in the present study, may have detrimental effects on oocyte maturation and developmental competence in cattle with low follicle numbers.


961

FIG. 6. Abundance of CTSB (cathepsin B) mRNA in cumulus cells of F1– F3 follicles combined for cattle with a consistently low versus a high AFC during follicular waves. Results are expressed as copies of CTSB mRNA per copy of 18S rRNA. Each bar represents the mean 6 SEM for a total of 15 follicles (three follicles in each of five animals). Asterisks indicate statistical significance between means, ***P , 0.01.

FIG. 5. Abundance of mRNA for ESR1 (estrogen receptor a) and ESR2 (estrogen receptor b) for cumulus cells of F1–F3 follicles combined for cattle with a consistently low versus a high AFC during follicular waves. Results are depicted as copies of ESR1 or ESR2 mRNA per copy of 18S rRNA. Each bar represents the mean 6 SEM for a total of 15 follicles (three follicles in each of five animals). Asterisks indicate statistical significance between means, ***P , 0.01, **P , 0.05.

The novel findings in the present study that animals with relatively low follicle numbers and high intrafollicular estradiol concentrations also have higher expression of the genes for CTSB and CTSS (tendency) in cumulus cells, but much lower expression of AMH and TBC1D1 in granulosal and thecal cells, further support the possibility that follicular function and oocyte maturation and quality are diminished in healthy young adult cattle with chronically low versus high follicle numbers. For example, cathepsins are lysosomal cysteine proteinases involved in a variety of cell processes, pathologies, and apoptosis [91] and, most importantly, recent studies from our laboratory show that gene transcripts for several cathepsins are highly expressed in cumulus cells surrounding oocytes from prepubertal (a model of poor oocyte quality) versus adult cattle [72]. Moreover, these same mRNA transcripts are more abundant in the cumulus cells for adult metaphase II bovine oocytes that develop to the blastocyst stage at a low percentage following parthenogenetic activation, and inhibition of cathepsin activity during meiotic maturation results in enhanced rates of blastocyst development following in vitro fertilization [72]. Thus, relatively high expression of cathepsins in cumulus cells, as observed for animals with low follicle numbers in the present study, implies that oocyte quality may be compro-

mised. Although it is unknown whether the much higher intrafollicular estradiol observed for animals with low versus high follicle numbers in the present study also explains the enhanced expression of the cathepsin transcripts in cumulus cells, previous studies in rodents support a potentially positive role for estradiol in cathepsin regulation [92, 93]. AMH is a member of the TGFb superfamily of growth factors that is produced in females only by healthy granulosal and cumulus cells of growing follicles [94, 95], and AMH concentrations are associated positively not only with follicle numbers but also oocyte quality and fertility in humans [65, 95]. Although circulating FSH and AMH concentrations were not measured in the present study, young adult cattle with relatively low follicle numbers also have correspondingly 10% to 50% higher circulating FSH [17, 22–24] but 80% lower circulating AMH concentrations compared with age-matched animals with high follicle numbers during follicle waves [25]. These observations, coupled with the much lower expression of AMH transcripts in granulosal cells of cattle with low follicle numbers observed in the present study, further support the possibility that oocyte quality and perhaps fertility are lower for cattle with low follicle numbers. However, although FSH action is enhanced in AMH knockout mice [95], it is unknown whether the significant inverse relationship between circulating FSH and AMH concentrations for cattle with chronically low follicle numbers during follicle waves [17, 22–25] causes or contributes to the detrimental effects of high physiological concentrations of FSH and estradiol on oocyte maturation and competence reported by others [63, 87–90]. TBC1 (tre-2/USP6, BUB2, cdc 16) domain family, member 1 (TBC1D1) is the founding member of a protein family containing TBC domains that regulate Rab GTPases. Rab

962

IRELAND ET AL.

GTPases have a significant regulatory role in vesicle trafficking [96] and cell signaling, differentiation, and growth [97, 98], and Rab GTPases are upregulated in human granulosal cells by gonadotropins [99]. Genetic variation in the TBC1D1 gene is associated with risk for severe obesity in women [100]. Moreover, the TBC1D1 protein is structurally similar to AS160, a Rab GTPase-activating protein (GAP) and known regulator of insulin-mediated Glut4 translocation in adipocytes and muscle. TBC1D1 may therefore mediate glucose or fatty acid transporters, especially in insulin-sensitive tissues, and in turn be important for overall energy balance [98, 100, 101]. Although the regulation and role of TBC1D1 in the ovary are unknown, the TBC1D1 transcript is highly expressed in thecal cells, which are insulin sensitive [102], throughout the development of dominant follicles compared with subordinate follicles during follicular waves in the bovine [57]. This finding implies that the lower expression of TBC1D1 in thecal cells observed in the present study may have a negative impact on thecal cell function and survival of the largest-growing follicles during follicular waves in cattle with low versus high AFCs. In conclusion, results demonstrate that the inherently high variation in follicle numbers during follicular waves is linked with significant alterations not only in intrafollicular estradiol production, but also expression of key genes involved in granulosal cell estradiol production (CYP19A1), and potentially in regulation of FSH action (AMH), differentiation and function of thecal cells (TBC1D1), estradiol responsiveness (ESR1, ESR2), and determinants of oocyte quality in cumulus cells (CTSB). More direct studies will be necessary to verify the potential positive association of the intrinsic variation in follicle numbers with oocyte competence, as well as the potential functional roles for estradiol, estrogen receptors, aromatase, cathepsins, AMH, and TBC1D1 in dominant follicle function and/or oocyte quality. The bovine model is uniquely appropriate to resolve these important mechanistic questions because the results of the present study provide new insights into differential gene expression in various follicular cell compartments (measured early in a follicular wave) of animals with a low versus a high AFC and size of the ovarian reserve, and hence the potential mechanisms whereby variation in follicle numbers may have an impact on follicular differentiation, function, and oocyte quality. REFERENCES 1. Block E. A quantitative morphological investigation of the follicular system in newborn female infants. Acta Anat 1953; 17:201–206. 2. Forabosco A, Sforza C, De Pol A, Vizzotto L, Marzona L, Ferrario VF. Morphometric study of the human neonatal ovary. Anat Rec 1991; 231: 201–208. 3. Gougeon A, Ecochard P, Christophe TJ. Age-related changes of the population of human ovarian follicles: increase in the disappearance rate of non-growing and early-growing follicles in aging women. Biol Reprod 1994; 50:653–663. 4. Erickson BH. Development and senescence of the postnatal bovine ovary. J Anim Sci 1966; 25:800–805. 5. Erickson BH. Development and radio-response of the prenatal bovine ovary. J Reprod Fertil 1966; 10:97–105. 6. Richardson SJ, Senikas V, Nelson JF. Follicular depletion during the menopausal transition: evidence for accelerated loss and ultimate exhaustion. J Clin Endocrinol Metab 1987; 65:1231–1237. 7. Lass A, Silye R, Abrams DC, Krausz T, Hovatta O, Margara R, Winston RM. Follicular density in ovarian biopsy of infertile women: a novel method to assess ovarian reserve. Hum Reprod 1997; 12:1028–1031. 8. Chang MY, Chiang CH, Hsieh TT, Soong YK, Hsu KH. Use of the antral follicle count to predict the outcome of assisted reproductive technologies. Fertil Steril 1998; 69:505–510. 9. Scheffer GJ, Broekmans FJM, Dorland M, Habbema JDF, Looman CWN, te Velde ER. Antral follicle counts by transvaginal ultrasonog-

10.

11.

12.

13.

14.

15.

16.

17.

18. 19.

20. 21.

22.

23.

24.

25.

26. 27.

28. 29.

30.

raphy are related to age in women with proven natural fertility. Fertil Steril 1999; 72:845–851. Huang FJ, Chang SY, Tsai MY, Kung FT, Wu JF, Chang HW. Determination of the efficiency of controlled ovarian hyperstimulation in the gonadotropin-releasing hormone agonist-suppression cycle using the initial follicle count during gonadotropin stimulation. J Assist Reprod Genet 2001; 18:91–96. Beckers NGM, Macklon NS, Eijkemans MJC, Fauser BCJM. Women with regular menstrual cycles and a poor response to ovarian hyperstimulation for in vitro fertilization exhibit follicular phase characteristics suggestive of ovarian aging. Fertil Steril 2002; 78:291– 297. Scheffer GJ, Broekmans FJM, Looman CWN, Blankenstein M, Fauser BCJM, De Jong FH, te Velde ER. The number of antral follicles in normal women with proven fertility is the best reflection of reproductive age. Hum Reprod 2003; 18:700–706. Erickson BH, Reynolds RA, Murphree RL. Ovarian characteristics and reproductive performance of the aged cows. Biol Reprod 1976; 15:555– 560. Oliveira JF, Neves JP, Moraes JC, Goncalves PB, Bahr JM, Hernandez AG, Costa LF. Follicular development and steroid concentrations in cows with different levels of fertility raised under nutritional stress. Anim Reprod Sci 2002; 73:1–10. Cushman RA, DeSouza JC, Hedgpeth VS, Britt JH. Superovulatory response of one ovary is related to the micro- and macroscopic population of follicles in the contralateral ovary of the cow. Biol Reprod 1999; 60:349–354. Taneja M, Bols PE, Van de Velde A, Ju JC, Schreiber D, Tripp MW, Levine H, Echelard Y, Riesen J, Yang X. Developmental competence of juvenile calf oocytes in vitro and in vivo: influence of donor animal variation and repeated gonadotropin stimulation. Biol Reprod 2000; 62: 206–213. Singh J, Dominguez M, Jaiswal R, Adams GP. A simple ultrasound test to predict the superstimulatory response in cattle. Theriogenology 2004; 62:227–243. Ginther OJ, Wiltbank MC, Fricke PM, Gibbons JR, Kot K. Selection of the dominant follicle in cattle. Biol Reprod 1996; 55:1187–1194. Ireland JJ, Mihm M, Austin E, Diskin MG, Roche JF. Historical perspective of turnover of dominant follicles during the bovine estrous cycle: key concepts, studies, advancements, and terms. J Dairy Sci 2000; 83:1648–1658. Baerwald AR, Adams GP, Pierson RA. Characterization of ovarian follicular waves in women. Biol Reprod 2003; 69:1023–1031. Baerwald AR, Adams GP, Pierson RA. A new model for ovarian follicular development during the human menstrual cycle. Fertil Steril 2003; 80:116–122. Burns DS, Jimenez-Krassel FJ, Ireland JLH, Knight PG, Ireland JJ. Numbers of antral follicles during follicular waves in cattle: evidence for high variation among animals, very high repeatability in individuals, and an inverse association with serum follicle-stimulating hormone concentrations. Biol Reprod 2005; 73:54–62. Ireland JJ, Ward F, Jimenez-Krassel F, Ireland JLH, Smith GW, Lonergan P, Evans ACO. Follicle numbers are highly repeatable within individual animals but are inversely correlated with FSH concentrations and the proportion of good-quality embryos after ovarian stimulation in cattle. Hum Reprod 2007; 22:1687–1695. Haughian JM, Ginther OJ, Kot K, Wiltbank MC. Relationship between FSH patterns and follicular dynamics and the temporal associations among hormones in natural and GnRH-induced gonadotropin surges in heifers. Reproduction 2004; 127:23–33. Ireland JLH, Scheetz D, Jimenez-Krassel F, Themmen APN, Ward F, Lonergan P, Smith GW, Perez GI, Evans ACO, Ireland JJ. Antral follicle count reliably predicts number of morphologically healthy oocytes and follicles in ovaries of young adult cattle. Biol Reprod 2008; 79:1219– 1225. Block E. A quantitative morphological investigation of the follicular system in newborn female infants. Acta Anatomica 1953; 17:201–206. Tomas C, Nuojua-Huttunen S, Martikainen H. Pretreatment transvaginal ultrasound examination predicts ovarian responsiveness to gonadotropins in in-vitro fertilization. Hum Reprod 1997; 12:220–223. Prior JC. Perimenopause: the complex endocrinology of the menopausal transition. Endocr Rev 1998; 19:397–428. Kupesic S, Kurjak A, Bjelos D, Vujisic S. Three-dimensional ultrasonographic ovarian measurements and in vitro fertilization outcome are related to age. Fertil Steril 2003; 79:190–197. van Rooij IA, Broekmans FJ, te Velde ER, Fauser BC, Bancsi LF, de


31.

32.

33.

34.

35.

36.

37.

38.

39.

40.

41.

42. 43.

44.

45.

46.

47.

48.

49. 50.

51.

52.

Jong FH, Themmen AP. Serum anti-Mullerian hormone levels: a novel measure of ovarian reserve. Hum Reprod 2002; 17:3065–3071. Bryner RW, Garcia-Winder M, Lewis PE, Inskeep EK, Butcher RL. Changes in hormonal profiles during the estrous cycle in old lactating beef cows. Dom Anim Endocrinol 1990; 7:181–190. Kawamata M. Relationship between the number of small follicles prior to superovulatory treatment and superovulatory response in Holstein cows. J Vet Med Sci 1994; 56:965–970. Wolfenson D, Inbar G, Roth Z, Kaim M, Bloch A, Braw-Tal R. Follicular dynamics and concentrations of steroids and gonadotropins in lactating cows and nulliparous heifers. Theriogenology 2004; 62:1042– 1055. Malhi PS, Adams GP, Singh J. Bovine model for the study of reproductive aging in women: follicular, luteal, and endocrine characteristics. Biol Reprod 2005; 73:45–53. El-Toukhy T, Khalaf Y, Hart R, Taylor A, Braude P. Young age does not protect against the adverse effects of reduced ovarian reserve—an eight year study. Hum Reprod 2002; 17:1519–1524. Ibanez L, Potau N, Ferrer A, Rodriguez-Hierro F, Marcos MV, de Zegher F. Reduced ovulation rate in adolescent girls born small for gestational age. J Clin Endocrinol Metab 2002; 87:3391–3393. Ibanez L, Potau N, Enriquez G, Marcos MV, de Zegher F. Hypergonadotrophinaemia with reduced uterine and ovarian size in women born small-for-gestational-age. Hum Reprod 2003; 18:1565–1569. Xu Z, Garverick HA, Smith GW, Smith MF, Hamilton SA, Youngquist RS. Expression of follicle-stimulating hormone and luteinizing hormone receptor messenger ribonucleic acids in bovine follicles during the first follicular wave. Biol Reprod 1995; 53:951–957. Tisdall DJ, Watanabe K, Hudson NL, Smith P, McNatty KP. FSH receptor gene expression during ovarian follicle development in sheep. J Mol Endocrinol 1995; 15:273–281. Kumar TR, Wang Y, Lu N, Matzuk MM. Follicle stimulating hormone is required for ovarian follicle maturation but not male fertility. Nat Genet 1997; 15:201–204. Turzillo AM, Fortune JE. Suppression of the secondary FSH surge with bovine follicular fluid is associated with delayed ovarian follicular development in heifers. J Reprod Fertil 1990; 89:643–653. Turzillo AM, Fortune JE. Effects of suppressing plasma FSH on ovarian follicular dominance in cattle. J Reprod Fertil 1993; 98:113–119. Mihm M, Good TEM, Ireland JLH, Ireland JJ, Knight PG, Roche JF. Decline in serum follicle-stimulating hormone concentrations alters key intrafollicular growth factors involved in selection of the dominant follicle in heifers. Biol Reprod 1997; 57:1328–1337. Baarends WM, Uilenbroek JT, Kramer P, Hoogerbrugge JW, van Leeuwen EC, Themmen AP, Grootegoed JA. Anti-mullerian hormone and anti-mullerian hormone type II receptor messenger ribonucleic acid expression in rat ovaries during postnatal development, the estrous cycle, and gonadotropin-induced follicle growth. Endocrinology 1995; 136: 4951–4962. Durlinger AL, Gruijters MJ, Kramer P, Karels B, Kumar TR, Matzuk MM, Rose UM, deJong FH, Uilenbroek JT, Grootegoed JA, Themmen AP. Anti-Mullerian hormone attenuates the effects of FSH on follicle development in the mouse ovary. Endocrinology 2001; 142:4891–4899. Sunderland SJ, Crowe MA, Boland MP, Roche JF, Ireland JJ. Selection, dominance and atresia of follicles during the oestrous cycle of heifers. J Reprod Fertil 1994; 101:547–555. Evans ACO, Ireland JLH, Winn ME, Lonergan P, Smith GW, Coussens PM, Ireland JJ. Identification of genes involved in apoptosis and dominant follicle development during follicular waves in cattle. Biol Reprod 2004; 70:1475–1484. Bettegowda A, Patel OV, Ireland JJ, Smith GW. Quantitative analysis of messenger RNA abundance for ribosomal protein L-15, cyclophilin-A, phosphoglycerokinase, glucuronidase, glyceraldehyde 3-phosphate dehydrogenase, b-actin and histone H2A during bovine oocyte maturation and early embryogenesis in vitro. Mol Reprod Dev 2006; 73:267–278. Sample WF, Lippe BM, Gyepes MT. Gray-scale ultrasonography of the normal female pelvis. Radiology 1977; 125:477–483. Ireland JJ, Roche JF. Development of nonovulatory antral follicles in heifers: changes in steroids in follicular fluid and receptors for gonadotropins. Endocrinology 1983; 112:150–156. Ireland JJ, Roche JF. Growth and differentiation of large antral follicles after spontaneous luteolysis in heifers: changes in concentration of hormones in follicular fluid and specific binding of gonadotropins to follicles. J Anim Sci 1983; 57:157–167. Forde N, Mihm M, Canty MJ, Zielak AE, Baker PJ, Park S, Lonergan P, Smith GW, Coussens PM, Ireland JJ, Evans ACO. Differential expression of signal transduction factors in ovarian follicle development:

53.

54. 55.

56.

57.

58.

59.

60.

61.

62.

63.

64.

65.

66.

67.

68.

69.

70.

71.

963

a functional role for betaglycan and FIBP in granulosa cells in cattle. Physiol Genomics 2008; 33:193–204. Dupont S, Krust A, Gansmuller A, Dierich A, Chambon P, Mark M. Effect of single and compound knockouts of estrogen receptors alpha (ERalpha) and beta (ERbeta) on mouse reproductive phenotypes. Development 2000; 127:4277–4291. Schams D, Berisha B. Steroids as local regulators of ovarian activity in domestic animals. Domest Anim Endocrinol 2002; 23:53–65. Bao B, Garverick HA, Smith GW, Smith MF, Salfen BE, Youngquist RS. Changes in messenger ribonucleic acid encoding luteinizing hormone receptor, cytochrome P450-side chain cleavage, and aromatase are associated with recruitment and selection of bovine ovarian follicles. Biol Reprod 1997; 56:1158–1168. Modi D, Bhartiya D, Puri C. Developmental expression and cellular distribution of Mullerian inhibiting substance in the primate ovary. Reproduction 2006; 132:443–453. Zielak AE, Forde N, Park SDE, Doohan F, Coussens PM, Smith GW, Ireland JJ, Lonergan P, Evans ACO. Identification of novel genes associated with dominant follicle development in cattle. Reprod Fertil Dev 2007; 19:967–975. Jakimiuk AJ, Weitsman SR, Yen HW, Bogusiewicz M, Magoffin DA. Estrogen receptor alpha and beta expression in theca and granulosa cells from women with polycystic ovary syndrome. J Clin Endocrinol Metab 2002; 87:5532–5538. Rosenfeld CS, Yuan X, Manikkam M, Calder MD, Garverick HA, Lubahn DB. Cloning, sequencing, and localization of bovine estrogen receptor-b within the ovarian follicle. Biol Reprod 1999; 60:691–697. Forde N, Rogers M, Canty MJ, Lonergan P, Smith GW, Coussens PM, Ireland JJ, Evans ACO. Association of the prion protein and its expression with ovarian follicle development in cattle. Mol Reprod Dev 2008; 75:243–249. Zielak AE, Canty MJ, Forde N, Coussens PM, Smith GW, Lonergan P, Ireland JJ, Evans ACO. Differential expression of genes for transcription factors in theca and granulosa cells following selection of a dominant follicle in cattle. Mol Reprod Dev 2008; 75:904–914. Van Tol HTA, Van Eijk MJT, Mummery CI, Van Den Hurk R, Bevers MM. Influence of FSH and hCG on the resumption of meiosis of bovine oocytes surrounded by cumulus cells connected to membrana granulosa. Mol Reprod Dev 1996; 45:218–224. Beker-van Woudenberg AR, van Tol HTA, Roelen BAJ, Colenbrander B, Bevers MM. Estradiol and its membrane-impermeable conjugate (estradiol-bovine serum albumin) during in vitro maturation of bovine oocytes: effects on nuclear and cytoplasmic maturation, cytoskeleton, and embryo quality. Biol Reprod 2004; 70:1465–1474. Austin EJ, Mihm M, Evans ACO, Knight PG, Ireland JLH, Ireland JJ, Roche JF. Alterations in intrafollicular regulatory factors and apoptosis during selection of follicles in the first follicular wave of the bovine estrous cycle. Biol Reprod 2001; 64:839–848. Fanchin R, Mendez Lozano DH, Frydman N, Gougeon A, di Clemente N, Frydman R, Taieb J. Anti-Mullerian hormone concentrations in the follicular fluid of the preovulatory follicle are predictive of the implantation potential of the ensuing embryo obtained by in vitro fertilization. J Clin Endocrinol Metab 2007: 97:1796–1802. Kobayashi Y, Jimenez-Krassel F, Li Q, Yao J, Huang R, Ireland JJ, Coussens PM, Smith GW. Evidence that cocaine- and amphetamineregulated transcript is a novel intraovarian regulator of follicular atresia. Endocrinology 2004; 145:5373–5383. Ireland JLH, Good TEM, Knight PG, Ireland JJ. Alterations in amounts of different forms of inhibin during follicular atresia. Biol Reprod 1994; 50:1265–1276. Jimenez-Krassel F, Winn ME, Burns D, Ireland JLH, Ireland JJ. Evidence for a negative intrafollicular role for inhibin in regulation of estradiol production by granulosa cells. Endocrinology 2003; 144:1876– 1886. Bettegowda A, Yao J, Sen A, Li Q, Lee K-B, Kobayashi Y, Patel OV, Coussens PM, Ireland JJ, Smith GW. JY-1, an oocyte-specific gene, regulates granulosa cell function and early embryonic development in cattle. Proc Natl Acad Sci U S A 2007; 104:17602–17607. Silva CC, Groome NP, Knight PG. Immunohistochemical localization of inhibin/activin alpha, betaA and betaB subunits and follistatin in bovine oocytes during in vitro maturation and fertilization. Reproduction 2003; 125:33–42. Patel VP, Bettegowda A, Ireland JJ, Coussens PM, Lonergan P, Smith GW. Functional genomics studies of oocyte competence: evidence that reduced transcript abundance for follistatin is associated with poor developmental competence of bovine oocytes. Reproduction 2006; 132: 1–13.

964

IRELAND ET AL.

72. Bettegowda A, Patel OV, Lee K-B, Park KE, Salem M, Yao J, Ireland JJ, Smith GW. Identification of novel bovine cumulus cell molecular markers predictive of oocyte competence: functional and diagnostic implications. Biol Reprod 2008; 79:301–309. 73. Prendiville DJ, Enright WJ, Crowe MA, Finnerty M, Hynes N, Roche JF. Immunization of heifers against gonadotropin-releasing hormone: Antibody titers, ovarian function, body growth and carcass characteristics. J Anim Sci 1995; 73:2382–2389. 74. Bleach ECL, Glencross RG, Feist SA, Groome NP, Knight PG. Plasma inhibin A in heifers: relationship with follicle dynamics, gonadotropins, and steroids during the estrous cycle and after treatment with bovine follicular fluid. Biol Reprod 2001; 64:743–752. 75. Knight PG, Castillo RJ, Glencross RG, Beard AJ, Wrathall JHM. Isolation of bovine ovarian inhibin, its immunoneutralization in vitro and immunolocalization in bovine ovary. Domest Anim Endocrinol 1990; 7: 299–313. 76. Knight PG, Muttukrishna S, Groome NP. Development and application of a two-site enzyme immunoassay for the determination of ‘‘total’’ activin-A concentrations in serum and follicular fluid. J Endocrinol 1996; 148:267–279. 77. Tannetta DS, Feist SA, Bleach ECL, Groome NP, Evans LW, Knight PG. Effects of active immunization of sheep against an amino terminal peptide of the inhibin alpha-c subunit on intrafollicular levels of activin A, inhibin A and follistatin. J Endocrinol 1998; 157:157–168. 78. Kevenaar ME, Meerasahib MF, Kramer P, van de Lang-Born BMN, de Jong FH, Groome NP, Themmen APN, Visser JA. Serum AMH levels reflect the size of the primordial follicle pool in mice. Endocrinology 2006; 147:3228–3234. 79. Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real-time quantitative PCR and the 2-DDCT method. Methods 2001; 25:402–408. 80. Whelan JA, Russell NB, Whelan MA. A method for the absolute quantification of cDNA using real-time PCR. J Immunol Methods 2003; 278:261–269. 81. Sen A, Bettegowda A, Jimenez-Krassel F, Ireland JJ, Smith GW. Cocaine- and amphetamine-regulated transcript regulation of folliclestimulating hormone signal transduction in bovine granulosa cells. Endocrinology 2007; 148:4400–4410. 82. SAS Institute Inc. SAS User’s Guide: Statistics. Cary, NC: SAS Institute Inc; 1995. 83. Silva JM, Price CA. Effect of follicle-stimulating hormone on steroid secretion and messenger ribonucleic acids encoding cytochromes P450 aromatase and cholesterol side-chain cleavage in bovine granulosa cells in vitro. Biol Reprod 2000; 62:186–191. 84. Richards JS. Estradiol receptor content in rat granulosa cells during follicular development: modification by estradiol and gonadotropins. Endocrinology 1975; 97:1174–1184. 85. Peng XR, Hsueh AJ, LaPolt PS, Bjersing L, Ny T. Localization of luteinizing hormone receptor messenger ribonucleic acid expression in ovarian cell types during follicle development and ovulation. Endocrinology 1991; 129:3200–3207. 86. Li R, Norman RJ, Armstrong DT, Gilchrist RB. Oocyte-secreted factor(s) determine functional differences between bovine mural granulosa cells and cumulus cells. Biol Reprod 2000; 63:839–845.

87. Eppig JJ, O’Brien MJ, Pendola FL, Watanabe S. Factors affecting the developmental competence of mouse oocytes grown in vitro: folliclestimulating hormone and insulin. Biol Reprod 1998; 59:1445–1453. 88. McTavish KJ, Jimenez M, Walters KA, Spaliviero J, Groome NP, Themmen AP, Visser JA, Handelsman DJ, Allan CM. Rising folliclestimulating hormone levels with age accelerate female reproductive failure. Endocrinology 2007; 148:4432–4439. 89. Lonergan P, Monaghan P, Rizos D, Boland M, Gordon I. Effect of follicle size on bovine oocyte quality and developmental competence following maturation, fertilization and culture in vitro. Mol Reprod Dev 1994; 37:48–53. 90. Blondin P, Coenen K, Guilbault LA, Sirard MA. Superovulation can reduce the developmental competence of bovine embryos. Theriogenology 1996; 46:1191–1203. 91. Turk V, Turk B, Turk D. Lysosomal cysteine proteases: facts and opportunities. EMBO J 2001; 20:4628–4633. 92. Waters KM, Safe S, Gaido KW. Differential gene expression in response to methoxychlor and estradiol through ER(alpha), ER(beta), and AR in reproductive tissues of female mice. Toxicol Sci 2001; 63:47–56. 93. Oksjoki S, Soderstrom M, Vuorio E, Anttila L. Differential expression patterns of cathepsins B, H, K, L and S in the mouse ovary. Mol Hum Reprod 2001; 7:27–34. 94. Rey R, Lukas-Croisier C, Lasala C, Bedecarra´s P. AMH/MIS: what we know already about the gene, the protein and its regulation. Mol Cell Endocrinol 2003; 211:21–31. 95. Visser JA, de Jong FH, Laven JSE, Themmen APN. Anti-Mullerian hormone: a new marker for ovarian function. Reproduction 2006; 131: 1–9. 96. Roach WG, Chavez JA, Mıˆinea CP, Lienhard GE. Substrate specificity and effect on GLUT4 translocation of the Rab GTPase-activating protein Tbc1d1. Biochem J 2007; 403:353–358. 97. Schwartz SL, Cao C, Pylypenko O, Rak A, Wandinger-Ness A. Rab GTPases at a glance. J Cell Sci 2007; 120:3905–3910. 98. Chavez JA, Roach WG, Keller SR, Lane WS, Lienhard GE. Inhibition of GLUT4 translocation by Tbc1d1, a Rab GTPase-activating protein abundant in skeletal muscle, is partially relieved by AMP-activated protein kinase activation. J Biol Chem 2008; 283:9187–9195. 99. Sasson R, Rimon E, Dantes A, Cohen T, Shinder V, Land-Bracha A, Amsterdam A. Gonadotrophin-induced gene regulation in human granulosa cells obtained from IVF patients. Modulation of steroidogenic genes, cytoskeletal genes and genes coding for apoptotic signalling and protein kinases. Mol Hum Reprod 2004; 10:299–311. 100. Stone S, Abkevich V, Russell DL, Riley R, Timms K, Tran T, Trem D, Frank D, Jammulapati S, Neff CD, Iliev D, Gress R, et al. TBC1D1 is a candidate for a severe obesity gene and evidence for a gene/gene interaction in obesity predisposition. Hum Mol Genet 2006; 15:2709– 2720. 101. Taylor EB, An D, Kramer HF, Yu H, Fujii NL, Roeckl KSC, Bowles N, Hirshman MF, Xie J, Feener EP, Goodyear LJ. Discovery of TBC1D1 as an insulin-, AICAR-, and contraction-stimulated signaling nexus in mouse skeletal muscle. J Biol Chem 2008; 283:9787–9796. 102. Spicer LJ, Francisco CC. Adipose obese gene product, leptin, inhibits bovine ovarian thecal cell steroidogenesis. Biol Reprod 1998; 58:207– 212.