Follistatin: Essential Role for the N-terminal Domain in Activin Binding and Neutralization*

Yisrael SidisDagger , Alan L. SchneyerDagger , Patrick M. SlussDagger , Leslie N. Johnson§, and Henry T. Keutmann§

From the Dagger  Reproductive Endocrine Unit and National Center for Infertility Research and the § Endocrine Unit, Massachusetts General Hospital and Harvard Medical School, Boston, Massachusetts 02114

Received for publication, January 25, 2001, and in revised form, March 14, 2001


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Follistatin is recognized to be an important regulator of cellular differentiation and secretion through its potent ability to bind and bioneutralize activin with which it is colocalized in many tissue systems. The 288-residue follistatin molecule is comprised of a 63-residue N-terminal segment followed by three repeating 10-cysteine "follistatin domains" also represented in several extracellular matrix proteins. We have used chemical modifications and mutational analyses to define structural requirements for follistatin bioactivity that previously have not been investigated systematically. Mutant follistatins were stably expressed from Chinese hamster ovary cell cultures and assayed for activin binding in a solid-phase competition assay. Biological activities were determined by inhibition of activin-mediated transcriptional activity and by suppression of follicle-stimulating hormone secretion by cultured anterior pituitary cells. Deletion of the entire N-terminal domain, disruption of N-terminal disulfides, and deletion of the first two residues each reduced activin binding to <5 % of expressed wild-type follistatin and abolished the ability of the respective mutants to suppress activin-mediated responses in both bioassay systems. Hence, the three follistatin domains inherently lack the ability to bind or neutralize activin. Activin binding was impaired after oxidation of at least one tryptophan, at position 4, in FS-288. Mutation of Trp to Ala or Asp at either positions 4 or 36 eliminated activin binding and bioactivity. Mutation of a third hydrophobic residue, Phe-52, reduced binding to 20%, whereas substitutions for the individual Lys and Arg residues in the N-terminal region were tolerated. These results establish that hydrophobic residues within the N-terminal domain constitute essential activin-binding determinants in the follistatin molecule. The correlation among the effects of mutation on activin binding, activin transcriptional responses, and follicle-stimulating hormone secretion substantiates the concept that, at least in the pituitary, the biological activity of follistatin is attributable to its ability to bind and bioneutralize activin.


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Follistatin (FS)1 has gained recognition as an important mediator of cell secretion, development, and differentiation in a number of tissue and organ systems. Follistatin was first isolated from ovarian follicular fluid as a protein factor capable of suppressing FSH secretion by pituitary cells in culture in a manner similar to inhibin (reviewed in Refs. 1-4). Cloning and sequencing (5) showed it to be a protein of 288 amino acids (FS-288), unrelated to inhibin, with a C-terminal-extended form (FS-315) derived from alternative splicing. No "receptor" for follistatin has been found, but its mode of action in the pituitary became clear with the demonstration (6) that the protein binds the activin A homodimer with high affinity, approaching irreversibility because of its slow dissociation rate (7). Multiple lines of evidence have now shown that, rather than "presenting" activin to its receptor as in the case of certain circulating binding proteins, follistatin sequesters activin to prevent stimulation of FSH secretion (8, 9). More recently, follistatin has been reported to accelerate endocytosis and degradation of activin (10). Insights into follistatin's importance have paralleled the steadily unfolding evidence for multiple roles played by activin and its relatives in the transforming growth factor-beta family of regulatory factors (2, 3).

Localization appears to be facilitated through interaction with cell surface proteoglycans through at least one heparin binding site (11). Hence, earlier emphasis on follistatin as a circulating factor has been largely superseded by evidence for its role as a local cellular regulator with structural similarities to a number of extracellular matrix proteins involved in cellular regulation and development. Although most abundant in pituitary, ovary, testis, and kidney, follistatin is widely distributed among all tissues in which activin is also present (2). In fact, the lethal effects found in follistatin-null animals are attributable to skeletal and cutaneous abnormalities (12).

The domain structure of follistatin is characteristic of a large number of proteins derived originally through a process of exon shuffling. Following a signal peptide and a 63-residue N-terminal segment, the remainder of the molecule (residues 64-288) consists of three successive 73-77 residue domains, precisely defined by exon-intron junctions, which are clearly related by alignment of their ten cysteine residues (Fig. 1). These repeats were likened initially to the epidermal growth factor-like sequences found in many proteins, as well as to the Kazal or ovomucoid family of protease inhibitors. However, the cysteines in these sequences align only partially, and in the case of the ovomucoids, intron phasing of these repeats do not match those found in follistatin (13). With the appearance of similar ten-cysteine sequences in osteonectin (SPARC/BM40), agrin and an increasing number of other extracellular matrix proteins, it has become clear that this repeating motif represents a distinct "follistatin-like" domain (13). Each follistatin domain forms an autonomous folding unit, as confirmed by the crystal structure of the single follistatin domain from SPARC/BM40 (14) localizing all disulfide linkages exclusively to intradomain cysteines.

Follistatin domains have been proposed or shown to interact with growth factors and other ligands in several extracellular or transmembrane proteins (13-16), as well as in a recently described activin-binding follistatin-related gene product (FLRG) (17). However, the structural requirements for activin binding by follistatin itself have not been investigated systematically.

The (1-63) N-terminal domain differs markedly from the follistatin domains in its length, amino acid sequence, and the alignment of its six cysteine residues. Its functional importance has been suggested by our own results (18) showing direct binding of activin by two synthetic peptides representing discontinuous sequences from this region, together with an earlier mutagenesis experiment (19) in which insertion of two residues at the N terminus abolished activin binding. The chemical modifications and mutational analyses of FS-288 described in this report establish the essential role of an intact N-terminal domain in activin binding and in the transcriptional and biological effects of follistatin-activin interaction. A striking requirement for hydrophobic residues within this domain suggests a mechanism for activin neutralization through competition with essential hydrophobic residues (20) in the type II activin receptor binding site.

    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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Reagents-- Pure recombinant human follistatin-288 was obtained courtesy of the National Hormone and Pituitary Project, NIDDK, National Institutes of Health, Bethesda, MD. Partially purified follistatin for coating of plates in the binding assays was prepared by affinity chromatography of expressed FS-288 on a solid support containing polyclonal anti-FS antibody 7FS30 (21). Recombinant human activin A for iodination was purchased from R&D Systems, Minneapolis, MN. Activin A for treating cells was prepared by transfection of human embryonic kidney-293 cells with an expression vector containing the human inhibin beta A-subunit cDNA as described by Delbaere et al. (22).

Preparation of Mutant Follistatins-- The follistatin-288 coding sequence was removed from pHTF302R (a gift of Dr. S. Shimasaki, School of Medicine, University of California, San Diego) and subcloned into the mammalian expression vector pcDNA3.1/myc-His (Invitrogen, Carlsbad CA). The resulting construct (pFS288mycHis) was then used as a template for site-directed mutagenesis using the QuikChange kit (Stratagene, La Jolla, CA) following the manufacturer's recommendation. To make the N-terminal deletion construct (Delta NTD), the follistatin signal peptide sequence (exon 1) was fused to the first FS domain (exon 3) by two polymerase chain reaction amplification steps. In the first step, two partially overlapping FS fragments were generated using the fused sequence oligonucleotide CCCCAACTGCATCCCCTGTAAAAAGACTTGTCGGGATGTTTTCTGTCC as a forward primer with a pcDNA3.1/bGH reverse primer in one reaction and a T7 primer with the complementary oligonucleotide as a reverse primer in a separate concurrent reaction. The two overlapping products were fused and amplified using the T7 and pcDNA3.1/bGH reverse primers in the second polymerase chain reaction step. Following restriction digestion, the final mutated polymerase chain reaction product was purified and cloned back into pcDNA3.1/myc-His. Mutant sequences were verified by bidirectional sequencing at the DNA sequencing core facility of Massachusetts General Hospital.

Expression of Recombinant Follistatins-- The pFS288mycHis vectors bearing mutant or wild-type follistatins were transfected into CHO cells using polybrene (Specialty Media, Phillipsburg, NJ) and stably secreting cells were selected using geneticin. Secretion was monitored by immunoassay (below), and screened for activin binding by solid-phase assay of conditioned medium. Follistatins were isolated from medium by binding to nickel-Sepharose affinity columns (Qiagen, Valencia, CA) via the C-terminal poly(His) tag. Following stepwise elution with imidazole, products (typically eluting between 50 and 150 mM imidazole at pH 6.8) were concentrated and exchanged into activin binding assay buffer by filter centrifugation (Centriprep-10 tubes; Amicon, Bedford, MA). Conditioned medium from nontransfected CHO cells was processed similarly for use as a control preparation in all assays.

Quantitation of Secreted Follistatins-- Follistatin concentrations in medium and affinity eluates were established by two independent immunological assays: (a) a two-site solid-phase immunochemiluminescent assay (SPICA) using a monoclonal detection antibody (7FS-30) specific to an epitope (residues 168-178; Fig. 1A) within FS domain II, as previously described (21, 23) and (b) a solution-phase assay directed toward the C-terminal Myc tag. The synthetic peptide (YGGGGEQKLISEEDLN), incorporating the Myc epitope linked by a poly(Gly) spacer to an N-terminal tyrosine for 125I labeling, was used as radioligand and reference standard. Sample (0.3-100 nM) and radioligand were incubated in phosphate-buffered saline, 0.1% bovine serum albumin buffer under equilibrium conditions for 20 h at 20 °C with a rabbit polyclonal anti-Myc antibody (Upstate Biotechnology, Lake Placid, NY) at a final concentration of 1:2400 in a total assay volume of 500 µl. Tracer-bound antibody was precipitated for counting by addition of 100 µl of a 1:12 dilution of ovine anti-rabbit gamma -globulin prepared in the Reproductive Endocrine Unit at MGH. Content of Myc-tagged follistatin was computed from the Myc-peptide standard curve and compared with the concentrations based on the SPICA assay (above).

Activin Binding Assay-- Binding of expressed follistatins to labeled activin was determined by competition assay as previously described (7). Mutant or wild-type follistatins were incubated with 125I-labeled activin in binding buffer (10 mM phosphate-buffered saline, 0.1% gelatin, 0.05% Tween; 200 µl) for 2 h at 20 °C and then added to 96-well plates (Immulon-2; Dynatech Laboratories, Chantilly, VA) coated with 25 ng of affinity-purified FS288. After incubation at 20o for 90 min, wells were washed and counted in a gamma counter. Each mutant preparation was assayed in at least three independent experiments. Relative potencies were calculated by comparison of half-maximal inhibiton of labeled activin binding to the solid-phase follistatin by mutant and wild-type follistatins, respectively.

Cyanogen Bromide Cleavage and Peroxide Oxidation-- Methionine residues were cleaved by incubating a 50 µg-aliquot of pure FS-288 with 130 mM cyanogen bromide in 70% trifluoroacetic acid/H2O (18 h, 20 °C) followed by lyophilization for binding assay. Cleavage was confirmed by Edman amino acid sequence analysis of modified FS aliquots using the Applied Biosystems 477A gas/liquid-phase microsequencer. Mild oxidation of methionine and tryptophan was performed by incubation of pure FS-288 in a 1:50 dilution of 30% H2O2 for 45 min at 37 °C followed by lyophilization and reconstitution in binding assay buffer.

Assay for Transcriptional Response to Activin-- HEK-293 cells, maintained in RPMI medium supplemented with 10% fetal calf serum, were plated in 24-well trays at 105 cells per well. When 60-70% confluent, cells were cotransfected (Effectene; Qiagen) with 100 ng of ARE-GFP-Lux (22), 80 ng of pFAST-1 expression vector (a gift of Dr. Malcolm Whitman, Harvard Medical School), and 20 ng of pRL-TK (Promega, Madison WI) for normalizing responses based on Renilla activity. The construction of the ARE-GFP-Lux and specificity of the ARE-FAST-1 reporter system for activin has been previously described (22). 16 h post-transfection, cells were treated with fresh medium containing 5 ng/ml (0.15 nM) activin, alone or preincubated (60 min) with 50 ng/ml (1.5 nM) of various FS preparations for an additional 24 h in triplicate. Cell extracts were assayed for luciferase activity using the Dual-Luciferase Reporter Assay system from Promega. Experiments were performed at least twice, and the mean ± S.E. of triplicate wells from a representative experiment is reported.

Bioassay for Pituitary FSH Secretion-- Assay for suppression by follistatin of basal FSH secretion in cultured rat anterior pituitary cells was based on the method of Scott et al. (24). The anterior pituitary glands of adult male Sprague Dawley rats (Pel Freez Biologicals, Rogers AK) were mechanically and enzymatically dispersed with 0.4% trypsin and 0.25% DNase and plated at 2.5 × 105 cells/0.5 ml well in 48-well trays in alpha -minimum essential medium (alpha MEM) containing 21 mM NaHCO3, 10% heat-inactivated fetal bovine serum, and 10% penicillin/streptomycin solution, pH 7.4. Following incubation at 37 °C in 95% air, 5% CO2 for 72 h, the monolayers were washed with phosphate-buffered saline and reincubated in 0.5 ml of fresh medium containing the various follistatin and control preparations at the specified concentrations. After 72 h, the conditioned medium was assayed for rat FSH using reagents and protocols provided by Dr. A. F. Parlow through the National Hormone and Pituitary Program, NIDDK, National Institutes of Health.

    RESULTS
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INTRODUCTION
MATERIALS AND METHODS
RESULTS
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Characterization of Secreted Follistatins-- Concentrations in conditioned medium of the various mutant follistatins expressed from CHO cells were typically 150-300 ng/ml based on the SPICA assay for free follistatin (23). After partial purification by metal affinity chromatography and exchange into assay buffer, concentrations ranged from 1-5 µg/ml. The principal epitope in the SPICA assay is a well defined sequence (residues 168-178; Fig. 1A) within the second follistatin domain (18). Although this region was not directly involved in the mutations employed here, we confirmed that the mutations did not disrupt quantitation by using a second assay directed toward the Myc epitope provided at the C terminus of each preparation. The concentrations obtained by the two methods were in agreement for all N-domain mutants and deletion products. This also implies that any loss of binding activity after mutation cannot be accounted for by a conformational change within the domain II epitope. By competition assay for activin binding, the expressed C-terminal Myc-poly(His) wild-type sequence inhibited labeled activin binding with a dose-response profile that coincided with the purified NIH FS-288 preparation. The wild-type expression product was thus used as reference preparation for all comparisons with mutant follistatins.


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Fig. 1.   Primary sequences of follistatins and homologs. A, amino acid sequence of human follistatin-288. The three follistatin domains (FSD) are aligned at their cysteine residues. The heparin binding sequence in FSD-1 is single-underlined and the recognition site for antibody 7FS30 in FSD-2 is double-underlined. B, comparison of N-terminal domain sequences from follistatin and its homologs. Mammalian follistatins are numbered as the mature protein, aligned with the corresponding segments of homologous proteins as deduced from their respective cDNA sequences. The fibrillin sequence is representative from among several repeats in the full-length fibrillin-2 molecule and its homologs. Arrows denote functionally important hydrophobic residues described in this report. GenBankTM/EBI accession numbers for species and homologs are: human (P19883), porcine (AAA31037), bovine (2101261A), ovine (P31514), equine (O62650), rat (NP  036693), murine (CAA58291), chicken (AAB35335), Xenopus(A53502), zebrafish (AAD09175), human FLRG (NP_005851), Drosophila gene product CG12956 (AAF58157), human fibrillin-2 (NP_001990).

Deletion of the N-terminal Domain-- Fig. 1A summarizes the domain structure of human FS-288. Among vertebrate species, the molecule is highly conserved throughout, including the N-terminal (1-63) domain as shown in Fig. 1B; cysteines and several other residues also are common to related gene products in human (FLRG) and Drosophila (GG1596). Initial mutational analyses of FS-288 were designed to evaluate tolerance for deletion or structural disruption of the N-terminal domain. A molecule comprising exclusively the three follistatin domains (residues 64-288), devoid of the N-terminal domain, was expressed from mammalian cells in concentrations comparable with full-length FS-288. By competition assay, the affinity-processed product was found to bind activin with a potency <5% of expressed wild-type (Fig. 2). This response was comparable with equivalent volumes of control medium processed from nontransfected cells.


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Fig. 2.   Binding inhibition curves for FS-288 expression products with deleted (Delta NTD), truncated (Delta G1N2), and disulfide-disrupted (CC(26,27)AA) N-terminal domain in a competition assay for labeled activin binding (7). Mutations diminished binding potency to 5% or less compared with expressed wild type (WT).

Truncation at the N Terminus-- Deletion of the first two residues (Gly-Asn) from the N terminus of FS-288 resulted in a non-parallel dose-dilution curve from which a relative potency estimate of 5-10% of wild-type FS-288 could be estimated (Fig. 2). Substitution of the two N-terminal residues with alanine restored binding activity to 45% of wild-type (Table I).

                              
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Table I
Comparative activin binding activity of follistatin-288 mutants
Relative potencies (mean ± S.E.) in competition binding assay, expressed as ratio of half-maximal binding inhibition by mutant compared with expressed Myc-tagged wild-type FS-288 (1.00; IC50 = 0.30 nM). Data based on three or more assays for each mutant.

N-terminal Domain Disulfide Disruption-- In our previous study (18), activin binding was abolished after reduction of disulfide linkages in full-length FS-288. Disulfide disruption limited to the N-terminal domain was replicated through expression of a construct replacing the adjacent cysteines at positions 26 and 27 with alanine. This would represent disruption of two intrinsic N-domain disulfide linkages because (a) adjacent cysteines normally do not link and (b) follistatin domain disulfides occur exclusively within each individual domain (14). The Cys-substituted product was expressed at levels comparable with wild-type and migrated identically to the wild-type by polyacrylamide gel electrophoresis (data not shown). Activin binding was reduced to <5% of the wild-type, an effect approaching that of outright deletion of the N-terminal domain (Fig. 2).

Modification and Mutation of Methionine-- We first used chemical modification in conjunction with site-directed mutagenesis to define the importance of methionine and tryptophan throughout FS-288, followed by additional mutations targeting more explicitly other residues within the N-terminal domain. The three methionine residues, including Met-50 in the N-domain as well as Met-79 and Met-268 in follistatin domains I and III respectively (Fig. 1), were modified by cyanogen bromide treatment which cleaves the peptide chain leaving methionine as a C-terminal homoserine lactone. Sequence analysis (data not shown) confirmed that cleavage was limited to the predicted sites. As shown in Fig. 3A, activin binding was comparable with that observed after incubation with reaction solvent alone, suggesting that none of the methionines in follistatin are critical for binding. This was confirmed for Met-50 by point mutation to glutamic acid, a residue closely replicating the methionine sulfoxide oxidation product implicated in loss of activity of some native enzymes and hormones (25). In this case, the M50E product showed the full binding activity of expressed wild-type FS-288 (Fig. 3B).


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Fig. 3.   Modification of methionine residues in FS-288. A, inhibition curves showing retention of activin binding activity following cyanogen bromide cleavage/modification of the three methionines in purified FS-288. The control FS-288 preparation was incubated with reaction solvent (70% trifluoroacetic acid/H20) alone under otherwise identical conditions (18 h, 20 °C). B, activin binding by expressed FS-288 after replacement of the single N-domain methionine (Met-50) by glutamic acid. As predicted by the cyanogen bromide study (A), binding inhibition by the M50E mutant was comparable with expressed wild-type FS-288. C, effect of hydrogen peroxide oxidation on activin binding by purified FS-288. Sequence analyses showed that, in addition to methionine, at least one tryptophan (position 4) was modified; the oxidized product was inactive in the competition assay for labeled activin binding. Activity of FS-288 was unaffected by incubation with oxidation solvent alone (H2O; 37 °C, 45 min).

Role of Tryptophan and Hydrophobic Residues-- In evaluating the role of methionine (above), mild oxidation with hydrogen peroxide was employed as an alternative form of modification. Unexpectedly, the oxidized product did not bind activin (Fig. 3C). Sequence analysis showed loss of the tryptophan residue at position 4, replaced by a more hydrophilic product eluting between Tyr and Pro in the phenylthiohydantoin HPLC profile; similar oxidative changes in tryptophan have been described previously (26). Hence, at least one tryptophan in FS-288 appeared intolerant to modification to an oxidized form. Systematic mutation of individual tryptophans to Ala or Asp reduced binding activity to 2-5% of the wild-type expression product after substitution for either Trp-4 or Trp-36 within the N-terminal domain (Fig. 4A). Substitution by Phe restored 50-60% activity to position 4 and full activity to position 36 (Fig. 4B), consistent with a requirement for a large hydrophobic residue, not specifically tryptophan, at these positions.


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Fig. 4.   Activin binding inhibition curves for key N-terminal domain point mutations. A, alanine substitution for tryptophan at positions 4 or 36 reduced binding to 2-5% of expressed wild type (WT) FS-288. Replacement of Lys-23 decreased binding to 30%, the only N-domain basic residue affected by mutation. B, binding was restored to >50% after substitution of Trp-4 and Trp-36 with another hydrophobic residue, phenylalanine. The W258A mutation in the third FS domain did not affect activin binding but did impair immunoreactivity in the follistatin immunochemiluminescent assay (SPICA) as described in text.

Alanine replacement of Trp-49 within the N-domain, Trp-98 in follistatin domain I, or 258 in follistatin domain III did not reduce activin binding, as summarized in Table I. Interestingly, the W258A mutation did impair immunoreactivity in the SPICA assay, which detected a concentration only one-fifth that of the Myc-tag radioimmunoassay. This mutation apparently induced conformational changes within the follistatin domains that nonetheless did not affect association with activin (Fig. 4B). Among other N-domain hydrophobic residues, alanine replacement of Phe-52 resulted in a decrease in activin binding to 19% of wild-type (Table I). On the other hand, Phe-47, as well as Leu-32 and -46, were tolerant to mutation. Minor reductions of approximately 2-fold were found after acidic substitutions for the conserved residues Leu-5 and Gln-7.

Mutation of Charged Residues-- The several basic (Lys, Arg) residues in the follistatin N-domain were tolerant to alanine mutation with the single exception of Lys-23, which showed a partial reduction in activin binding to 30% of expressed wild-type (Fig. 4A; Table I). Replacement of Glu-25 was also without effect despite its conservation among all FS-related proteins (Table I). Hence, charge interactions involving the N-terminal domain appear less important than hydrophobic contacts in mediating association between follistatin and activin.

Activin-mediated Transcriptional Response-- If activin binding is relevant to the physiological function of follistatin, mutants lacking activin binding should also be impaired in their ability to suppress activin-mediated biological responses. This was evaluated using assays representing two different aspects of the activin response pathway. In HEK-293 cells transfected with a luc-coupled activin response element (22), a 3-fold transcriptional response to 5 ng/ml (0.15 nM) activin was reduced to basal levels by a 1.5 nM concentration of either NIH FS-288 or expressed Myc-tagged wild-type FS-288 (Fig. 5). Mutants deficient in activin binding failed to suppress activin-stimulated activity; these included the Trp-4 and -36 substitutions, as well as the N-domain-deleted and N-domain disulfide-disrupted follistatins (Fig. 5). Mutations that did not affect activin binding antagonized activin as effectively as wild-type FS-288.


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Fig. 5.   Suppression of activin-mediated transcriptional responses in cultured HEK-293 cells transfected with a luciferase-coupled activin-response element. Activin (5 ng/ml; 0.15 nM) was incubated with 50 ng/ml (1.5 nM) wild-type or mutant follistatins for 60 min before addition to cells. After 24 h, cells were harvested, extracted, and assayed for luciferase activity. Follistatin mutants deficient in activin binding (see Figs. 2, 4) did not suppress activin-stimulated luciferase activity whereas wild-type FS-288 and tolerated mutants (R6A, K9A, W4F) depressed activity to levels comparable with cells untreated with activin or follistatin (basal).

Pituitary Cell FSH Secretion-- The effect of activin on pituitary gonadotrophs to increase FSH release is well documented, as is the ability of follistatin to suppress secretion sustained by endogenously produced activin (1, 3, 4). Incubation of 100 ng/ml (3.2 nM) expressed wild-type follistatin for 72 h in primary pituitary cell culture diminished FSH secretion to 20% of levels observed in the presence of control medium from nontransfected CHO cells (Fig. 6A). Dose-dependent inhibition comparable with wild-type was observed with the "active" mutants R6A and K9A (Fig. 6B). The N-domain-deleted protein comprising only the follistatin domains, and the several N-domain mutants with impaired activin binding were devoid of FSH-suppressing activity at doses up to 100-fold higher than those eliciting wild-type responses.


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Fig. 6.   Effects of FS-288 mutation on suppression of endogenous FSH secretion in cultured rat pituitary cells. Cells were dispersed and incubated in alpha MEM for 72 h, washed, and treated with fresh medium containing follistatin preparations for an additional 72 h. Medium was harvested and assayed for FSH by radioimmunoassay using reagents provided by NIDDK, National Institutes of Health. A, FSH secretion in presence of 100 ng/ml (3.2 nM) doses of wild-type and mutant follistatins. FSH secretion remained at >80% of basal levels in presence of all mutants deficient in activin binding. B, dose-response curves for FSH suppression by wild-type (WT) and representative mutant follistatins with normal (R6A, K9A) and impaired activin binding (W4A, N-terminal domain (NTD) deletion).

As summarized by the comparative responses shown in Fig. 7, the respective assays show a strong correlation between activin binding and downstream biological effects among follistatins with deleted, disrupted, or modified N-terminal domain sequences. In addition, these results support the concept that follistatin's biological activity in the pituitary is attributable to its ability to bind activin.


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Fig. 7.   Comparison of the binding and biological activities of key mutant follistatins. Mean responses in the assays for activin binding, inhibition of activin-mediated transcriptional activity (ARE-luc) and suppression of pituitary cell FSH secretion are plotted relative to the respective wild-type (100%) responses. Close correlation among the three assay systems shows that mutations impairing activin binding also abolish follistatin's ability to inhibit activin-dependent biological activities.


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ABSTRACT
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The structure-function studies reported here clearly document the importance of the 63-residue N-terminal domain in the biological actions of follistatin. Deletion of the N-domain diminished activin binding below the lower limits of detection (<5% of wild-type) and abolished bioactivity in assays representing two aspects of the activin response pathway. Thus, the three follistatin domains comprising the majority of the molecule (residues 64-288) inherently lack the necessary structural determinants for activin binding and neutralization, which is at present the only confirmed biological activity for follistatin.

Our results show the structural integrity of the N-terminal domain to be functionally critical. Because each follistatin domain is internally disulfide-linked (14), the N-terminal domain must likewise be a separate folding unit in which all six cysteines are involved in disulfide linkages (18). Disruption of two of the three intradomain disulfides through the mutation of Cys-26 and -27 to alanine was comparable with outright deletion of the domain in its effect on binding and bioactivity. Besides intact disulfides, a complete N terminus is required because deletion of the first two residues (Gly-Asn) reduced binding activity to <10% of wild-type FS-288. This may represent a stringent length requirement, as an early mutagenesis study (19) found that lengthening the N terminus by insertion of two residues between positions 2 and 3 also impairs activin binding. Replacement of residues 1 and 2 with alanine partially, though not completely, restored binding activity; hence alanine may not entirely satisfy the size requirement fulfilled optimally by the smaller glycine residue. Alternatively, the Gly-Asn deletion may disrupt folding or disulfide formation when Cys-3 is left as the N-terminal residue. In support of this is the previous observation (18) that the synthetic (1-26) peptide still binds activin after shortening to (3-26). We found expression levels of the truncated molecule to be lower than most full-length follistatin proteins, but SPICA immunoreactivity is retained suggesting that at least the follistatin domains fold normally.

Among point mutations in the N-terminal domain, the striking effect of tryptophan replacement by alanine or aspartic acid at positions 4 and/or 36 on binding and bioactivity indicates the importance of hydrophobic interactions in the high affinity association of follistatin with activin. This requirement is met by a bulky hydrophobic residue, rather than any specific attribute of tryptophan itself, because substitution at either position by phenylalanine substantially restored activity. A third hydrophobic residue, Phe-52, appears also to be important because its replacement by alanine reduced binding potency 5-fold. By contrast, numerous other residues were remarkably tolerant to mutation given the strong conservation among follistatins from different species (Fig. 1). Among the several basic residues, only Lys-23 showed decreased binding activity after mutation, but even this could represent a hydrophobic contact because lysine can act as a nonpolar rather than charged residue by virtue of its long methylene side-chain (27).

Hydrophobic interactions are perhaps most notably illustrated by the association between growth hormone and its receptor (28). Significantly, in the crystal structure of the extracellular region of the activin type II receptor (29), at least three hydrophobic residues have been identified by Gray et al. (20) to be important components of a binding pocket for activin. Follistatin thus might neutralize activin by competition for hydrophobic sites on the activin molecule that would otherwise interact with receptor.

Receptor contact sites on activin A have been investigated extensively by site-directed mutagenesis (30). Sites of follistatin binding have been mapped by a more limited number of mutations (30) and by analysis of direct binding by a series of overlapping synthetic peptides covering the full-length of the beta A subunit (7). One binding peptide (residues 15-29) is rich in hydrophobic residues; these have not been studied by mutagenesis, although substitutions for Asp-27 have been found to affect receptor interaction. The two other follistatin-binding peptides both included Lys-102, another well documented contact residue in activin whose positive charge appears essential for binding to its receptor but not to follistatin (20, 30). Thus, not all contact sites are shared by receptor and follistatin, and it has been suggested that follistatin may act at least in part by masking Lys-102 (30). It must be emphasized, however, that activin mutations studied to date have focused on charged rather than hydrophobic residues. It is therefore likely that individual hydrophobic residues on activin that contact both receptor and follistatin remain to be identified.

Recently, a homologous protein called follistatin-related gene product (FLRG) (17, 31), also referred to as follistatin-related protein (FSRP) (32), has been cloned and localized to a wide range of tissues and cell types. The FLRG/FSRP molecule contains only two follistatin domains, preceded by a 6-cysteine N-terminal domain as in follistatin itself. Significantly, the important hydrophobic residues (Trp-4, Trp-36, and Phe-52) are conserved in FLRG/FSRP despite numerous sequence differences elsewhere. Because FLRG/FSRP binds activin and inhibits its transcriptional activity in reporter assays (17, 32), it is likely that these conserved hydrophobic residues contribute to interactions with activin in this molecule as well.

Other than in FLRG, the follistatin N-domain has no homologs except for several repeats in the fibrillin family of elastin components, that can be aligned with the first four cysteines and several other residues in the N-terminal region (Fig. 1). There are no evident functional relationships, as fibrillin is not known to bind any ligand. However, the matches do suggest a distant common ancester in which the first four cysteines may be linked to form a pair of loops comparable with those we have disrupted, with loss of bioactivity, after alanine replacement of Cys-26 and -27 in follistatin.

Although the present mutagenesis data and our earlier survey of activin-binding peptides (18) are mutually supportive in localizing binding to the N-terminal domain, certain differences involving specific residues or sequences are evident. These may provide clues as to the structural role of a residue or segment, but also may reflect the significant differences in assay format between the two types of experiment. The peptide assays measure activin binding directly to peptide immobilized on the plate and do not take into account conformational interactions present in the whole molecule in solution. Moreover, because of the high concentrations of immobilized peptide, it is difficult to distinguish low-affinity from high-affinity interactions, in contrast to mutant follistatins competing in solution for activin binding. Hence, for example, the peptide (27-43) including Trp-36 did not bind activin (18). This may mean that the Trp residue in the peptide was buried through hydrophobic interactions with the plate itself but also could suggest that the tryptophan interacts with other residues in whole follistatin rather than directly contacting activin. Conversely, the effect of deletion of N-domain residues 1 and 2 was not evident in the peptide studies in which (1-26) and (3-26) both bound activin. This may represent simply a low-affinity binding interaction by (3-36) or alternatively a length requirement for the whole molecule in solution that might not be evident in a linear peptide. Thirdly, it could indicate that an N-terminal cysteine disrupts folding of the truncated full-length follistatin as mentioned earlier. Conformational differences between the two formats may also explain why activin binding is retained after substitution for residues 46-47 but not after outright deletion from the peptide fragment (44-59). These issues emphasize how, as in studies with other proteins, the peptide approach is most useful for pointing out regions for more detailed examination by site-directed mutagenesis.

The inability of the follistatin domains comprising the remainder of the FS-288 molecule to bind activin on their own contrasts with the results of Tsuchida et al. (17) showing activin binding by the second of the two FS domains in human and murine FLRG/FSRP. This analysis was based on radiolabeled ligand binding after gel electrophoresis, so quantitative comparisons of binding or estimates of bioactivity were not available, and these authors did not rule out some contribution by the N-domain. In our study of activin binding by follistatin peptides (18), we did not find activin-binding peptides from outside the N-domain, and our limited mutational and chemical modifications to date within the FS domains have not affected activity. For example, a sequence including Trp-98 within domain I had been proposed as a potential binding site based on its homology to the inhibitory region of the Kazal family of protease inhibitors (14). This appears unlikely, as we found full binding activity to be retained after alanine mutation of Trp-98.

Nevertheless, the essential determinants identified in the N-terminal domain may still not be sufficient for high-affinity binding without participation of one or more of the follistatin domains acting through, for example, conformational influences or even direct contact with the N-domain to facilitate or augment activin association. Our observation that mutation of Trp-258 in FS domain III may disturb an antigenic site in domain II, and the finding by Wang et al. (18) that activin binding alters recognition by a monoclonal antibody directed toward an epitope in domain III, are indicators that the various domains can influence one another in FS-288.

Evidence for functional interactions between follistatin domains and neighboring domains are also provided by several studies with SPARC/BM40. A peptide sequence from the single FS domain has been found to replicate the effects of the full-length molecule in disrupting endothelial-cell focal adhesion sites (33), and in inhibiting vascular endothelial growth factor-stimulated DNA synthesis through a direct binding interaction with vascular endothelial growth factor (34). At least one other peptide sequence, within the adjacent extracellular calcium-binding (EC) domain, has similar effects, suggesting a cooperative interaction between the domains. This is supported more directly by recent structural and mutagenesis data (35) showing a close contact between a segment of the FS domain and an EF-hand calcium-binding region in the EC domain. This association appears essential for sustaining high-affinity calcium binding and could represent a precedent for N-domain/FS domain interactions in follistatin as well.

Another likely contribution of the follistatin domain region to bioactivity is its ability to bind to cell surfaces via heparin-binding sequence(s), especially the (75-86) segment within the FS domain 1 (Fig. 1A) (11). Despite their outwardly diverse actions, most of the FS domain-containing extracellular matrix proteins exhibit some mechanism for cell surface adhesion. Agrin and testican themselves contain glycosaminoglycan side-chains capable of recognizing binding sequences on cell surfaces (36, 37), and the hypothalamic protein X-7365 is anchored through a membrane-spanning domain close to the C terminus (38). An N-terminal laminin-binding site in agrin is essential for its association with synaptic basal lamina to induce formation of neuromuscular junctions (39, 40). In the case of follistatin, heparan sulfate proteoglycan binding may serve to maintain high local concentrations at cell surfaces to regulate activin access to its receptor in an autocrine or paracrine setting (9) and to provide a barrier to access by exogenous activin (22).

Absence from FLRG of a follistatin domain containing a consensus heparan sulfate binding sequence may account for at least one difference in bioactivity between FLRG and follistatin. Although, like follistatin, FLRG binds activin and inhibits activin-mediated transcriptional activity (17), it appears not to suppress pituitary FSH secretion (32). Thus, follistatin's biological effects may involve structural attributes beyond those simply mediating activin binding.

Whereas these investigations have focused on interactions with activin, follistatin has also been observed to bind other transforming growth factor-beta -related growth factors, including several of the BMPs (41-44) and also certain members of the growth differentiation factor (GDF) family (45, 46). Binding affinities for the BMPs are substantially lower than for activin (41, 43), but local concentrations of follistatin may be sufficient to enable it to act as a regulator in the numerous systems expressing these factors. Hence, follistatin has been shown to neutralize BMP action in embryogenesis (42, 43) and may potentially influence cartilage maturation and bone development through interactions with BMP-7, among others (47, 48). In cultured prostate tumor cells, follistatin reverses the antiproliferative effects of both BMP-6 and BMP-7 (49). A recent report (50) suggests that BMP-6 and BMP-7 and potentially their interactions with follistatin are important components in the control of pituitary FSH secretion.

The relatively low binding affinities for BMPs make comparisons among mutant follistatins difficult, pending availability of assays in specific BMP-dependent systems. Neutralizing effects of follistatin on BMPs and GDFs may involve the same determinants as we have found for activin binding, with the extensive sequence differences among these ligands accounting for their lower potencies. However, the involvement of different contact sites on follistatin itself cannot be ruled out. This is particularly true in the reported instances (43, 44, 46) in which follistatin does not neutralize BMP or GDF receptor interaction, suggesting instead a tripartite complex between ligand, its receptor, and follistatin that might indirectly modulate or even enhance growth factor activity.

Nevertheless, the correlation between activin binding, activin transcriptional activity and pituitary FSH suppression among the mutants we have tested supports strongly the neutralization of activin as a primary mechanism for follistatin's biological and physiological actions. Transcriptional assays, such as ours using a luc-coupled ARE responsive to a Smad-2/FAST-1 signaling pathway, are intended to replicate, at least in part, transcriptional responses in whole pituitary cells as used in measurement of FSH suppression. However, the effects of activin in pituitary cells inevitably involve additional mediators and their associated transcription factors.

Furthermore, activin increases not only the transcription of FSHbeta but also the secretion of the FSH heterodimer by the gonadotroph, and further facilitates FSH production in vivo by increasing GnRH receptor levels (3, 4). Follistatin conceivably may affect these multiple activin effects differentially, especially if its actions involve other ligands or even association with some as-yet-undefined "receptor." Use of the mutant follistatins described here in additonal assay systems, and ultimately in whole animals through transgenic methodology, offers another means to explore possible unrecognized effects of follistatin beyond those attributable to the binding and neutralization of activin.

    ACKNOWLEDGEMENTS

We thank Dr. QiFa Wang for helpful discussions and advice throughout the course of this work and Ashok Khatri and Teofy Ostrea of the MGH Endocrine/Reproductive Chemistry Core Facility for expert sequencing and analyses of the modified native follistatin preparations and synthesis of the peptide ligands used in our radioimmunoassays. We also acknowledge the National Hormone and Pituitary Program, NIDDK, directed by Dr. A. F. Parlow, for preparing and providing the reagents for the rat FSH radioimmunoassay.

    FOOTNOTES

* Supported by Grant DK-53828 from the National Institutes of Health.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

To whom correspondence should be addressed: Endocrine Unit, Wellman 501, Massachusetts General Hospital, Boston, MA 02114. Tel.: 617-726-3966; Fax: 617-726-7543; E-mail: Keutmann@helix.mgh.harvard.edu.

Published, JBC Papers in Press, March 14, 2001, DOI 10.1074/jbc.M100736200

    ABBREVIATIONS

The abbreviations used are: FS, follistatin; FSH, follicle-stimulating hormone; BMP, bone morphogenic protein; CHO, Chinese hamster ovary; FLRG, follistatin-related gene product; FSRP, follistatin-related protein; GDF, growth differentiation factor; SPARC, secreted protein, acidic and rich in cysteine; SPICA, solid-phase immunochemiluminescent assay; HPLC, high performance liquid chromatography.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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