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INTRODUCTION |
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-
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largely superseded by evidence for its role as a local cellular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.
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MATERIALS AND METHODS |
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
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 (
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
-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
-minimum
essential medium (
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.
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RESULTS |
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).
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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
( NTD), truncated
( 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).
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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.
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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).
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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.
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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).
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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 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).
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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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DISCUSSION |
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
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-
-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 FSH
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.