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Regular Article |
a Departments of Animal Health and Biomedical Sciences and Dairy Science,
b University of Wisconsin, Madison, Wisconsin 53706
ABSTRACT
Intrafollicular changes in the largest follicle (F1) and second-largest (F2) follicle were examined in relation to follicle diameter deviation. Deviation is characterized by continued growth of the largest follicle and the cessation of growth of the smaller follicles. Granulosa cells and follicular fluid were obtained from slaughterhouse ovaries (n = 95 pairs, experiment 1), and follicular fluid was collected in vivo (n = 28 heifers, experiment 2). Several ranges in the diameter of F1 were used to represent the progressive growth of the follicle. The diameter range with the first significant increase in the difference between F1 and F2 was determined for each end point and was used as an indicator of the sequence of events associated with diameter deviation. An increased difference for diameter and for estradiol concentration occurred (P < 0.05) simultaneously at the 8.5- to 8.9-mm range in both experiments. In experiment 1, the increased difference between F1 and F2 in LH receptor (LHr) mRNA expression occurred (P < 0.05) at the 8.0- and 8.4-mm range. In F2 of experiment 2, there was a progressive decrease (P < 0.05) in free insulin-like growth factor (IGF)-1 and a progressive increase (P < 0.05) in IGF binding protein (BP)-2 across the follicle-diameter ranges (7.5-11.2 mm). No differences were detected between F1 and F2 for 3ß-hydroxysteroid dehydrogenase mRNA expression in experiment 1 and testosterone, total inhibin, and dimeric inhibin-A concentrations in experiment 2. The results indicated that the acquisition of granulosa cell LHrs by F1, as indicated by increased LHr mRNA expression, occurred one diameter range before an increased difference between F1 and F2 for diameter or estradiol concentrations. On a temporal basis, it is concluded that LHr acquisition plays a role in the establishment of diameter deviation. In addition, the reduced growth of F2 may have involved the reduced bioavailability of IGF-1 in association with elevated IGFBPs.
estradiol, follicular development, granulosa cells, growth factors, luteinizing hormone, ovary
INTRODUCTION
Follicular growth occurs in two or three waves during the bovine estrous cycle [1, 2]. Each wave begins with a cohort of growing follicles (4.0 mm in diameter) from which a single follicle continues growth and becomes dominant; other follicles in the cohort cease growth and regress [3]. The beginning of the difference in growth rates between the two largest follicles of a wave has been termed the beginning of diameter deviation [4]. From several studies in our laboratory (reviewed by Ginther [5]), diameter deviation began when the largest follicle of the wave reached a mean of 8.5 mm, about 2.5 days after follicle emergence at 4 mm. Although the growth rate of the two largest follicles was similar from emergence to beginning of deviation (parallel-growth phase), mean diameter of the second-largest follicle lagged behind the diameter of the largest follicle by about 0.5 mm or by an equivalent of 8 h.
The mechanism of follicle deviation is unknown; however, it has been postulated that deviation involves a close two-way functional coupling between changes in FSH concentration and follicle growth [6]. Temporally, the peak in FSH concentration in association with wave emergence is followed by a decline during the parallel growth phase. Despite the reduced growth rate of all but the largest follicle at the end of the parallel phase or the beginning of deviation, FSH concentration continues to decrease and reaches a nadir shortly thereafter [5]. Recently, the decline of FSH to basal concentrations after deviation was attributed, at least partly, to estradiol emanating from the future dominant follicle [7]. Ablation of the largest follicle when it reached 8.5 mm resulted in a decrease in systemic estradiol concentrations and an increase in FSH concentrations. Moreover, a minimal single dose of exogenous estradiol interfered with the ablation-induced FSH increase [7]. Apart from the functional role of estradiol in follicle deviation, other biochemical events have been studied in relation to follicle selection in cattle but have not focused on the time of diameter deviation.
It has been proposed that expression of LH receptor (LHr) mRNA may be associated with follicle selection [8]. The expression of LHr mRNA was first detected in the granulosa cells of follicles 8 mm in diameter and was significantly higher in the dominant follicle at a mean of 10.8 mm; LHr mRNA expression was not detected in granulosa cells of follicles <8 mm or in subordinate follicles. Moreover, LHr mRNA expression was detected in bovine granulosa cells of slaughterhouse ovaries from follicles <4 mm [9]. Qualitatively, therefore, it appears that LHr mRNA is present in granulosa cells of antral follicles early in development. Quantitatively, however, it is not known whether the amounts of LHr mRNA differ among follicles in association with follicle deviation, especially with respect to the future dominant and subordinate follicles.
Several studies have shown that insulin-like growth factor-1 (IGF-1) plays a role in ovarian folliculogenesis in cattle and other species through paracrine-autocrine regulation [10]. The bioactivity of IGF-1 is controlled by its association with at least six IGF binding proteins (IGFBPs). The high affinity of these binding proteins to IGF-1 ensures that most (90–95%) of the IGF-1 in the body is present in bound form [11]. It has been proposed that the unbound or the free fraction of the IGF-1 may be the dominant biologically active form [12]. Differences in total IGF-1 levels have been found in follicular fluid among large (8 mm) estrogen-active follicles, large estrogen-inactive follicles, and small (<5 mm) follicles [13]. However, others did not find differences in total IGF-1 in follicular fluid of corresponding dominant, large, and small follicles [14] or between dominant and subordinate follicles [15]. Reportedly [13], growing follicles have decreasing amounts of IGFBP-2 and other low molecular weight IGFBPs; therefore, more IGF-1 would be available for binding with the IGF-1 receptors. In this regard, mRNA expression of IGF-1 and IGFBP-2 increased and decreased, respectively, in dominant follicles of wave 1 [16].
Two experiments were done to test the hypothesis that the increase in the difference between the two largest follicles occurred at the same time for diameter, follicular-fluid concentrations of estradiol, and expression of mRNA for LHr and 3ß-hydroxysteroid dehydrogenase (HSD). For observational purposes, differential changes in follicular fluid were also examined for concentrations of testosterone, progesterone, free IGF-1, IGFBP-2, total inhibin, and dimeric inhibin-A.
MATERIALS AND METHODS
Experiment 1
Paired ovaries with an apparent immature corpus luteum (CL) were collected from a slaughterhouse. Immaturity of the CL was based on gross appearance and a diameter of 16 mm or less [17]. The cattle were primarily Holstein Friesians. Age and reproductive status were unknown, except that ovaries from pregnant animals, based on external inspection of the uterus, and apparent abnormal reproductive organs were not used. Each pair of ovaries was placed in a separate bag, placed on ice, and delivered to the laboratory in <3 h. In the laboratory, each ovary of a pair was washed with PBS and scanned with an ultrasound machine (Aloka SSD-500V; Aloka Co., Wallingford, CT) equipped with a 7.5-MHz linear-array transducer. The scanning was done in a water bath as described [18]. The distance from the transducer face to the specimen was approximately the same as for transrectal scanning to favor similar resolution between experiments 1 and 2. Briefly, an ovary was pinned to a standoff pad and follicular and luteal measurements were made. The mean of the maximum diameter of width and height was used as the diameter of the follicle or CL. The three largest follicles (largest, F1; second-largest, F2; third-largest follicle, F3) were identified, and collection of follicular contents was done using an 18-gauge needle attached to a 1-ml syringe. After removal of follicular fluid, each follicle was flushed twice with PBS to collect the granulosa cells.
Granulosa cells were separated from the follicular fluid and PBS by centrifuging at 500 x g for 10 min and immediately lysed in homogenization buffer (4 M guanidinium isothiocyanate, 10 mM Tris-HCl, pH 8.0, 0.5% SDS, and 1% dithiothreitol) for isolation of mRNA and stored at -80°C. The methodology for isolation of mRNA has been described in detail [19]. The follicular fluid was stored at -20°C until assay. Because ovarian status was not known, morphologic and functional restraints were imposed to optimize the study of follicles of wave 1, encompassing the expected time of deviation. The morphologic conditions were based on a previous report [17] and involved using only pairs of ovaries having an F1 ranging between 7.0 mm and 10.0 mm and a new CL of 16 mm or less. The 10.0-mm limit on F1 was intended to exclude cattle with a mature dominant follicle, and the small developing CL helped define the follicles as being part of wave 1. Inspection of previous data [17] indicated that regressing follicles from a previous wave were seldom as large as 7.0 mm in the presence of an immature CL. For functional restraint, follicles were excluded if follicular-fluid estradiol concentrations were <3.0 ng/ml. Inspection of data of recent experiments in our laboratory involving follicles known to be in the growing phase and as large as 7.0 mm did not have intrafollicular concentrations of estradiol <3.0 ng/ml. Thus, an F1 with <3.0 ng/ml of estradiol was replaced for all end points with F2 and, F2 was replaced with F3 if F2 and F3 had >3.0 ng/ml of estradiol. Follicle diameter (mm) ranges were established for F1 and the corresponding F2 based on the F1 diameter as follows: 7.0–7.4, 7.5–7.9, 8.0–8.4, 8.5–8.9, 9.0–9.4, and 9.5–10.0. An F2 that attained 9.0 mm or larger diameter in the 9.0- to 9.4-mm and 9.5- to 10.0-mm ranges was considered to be a codominant follicle, and therefore, F3 was used to replace F2; the codominant F2 was analyzed separately. A total of 104 pairs of ovaries were scanned and after applying the functional restraints, 95 F1s and 78 F2s were used for the study.
Experiment 2
Twenty-eight Holstein heifers between 24 and 36 mo of age and weighing 490–680 kg were used. The feeding program and the prostaglandin F2
(Lutalyse; Pharmacia and Upjohn Co., Kalamazoo, MI) protocol for inducing luteolysis to schedule ovulation have been described [20]. The same ultrasound machine and transducer as used in experiment 1 was used transrectally to scan the ovaries and measure follicles [21, 22]. Follicular growth was monitored daily until the largest follicle of the postovulatory wave (wave 1) reached 6.5 mm; thereafter, examinations were done every 8 h (0600, 1400, and 2200 h). Follicular fluid was aspirated from the two largest follicles (F1 and F2) when F1 reached 7.5, 8.5, 9.5, or 10.5 mm (n = 7 heifers/group). Follicular fluid samples (50–300 µl) were collected using a 5.0-MHz convex-array ultrasound transducer equipped with a 50-cm extension for intravaginal access as described [20]. The follicular fluid was taken to the laboratory on ice, centrifuged at 500 x g for 10 min, and decanted and stored at -20°C until hormone analyses. The cell pellet was discarded. If F2 reached a diameter of 9.0 mm in the 9.5- and 10.5-mm groups, it was considered a codominant follicle. A codominant follicle (F2) that had either an extreme diameter or follicular-fluid estradiol concentration that was a statistical outlier (described later [23]) was excluded from analysis for all end points.
Messenger RNA Quantification
The detailed procedure of standard curve, quantitative, competitive, reverse transcriptase-polymerase chain reaction (RT-PCR) using granulosa cell mRNA isolated by Magnetight Oligo(dt) beads (Novagen, Madison, WI) has been described [19, 24]. Native and competitor cDNA for LHr and 3ß-HSD were prepared from plasmids containing the gene-specific constructs, amplified by PCR using primers flanking the T7 promoter site, and transcribed in vitro using T7 RNA polymerase [24, 25]. The cRNA was aliquoted and stored at -80°C. Each aliquot was used only once to reduce variation due to potential degradation of RNA after freezing and thawing. The RNA was reverse transcribed at 42°C for 1 h followed by heating to 95°C for 10 min and quick chilling to 4°C in a programmable thermocycler (PTC-100; MJ Research, Watertown, MA). Five microliters of the RT product in the presence of 15 µl of PCR mix were subjected to 33 cycles of amplification by PCR (30-sec denaturation at 95°C, 30-sec annealing at 57°C, and 30-sec extension at 72°C) followed by final extension at 72°C for 5 min. The PCR products (10 µl) were directly separated on a 5% acrylamide gel with 1x TBE (0.09 M Tris, 0.09 M boric acid, 0.001 M EDTA, pH 8.0) buffer at 110 V for 40 min using a Miniprotean-II electrophoresis system (BioRad, Richmond, CA). The gel was stained with ethidium bromide and viewed under a UV transilluminator. The UV image was analyzed by Collage software (Fotodyne, Hartland, WI). A ratio of the pixel intensity of the native:competitor was calculated from each band in respective lanes of the gel. The logarithmic ratio of native:competitor cRNA was plotted against the logarithmic transformation of known amounts of native cRNA to produce a standard curve. The concentration (copies/cell) of specific transcripts for LHr and 3ß-HSD mRNA of experimental samples was obtained by comparison to the standard curve [26]. For quality controls, mRNA was prepared from bovine luteal tissue, aliquoted, and stored at -80°C. With each mRNA assay an equal volume of quality control mRNA was run to calculate intra- and interassay coefficients of variation (CV). The intra- and interassay CVs were 17.0% and 22.3% for LHr mRNA assay and 15.8% and 6.5% for 3ß-HSD mRNA, respectively.
Hormone Assays
Follicular-fluid concentrations of estradiol were determined using an RIA kit (double antibody estradiol; DPC, Los Angeles, CA) with modifications and validated for use in cattle [27]. The details of the method as used in our laboratory for bovine follicular fluid have been reported [28]. A working dilution of 1:1000 in assay buffer was used for follicular fluid samples. The intra- and interassay CV for quality control samples were 8.8% and 15.5% for low concentration and 15.8% and 7.4% for high concentration, respectively. The sensitivity was 2.1 pg/ml (n = 7 assays) as calculated from 2 SD below the mean maximum percent binding.
Follicular-fluid concentrations of progesterone were determined using a competitive ELISA described for use in cattle [18, 29] in which the color intensity of the enzyme substrate was inversely proportional to the concentration. A protein-based assay buffer containing 1% (v/v) steroid-reduced bovine follicular fluid was used to prepare the standards (0.16-10 ng/ml), and buffer alone was used for dilution of follicular fluid and quality control samples. Serial dilutions (0.25-5 µl) of a pool of bovine follicular fluid in a total volume of 100 µl resulted in a displacement curve that was similar to the standard curve. A working dilution of 1:100 was used for the experimental follicular fluid samples because 1 µl of pooled follicular fluid resulted in a percent binding that was central to the range of the standard curve. The intra- and interassay CVs for quality control samples were 7.3% and 4.7% for low concentration and 8.7% and 4.6% for high concentration, respectively. The assay sensitivity was 0.1 ng/ml (n = 3 assays) as calculated from 2 SD below the mean optical density (OD) of the mean maximum percent binding.
Follicular-fluid testosterone concentrations were determined using a double antibody RIA kit (DSL-4100; Diagnostic Systems Laboratories, Inc., Webster, TX). The kit was developed for use with human serum, but was adapted and validated for use with bovine follicular fluid in our laboratory. The RIA assay buffer (PBS with 0.1% gelatin) containing 1% (v/v) steroid-reduced bovine follicular fluid was used to prepare testosterone (Sigma Chemical Co., St. Louis, MO) standards (0.05-25 ng/ml), and buffer alone was used for dilution of follicular fluid and quality control samples. Serial dilutions (0.125-10 µl) of pooled follicular fluid in a total volume of 50 µl resulted in a displacement curve that was similar to the standard curve. A working dilution of 1:100 was used for experimental follicular-fluid samples because 0.5 µl of pooled follicular fluid resulted in a percent binding that was central to the range of the standard curve. According to the manufacturer, the cross-reactivity of the antiserum was minimal or nondetectable with other steroids (5-dihydrotestosterone, 6%; 5-androstane-3ß,17ß-diol, 2.2%; 11-oxotestosterone, 1.8%; androstenedione, 0.9%; 5ß-dihyrotestosterone, 0.6%; 5ß-androstane-3ß,17ß-diol, 0.5%; estradiol-17ß, 0.4%; 5-androstane-3-ol-17-one, 0.2%). The intra-assay CVs for quality control samples were 14.1% and 7.1% for low and high concentration, respectively. The sensitivity of the assay was 0.02 ng/ml as determined by 2 SD below the mean maximum percent binding.
Follicular-fluid concentrations of IGFBP-2 were determined using a double antibody RIA kit (DSL-7100; Diagnostic Systems Laboratories) that was developed for use with human serum but was adapted and validated for use with bovine follicular fluid in our laboratory. This approach contrasts with the previous studies in which IGFBPs were analyzed using qualitative or semiquantitative ligand blot assay [30]. The kit-supplied standards (2.5-100 ng/ml) and quality-control samples were reconstituted with distilled water, and follicular-fluid samples were diluted with the protein-based zero standard supplied in the kit. Serial dilutions (2-8 µl) of a pool of bovine follicular fluid in a total volume of 200 µl resulted in a displacement curve that was similar to the standard curve. A working dilution of 1:50 was used for the experimental follicular-fluid samples because 4 µl resulted in a percent binding that was central to the range of the standard curve. According to the manufacturer, the cross-reactivity of this assay with IGFBP-3, -4, -5, and -6 was not detectable at 0.5 µg/tube. Correspondingly, IGF-1 and IGF-2 were not detectable at 1 µg/tube. All the experimental samples were assayed in a single assay and the intra-assay CVs for quality control samples were 9.4% and 15.5% for low and high-concentration, respectively. The assay sensitivity was 0.32 ng/ml as determined by 2 SD below the mean maximum percent binding.
Follicular-fluid concentrations of free IGF-I were determined using a sandwich-type ELISA kit (DSL 10-9400; Diagnostic Systems Laboratories) that was developed for use with human serum but was adapted and validated for use with bovine follicular fluid in our laboratory, because the homology of the amino-acid sequence of human and bovine IGF-1 has been reported to be 100% [31]. Unlike the ELISA for progesterone, the color intensity of the enzyme substrate was directly proportional to the free IGF-I concentration. The standards from the kit (0.06-2.7 ng/ml) and quality control samples were reconstituted with distilled water while follicular-fluid samples were diluted with the protein-based zero standard containing BSA supplied in the kit. Serial dilutions (3-25 µl) of a pool of bovine follicular fluid in a total volume of 50 µl resulted in a dose-response curve that was similar to the standard curve. A working dilution of 1:10 was used for the experimental follicular-fluid samples because 5 µl resulted in an OD that was central to the range of the standard curve. According to the manufacturer, the addition of IGFBP-3 to IGF-1 resulted in a dose-dependent decrease in the concentration of free IGF-1 detected by the assay. In addition, the manufacturer indicated that the cross-reactivity of this assay with IGF-2, insulin, proinsulin, and growth hormone was not detectable at 0.2 µg/tube. The intra-assay CVs for quality control samples were 13.2% and 15.1% CV for low and high concentration, respectively. The assay sensitivity was 0.04 ng/ml as determined by 2 SD above the mean OD of the zero standard.
Follicular-fluid concentrations of total inhibin were determined using an RIA kit (Institute of Reproduction and Development; Monash Medical Center, Clayton, Victoria, Australia). The kit included inhibin as a 32-kDa fraction of bovine follicular fluid for iodination and anti-inhibin (no. 1989, pool B) that was generated against a 31-kDa fraction of bovine follicular fluid. As reported [32], the anti-serum cross-reacts with dimeric inhibin-A and -B, higher molecular weight forms of inhibin, -N--C and pro--C. Despite minimal cross-reactivity (<1.0%) with -N and the ßA subunits, the assay was considered to detect total inhibin. Follicular-fluid concentrations of total inhibin were expressed relative to a recombinant preparation of the 32-kDa form of bovine inhibin (IP-1095; Peninsula Laboratories Europe Ltd., St. Helens, Merseyside, England). Procedures for the assay were similar to those previously described [32], except that the iodination was done using Iodogen [33]. Serial dilutions of the standard (2.5-250 ng/ml) and a pool of bovine follicular fluid (0.025-0.4 µl) in a total volume of 100 µl resulted in displacement curves that were similar. A working dilution of 1:4000 was used for the follicular-fluid samples because 0.025 µl resulted in a percent binding that was central to the range of standard curve. The intra-assay CV was 13.5% for quality control samples prepared from a pool of bovine follicular fluid. The assay sensitivity was 5.2 ng/ml as determined by 2 SD below the mean maximum percent binding.
Follicular-fluid concentrations of dimeric inhibin-A were determined using a sandwich-type ELISA kit (MCA 950KZZ; Serotec Inc., Raleigh, NC) that was developed for use with human serum. Monoclonal antibodies were directed against a native portion of the ßA subunit for capture and against a synthetic portion of the -C subunit for detection. Color intensity of the enzyme substrate was directly proportional to the concentration. The kit has been validated for use with bovine follicular fluid [34] and was adapted for use in our laboratory. The standards from the kit (3.9-500 pg/ml) were reconstituted with fetal calf serum that also served as the zero standard. Serial dilutions (0.025-0.2 µl) of a pool of bovine follicular fluid in a total volume of 100 µl resulted in a dose-response curve that was similar to the standard curve. A working dilution of 1:2000 was used for experimental follicular fluid samples because 0.05 µl of pooled follicular fluid resulted in an OD that was central to the range of standard curve. According to the manufacturer, the cross-reactivity of the assay with pro-alpha C subunit, inhibin-B, and activin was minimal. The intra- and interassay CVs for quality control samples were 2.5% and <1.0%, respectively. Determinations were made from a pool of bovine follicular fluid. The sensitivity was 9.2 pg/ml (n = 2 assays) as determined by 2 SD above the mean OD of the zero standard.
Statistical Analyses
A suspected outlier for each end point within a follicle-diameter range was challenged using Dixons outlier test [23]. If the calculated r-value exceeded the critical r-value at P < 0.05, the extreme observation was excluded from the analyses. Follicle diameter, follicular-fluid concentrations of estradiol, progesterone, testosterone, IGFBP-2, free IGF-1, total inhibin, dimeric inhibin-A, and LHr and 3ß-HSD mRNA were analyzed with a general linear model using the statistical analysis system (SAS) to determine the effects of follicle (F1 and F2), follicle-diameter range, and their interactions [35]. A significant (P < 0.05) effect of follicle or interaction was further analyzed by Duncans multiple range test to locate the differences among follicle-diameter ranges. One-sided, unpaired t-tests were used for single-point measurements, comparing follicles within or between diameter ranges. Pearsons correlation coefficients were used to examine relationships among end points.
RESULTS
Experiment 1
Of the initial 173 observations (95 F1s and 78 F2s) for each end point, 169, 166, and 157 observations for estradiol, LHr mRNA, and 3ß-HSD mRNA, respectively, were available for analysis after removing extreme values (4.0–9.2%) as statistical outliers; all 173 observations were available for follicle diameter.
A total of eight apparent codominant follicles (8.4%, F2 >9.0 mm) were detected and compared to the corresponding F1 for diameter, follicular-fluid estradiol concentration, and granulosa cell mRNA content of LHr and 3ß-HSD. There were no significant differences between F1 and codominant F2, except for diameter (P < 0.001).
Mean changes in the diameter, follicular-fluid estradiol concentration, and granulosa-cell mRNA content of LHr and 3ß-HSD associated with the two largest follicles are shown with the results of statistical analyses (Fig. 1). There was a progressive increase in mean diameter and estradiol concentration of F1 over all the follicle-diameter ranges. Conversely, the diameter of F2 increased (P < 0.04) between the 7.0- to 7.4-mm and 8.0- to 8.4-mm ranges, and estradiol increased (P < 0.04) between the 7.5- to 7.9-mm and 8.0- to 8.4-mm ranges; thereafter, diameter and estradiol of F2 remained unchanged. The magnitude of diameter difference and estradiol difference between F1 and F2 was greater (P < 0.03) for the 8.5- to 8.9-mm range than for the 8.0- to 8.4-mm range (difference between F1 and F2: 4.6 ± 2.2 vs 23.3 ± 10.3 ng/ml and 0.7 ± 0.1 vs 1.1 ± 0.2 mm, respectively). This resulted in follicle (F1 vs. F2) by diameter or follicle by estradiol interactions.
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Averaged over all follicle-diameter ranges, a significant main effect of diameter range for granulosa cell LHr mRNA resulted from a greater (P < 0.05) amount of mRNA in F1 (49.2 ± 6.5 copies/cell) than in F2 (16.7 ± 2.5 copies/cell). Within F1, the LHr mRNA values were greater (P < 0.05) for the 8.5– to 8.9-mm range than for the 7.5- to 7.9-mm range. The magnitude of the difference for LHr mRNA between F1 and F2 was greater (P < 0.03) for the 8.0- to 8.4-mm range than for the 7.0- to 7.4-mm range (difference between F1 and F2: 16.6 ± 8.3 vs -2.4 ± 5.3 copies/cell, respectively). The significant main effect of follicle-diameter range for granulosa cell 3ß-HSD mRNA was due primarily to higher (P < 0.05) amounts of mRNA in both F1 and F2 in the 9.0- to 9.4-mm range compared to the two preceding ranges.
Combined for F1 and F2, positive correlations were detected between diameter and estradiol (r = 0.44; P < 0.0001), diameter and LHr mRNA (r = 0.38; P < 0.0001), diameter and 3ß-HSD mRNA (r = 0.16; P < 0.04), estradiol and LHr mRNA (r = 0.39; P < 0.0001), estradiol and 3ß-HSD mRNA (r = 0.22; P < 0.008), and between LHr and 3ß-HSD mRNA (r = 0.22; P < 0.007).
Experiment 2
Of the total 56 observations, six follicles were identified as codominant (F2 >9.0 mm); however, only two were detected as statistical outliers based on intrafollicular concentrations of estradiol, and these two were excluded from analysis of all end points. Of the remaining 54 observations for each end point, 48-54 observations were available for analysis after removing the extreme values (1.9%-5.6%) as statistical outliers, except that all 54 observations were available for IGFBP-2.
Mean changes in diameter and follicular-fluid steroid hormone, free IGF-1, and IGFBP-2 concentrations of the two largest follicles are shown with results of the statistical analyses (Fig. 2). Actual diameters of F1 when it reached at least 7.5, 8.5, 9.5, or 10.5 mm ranged from 7.5–8.0, 8.5–8.9, 9.5–9.9, and 10.5–11.2 mm, respectively. In this regard, the identity of the follicle groups was changed to follicle-diameter ranges to reflect the actual diameters for conformity with experiment 1. Diameter of both F1 and F2 increased (P < 0.05) between the 7.5- to 8.0-mm and 9.5- to 9.9-mm ranges, and thereafter, F1 continued to increase (P < 0.05) but F2 did not (P > 0.05). Estradiol was similar between F1 and F2 at the 7.5- to 8.0-mm range and, thereafter, increased (P < 0.05) in F1 and decreased (P < 0.05) in F2. As a result of the increase in F1 for diameter and estradiol concentration, the first increase (P < 0.04) in the difference between F1 and F2 occurred at the 8.5- to 8.9-mm range for both end points (differences for the two adjacent ranges, 0.5 ± 0.2 vs. 1.4 ± 0.4 mm and 48.8 ± 51.9 vs. 338.2 ± 130.2 ng/ml, respectively).
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There were no significant changes in F1 in intrafollicular concentrations of free IGF-1 and IGFBP-2 between the 7.5- to 8.0-mm and 10.5- to 11.2-mm follicle-diameter ranges. Conversely, free IGF-1 decreased (P < 0.05) and IGFBP-2 increased (P < 0.05) in F2 during the corresponding interval. Averaged over all follicle-diameter ranges, intrafollicular concentrations of progesterone were significantly greater in F1 than in F2. There were no significant changes in intrafollicular concentrations of testosterone between follicles and among ranges.
Mean changes in the ratios of follicular-fluid steroid hormones associated with the two largest follicles are shown with the results of the statistical analyses (Fig. 3). Averaged over all follicle-diameter ranges, the ratios of estradiol:progesterone and estradiol:testosterone were significantly greater for F1, whereas the ratio of testosterone:progesterone was significantly greater for F2.
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There were no significant main effects of follicle (F1 vs. F2) and diameter range or corresponding interactions for the mean changes in follicular-fluid concentrations of total inhibin and dimeric inhibin-A (Fig. 4). A positive correlation was detected between total inhibin and dimeric inhibin-A (r = 0.64; P < 0.0007).
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Combined for F1 and F2, positive correlations were detected between diameter and estradiol (r = 0.64; P < 0.0001), estradiol and testosterone (r = 0.40; P < 0.004), and estradiol and free IGF-1 (r = 0.50; P < 0.001), whereas a negative correlation was detected between free IGF-1 and IGFBP-2 (r = -0.50; P < 0.0007).
DISCUSSION
It was expected that the results of the present experiments would provide an initial indication of the sequence of events centered around and perhaps involved in follicle diameter deviation. In previous studies (reviewed by Ginther et al. [36]), diameter deviation was related to the time of occurrence (e.g., days or hours after wave emergence). In the present experiments, the ranges in diameter for the F1 were used to represent time. The first diameter range with a significant change in the differences between F1 and F2 was used to allow comparisons among end points.
The range of means for concentrations of estradiol in the follicular fluid in the present study were comparable to earlier reports for slaughterhouse ovaries [37] and for follicular fluid collected in vivo [20]. The hypothesis that the first increase in the difference between F1 and F2 for diameter and estradiol occurred at the same time was supported; the two events occurred at the 8.5- to 8.9-mm range in both experiments. These results agree with the those of previous studies; increase in the difference between F1 and F2 in diameter and follicular-fluid estradiol began on the same day, based on an in vivo follicular-fluid collection procedure [20] and on systemic blood sampling [38]. Furthermore, based on sampling of blood from the caudal vena cava cranial to the entry of ovarian venous blood, estradiol concentrations began to increase when the largest follicle was 8.5 mm (expected time of deviation; [7]); ablation of the 8.5-mm follicle prevented the estradiol increase. Future attempts to determine whether the diameter or estradiol change precedes the other will likely require more frequent assessments.
Temporal support for a role of LH in diameter deviation is the occurrence of an increase and subsequent decrease in circulating LH concentrations, so that elevated concentrations encompass the beginning of deviation in heifers and ponies (reviewed by Ginther [5]). The role of LH on follicle development has also been approached by others by studying expression of LHr mRNA and time of acquisition of LHr by the granulosa cells. Luteinizing hormone receptor mRNA expression occurred in granulosa cells of <4-mm follicles [9] and >8-mm follicles [8] in cattle. The LHr protein was present in granulosa cells of 5-mm follicles in mares [39] and in the largest and second-largest follicles when the largest follicle was 8.5 mm in heifers [18]. Conversely, LHr mRNA expression was minimal in the largest, second-largest, and third-largest follicles 2-3 days after estrus [40]. This period corresponds to 1-2 days after ovulation and, if so, would presumably be before the expected occurrence of diameter deviation [38]. The failure to detect LHr mRNA in granulosa cells of follicles <8.0 mm, subordinate follicles [8], and follicles collected on Days 2 and 3 after estrus [40] may relate to the sensitivity of the assays used. Moreover, the first significant increase in LHr mRNA in the largest follicle in cattle was not detected until the follicle reached a mean of 10.8 mm [8], which would be well past the expected beginning of deviation. Similarly, granulosa cells of cattle expressed LHr mRNA between 2 and 4 days after wave emergence; the mRNA was expressed in follicles >9.0 mm [41]. Despite the discrepant results among previous studies, it appears that LHr mRNA and protein are present in granulosa cells before and after deviation. However, previous receptor studies have not focused on the differential (F1 vs. F2) expression of LHr mRNA in bovine granulosa cells in association with follicle deviation.
In the present study (experiment 1), the expression of granulosa cell LHr mRNA significantly increased between the 7.5- to 7.9-mm and 8.5- to 8.9-mm follicle-diameter ranges in F1 but did not increase in F2. The first increase in the differences between F1 and F2 in LHr mRNA expression in granulosa cells occurred in the 8.0- to 8.4-mm range. The hypothesis that the change in LHr mRNA would occur at the same time as the changes in diameter and follicular-fluid estradiol concentrations was not supported. A significant increase in the difference between F1 and F2 in LHr mRNA expression occurred one follicle-diameter range before the first increase in diameter and estradiol concentration. The difference in diameter ranges between the increase in LHr mRNA and the other two end points is equivalent to a diameter difference of 0.5 mm or 8 h of growth [38]. Assuming that increased LHr mRNA expression represents increased LHr in the granulosa cells, the LHr is in a temporal position to play a role in the events associated with follicle diameter deviation.
In bovine follicles, the 
5-pathway is preferred for conversion of pregnenolone to androstenedione by 17-hydroxylase and 3ß-HSD enzymes [42]. Expression of 3ß-HSD mRNA in granulosa cells has been suggested to play a role in follicle selection [43]; expression of 3ß-HSD mRNA was first detected in follicles 
8.0 mm and significantly increased as follicles grew to a mean diameter of 10.2 mm. Granulosa cells of follicles <8.0 mm or subordinate follicles did not express 3ß-HSD mRNA. Thus, the increased 3ß-HSD expression apparently occurred after the beginning of diameter deviation. In another study, no difference was shown in 3ß-HSD enzyme activity in granulosa cells between growing dominant and nongrowing dominant follicles collected on Days 5-18 of the estrous cycle [44]. Immunohistochemical localization did not detect any 3ß-HSD enzyme in granulosa cells of sheep and pigs at any stage of follicle development but found the enzyme in bovine granulosa cells of preovulatory follicles [45]. In experiment 1, there was no difference in granulosa cell 3ß-HSD mRNA expression between F1 and F2 at any diameter range; the hypothesis of a relationship between 3ß-HSD mRNA changes and changes in diameter, estradiol, and LHr mRNA was not supported. The 3ß-HSD mRNA increased significantly in both F1 and F2 between the 8.5- to 8.9-mm and 9.0- to 9.4-mm diameter ranges. This was after the beginning of the diameter deviation. The observation that granulosa cell LHr mRNA and 3ß-HSD mRNA are expressed in a similar pattern in the selected follicle [43] was not supported by our findings. In contrast to a report of not detecting correlations between estradiol concentration in follicular fluid and 3ß-HSD activity in growing dominant, nongrowing dominant, and nongrowing nondominant follicles [44], a positive correlation between estradiol and 3ß-HSD mRNA content was found in experiment 1. Further studies will be needed to clarify these apparently contradictory results and to determine whether 3ß-HSD has a role in follicle deviation.
Relationships between follicular-fluid concentration of total IGF-1 and the stage of follicle development have been studied in many species, including cattle [13, 15, 46, 47] and sheep [48]. Higher total IGF-1 concentrations were detected in the follicular-fluid of large estrogen-active follicles (>8 mm) than in the fluid of either large estrogen-inactive follicles (>8 mm) or pools of follicular fluid from medium (5–7 mm) and small (<5 mm) follicles [13]. Intrafollicular concentrations of total IGF-1 were not different between dominant and subordinate follicles at a mean 4.8 days after estrus; however, concentrations were higher compared to follicles at 3 days after estrus [15]. Conversely, no difference in total IGF-1 levels in bovine follicular fluid was found between dominant (mean, 11.9 mm), large (mean, 8.5 mm), or small follicles (<6 mm) during 4–6 days or 8–12 days after estrus [14] or between dominant and subordinate follicles at 5, 8, and 12 days after estrus [46]. Reasons for the apparent conflicting results among reports with respect to follicular-fluid concentrations of total IGF-1 in cattle are not known but may be related to different methods of selecting follicles for sampling and assay techniques.
In the present study, follicular-fluid samples were collected in vivo and assayed for free IGF-1 using a commercially available ELISA kit. The validity of the kit to detect changes in follicular fluid was supported by similar dose-response curves for standard concentrations and pooled follicular-fluid volumes. The pool of follicular fluid was collected in vivo from the two largest follicles of wave 1 in several contemporary animals, and therefore, the pool was representative of the experimental samples. Furthermore, the concentrations of free IGF-1 reported herein support the approximation of the free IGF-1 fraction as 1–5% of the total IGF-1 [49]. The lowest and highest means of free IGF-1 selected from the follicle-diameter ranges within F2 and F1 were 1.06-3.61 ng/ml, respectively. Comparatively, these concentrations were about 1.2% of the mean of the total IGF-1 concentrations ranging from 90 to 293 ng/ml reported by others [14, 15].
In experiment 2, follicular-fluid concentrations of free IGF-1 were significantly lower in F2 compared to F1 averaged over all follicle-diameter ranges. The difference between follicles reflected a progressive decrease in F2; concentrations were significantly lower in the 9.5- to 9.9-mm range compared to the 7.5- to 8.0-mm range. It has been suggested that absolute intrafollicular concentrations of total IGF-1 may remain constant and that an increase in concentration of follicular-fluid IGFBPs may be one mechanism by which the bioavailability of IGF-1 is reduced in subordinate follicles [46]. Reportedly, the bioactivity of IGF-1 is controlled by its association with at least six IGFBPs [11]. In cattle, an increase in IGFBP-2, -4, and -5 has been related to follicular atresia [13–15, 46]. In the present study, mean follicular fluid concentrations of IGFBP-2 progressively increased during the time that free IGF-1 decreased in F2. Despite the lack of significant changes of free IGF-1 and IGFBP-2 in F1, the combined results from the two follicles showed a significant negative correlation between the two factors. Although the profile of IGFBP-2 across the follicle-diameter ranges is similar to published reports [13–15], the comparison of quantitative results in the present study with qualitative results in the previous studies may be of limited value. Nonetheless, the concept that high levels of IGFBP-2, and perhaps other low molecular weight IGFBPs, limit the bioavailability of IGF-1 in subordinate follicles is further supported by our findings.
The role of IGF-1 in follicular development in cattle has been studied in vitro and in vivo. The results of in vitro studies have indicated that IGF-1 stimulates mitosis of granulosa [50] and theca cells [51, 52] and increases the synthesis of estradiol, progesterone [53], and androgen [51] from bovine granulosa and theca cells. Correspondingly, in vivo studies have indicated that the largest follicle was greater in diameter in cattle that received an intraovarian infusion of IGF-1 compared to controls (mean, 17.6 vs. 13.7 mm; [54]). In addition, higher intrafollicular concentrations of IGF-1 were detected in the two largest follicles in animals selected for twinning [55]. Thus, the present results of the intrafollicular IGF-1 and IGFBP-2 in association with follicle development substantiate the role of the IGF system in ovarian function; however, the role of the IGF system in association with follicle deviation needs further study.
In experiment 2, changes were not detected in follicular-fluid concentrations of progesterone among the follicle-diameter ranges, but concentrations were significantly higher in F1 than in F2. Reportedly, intrafollicular concentrations of progesterone do not vary between follicles during development of the first follicular wave in cattle [14, 15]. In our study, follicular fluid was collected in vivo compared to the previous studies that collected follicular fluid from excised ovaries and, therefore, the difference in collection methods may have contributed to the difference between the present and previous results. Nevertheless, the higher progesterone concentrations in F1 reflected the higher steroidogenic activity in F1 compared to F2. The estrogen:progesterone (E:P) ratio significantly increased after the beginning of deviation (8.5- to 8.9-mm follicle-diameter range), whereas the E:P ratio of F2 remained unchanged. Similar increase in the E:P ratio associated with the largest follicle of wave 1 has been reported [14, 15, 18] but not in relation to the time of deviation.
In experiment 2, concentrations of total inhibin and dimeric inhibin-A were not significantly different between F1 and F2 or among the follicle-diameter ranges. It has been reported [56] that the concentrations of bioactive inhibin were higher in dominant follicles than in atretic follicles and that the concentrations of the immunoreactive inhibin and 34-kDa dimeric inhibin did not change in follicular fluid of the largest follicle between 1 and 3 days after estrus [57]. In the former study, the follicles were defined according to E:P ratio, and therefore, the atretic follicles may have been from a previous wave. Moreover, in contrast to our results, a previous study [15] detected lower intrafollicular concentrations of inhibin-A in the dominant follicle a mean of 4.8 days after estrus compared to intrafollicular concentrations 3 days after estrus. Apparently, the mean diameter of the dominant follicle was 11.9 mm, a size corresponding to a stage of development later than that in the present study (10.8 mm). Thus, considering the lack of significant changes and the lack of any correlation with other follicular-fluid factors, the present study did not suggest a role of follicular-fluid inhibin in follicle deviation.
In conclusion, LHr mRNA was expressed in granulosa cells of the two largest follicles before the beginning of follicle deviation. The first increase in the difference in LHr mRNA between the two largest follicles occurred one diameter-range before the corresponding difference in either diameter or estradiol concentration. A significant increase in the LHr mRNA occurred in the largest follicle but not the second largest. On a temporal basis, these results indicate that differential acquisition of LH receptors by the granulosa cells of largest the follicle, indicated by increased LHr mRNA expression, occurs before diameter deviation or a change in estradiol concentrations. The levels of free IGF-1 progressively decreased while levels of IGFBP-2 progressively increased in the second-largest follicle. Decreased intrafollicular concentration of free IGF-1 and increased IGFBP-2 may deprive the second-largest follicle of the tropic effects of IGF-1, thus contributing to the cessation of growth.
ACKNOWLEDGMENTS
The authors thank the Pharmacia and Upjohn Company for the gift of Lutalyse and S.C. Jensen for technical assistance.
FOOTNOTES
1 Research supported by the University of Wisconsin, Madison, and by Equiservices publishing and the Eutherian Foundation, Cross Plains, WI. 
2 Correspondence: O.J. Ginther, Department of Animal Health and Biomedical Sciences, 1656 Linden Drive, University of Wisconsin-Madison, Madison, WI 53706. FAX: 608 262 7420;ojg{at}ahabs.wisc.edu
Accepted: September 11, 2000.
Received: July 5, 2000.
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