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Biology of Reproduction 59, 561-570 (1998)
©Copyright 1998 Society for the Study of Reproduction, Inc.

Immunohistochemical Distribution of Follistatin in Dominant and Subordinate Follicles and the Corpus Luteum of Cattle1

Jaswant Singh and a, , and Gregg P. Adams2,a

a Department of Veterinary Anatomy, University of Saskatchewan, Saskatoon, Saskatchewan, Canada 37N 5B3


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The study was done to quantitatively characterize the distribution of follistatin in ovarian follicles and corpora lutea at specific stages of development. Transrectal ultrasonography was used to monitor the growth of individually identified follicles from 2 days before ovulation until the day of ovariectomy on Day 3 of wave 1 (n = 8), Day 6 of wave 1 (n = 6), Day 1 of wave 2 (n = 6), or after onset of proestrus, at least 17 days postovulation (n = 7). Days of ovariectomy represent the growing, early-static, late-static, and regressing phases of the dominant follicle of wave 1, respectively. Subordinate (n = 24), preselection (n = 15), and preovulatory (n = 6) follicles and corpora lutea (n = 31) were also analyzed. Follistatin was localized using immunohistochemical labeling of paraffin sections, and relative amounts were quantitated using densitometric analysis. Follistatin was distributed in the perinuclear cytoplasm of granulosa and luteal cells but not in theca cells. Dominant follicles contained more (p < 0.05) follistatin than corresponding subordinate follicles. The amount of follistatin was maximal during the mid-growing phase of the dominant follicle and decreased thereafter (p < 0.05). Among the corpora lutea, the maximal amount was detected at mid-diestrus (Day 11 postovulation). Less than half of luteal cells displayed the stain for follistatin; positively stained luteal cells were located in close proximity to blood capillaries. Follistatin was not detectable in the corpus luteum during metestrus (Day 3 postovulation) or proestrus (Day >= 17 postovulation). In summary, the degree of immunohistochemical expression of follistatin was phase specific for both follicles and corpora lutea. The most intense staining in follicles was associated with the period of functional dominance and in corpora lutea was seen during the phase of maximal development. Significant phase-related differences in follistatin expression provide rationale for the hypothesis that follistatin is involved in the final stages of follicle and luteal gland development in cattle.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Follistatin was first identified in bovine and porcine follicular fluid by its ability to suppress FSH secretion from pituitary cell cultures [1, 2]. Follistatin is a glycosylated single-chain protein that occurs in at least 6 forms ranging from 31- to 39-kDa molecular mass [3]; smaller forms represent a carboxyl terminal truncation of the larger precursor [4]. Although follistatin is known to be expressed in a number of tissues, a major site of production is the granulosa cells of ovarian follicles [5, 6]. In cattle, follistatin mRNA expression increases as follicle size increases; the strongest signal was observed in preovulatory follicles [7]. Based on data obtained in vitro, it has been postulated that follistatin modulates granulosa cell function in an autocrine manner and that its actions favor luteinization or atresia [8]. Follistatin is a specific binding protein for activin [9], and its actions are believed to be mediated mainly through neutralizing the effects of activin. Follistatin has been measured in peripheral blood of humans [10] and sheep [11] with the supposition that it might act as a circulating hormone; however, concentrations of follistatin varied little during the estrous cycle of the ewe, and there was little or no reduction after ovariectomy [11]. Although the regulation, production, and actions of follistatin have been studied extensively in vitro, data on its role in ovarian function in vivo are insufficient and confusing.

The growth and atresia of individual antral bovine follicles have been monitored using sequential ultrasonography, and it has been established that growth of follicles >= 4 mm takes place in 2 or 3 waves during the estrous cycle [12–15]. Emergence of successive follicular waves during the estrous cycle was associated with increases in circulating concentrations of FSH that preceded wave emergence by 1 day [16]. During an anovulatory wave, the development of each follicle has been subdivided into growing (increasing diameter), static (no change in diameter), and regressing (decreasing diameter) phases [14]. These ultrasonically classified phases have been shown to be closely correlated with the follicle's ability to produce steroid and protein hormones (estrogen:progesterone and estradiol:androstenedione ratios, {alpha}-inhibin and dimeric inhibin concentrations) indicative of follicle health or atresia [17–19]. The dynamics of follistatin expression in bovine follicles and corpora lutea relative to follicular wave status and the stage of the estrous cycle have apparently not been documented.

The present study was designed to characterize the relative amount and distribution of follistatin in dominant and subordinate follicles during the growing, static, and regressing phases; to compare the features of follicles of the anovulatory versus ovulatory waves; and to characterize the changes in follistatin distribution during the immature, mature, and postmature phases of the corpus luteum in cattle.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animal Grouping and Ultrasonography

The study was conducted on 16- to 18-mo-old sexually mature, nulliparous, crossbred beef heifers (primarily Hereford). Heifers (n = 27) were maintained in a single corral and fed to gain approximately 1.3 kg weight per day. Ovarian follicle development was monitored daily by transrectal ultrasonography using a 7.5-MHz linear-array transducer (Aloka Co. Ltd., Tokyo, Japan). Ultrasound examinations commenced at least 2 days before the ovulation preceding the estrous cycle under study and continued until the day of ovariectomy in order to monitor the development of follicles >= 4 mm in diameter. Topographic location and diameter of individually identified follicles and corpora lutea were recorded on a daily basis [13]. The day of wave emergence (Day 0) was determined retrospectively and defined as the day on which the dominant follicle of a wave was first detected at a diameter of 4–5 mm [14, 15, 20]. The dominant follicle was defined as the largest follicle of a wave, and subordinate follicles were defined as those that appeared to originate from the same pool of follicles [13, 14]. As the heifers ovulated, they were randomly designated to be ovariectomized on Day 3 of wave 1 (D3W1, n = 8), Day 6 of wave 1 (D6W1, n = 6), Day 1 of wave 2 (D1W2; n = 1 on Day 9, 2 on Day 10, 2 on Day 11, 1 on Day 13 after ovulation), or in the immediate preovulatory period >= 17 days after ovulation (D>=17, n = 2 on Day 17, 3 on Day 18, 1 on Day 19, 1 on Day 21 after ovulation). Heifers in group D>=17 were ovariectomized 1 day after detection of proestrus; proestrus was defined as the day when any 3 of 4 estrus-like characteristics (i.e., high uterine tone, edematous echotexture, intrauterine fluid collection, mucous discharge) were detected [21]. Days of ovariectomy were selected on the basis of previous studies [14–16] to represent the growing (D3W1), early-static (D6W1), late-static (D1W2), and regressing (D>=17) phases of the dominant follicle of wave 1 (Fig. 1). The design of the experiment also allowed collection of subordinate follicles of wave 1 and the ovulatory wave; preselection follicles of wave 2; preovulatory dominant follicles; and corpora lutea during metestrus (D3W1, Day 3 after ovulation), early diestrus (D6W1; Day 6 after ovulation), mid-diestrus (D1W2; mean Day 11 after ovulation), and proestrus (D>=17, mean Day 18 after ovulation; Fig. 1). Regressed corpora lutea of the previous estrous cycle were also collected on D3W1 (n = 3) and D6W1 (n = 2).



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  		FIG. 1. Diameter profiles (mean ± SEM) of the corpus luteum and dominant, largest subordinate, and second-largest subordinate follicles of wave 1 (anovulatory) and wave 2 (ovulatory); the dotted lines indicate the mean days of ovariectomy on Day 3 of wave 1 (D3W1), Day 6 of wave 1 (D6W1), Day 1 of wave 2 (D1W2), and during proestrus >= 17 days postovulation (D>=17). The numbers in parentheses are the number of corpora lutea and follicles of each type collected for each period. Breaks in the profiles indicate that data were included only from those heifers that were ovariectomized at the end of that segment.

Ovariectomies and Tissue Processing

Heifers were ovariectomized via colpotomy [22] in the standing position, under caudal epidural anesthesia using 2% w:v lidocaine HCl with 0.001% w:v epinephrine. Clenbuterol (6 mg/10 kg BW i.v., Ventipulmin; Boehringer Ingelheim Ltd., ON, Canada) was given 10 min before colpotomy to induce relaxation of the uterine and broad ligament musculature [23]. After manual compression of the mesovarium with a lidocaine-soaked gauze, both ovaries were removed through a single incision in the dorsolateral aspect of the vaginal fornix using a chain écraseur. Heifers were treated postoperatively with procaine penicillin G for 4 days. Immediately upon removal, the ovarian artery or its branches were cannulated, flushed with 15–20 ml PBS (0.1 M phosphate buffer, 0.9% sodium chloride, pH 7.2–7.4) to remove blood, and then perfused with 30 ml of Karnovsky's fixative [24] containing 4% paraformaldehyde and 0.1% glutaraldehyde in 0.1 M phosphate buffer (pH 7.4) at a flow rate of 2 ml/min. The fixation procedure was completed within 45 min of ovariectomy. The ovaries were immersed in Karnovsky's fixative for 3 h, after which each individually identified dominant follicle, subordinate follicle, and corpus luteum was sectioned through its largest diameter. Slices of follicles and corpora lutea were fixed for another 3 h by immersion in aqueous Bouin's fixative for light microscopy. The double-fixation procedure permitted use of the same tissues for light microscopy, scanning and transmission electron microscopy, and immunohistochemistry. Initial perfusion fixation helped to maintain the structural integrity of individual follicles, thus preventing their collapse and artifactual loss of adluminal granulosa cells due to coagulation of follicular fluid. In addition, perfusion fixation resulted in retention of a thin layer of coagulated follicular fluid along the antral wall throughout tissue processing, thus permitting assessment of sloughed cells. Tissues were processed as described above to obtain 5-mm paraffin sections on poly-L-lysine-coated glass slides. In addition to follistatin immunohistochemical staining, adjacent sections were stained with hematoxylin and eosin to confirm the architecture of tissues under study.

On the basis of daily ultrasonographic records, follicles were identified as the dominant, largest subordinate, and second-largest subordinate on the day of ovariectomy. Regarding the dominant and largest subordinate follicles, only those that were individually identified were collected and analyzed. If on the day of ovariectomy the second-largest subordinate follicle could not be distinguished from other subordinates (i.e., similar diameter; D6W1, D>=17), one was chosen arbitrarily and included in the second-largest subordinate category. Hence, the second-largest subordinate category included follicles that were individually identified as well as those that could not be individually identified. On D1W2, the three largest follicles of wave 2 were collected to represent preselection follicles because dominance was not yet apparent.

Anti-Follistatin Antibody

Polyclonal rabbit antiserum (Rb-32, bleed date 02–23–90) raised against native porcine follistatin [10] was generously provided by Dr. Nicholas Ling (Whittier Institute for Diabetes and Endocrinology, La Jolla, CA) and was used as the primary antibody for immunohistochemical localization of follistatin. The antiserum has been characterized and does not cross-react with activin A or inhibin A [10]. The binding ability of antiserum was tested using Western blotting, and it detected recombinant human follistatin (rhFS-288; Code #B3904 National Hormone and Pituitary Program, NIDDK, Rockville, MD) and recombinant porcine follistatin (gift from C.R. Christensen, University of Saskatchewan, Saskatoon, SK, Canada). Lyophilized primary antiserum was stored at -20°C until use and diluted 1:1000 with 4% normal goat serum prepared in PBS (0.1 M phosphate buffer, 0.9% NaCl, pH 7.2–7.4). Excess of rhFS-288 was added to half of the diluted primary antiserum (10 mg rhFS-288/ml of 1:1000 diluted antiserum) and allowed to stand overnight at 4°C to adsorb the primary antibody. The adsorbed and nonadsorbed antisera were then centrifuged for 15 min at 3000 x g to remove any particulate material. Follistatin-adsorbed antiserum was used to measure the degree of nonspecific staining for quantitative immunohistochemistry.

Immunohistochemical Staining for Follistatin

A pair of glass slides containing adjacent sections of tissue were incubated overnight with 1:1000 dilution of primary antiserum and follistatin-adsorbed primary antiserum (one slide each) to localize the follistatin and nonspecific reaction, respectively. The avidin-biotin peroxidase technique was used as described previously [25, 26] using a Vectastain ABC kit (Vector Laboratories, Burlingame, CA). Microscope slides of all follicles under study were stained as a single batch using a Code-On-Histomatic slide-staining station (Fisher Scientific Co., Edmonton, AB, Canada) to minimize the variation in reaction due to difference in staining batches. Similarly, all corpora lutea were stained as a single batch. Briefly, endogenous peroxidase was blocked with 2% v:v hydrogen peroxide in methanol for 10 min, and proteolytic enzyme digestion was done with 0.05% w:v protease XIV (catalogue no. P5147; Sigma Chemical Company, St. Louis, MO) for 30 min [26]. Slides were treated with 4% normal goat serum (Gibco BRL, Life Technologies, Gaithersburg, MD) containing 0.05% v:v Tween 20 (Aldrich Chemical Co., Milwaukee, WI) for another 30 min to minimize nonspecific binding of the secondary antibody. Slides were incubated overnight at 4°C with primary antiserum or follistatin-adsorbed primary antiserum. The next day, slides were incubated for 30 min with 1:400 dilution of biotinylated goat anti-rabbit IgG antibody (catalogue no. BA-1000; Vector Laboratories) prepared in 4% normal goat serum. The rest of the staining was carried out according to instructions provided in the Vectastain ABC kit. Slides were then dehydrated and coverslips were applied. For purposes of illustration, coverslips were removed from representative slides after densitometric quantitation, sections were hydrated, counterstained with 1:1 diluted Harris hematoxylin for 20 sec, and dehydrated; the coverslips were then reapplied.

Gray-Scale Densitometric Analysis

Gray-scale densitometric analysis of follicles and corpora lutea was performed using a Carl Zeiss (Thornwood, NY) Ultraphot microscope, a high-resolution black and white microscope video camera (Hamamatsu CCD XC-77; Hamamatsu Photonics KK, Hamamatsu City, Japan) and camera control unit (C2400–60; Hamamatsu Photonics KK), and a computer-based, commercially available image analysis system (Image 1/AT; Universal Imaging Corporation, West Chester, PA). Microscope illumination, condenser position, diaphragm control, objective lens, projective lens, and video camera contrast enhancement and offset controls were standardized and remained unchanged throughout the analysis period. An objective lens of x40, a projective lens of x1.25, and a tube factor of x6.12 were used to obtain a final magnification of x306 on the video monitor. The position of the glass slide was fixed on the microscope stage, and each gray-scale image, consisting of 512 x 480 picture elements (pixels), was obtained for computer analysis by averaging 32 successive video frames from the video camera according to the manufacturer's instructions. Illumination was set for optimal uniform brightness throughout the microscopic field of view and was unchanged throughout the analysis period. In addition, the background subtraction technique was used to account for any visually undetectable nonuniformity of light in different areas of the microscopic field. Moreover, the position of the measuring box was kept constant in relation to the microscopic field of view so that the same part of the field was measured for all images. Gray-scale values (mean and standard deviation) were measured on a scale ranging from 0 (pure black) to 255 (pure white) in a 140 x 140 pixel-area box representing 0.25 mm2 of the original section for the corpus luteum, and in a 34 x 167 pixel-area box representing 0.06 mm2 of the original area for the follicle wall (Fig. 2). Mean gray-scale values were used to calculate the absorptive index, and standard deviation values were used to calculate heterogeneity. Values were recorded at 4 different locations within the same section that exhibited the positive reaction (corpus luteum or granulosa layer of follicles). Values were also taken from another 4 locations from the same section that exhibited no specific reaction (ovarian stroma for corpus luteum, theca interna for follicles). Areas for measurement were selected based on the uniform random sampling technique [27].



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  		FIG. 2. The technique for densitometric measurement of the follistatin reaction in the granulosa layer of ovarian follicles. Positive reaction was localized in the granulosa layer (a); the theca interna was used as a nonreactive reference area (b). Mean gray-scale values were measured in a box (values below the box on each figure; black = 0, white = 255) encompassing 34 x 167 picture elements (0.06-mm2 area) from the granulosa layer (a, c) and theca interna (b, d) of sections stained with the primary antiserum (a, b) and those stained with follistatin-adsorbed primary antiserum (c, d). Total absorptive index was calculated by using the values of reactive area versus nonreactive area (a vs. b), and nonspecific absorptive index (calculated from c vs. d) was subtracted to obtain the specific absorptive index for the granulosa layer.

The mean gray-scale value through a clear glass slide (with coverslip) was considered 100% transmittance, and the mean gray-scale value obtained by blocking the light path of the microscope was considered 0% transmittance. Mean gray-scale values obtained for follicles and corpora lutea were then converted to percentage transmittance. The values for percentage transmittance were averaged for the 4 samples and transformed to absorbance (negative value of log of percentage transmittance), and absorptive index was calculated [28] by obtaining the ratio of absorbance of reactive to nonreactive areas. This method of reporting the results provides considerable advantage over other methods because the absorptive index value is independent of section thickness [29]. The specific absorptive index for each of the follicles and corpora lutea was calculated by subtracting the absorptive index of the section treated with follistatin-adsorbed primary antiserum (nonspecific and unknown cross-reactions of the antiserum) from the absorptive index of sections treated with nonadsorbed primary antiserum. Similarly, the gray-scale standard deviation values for the positive reaction were converted to a scale of 0–100 and were defined as heterogeneity. Intra- and inter-day coefficients of variation for gray-scale densitometry were 6.8% and 8.5%, respectively. Used in this fashion, the absorptive index is an indicator of the amount (concentration) of follistatin in the tissue but is independent of the pattern of distribution of follistatin within the sample. Heterogeneity indicates the relative difference in distribution at the tissue level, that is, how evenly the reaction product is distributed within the tissue.

Statistical Analysis

Data for the corpus luteum were examined by ANOVA to detect a day effect. Data for follicles were examined by 2-factor ANOVA to determine the effect of follicle type, day of wave, and follicle-by-day interaction. If the analysis resulted in a probability value of <= 0.05, multiple comparisons were made using the method of least significant difference. Initially, data for absorptive index and heterogeneity were analyzed for differences among dominant, largest subordinate, and second-largest subordinate follicles (Fig. 1); however, as no statistical differences were detected between largest and second-largest subordinate follicles, the data were reanalyzed after subordinate follicles were combined into a single group, and results are presented as such.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
The diameter profiles of follicles and corpora lutea are illustrated in Figure 1, and the method for densitometric quantitative analysis is illustrated in Figure 2. The specificity of the immunohistochemical staining was confirmed by adsorbing primary antiserum with excess of follistatin and by omitting the secondary antibody treatment during the staining procedure (Fig. 3). A positive reaction (brown staining) detected after primary antiserum treatment of follicles and corpora lutea was undetectable after the antiserum was adsorbed with follistatin or the secondary antibody was omitted from the staining procedure (Fig. 3). Data and statistical results are summarized separately for follicles (see Figs. 4, 6, and 7) and corpora lutea (see Figs. 5 and 8).



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  		FIG. 3. Micrographs showing the specificity of follistatin immunohistochemical staining (brown color) of a growing dominant follicle (a–c) and a mid-diestrous corpus luteum (d–f). When primary antiserum was used, intense staining was detected in the granulosa layer of the follicle (a) and in the luteal cells of the corpus luteum (d). The labeling was not detected when the same tissues were stained with follistatin-adsorbed antiserum (b, e) or when the secondary antibody was omitted from the staining procedure (c, f). Bar = 50 mm.



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  		FIG. 4. Intensity (absorptive index, a) and distribution (heterogeneity, b) of follistatin staining (mean ± SEM) measured by densitometric analysis of dominant and subordinate follicles. The data have been arranged to illustrate the growing (D3W1), early-static (D6W1), late-static (D1W2), and regressing (D>=17) phases of the dominant follicle of wave 1; preselection follicles of wave 2 one day after wave emergence (D1W2); and follicles of the ovulatory (Ov.) wave during proestrus (D>=17). Numbers in parentheses below the x-axis indicate the number of follicles analyzed for each follicle type. Bars with no common letters indicate statistical difference (p < 0.05).



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  		FIG. 6. Immunohistochemical distribution of follistatin (brown color) in the wall of dominant (a, c, e, g) and subordinate (b, d, f) follicles of wave 1 collected on Day 3 of wave 1 (a, b), Day 6 of wave 1 (c, d), Day 1 of wave 2 (e, f), and Day >= 17 after ovulation (g). Note that follistatin staining is confined to the granulosa layer and that a progressive decrease in staining intensity is evident from the growing (a) to the regressing (g) phase. A similar decrease occurred in subordinate follicles. The antral region of the granulosa layer of an early-static follicle has a stronger reaction than the basal region (c). Bar = 50 mm.



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  		FIG. 7. Immunohistochemical distribution of follistatin (brown color) in the wall of the preselection follicles of wave 2 (a, largest; b, second largest) and in follicles of the ovulatory wave (c, preovulatory dominant; d, subordinate). The granulosa layers of preselection follicles (a, b) have similar staining intensity, while the preovulatory follicle (c) displays an intense reaction in contrast to its subordinate counterpart (d). Bar = 50 mm.



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  		FIG. 5. Intensity (absorptive index, a) and distribution (heterogeneity, b) of follistatin staining (mean ± SEM) measured by densitometric analysis of bovine corpora lutea during metestrus (3 days after ovulation), early diestrus (6 days after ovulation), mid-diestrus (between 9 to 13 days after ovulation), and proestrus (between 17 to 21 days after ovulation) and of regressed corpora lutea of a previous estrous cycle collected between Days 3 and 6 after ovulation. Numbers in parentheses below the x-axis indicate the number of corpora lutea analyzed for each phase. Bars with no common letters indicate statistical difference (p < 0.05).



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  		FIG. 8. Immunohistochemical distribution of follistatin (brown color) in the corpus luteum during metestrus (a), early diestrus (b), mid-diestrus (c), and proestrus (d). Luteal cells stained positively for follistatin only during diestrus (b, c). Note the stronger reaction in the part of cytoplasm facing a blood capillary (inset in c). Bar = 50 mm.

Follicles

As evidenced by diameter profiles (Fig. 1), all dominant follicles of wave 1 were in the growing phase on D3W1, the early-static phase on D6W1, the late-static phase on D1W2, and the regressing phase on D>=17. Subordinate follicles of wave 1 were in the early-static phase on D3W1 and the regressing phase on D6W1. All preselection follicles of wave 2 on D1W2, and the preovulatory follicles on D>=17, were in the growing phase. Subordinate follicles of the ovulatory wave were regressing on D>=17.

A discrete brown-colored granular reaction for follistatin was detected only in the granulosa layer; the theca interna layer was devoid of staining regardless of follicle type or phase. A day effect (p < 0.001) and a follicle effect (p < 0.001) were detected for absorptive index. A decrease in the staining intensity for the dominant follicle over time (Fig. 6) was reflected in a gradual decrease in the absorptive index (Fig. 4). Subordinate follicles displayed a trend similar to that of dominant follicles, but the absorptive index was higher (p < 0.05) for dominant than for subordinate follicles. Heterogeneity values did not differ among days, but they were higher (p < 0.012) in dominant than in subordinate follicles. The follistatin reaction was stronger in the granulosa cells near the antral surface in 4 of 13 growing or early-static phase dominant follicles (Fig. 6) but otherwise appeared to be distributed evenly throughout the granulosa layer. Staining appeared more intense in the perinuclear area of the granulosa cell cytoplasm.

No differences were detected (p > 0.5) in the absorptive index (Fig. 4) and staining pattern (Fig. 7) among the preselection follicles (D1W2). After selection, the absorptive index was markedly higher (p < 0.05) in the granulosa layer of dominant follicles than in the subordinate follicles (D3W1 and the ovulatory wave). The absorptive index (Fig. 4) and staining pattern (Fig. 7) of dominant follicles of the ovulatory wave were similar to those of the growing dominant follicles of wave 1 (D3W1). In sequence of follicle maturation (both dominant and subordinate), follistatin expression was maximal during the mid-growing phase and decreased progressively through the static and regressing phases.

Corpus Luteum

Follistatin was detected (Fig. 8) in the luteal cells during early diestrus (6 days after ovulation) and mid-diestrus (between 9 to 13 days after ovulation), but not during metestrus (3 days after ovulation) or proestrus (between 17 to 21 days after ovulation). Not all luteal cells displayed staining for follistatin. Cells exhibiting the positive reaction were present in close proximity to blood capillaries (Fig. 8c, inset). The granular reaction was strongest in the perinuclear part of the luteal cell cytoplasm, and in most cases the reaction was localized more on one side of the nucleus (Fig. 8c, inset). A higher absorptive index (p < 0.05) was detected in mid-diestrus than in all the other stages. The heterogeneity increased progressively (p < 0.05) from metestrus to mid-diestrus and decreased thereafter (p < 0.05). Regressed corpora lutea from the previous estrous cycle (collected 3–6 days after ovulation) did not display any follistatin reaction.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Phase-specific changes in follistatin expression in granulosa and luteal cells documented in the present study are suggestive of a functional role for follistatin in follicular and luteal gland development in cattle. Although follistatin expression was numerically greater in the largest follicle as early as 1 day after wave emergence (i.e., D1W2), the difference among follicles did not reach significance until 3 days after wave emergence (i.e., D3W1), when dominance was clearly established. Dominant follicles maintained higher amounts of follistatin than their subordinate counterparts, regardless of phase, and follistatin was expressed maximally in the mid-growing and preovulatory phases. Loss of functional dominance of the wave 1 follicle, indicated by emergence of a new wave [30–33], was marked by a dramatic decrease in immunostained follistatin in the granulosa cells. After ovulation, follistatin was not visually detectable in the luteinizing cells of the follicle wall until early diestrus; and by assessment of absorptive index, maximal levels of follistatin were not expressed in luteal cells until mid-diestrus.

Follistatin immunostaining was localized only in the granulosa layer of all follicles in the cattle studied, a finding consistent with the results of earlier studies in the rat and human [5, 34–37]. Follistatin was not detected in the theca interna regardless of follicle type or phase, consistent with an earlier report that theca interna tissues obtained from bovine follicles did not produce follistatin in vitro [6].

Although follicle wave-related changes in follistatin production have apparently not been reported previously, mRNA for follistatin in rat granulosa cells was higher in growing secondary and tertiary follicles than in other follicles [38]. The mRNA signal for follistatin apparently decreased in mature preovulatory follicles of the rat, but immunostained follistatin was most intense in graafian stage follicles [34]. In bovine granulosa cells in vitro, the strongest mRNA signal for follistatin was found in preovulatory follicles [7]. Ovaries of cows given superstimulatory treatment also contained high levels of follistatin mRNA [39]. Results of the present in vivo study, however, revealed that follistatin expression was equally high in the mid-growing phase and preovulatory phase of dominant follicle development.

Relative to follicle wave dynamics, there appears to be a temporal relationship between the expression of follistatin (present study), estradiol production [19], and morphological features [40] of the dominant and subordinate follicles of cattle. Immediately after divergence in the diameter profiles of dominant and subordinate follicles (i.e., selection of dominant follicle), differential expression of both follistatin (present study) and estradiol production [19] was observed. The granulosa layer of only morphologically healthy follicles produced follistatin, and maxima in follistatin expression in granulosa cells were coincident with maxima in intrafollicular estradiol concentrations. Follistatin and estradiol may simply be increasing coincidentally as a result of accelerated cell function, or perhaps follistatin is involved in the process of selection of the dominant follicle by enhancing differentiation of the granulosa and theca cells. In contrast to the in vivo temporal relationship with estradiol, follistatin has been shown to enhance progesterone production in vitro from bovine granulosa cells obtained from 5- to 10-mm follicles, but not from the preovulatory follicles [41]. Follistatin also increased LH-induced progesterone production, but not androstenedione production, from in vitro-cultured bovine theca cells obtained from 5- to 10-mm follicles [42]. In the rat and the human, activin has an inhibitory effect on androgen synthesis by theca cells in vitro [43, 44]; hence, follistatin was expected to have a stimulatory role. Thus, granulosa-derived activin and follistatin may have selective paracrine modulatory roles on thecal androgen synthesis and, in turn, affect estradiol production by the granulosa cells. It is not known yet whether higher estradiol production by the granulosa cells caused them to express more follistatin or vice versa. Although existence of a cause-and-effect relationship is speculative at this stage, temporal relationships discovered during this study provide a basis for the postulate that greater expression of follistatin promotes follicle dominance and lesser expression will lead to continued growth of the subordinates.

Our results demonstrated stage-specific changes in the amount of follistatin in the granulosa cells of dominant follicles. In contrast, the concentration of follistatin in human follicular fluid did not vary among healthy, atretic, and preovulatory follicles in women undergoing in vitro fertilization, nor were differences found in abnormal follicles obtained from patients with polycystic ovarian syndrome [45]. Differences in results may be attributed to species differences or to the possibility that the amount of follistatin detected in the follicular fluid may not truly reflect the amount of follistatin produced by granulosa cells. In this regard, follistatin has a high affinity for sulfated proteoglycans, particularly heparin sulfate located in follicular fluid and on the surface of granulosa cells [46]. In addition, the localization of follistatin on the surface of cells is not necessarily correlated with the production of follistatin by the same cell type. However, in the present study, we observed specific perinuclear localization of follistatin with aggregates close to the nucleus of both granulosa and luteal cells, presumably in the Golgi region of the cells.

Similar to follicles, the bovine corpus luteum also displayed stage-specific changes in the follistatin levels. When the corpus luteum was developing but not yet fully functional, immature luteal cells did not display any immunostaining for follistatin; the amount of staining increased with maturity of the corpus luteum (maximum at mid-diestrus) and decreased again during regression. Our results are consistent with earlier reports in which expression of follistatin mRNA or protein was detected in the corpus luteum of the cow, ewe, and human [7, 37, 47, 48]. In the rat, the primary gonadotropin surge appeared to suppress the expression of follistatin [36] and follistatin mRNA, and the protein declined sometime prior to, or immediately after, ovulation [34, 38, 49]. Similar changes may also occur in bovine granulosa cells in which follistatin expression ceases during the late preovulatory stage. We found that not all luteal cells displayed follistatin immunostaining, and the cells that were positively stained were in close proximity to blood capillaries. In the present study, no attempt was made to distinguish between the large and small luteal cells, but results provide rationale for the hypothesis that luteal cells that are derived from the granulosa layer synthesize follistatin. It is not yet clear what specific role is played by the follistatin in the luteal cells.

In summary, follistatin was detected in the granulosa layer of healthy bovine follicles and in luteal cells of diestrous corpora lutea. The degree of immunohistochemical expression of follistatin was phase specific for both follicles and corpora lutea. The most intense staining was associated with the period of functional dominance (growing, early-static, and preovulatory phases) in dominant follicles and during the phase of maximal luteal gland development in diestrus. Results suggest that follistatin may be involved in the control of the final stages of follicle and luteal gland development in cattle, and that production or secretion involves an endocrine route.


   ACKNOWLEDGMENTS
 
We thank Carla Becker, Kelly Lightfoot, and Joseph Tom for assistance in animal handling; Bill Kerr and his staff at the Goodale Research Farm for animal maintenance; Brian Chelack, Jim Gibbons, Ken Gummeson, and Debby Haines for assistance with tissue preparation; and Ian Shirley for assistance with densitometric image analysis.


   FOOTNOTES
 
1 Supported by grants from the Natural Sciences and Engineering Research Council of Canada. J.S. was supported by a Canadian Commonwealth Scholarship. Back

2 Correspondence: Gregg P. Adams, Department of Veterinary Anatomy, University of Saskatchewan, 52-Campus Drive, Saskatoon, SK, Canada S7N 5B4. FAX: (306) 966–7405; adams{at}usask.ca Back

Accepted: April 20, 1998.

Received: April 15, 1997.


   REFERENCES
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 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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