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Biology of Reproduction 66, 1395-1402 (2002)
© 2002 Society for the Study of Reproduction, Inc.

Regular Article

Neuroendocrine Control of Follicle-Stimulating Hormone (FSH) Secretion: II. Is Follistatin-Induced Suppression of FSH Secretion Mediated via Changes in Activin Availability and Does It Involve Changes in Gonadotropin-Releasing Hormone Secretion?1

Vasantha Padmanabhan2,,a, Deborah Battagliaa, Morton B. Browna, Fred J. Karscha, James S. Leea, Wenqin Pana, David J. Phillipsb, and Judith Van Cleeff3,,a

a Departments of Pediatrics, Physiology, and Biostatistics and the Reproductive Sciences Program, University of Michigan, Ann Arbor, Michigan 48109-0404 b Monash Institute of Reproduction and Development, Monash University, Clayton, Victoria 3168, Australia


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The objective of the present study was to determine to what extent activin participates in setting the level of FSH secretion and if this regulation includes mediation via changes in GnRH secretion. We administered follistatin, the high-affinity binding protein for activin, to five ovariectomized sheep; we reasoned that the resultant binding of follistatin to activin should lower activin bioavailability and FSH secretion. Hypophyseal portal and peripheral blood samples were collected simultaneously at 10-min intervals for 18 h to measure GnRH, LH, FSH, and both activin-free and total follistatin. Six hours into collection, each ewe received 150 µg/kg i.v. of recombinant human follistatin-288. A week later, the same ewes were subjected to a second series of blood collections of similar length (time control). The FSH levels in pituitary portal blood were approximately 8-fold higher than those in the peripheral circulation. The FSH secretory patterns changed minimally during the time-control period. In contrast, follistatin had profound suppressive effects on FSH secretion. Maximal FSH suppression after FS-288 administration occurred at 5–6 h in the pituitary portal (65% suppression) and 9–10 h in the peripheral (48% suppression) circulation. Follistatin had no effect on GnRH or LH secretory patterns. Disappearance of total follistatin (i.e., free follistatin plus activin-bound follistatin) from the circulation was slower (P < 0.05) than that of free follistatin alone, suggesting that some of the follistatin was complexed with circulating activin, thus reducing the bioavailability of activin. The slower clearance of total follistatin and the lack of follistatin effects on GnRH secretion suggest that changes in activin bioavailability dictate the level of pituitary FSH secretion and that this is a pituitary-specific effect.

follicle-stimulating hormone, follistatin, gonadotropin-releasing hormone, luteinizing hormone, neuroendocrinology, pituitary


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Inhibin and activin, which are members of the transforming growth factor ß (TGFß) superfamily [1], are produced at both the ovarian and the pituitary level [2–4]. A large body of evidence supports a role for inhibin and activin in regulating the production of FSH, a key reproductive hormone involved in ovarian follicular development [5]. Variant forms of inhibin and activin are present, with the relevant ones in terms of FSH control being inhibin-A, inhibin-B, activin-A, and activin-B. Whereas inhibin can inhibit FSH through specific inhibin-binding proteins (receptors?) [6, 7] that interfere with activin's signal transduction, inhibin at high concentrations can also antagonize activin's stimulatory action on FSH secretion by competing for the activin receptors [8, 9]. Follistatin, the third of a triumvirate of proposed nonsteroidal regulators of FSH, is a high-affinity binding protein of activin with neutralizing activities [10–12]. Thus, inhibin and follistatin appear to inhibit FSH secretion by blockade of activin action.

The majority of evidence supporting a role for these regulators in mediating FSH secretion comes from in vitro pituitary cell culture studies [1] or from determination of FSH in the peripheral circulation following either administration of recombinantly derived activin-A, inhibin-A, or follistatin or neutralization of inhibin in vivo [4, 13]. In these instances, the long circulatory half-life of FSH [14, 15] has precluded assessment regarding the true impact of these FSH-regulatory proteins on FSH secretory dynamics in vivo. Recently, this limitation has been overcome [16, 17] by the availability of a technology for the collection of hypophyseal portal blood, which was originally developed for monitoring the release of hypophysiotropic hormones [18]. Because hypophyseal portal vessels are lesioned at the surface of the pituitary, this approach provides access to the secondary capillary plexus and, for the first time, an opportunity to address the true impact of these FSH-regulatory proteins on FSH secretion at the in vivo level.

Whereas the general consensus is that these regulators modulate FSH secretion by acting directly at the pituitary level, some studies suggest that the effects of activin on FSH may also be mediated via changes in GnRH synthesis and secretion. Both GnRH-positive perikarya and fibers have been found in close association with activin ßA subunit-immunoreactive fibers as well as follistatin-positive cell bodies [19, 20]. Various in vitro studies have demonstrated the ability of activin to stimulate GnRH secretion [21–24]. Studies showing increases in hypothalamic GnRH mRNA expression, but not in testosterone secretion, following administration of activin-A to male rats [25] suggest that peripheral changes in activin may regulate FSH secretion by acting directly not only at the pituitary level but also at the hypothalamic level. Whether follistatin/activin can cross the blood-brain barrier is unknown. However, they do have the potential to affect hypothalamic secretions by acting directly at the median eminence, which is outside the blood-brain barrier, or by gaining entry into the hypothalamus via the organo vasculosum of the lamina terminalis, which is in close proximity to the preoptic area. To our knowledge, no information is available regarding the impact of these regulators in modulating GnRH secretion in vivo.

The present study addressed three questions: Does follistatin participate in the control of FSH release from the pituitary? Are the effects of follistatin mediated via changes in activin availability? Are part of the suppressive effects of follistatin mediated through a hypothalamic effect to modify GnRH secretion into hypophyseal portal blood? To answer these questions, we capitalized on the ability of follistatin to neutralize activin action. By simultaneously measuring pituitary FSH secretion and hypothalamic GnRH output, we were also able to assess if follistatin suppression of FSH is mediated, in part, via alterations in GnRH secretion.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Experimental Design

The present study addressed the impact of follistatin administration on GnRH, LH, and FSH secretion in ovariectomized Suffolk ewes maintained outdoors under natural photoperiod at the Sheep Research Facility, Ann Arbor, MI. Ewes (n = 5) were ovariectomized in early March and studied in June/July. Briefly, the animals were surgically fitted with an apparatus for collection of hypophyseal portal blood [26]. Ten days after surgery, integrated 10-min samples of hypophyseal portal (average blood volume collected from animal to animal ranged from 2.0 to 3.2 ml over 10 min) and jugular blood (3 ml over 10 min) were collected for 18 h from all ewes (study period 1). Six hours into the collection period, all ewes (body weight range, 61–69 kg; mean body weight ± SEM, 64.4 ± 1.8 kg) were injected i.v. with 150 µg/kg of recombinant human follistatin-288 (FS-288) through an indwelling jugular saline line, and sampling continued for an additional 12 h. The FS-288 (batch, B4384) used in these studies was made available through the National Hormone and Pituitary Program, National Institute of Diabetes and Digestive and Kidney Disease (NIDDK), Bethesda, MD. One week later, hypophyseal portal and peripheral samples were collected from the same animals to determine if hormonal secretory patterns varied across the 18-h sampling period (study period 2). The average volume of hypophyseal portal blood collected from animal to animal ranged from 1.2 to 4.2 ml. The second sampling required a second lesioning of the pituitary for reinitiating hypophyseal portal blood flow [26]. As such, comparisons of hormonal patterns between the two sampling periods are valid comparisons, but absolute hormone levels within the two time periods are not comparable. Blood volume limitations, possible risk of infections occurring at the lesioned site between sampling periods, scarcity of FS-288, and lack of information regarding the long-lasting effects of FS-288 prevented us from risking the ideal cross-over design. To assess the effects of FS-288 in the event that a second sampling was not possible, the first sampling period included both pre- and posttreatment periods.

Hypophyseal portal samples were collected in tubes containing 0.5 ml of 3 x 10-3 M bacitracin (Sigma Chemical Co., St. Louis, MO) in phosphate-buffered saline. At the end of collection, all ewes were killed with a barbiturate overdose (Beuthanasia; Schering-Plough Animal Health Corp., Kenilworth, NJ), and the site and extent of pituitary vasculature lesioning were determined. All experimental procedures were approved by the University of Michigan Committee on the Use and Care of Animals.

Radioimmunoassays

Hypophyseal portal GnRH, LH, and FSH as well as jugular LH and FSH were assayed in duplicate using well-validated radioimmunoassays [27–29]. For the measurement of GnRH, 750 µl of portal plasma were extracted in methanol, reconstituted in 125 µl of assay buffer, and assayed in duplicate 50-µl aliquots. Assay sensitivity (2 x SD of buffer control), 50% displacement point, and the intraassay coefficient of variation (CV) as determined by the median variance ratio of assay replicates of the GnRH assay averaged (± SEM) 0.18 ± 0.02 pg, 3.8 ± 0.53 pg, and 0.08 ± 0.01, respectively (n = 6 assays). The LH assay used an ovine serum-based standard, B1371, calibrated against the reference standard, NIH-LH-S12. For FSH assays, NIDDK-ovine FSH-1 was used as the reference standard. All jugular or portal samples from a given animal were measured in a single assay. Hypophyseal portal plasma samples were measured in a volume of 1 or 2 µl for LH and of 10 or 15 µl for FSH determinations. Peripheral samples were assayed at 25 µl and at 40 or 50 µl for LH and FSH, respectively. The FSH assay sensitivity and 50% displacement point of assays averaged 0.05 ± 0.01 and 0.38 ± 0.02 ng, respectively. The LH assay sensitivity and 50% displacement point of assays averaged 0.14 ± 0.01 and 0.66 ± 0.01 ng, respectively. Intraassay CVs at 80% and 20% displacement points averaged 8.58 ± 0.49% and 4.29 ± 0.24%, respectively, for LH and 10.78 ± 0.67% and 5.30 ± 0.33%, respectively, for FSH. The interassay CVs based on three quality-control pools averaged 7.90 ± 0.72% and 14.32 ± 0.33% for LH and FSH, respectively. The median variance ratio for LH and FSH assays averaged 0.08 ± 0.01 and 0.04 ± 0.004, respectively.

Differences in the temporal patterns of total follistatin (i.e., activin-bound and free follistatin) and of free follistatin can provide an indirect assessment of activin bioavailability in circulation. Circulating patterns of administered FS-288 were tracked with the aid of two validated follistatin assays, one that measures total follistatin levels and the other only free follistatin (i.e., does not detect activin-bound follistatin) [30, 31]. It should be recognized that this is an indirect approach to assessing activin availability. Assays that are sensitive enough to measure the various free forms of activin in circulation to obtain a direct estimate of activin availability are not available. Both follistatin assays utilized in this study are largely specific for human and minimally detect sheep follistatin; hence, they are ideal for tracking the administered FS-288. Total follistatin levels were measured in integrated 20-min hypophyseal and peripheral samples (by pooling equal volumes of plasma from two consecutive 10-min samples) at 90-min intervals before FS-288 administration (0–6 h) and in all the 20-min integrated samples (pooled as described above) starting from the time of FS-288 administration. For total follistatin measurements, both hypophyseal and peripheral plasma were assayed in duplicate 25-µl volumes. When necessary, samples were reassayed at dilutions ranging from 1:250 to 1:5000 (v/v), depending on the amount of total FS-288 contained in the sample. In addition, integrated 2-h samples (equal volume pooled from all 10-min samples within each 2-h period) from both hypophyseal portal and peripheral samples were measured in a free follistatin assay. Limited availability of samples and resources precluded more detailed assessment of free follistatin levels. The assay sensitivity and the intra- and interassay CVs averaged 0.031 ± 0.005 ng/ml, 5.4 ± 0.8%, and 6.2%, respectively, for the total follistatin assay (n = 15 assays). For the free follistatin measurements, hypophyseal and peripheral samples were measured in duplicate 100-µl aliquots. When samples were out-of-range high, they were rediluted with assay buffer and reassayed. Sensitivity of this assay averaged 1 ng/ml (n = 2 assays). The intraassay CV averaged less than 10% for concentrations greater than 3 ng/ml and less than 20% for concentrations between 1 and 3 ng/ml (mean CV for this series, 5.2%). The interassay CV based on three quality-control pools averaged 8%. In both assays, FS-288 was used as the standard.

Statistical Analysis

To facilitate direct comparison with FSH concentrations in peripheral samples (ng/ml), all FSH (as well as LH) measurements in hypophyseal portal plasma are reported as concentrations, as opposed to GnRH, which is reported as a collection rate. For determining the effect of follistatin on GnRH, LH, and FSH secretion, mean hormone concentrations in each hour were calculated by averaging the 10-min time points within that hour. Values were log-transformed before analysis by regression. All analyses included a factor for each ewe to allow for differences in hormone levels between ewes. The first 6-h period during both collection periods (corresponding to the time before FS-288 administration in study period 1) was assumed to be constant. For each hormone, linear regressions were fitted separately to the data in the FS-288 treatment and control periods. Each regression model was fitted to be constant for the first 6 h and then linear after that. The coefficient of the linear term was tested to see if it differed significantly from zero. In addition, tests were conducted to determine if the regression line was constant during the first 6 h (i.e., whether a linear term was necessary) and if the line was linear after that (i.e., whether a quadratic term was necessary). To distinguish time-related changes from true follistatin effects, regression slopes from the FS-288-treatment period (study period 1) and the time-control period (study period 2) were compared by a paired Student t-test.

The time course of follistatin effects on GnRH and on hypophyseal portal and peripheral LH and FSH levels was determined by subjecting percentage changes in hourly hormone levels from the baseline period to a repeated-measures ANOVA followed by a paired t-test. Pulse analyses on GnRH and hypophyseal portal LH values were performed using the Kushler pulse fit algorithm [32]. Because hormone series with 36 points (first 6 h) would have very little information for fitting, the entire hormone series (all 18 h) was fitted by pulse fit, and the pulse frequencies and amplitudes between the two time periods (0–6 and 6–18 h) were compared for each condition (study period 1 and study period 2). Because of difficulties associated with identification of FSH pulses during FS-288 suppression of FSH, the FSH values were not subjected to pulse analysis. The disappearance rate of both total and free FS-288 was estimated using a one-phase exponential model; preliminary analyses indicated that a two-phase exponential model was not appropriate. The median R2 value in fitting the disappearance curves was 0.996 (range, 0.943–0.999; n = 24 determinations).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Effect of FS-288 on GnRH and LH Secretion

The effects of FS-288 on GnRH in hypophyseal portal blood and on LH in hypophyseal portal and peripheral blood are summarized in Figures 1 (representative ewe) and 2 (group). Treatment with FS-288 had no effect on GnRH or LH secretion. Tests for linearity revealed a slight decline in GnRH/LH release as collection progressed during both study periods (FS-288-treatment and time-control periods). Slopes of decline were similar between time periods and averaged -0.015 and -0.004 for study periods 1 and 2, respectively, for GnRH; -0.047 and -0.043, respectively, for hypophyseal portal LH; and -0.021 and -0.024, respectively, for peripheral LH, confirming the lack of effect of FS-288 treatment on GnRH and LH (Fig. 2). Repeated measures ANOVA also revealed no effects of FS-288 on GnRH, hypophyseal portal LH, and peripheral LH levels.



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  		FIG. 1. Patterns of hypophyseal portal GnRH and hypophyseal portal and peripheral LH secretion in a representative ovariectomized sheep. Shaded area represents the period following follistatin injection. Note that patterns of GnRH and LH during the control period (right panels) were derived following a second lesion of the same sheep 1 wk later. Whereas comparison of absolute levels of LH achieved in peripheral circulation during the follistatin-treatment period (left) and the time-control period (right) is valid, the same does not hold true for the hypophyseal portal LH and GnRH measurements (see Materials and Methods for explanation)



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  		FIG. 2. Percentage change in hypophyseal portal GnRH and hypophyseal portal and peripheral LH secretion from the first 6-h baseline. Solid bars represent study period 1, when follistatin was administered at Time 0 (Hour 6 of collection). For comparison, changes in GnRH and LH achieved in hypophyseal portal circulation during study period 2 (time control) are expressed in a similar manner

During study period 1, GnRH pulse frequency averaged 0.8 ± 0.1 and 0.7 ± 0.1 pulses/h, respectively, before and after FS-288 administration and was not significantly different. The GnRH pulse frequency for the corresponding time points during study period 2 are not provided due to missing values in two animals (poor assay duplication and insufficient blood volume to repeat extraction and reassay). The LH pulse frequency in hypophyseal portal circulation before and after FS-288 administration averaged 1.07 ± 0.08 and 1.07 ± 0.05 pulses/h, respectively, during study period 1. In previous studies, we have shown a one-to-one concordance between hypophyseal portal and peripheral LH pulses [16, 17]. Whereas the pulse detection algorithm identified more LH than GnRH pulses, except for one or two LH pulses all identified LH pulses showed a corresponding, albeit small, increase in GnRH secretion that was not picked up by the pulse detection program. Lack of detection of GnRH pulses corresponding to the LH pulses in several instances may be a function of dilution of the GnRH signal between adjacent samples in concert with sample integration. The LH pulse frequency (i.e., a bioassay for GnRH pulse frequency) for corresponding time periods in study period 2 were 1.07 ± 0.8 and 1.07 ± 0.05 pulses/h, respectively. The GnRH pulse amplitude before and after FS-288 administration averaged 5.97 ± 1.94 and 5.26 ± 1.29 pg/min, respectively. Overall, FS-288 administration had no effect on hypophyseal portal GnRH/LH pulse frequency and pulse amplitude.

Effects of FS-288 on FSH Secretion

Effects of FS-288 on hypophyseal portal and peripheral FSH are summarized in Figures 3 (representative ewe) and 4 (group). Tests of slopes in the first 6 h were nonsignificant for hypophyseal portal and peripheral FSH during both study periods. Hypophyseal portal and peripheral FSH declined with time from 6 to 18 h (0–12 h in Fig. 4) during both collection periods. Comparison of the slopes of decline between the FS-288-treatment and time-control period (-0.25 and -0.016, respectively, for hypophyseal portal FSH and -0.06 and -0.012, respectively, for peripheral FSH during study periods 1 and 2) revealed that the slopes of decline of hypophyseal portal and peripheral FSH during the FS-288-treatment period were far greater than those during the time-control period (Z = -13.53 and -11.82 for hypophyseal portal and peripheral FSH, respectively; P < 0.0001). The quadratic term was also highly significant (P < 0.0001) for hypophyseal portal, but not for peripheral, FSH during the FS-288-treatment period, reflecting the rapid decline of FSH and subsequent establishment of a stable, but lower, baseline relative to the starting baseline (Figs. 3 and 4). The R2 estimates (i.e., a measure of the fit of the model) computed after adjusting for the effect of ewe were very high for hypophyseal portal and peripheral FSH (84.42% and 81.18%, respectively) during the FS-288-treatment period, thus substantiating the time course of follistatin suppression of FSH in both hypophyseal portal and peripheral FSH.



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  		FIG. 3. Patterns of hypophyseal portal and peripheral FSH secretion in a representative ovariectomized sheep. Shaded area represents the period following follistatin injection. Whereas comparison of the absolute levels of FSH achieved in peripheral circulation during the follistatin-treatment period (left) and the time-control period (right) is valid, the same does not hold true for the hypophyseal portal measurements (see Materials and Methods for explanation)



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  		FIG. 4. Percentage change in hypophyseal portal and peripheral FSH secretion from the first 6-h baseline. Solid bars represent study period 1, when follistatin was administered at Time 0 (Hour 6 of collection). For comparison, changes in GnRH and LH achieved in the hypophyseal portal circulation during study period 2 (time control) are expressed in a similar manner (open bars). Asterisks indicate significance (P < 0.05) compared to the pretreatment period. Downward arrows indicate time of maximal suppression; upward arrows indicate the time of recovery

Repeated-measures analysis followed by post hoc comparison confirmed that the time course of FSH suppression following FS-288 treatment differed in hypophyseal portal and peripheral blood. Specifically, suppressive effects of FS-288 on FSH were evident sooner in the hypophyseal portal than in the peripheral circulation. Additionally, FSH was significantly suppressed by 2–3 h in the hypophyseal portal circulation; suppression lagged by an hour in the peripheral circulation (Fig. 4). Maximal FSH suppression after FS-288 administration occurred at 5–6 h in the pituitary portal (65% suppression) and at 9–10 h in the peripheral (48% suppression) circulation. Levels of FSH started to recover from follistatin suppression 10–11 h after FS-288 administration (P < 0.05) in the hypophyseal portal, but not in the peripheral, circulation.

Time Course of Disappearance of Administered FS-288

To assess if follistatin binds circulating activin and reduces bioavailability of activin, the disappearance rates of FS-288 in the hypophyseal portal and peripheral circulation, as measured by a free follistatin assay (which does not recognize follistatin bound to activin), were compared with patterns measured by a total follistatin assay (which recognizes both free follistatin and follistatin bound to activin). The results revealed that the disappearance kinetics of total as well as free follistatin did not differ between the hypophyseal portal and peripheral circulation (Fig. 5). Decay curves of total follistatin assessed after averaging hypophyseal portal and peripheral measurements were slower (P < 0.05) than that of the free follistatin, suggesting that more of the administered FS-288 remained in the circulation complexed with activin. The median half-life was 108.3 min (range, 70.5–131.7 min) for total follistatin and 67.4 min (range, 46.7–106.7 min) for free follistatin.



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  		FIG. 5. Changes in hypophyseal (P) and peripheral (J) patterns of follistatin as measured in the total and free follistatin assays. Total follistatin levels (left panel) were measured at 90-min intervals (by pooling two 10-min samples) until the time of follistatin administration (study period 1) and at 20-min intervals (pools of two 10-min samples) in all samples from the time of follistatin administration (0–12 h). During the time-control period (study period 2), they were assessed only at 90-min intervals throughout the collection period. Restriction of sample volume limited assessment of free follistatin measurement in 2-h samples (pooling equal volume of plasma across a 2-h period) in both treatment and time-control series. The 2-h averages of total follistatin (computed by averaging values from the 20-min samples) are co-plotted along with free follistatin (right panel, dotted lines) to facilitate comparison between the slopes of the disappearance of free and total follistatin


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Results from the present study demonstrate that 1) FS-288 is a potent suppressor of FSH secretion in ovariectomized ewes; 2) suppression of FSH secretion is evident significantly earlier at the secretory (i.e., hypophyseal portal) level than in the peripheral circulation, which reflects the sum of secretion and clearance; 3) effects of follistatin are mediated through a pituitary-specific mechanism and do not involve changes in GnRH secretion; and 4) suppressive effects of follistatin are consistent with what is known regarding the effects of follistatin to bind activin and neutralize its availability and with the effect being mediated, at least in part, at the peripheral level. The first and last of these conclusions confirm previous observations, whereas the others extend these observations by providing new information regarding the nature of follistatin action, as discussed below.

Follistatin Suppression of FSH Secretion

In this study, the maximal degree of FSH suppression achieved after FS-288 administration was 48% at the peripheral level and 65% at the secretory level. In the study by Tilbrook et al. [33], administration of 25 µg/kg of FS-288 (one-sixth the dosage used in the present study) to wethers suppressed circulating levels of FSH by 20.6%. Other studies in rats [34, 35] have shown that administration of 80 µg of purified porcine follistatin (mix of all molecular variants of follistatin) per rat suppressed peripheral FSH by approximately 50%. We used much less FS-288 than the amount in these rat studies, but the greater effect in the present study may be explained by the greater biopotency of FS-288 compared with the longer follistatin variants contained in follicular fluid [36, 37]. In addition to the 288-amino acid form (i.e., FS-288), follistatin has been shown to exist as an alternate spliced variant of 315 amino acids (FS-315) [38] and a proteolytically cleaved, truncated form of FS-315 (~300 amino acids) [36]. A more relevant study in rats showed that administration of 30 µg of FS-288 over 6 h (on a body wt basis, these levels are comparable to those used in our study) resulted in a 45% suppression of circulating FSH and a 70% suppression of steady-state FSHß mRNA [39]. Our findings do not appear to result from toxic effects of administered follistatin on gonadotropes, because 1) no suppressive effects of follistatin were observed on LH secretion in spite of it being produced by the same gonadotrope [40], 2) FSH secretion started to recover from this suppression toward the end of the collection period, and 3) peripheral FSH levels observed a week later (time-control series) were of comparable magnitude to those of the pretreatment period during the first collection (follistatin-treatment series).

It is not possible to relate the circulating level of FS-288 that was achieved following administration of recombinant FS-288 to that which actually circulates in vivo. It is believed that the predominant circulating form of follistatin is not the FS-288 form but, rather, the alternatively spliced variant of follistatin, FS-315 [41]. However, recent assays that are specific for FS-288 [30] do detect measurable quantities of FS-288 in serum [42, 43]. Total follistatin is reported to be on the order of 8 ng/ml during the human menstrual cycle [44], which is several-fold lower than the levels achieved in the present and all the earlier studies [33–35, 39], but they are equivalent to circulating levels seen in women at 36 wk of gestation (3.6 µg/ml) [42], when circulating FSH levels are extremely low.

Time Lag Relationships Between Follistatin Administration and FSH Suppression

A time lag in peripheral FSH suppression has been reported following administration of FS-288 in wethers [33] and inhibin-A, which also works by antagonizing activin action, in monkeys [45]. In the present study, maximal suppression of circulating FSH was evident 9–10 h after FS-288 administration; this closely parallels the time lag of 12–15 h reported by Tilbrook et al. [33] in wethers. The long circulating half-life of FSH likely masks the actual time relationships of this suppression. This is evident from the estimate regarding the timing of maximal FSH suppression at the secretory level. Maximal suppression of FSH at the secretory level occurred at 5–6 h in the present study, which is several hours earlier than at the peripheral level. The delay in FSH response at the secretory level to FS-288 administration also supports the contention that the effects of FS-288 on FSH secretion are mediated via changes in FSH production as opposed to release, because effects on the release process would be expected to occur sooner. That a substantial portion of FSH is secreted in a constitutive manner and is not dependent on immediate stimulus secretion coupling is well-documented [46–48]. This component of FSH release appears to be dictated by the availability of translatable FSHß mRNA and consequent FSH production [13, 46–48]. Several studies have shown that FS-288 inhibits both FSH biosynthesis and consequent release [1–3].

Hypothalamic Versus Pituitary Site of Action by Follistatin in Mediating FSH Suppression

A lack of effect of FS-288 administration on GnRH or LH secretory dynamics argues against any hypothalamic contribution to the FSH suppression that followed FS-288 administration. Whether follistatin crosses the blood-brain barrier is unknown, but follistatin has the ability to modulate GnRH secretion by acting at the level of the median eminence or by gaining entry through the organum vasculosum of the lamina terminalis. Studies documenting increased GnRH transcription following peripheral administration of activin [25] provide support for this premise. Our studies strongly support the premise that the effects of FS-288 on FSH secretion are mediated in a pituitary-specific manner. Several pituitary cell types, including the gonadotropes, have been shown to express these regulators [49–52]. Furthermore, the majority of studies that have measured follistatin or activin-A in circulation have questioned an endocrine role for circulating follistatin and activin in the suppression of FSH secretion [31, 44, 53].

Follistatin Suppression of FSH Secretion and Activin Bioavailability

Follistatin is a high-affinity binding protein for activin, with Kd estimates ranging between 50 and 900 pM [10, 54, 55]. This represents nearly irreversible kinetics and is in the vicinity of activin binding to its receptor (100–400 pM) [54], thus enabling follistatin to be a potent neutralizer of activin [11, 12]. This has been predicted before on the basis of kinetic studies, but to our knowledge, the present study is the first report that likely documents the lowering of activin availability in vivo. The slower rate of disappearance of total (free plus activin-bound) FS-288 in relation to the activin-free follistatin from circulation suggests that a portion of administered FS-288 binds to circulating activin and, thus, for all practical purposes is "invisible" in the free follistatin assay, which does not recognize activin-bound FS-288. These findings suggest that the suppressive effects of follistatin are mediated, at least in part, in an endocrine manner. The recent finding that activin-FS-288 complexes associate with cell surface proteoglycans, are endocytosed, and then are broken down by lysosomal enzymes [56] raises the possibility that this degradative pathway may be involved in the disappearance of total FS-288 from the circulation.

Are the binding and disappearance kinetics of FS-288 in this study sufficient to say that the suppressive effects of FS-288 are mediated via activin neutralization? To address this question, we need to weigh the high-affinity, nearly irreversible binding of follistatin to activin against evidence that shows follistatin can bind other molecules, such as {alpha}2-macroglobulin [57], and to other members of the TGFß superfamily, such as bone morphogenetic protein (BMP)-4 and BMP-7 [58]. Interestingly, a very recent report suggests that BMP in microgram concentrations can stimulate FSH from ovine and rat pituitary cells in culture [59], and that the mouse pituitary expresses BMP-2, -4, and -7 mRNA [59]. However, the direct binding of follistatin to radiolabeled BMP is 20-fold lower than that of activin [60]. Additionally, in competition assays with labeled activin and unlabeled BMP-7, the affinity for BMP-7 appears to be approximately 500-fold lower, and activin appears to be capable of displacing BMP from follistatin complexes (A.L. Schneyer, personal communication). This then implies that, when both activin and BMP-7 are present together, activin would be the much-preferred ligand, and that approximately 500-fold greater concentrations of BMP would be required to overcome this preference. To our knowledge, no estimates of BMP levels in circulation are available to address if BMPs have a functional role in regulating FSH. Similarly, both activin and follistatin are known to bind to {alpha}2-macroglobulin [57, 61], but the relatively low affinity also suggests that this would represent a minor influence on the functional regulation of FSH secretion.

In summary, our study advances what is known regarding follistatin suppression of FSH in four fronts. First, the study provides clear documentation in vivo regarding the impact of follistatin suppression of FSH at the level of secretion (i.e., release as distinct from the balance between secretion and clearance). Second, the delay in observing the suppressive effects of follistatin at the systemic as compared to the secretory level highlights the difficulty encountered in delineating secretory changes in FSH from peripheral measurements due to the prolonged clearance of FSH. Third, the study provides evidence that all effects of administered follistatin are mediated in a pituitary-specific manner and do not involve changes in GnRH secretion. Fourth, demonstration of the slower disappearance of total (free plus activin-bound) than of free follistatin provides evidence for the presence of activin-bound follistatin and, thus, indirect evidence for the presence of free activin in the circulation, thereby supporting a role for peripheral activin in setting the level of FSH secretion in vivo.


   ACKNOWLEDGMENTS
 
We are grateful to Mr. Douglas D. Doop and Mr. Gary McCalla for providing quality care and maintenance of the sheep used in this study; Ms. Kristin McFadden and Mr. James Dearworth for their assistance with the LH and FSH assays; Dr. Patrick Sluss for the free follistatin service assays; Dr. Gordon Niswender and Dr. Leo E. Reichert for supplying LH assay reagents; Prof. Nigel Groome for supplying the total follistatin assay reagents; Dr. A.F. Parlow and the National Hormone and Pituitary Program for the generous gift of follistatin as well as the FSH standard and antisera; and Dr. Louis DePaolo, from the Reproductive Sciences Branch, NIH, for making possible the procurement of such large amounts of FS-288 for conducting studies of this magnitude in a large animal model.


   FOOTNOTES
 
First decision: 11 September 2001.

1 Supported by National Institutes of Health grant HD 34731 and National Health and Research Council of Australia Program grant 973218. Back

2 Correspondence: Vasantha Padmanabhan, Reproductive Sciences Program, 300 N. Ingalls Bldg., Rm. 1109 SW, Ann Arbor MI 48109-0404. FAX: 734 936 8620; vasantha{at}umich.edu Back

3 Current address: Department of Animal Sciences, University of Illinois, Urbana, IL 61801 Back

Accepted: December 3, 2001.

Received: August 9, 2001.


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 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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