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• W1996832276 abstract "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. 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. follistatin follicle-stimulating hormone bone morphogenic protein Chinese hamster ovary follistatin-related gene product follistatin-related protein growth differentiation factor secreted protein, acidic and rich in cysteine solid-phase immunochemiluminescent assay high performance liquid chromatography 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.1Vale W. Rivier C. Hsueh A. Campen C. Meunier H. Bicsak T. Recent Prog. Horm. Res. 1988; 44: 1-34PubMed Google Scholar, 2Michel U. Farnworth P. Findlay J.K. Mol. Cell. Endocrinol. 1993; 91: 1-11Crossref PubMed Scopus (136) Google Scholar, 3DePaolo L.V. Proc. Soc. Exp. Biol. Med. 1997; 214: 328-339Crossref PubMed Scopus (63) Google Scholar, 4Peng C. Mukai S.T. Biochem. Cell Biol. 2000; 78: 261-279Crossref PubMed Scopus (47) Google Scholar). Cloning and sequencing (5Shimasaki S. Koga M. Esch F. Cooksey K. Mercado M. Koba A. Ueno N. Ying S.-Y. Ling N. Guilleman R. Proc. Natl. Acad. Sci. U. S. A. 1988; 85: 4218-4222Crossref PubMed Scopus (222) Google Scholar) 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 (6Nakamura T. Takio K. Eto Y. Shibai H. Titani K. Sugino H. Science. 1990; 247: 836-838Crossref PubMed Scopus (792) Google Scholar) that the protein binds the activin A homodimer with high affinity, approaching irreversibility because of its slow dissociation rate (7Schneyer A.L. Rzucidlo D.A. Sluss P.M. Crowley W.F. Endocrinology. 1994; 135: 667-674Crossref PubMed Scopus (125) Google Scholar). 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 (8Meriggiola M. Dahl K.D. Mather J.P. Bremner W.J. Endocrinology. 1994; 134: 1967-1970Crossref PubMed Scopus (29) Google Scholar, 9deWinter J.P. ten Dijke P. de Vries C.J.M. van Achterberg T.A.E. Sugino H. de Waele P. Huylebroeck D. Verschueren K.,. van den Eijnden-van Raaij A.J.M. Mol. Cell. Endocrinol. 1996; 116: 105-114Crossref PubMed Scopus (173) Google Scholar). More recently, follistatin has been reported to accelerate endocytosis and degradation of activin (10Hashimoto O. Nakamura T. Shoji H. Shimasaki S. Hayashi Y. Sugino H. J. Biol. Chem. 1997; 272: 13835-13842Abstract Full Text Full Text PDF PubMed Scopus (170) Google Scholar). 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-β family of regulatory factors (2Michel U. Farnworth P. Findlay J.K. Mol. Cell. Endocrinol. 1993; 91: 1-11Crossref PubMed Scopus (136) Google Scholar, 3DePaolo L.V. Proc. Soc. Exp. Biol. Med. 1997; 214: 328-339Crossref PubMed Scopus (63) Google Scholar). Localization appears to be facilitated through interaction with cell surface proteoglycans through at least one heparin binding site (11Inouye S. Ling N. Shimasaki S. Mol. Cell. Endocrinol. 1992; 90: 1-6Crossref PubMed Scopus (52) Google Scholar). 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 (2Michel U. Farnworth P. Findlay J.K. Mol. Cell. Endocrinol. 1993; 91: 1-11Crossref PubMed Scopus (136) Google Scholar). In fact, the lethal effects found in follistatin-null animals are attributable to skeletal and cutaneous abnormalities (12Matzuk M.M. Llu N. Vogel H. Sellheyer K. Roop D.R. Jaenisch R. Bradley A. Nature. 1995; 374: 360-363Crossref PubMed Scopus (512) Google Scholar). 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 (13Patthy L. Nikolics K. Trends Neurosci. 1993; 16: 76-81Abstract Full Text PDF PubMed Scopus (104) Google Scholar). 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 (13Patthy L. Nikolics K. Trends Neurosci. 1993; 16: 76-81Abstract Full Text PDF PubMed Scopus (104) Google Scholar). Each follistatin domain forms an autonomous folding unit, as confirmed by the crystal structure of the single follistatin domain from SPARC/BM40 (14Hohenester E. Maurer P. Timpl R. EMBO J. 1997; 16: 3778-3786Crossref PubMed Scopus (137) Google Scholar) 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 (13Patthy L. Nikolics K. Trends Neurosci. 1993; 16: 76-81Abstract Full Text PDF PubMed Scopus (104) Google Scholar, 14Hohenester E. Maurer P. Timpl R. EMBO J. 1997; 16: 3778-3786Crossref PubMed Scopus (137) Google Scholar, 15Yan Q. Sage E.H. J. Histochem. Cytochem. 1999; 47: 1495-1505Crossref PubMed Scopus (309) Google Scholar, 16Uchida T. Wada K. Akamatsu T. Yonezawa M. Noguchi H. Mizoguchi A. Kasuga M. Sakamoto C. Biochem. Biophys. Res. Commun. 1999; 20: 593-602Crossref Scopus (96) Google Scholar), as well as in a recently described activin-binding follistatin-related gene product (FLRG) (17Tsuchida K. Arai K.Y. Kuramoto Y. Yamakawa N. Hasegawa Y. Sugino H. J. Biol. Chem. 2000; 275: 40788-40796Abstract Full Text Full Text PDF PubMed Scopus (171) Google Scholar). 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 (18Wang Q.F. Keutmann H.T. Schneyer A.L. Sluss P.M. Endocrinology. 2000; 141: 3183-3193Crossref PubMed Scopus (31) Google Scholar) showing direct binding of activin by two synthetic peptides representing discontinuous sequences from this region, together with an earlier mutagenesis experiment (19Inouye S. Guo Y. Ling N. Shimasaki S. Biochem. Biophys. Res. Commun. 1991; 179: 352-358Crossref PubMed Scopus (27) Google Scholar) 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 (20Gray P.C. Greenwald J. Blount A.L. Kunitake K.S. Donaldson C.J. Choe S. Vale W. J. Biol. Chem. 2000; 275: 3206-3212Abstract Full Text Full Text PDF PubMed Scopus (90) Google Scholar) in the type II activin receptor binding site. 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 (21Wang Q.F. Khoury R.H. Smith P.C. McConnell D.S. Padmanahban V. Midgley A.R. Schneyer A.L. Crowley W.F. Sluss P.M. J. Clin. Endocrinol. Metab. 1996; 81: 1434-1441Crossref PubMed Scopus (25) Google Scholar). 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 βA-subunit cDNA as described by Delbaere et al. (22Delbaere A. Sidis Y. Schneyer A.L. Endocrinology. 1999; 140: 2463-2470Crossref PubMed Scopus (27) Google Scholar). 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 (ΔNTD), 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. 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. 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.1 A) within FS domain II, as previously described (21Wang Q.F. Khoury R.H. Smith P.C. McConnell D.S. Padmanahban V. Midgley A.R. Schneyer A.L. Crowley W.F. Sluss P.M. J. Clin. Endocrinol. Metab. 1996; 81: 1434-1441Crossref PubMed Scopus (25) Google Scholar, 23McConnell D.S. Wang Q.F. Sluss P.M. Bolf N. Khoury R.H. Schneyer A.L. Midgley A.R. Reame N.E. Crowley W.F. Padmanabhan V. J. Clin. Endocrinol. Metab. 1998; 83: 851-858Crossref PubMed Scopus (72) Google Scholar) 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 γ-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). Binding of expressed follistatins to labeled activin was determined by competition assay as previously described (7Schneyer A.L. Rzucidlo D.A. Sluss P.M. Crowley W.F. Endocrinology. 1994; 135: 667-674Crossref PubMed Scopus (125) Google Scholar). Mutant or wild-type follistatins were incubated with125I-labeled activin in binding buffer (10 mmphosphate-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. 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. 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 (22Delbaere A. Sidis Y. Schneyer A.L. Endocrinology. 1999; 140: 2463-2470Crossref PubMed Scopus (27) Google Scholar), 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 onRenilla activity. The construction of the ARE-GFP-Lux and specificity of the ARE-FAST-1 reporter system for activin has been previously described (22Delbaere A. Sidis Y. Schneyer A.L. Endocrinology. 1999; 140: 2463-2470Crossref PubMed Scopus (27) Google Scholar). 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. Assay for suppression by follistatin of basal FSH secretion in cultured rat anterior pituitary cells was based on the method of Scott et al.(24Scott R.S. Burger H.G. Quigg H. Endocrinology. 1980; 107: 1536-1542Crossref PubMed Scopus (146) Google Scholar). 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 α-minimum essential medium (αMEM) containing 21 mmNaHCO3, 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. 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 (23McConnell D.S. Wang Q.F. Sluss P.M. Bolf N. Khoury R.H. Schneyer A.L. Midgley A.R. Reame N.E. Crowley W.F. Padmanabhan V. J. Clin. Endocrinol. Metab. 1998; 83: 851-858Crossref PubMed Scopus (72) Google Scholar). 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.1 A) within the second follistatin domain (18Wang Q.F. Keutmann H.T. Schneyer A.L. Sluss P.M. Endocrinology. 2000; 141: 3183-3193Crossref PubMed Scopus (31) Google Scholar). 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. Fig. 1 Asummarizes 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. 1 B; 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. 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 (TableI).Table IComparative activin binding activity of follistatin-288 mutantsTryptophan residuescharged residuesW4A<0.05R6A1.13 ± .09W4D<0.05K9A1.19 ± .08W4F0.56 ± .04R12A0.92 ± .17W36A0.05 ± .01K18A1.10 ± .17W36D0.07 ± .02K23A0.30 ± .05W36F1.18 ± .31E25A0.99 ± .07W49A0.91 ± .11R31A1.69 ± .26W98A1.21 ± .25K48A0.90 ± .21W258A1.35 ± .18K63A0.61 ± .03Other neutral residuesstructural disruptionL5D0.45 ± .04Δ(G1N2)0.06 ± .03Q7A0.84 ± .10GN(1,2)AA0.45 ± .05Q7E0.46 ± .03CC(26,27)AA<0.05L32D0.72 ± .05(1–63) deletion<0.05LF(46,47)AA1.27 ± .26M50E0.97 ± .03F52A0.19 ± .05Relative 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. Open table in a new tab 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. In our previous study (18Wang Q.F. Keutmann H.T. Schneyer A.L. Sluss P.M. Endocrinology. 2000; 141: 3183-3193Crossref PubMed Scopus (31) Google Scholar), 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 (14Hohenester E. Maurer P. Timpl R. EMBO J. 1997; 16: 3778-3786Crossref PubMed Scopus (137) Google Scholar). 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). 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. 3 A, 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 (25Neuman N.P. Moore S. Stein W.H. Biochemistry. 1962; 1: 68-75Crossref PubMed Scopus (82) Google Scholar). In this case, the M50E product showed the full binding activity of expressed wild-type FS-288 (Fig. 3 B). 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. 3 C). 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 (26Purnananda G. Balasubramanian D. Matsugo S. Saito I. Biochemistry. 1992; 31: 4296-4303Crossref PubMed Scopus (132) Google Scholar). 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. 4 A). Substitution by Phe restored 50–60% activity to position 4 and full activity to position 36 (Fig. 4 B), consistent with a requirement for a large hydrophobic residue, not specifically tryptophan, at these positions. 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. 4 B). 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. 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. 4 A; Table I). Replacement of Glu-25 was also without effect despite its conservation among a" @default.
• W1996832276 created "2016-06-24" @default.
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• W1996832276 date "2001-05-01" @default.
• W1996832276 modified "2023-10-11" @default.
• W1996832276 title "Follistatin: Essential Role for the N-terminal Domain in Activin Binding and Neutralization" @default.
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