Stromal Cell-derived Factor-1α Associates with Heparan Sulfates through the First β-Strand of the Chemokine
1999; Elsevier BV; Volume: 274; Issue: 34 Linguagem: Inglês
10.1074/jbc.274.34.23916
ISSN1083-351X
AutoresAli Amara, Olivier Lorthioir, Agustı́n Valenzuela-Fernández, Aude Magérus‐Chatinet, Marcus Thelen, Mônica Montes, Jean‐Louis Virelizier, Muriel Delepierre, Françoise Baleux, Hugues Lortat‐Jacob, Fernando Arenzana‐Seisdedos,
Tópico(s)Cell Adhesion Molecules Research
ResumoBiological properties of chemokines are believed to be influenced by their association with glycosaminoglycans. Surface plasmon resonance kinetic analysis shows that the CXC chemokine stromal cell-derived factor-1α (SDF-1α), which binds the CXCR4 receptor, associates with heparin with an affinity constant of 38.4 nm (kon = 2.16 × 106m−1 s−1 and koff = 0.083 × s−1). A modified SDF-1α (SDF-1 3/6) was generated by combined substitution of the basic cluster of residues Lys24, His25, and Lys27 by Ser. SDF-1 3/6 conserves the global native structure and functional properties of SDF-1α, but it is unable to interact with sensor chip-immobilized heparin. The biological relevance of these in vitro findings was investigated. SDF-1α was unable to bind in a CXCR4-independent manner on epithelial cells that were treated with heparan sulfate (HS)-degrading enzymes or constitutively lack HS expression. The inability of SDF-1 3/6 to bind to cells underlines the importance of the identified basic cluster for the physiological interactions of SDF-1α with HS. Importantly, the amino-terminal domain of SDF-1α which is required for binding to, and activation of, CXCR4 remains exposed after binding to HS and is recognized by a neutralizing monoclonal antibody directed against the first residues of the chemokine. Overall, these findings indicate that the Lys24, His25, and Lys27 cluster of residues forms, or is an essential part of, the HS-binding site which is distinct from that required for binding to, and signaling through, CXCR4. Biological properties of chemokines are believed to be influenced by their association with glycosaminoglycans. Surface plasmon resonance kinetic analysis shows that the CXC chemokine stromal cell-derived factor-1α (SDF-1α), which binds the CXCR4 receptor, associates with heparin with an affinity constant of 38.4 nm (kon = 2.16 × 106m−1 s−1 and koff = 0.083 × s−1). A modified SDF-1α (SDF-1 3/6) was generated by combined substitution of the basic cluster of residues Lys24, His25, and Lys27 by Ser. SDF-1 3/6 conserves the global native structure and functional properties of SDF-1α, but it is unable to interact with sensor chip-immobilized heparin. The biological relevance of these in vitro findings was investigated. SDF-1α was unable to bind in a CXCR4-independent manner on epithelial cells that were treated with heparan sulfate (HS)-degrading enzymes or constitutively lack HS expression. The inability of SDF-1 3/6 to bind to cells underlines the importance of the identified basic cluster for the physiological interactions of SDF-1α with HS. Importantly, the amino-terminal domain of SDF-1α which is required for binding to, and activation of, CXCR4 remains exposed after binding to HS and is recognized by a neutralizing monoclonal antibody directed against the first residues of the chemokine. Overall, these findings indicate that the Lys24, His25, and Lys27 cluster of residues forms, or is an essential part of, the HS-binding site which is distinct from that required for binding to, and signaling through, CXCR4. stromal cell-derived factor-1α glycosaminoglycans heparan sulfate macrophage inflammatory protein 1β interleukin-8 monoclonal antibody polyacrylamide gel electrophoresis resonance units surface plasmon resonance nuclear magnetic resonance nuclear Overhauser effect nuclear Overhauser effect spectroscopy total correlation fluoremethyloxycarbonyl phosphate-buffered saline Chinese hamster ovary regulated on activation normal T cell expressed and secreted fluorescence-activated cell sorter Based on the relative position of the two first cysteine residues, chemokines are classified in two main subfamilies, CC and CXC chemokines (1Baggiolini M. Dewald B. Moser B. Annu. Rev. Immunol. 1997; 15: 675-705Crossref PubMed Scopus (1977) Google Scholar). Stromal cell-derived factor-1 (SDF-1)1 also called pre-B cell-stimulating factor (2Nagasawa T. Kikutani H. Kishimoto T. Proc. Natl. Acad. Sci. U. S. A. 1994; 91: 2305-2309Crossref PubMed Scopus (705) Google Scholar) is a CXC chemokine originally purified from bone marrow cell supernatants (3Tashiro K. Tada H. Heilker R. Shirozu M. Nakano T. Honjo T. Science. 1993; 261: 600-603Crossref PubMed Scopus (636) Google Scholar). Two forms, α and β (68 and 72 amino acids, respectively), generated by alternative splicing from a unique sdf-1 gene, have been identified (4Shirozu M. Nakano T. Inazawa J. Tashiro K. Tada H. Shinohara T. Honjo T. Genomics. 1995; 28: 495-500Crossref PubMed Scopus (534) Google Scholar). The α form is the most abundant (4Shirozu M. Nakano T. Inazawa J. Tashiro K. Tada H. Shinohara T. Honjo T. Genomics. 1995; 28: 495-500Crossref PubMed Scopus (534) Google Scholar). Human and murine SDF-1α proteins differ by a single residue at position 18 (valine to isoleucine in the murine protein) (4Shirozu M. Nakano T. Inazawa J. Tashiro K. Tada H. Shinohara T. Honjo T. Genomics. 1995; 28: 495-500Crossref PubMed Scopus (534) Google Scholar). CXCR4 is the only identified receptor for SDF-1α. Furthermore, the interaction between SDF-1α and CXCR4 appears to be unique and non-promiscuous (5Nagasawa T. Hirota S. Tachibana K. Takakura N. Nishikawa S. Kitamura Y. Yoshida N. Kikutani H. Kishimoto T. Nature. 1996; 382: 635-638Crossref PubMed Scopus (1999) Google Scholar, 6Tachibana K. Hirota S. Iizasa H. Yoshida H. Kawabata K. Kataoka Y. Kitamura Y. Matsushima K. Yoshida N. Nishikawa S. Kishimoto T. Nagasawa T. Nature. 1998; 393: 591-594Crossref PubMed Scopus (1317) Google Scholar, 7Zou Y.R. Kottmann A.H. Kuroda M. Taniuchi I. Littman D.R. Nature. 1998; 393: 595-599Crossref PubMed Scopus (2114) Google Scholar). SDF-1α stimulates intracellular calcium flux and chemotaxis in monocytes, T lymphocytes, and neutrophils, a characteristic shared with other CXC chemokines (8Oberlin E. Amara A. Bachelerie F. Bessia C. Virelizier J.L. Arenzana-Seisdedos F. Schwartz O. Heard J.M. Clark-Lewis I. Legler D.F. Loetscher M. Baggiolini M. Moser B. Nature. 1996; 382: 833-835Crossref PubMed Scopus (1478) Google Scholar, 9Bleul C.C. Farzan M. Choe H. Parolin C. Clark-Lewis I. Sodroski J. Springer T.A. Nature. 1996; 382: 829-833Crossref PubMed Scopus (1749) Google Scholar). However, SDF-1α exhibits structural and molecular characteristics that make it a unique chemokine among members of CC and CXC families. SDF-1α possesses the peculiar capacity of attracting and promoting bone marrow engraftment of CD34+ CXCR4 hematopoietic stem cells (10Peled A. Petit I. Kollet O. Magid M. Ponomaryov T. Byk T. Nagler A. Ben-Hur H. Many A. Shultz L. Lider O. Alon R. Zipori D. Lapidot T. Science. 1999; 283: 845-848Crossref PubMed Scopus (1458) Google Scholar). In contrast to most chemokines, which are induced by cytokines or mitogenic stimuli, SDF-1α is constitutively expressed in a large number of tissues (4Shirozu M. Nakano T. Inazawa J. Tashiro K. Tada H. Shinohara T. Honjo T. Genomics. 1995; 28: 495-500Crossref PubMed Scopus (534) Google Scholar). Importantly, Sdf-1 gene knock-outs induce anomalies in hematopoiesis and the development of cardiovascular system provocating pre- or perinatal death of the embryos (5Nagasawa T. Hirota S. Tachibana K. Takakura N. Nishikawa S. Kitamura Y. Yoshida N. Kikutani H. Kishimoto T. Nature. 1996; 382: 635-638Crossref PubMed Scopus (1999) Google Scholar). Apart from these physiological functions, SDF-1α has the selective capacity to inhibit cell entry of CXCR4-dependent human immunodeficiency viruses by occupying and internalizing CXCR4 in T lymphocytes (8Oberlin E. Amara A. Bachelerie F. Bessia C. Virelizier J.L. Arenzana-Seisdedos F. Schwartz O. Heard J.M. Clark-Lewis I. Legler D.F. Loetscher M. Baggiolini M. Moser B. Nature. 1996; 382: 833-835Crossref PubMed Scopus (1478) Google Scholar, 9Bleul C.C. Farzan M. Choe H. Parolin C. Clark-Lewis I. Sodroski J. Springer T.A. Nature. 1996; 382: 829-833Crossref PubMed Scopus (1749) Google Scholar, 11Amara A. Legall S. Schwartz O. Salamero J. Montes M. Loetscher P. Baggiolini M. Virelizier J.L. Arenzana-Seisdedos F. J. Exp. Med. 1997; 186: 139-146Crossref PubMed Scopus (518) Google Scholar, 12Signoret N. Oldridge J. Pelchenmatthews A. Klasse P.J. Tran T. Brass L.F. Rosenkilde M.M. Schwartz T.W. Holmes W. Dallas W. Luther M.A. Wells T.N.C. Hoxie J.A. Marsh M. J. Cell Biol. 1997; 139: 651-664Crossref PubMed Scopus (336) Google Scholar). Overall, these findings indicate that SDF-1α and its receptor, both of which are expressed widely outside the lympho-hematopoietic system, accomplish important additional functions that are not typical for chemokines. The biological activities of chemokines are thought to be influenced by their association with cellular or extracellular matrix glycosaminoglycans (GAG). Usually attached to a core protein to form proteoglycans (13Iozzo R.V. Annu. Rev. Biochem. 1998; 67: 609-652Crossref PubMed Scopus (1335) Google Scholar), GAG are highly sulfated oligosaccharides characterized by a high degree of structural heterogeneity. The common GAG are heparin, heparan sulfate (HS), dermatan sulfate, chondroitin sulfate, and hyaluronic acid (14Kjellen L. Lindahl U. Annu. Rev. Biochem. 1991; 60: 443-475Crossref PubMed Scopus (1673) Google Scholar, 15Hardingham T.E. Fosang A.J. FASEB J. 1992; 6: 861-870Crossref PubMed Scopus (1009) Google Scholar). The interaction of cytokines with proteoglycans in the extracellular matrix or cell surface have important functional consequences in many biological systems (16Gordon M.Y. Riley G.P. Watt S.M. Greaves M.F. Nature. 1987; 326: 403-405Crossref PubMed Scopus (456) Google Scholar, 17Spivak-Kroizman T. Lemmon M.A. Dikic I. Ladbury J.E. Pinchasi D. Huang J. Jaye M. Crumley G. Schlessinger J. Lax I. Cell. 1994; 79: 1015-1024Abstract Full Text PDF PubMed Scopus (591) Google Scholar). Although the in vivo biological roles of chemokine-GAG complexes are not clear, an increasing body of evidence suggests that GAG immobilize and enhance local concentrations of the chemokines, promoting their oligomerization and facilitating their presentation to the receptors (18Tanaka Y. Adams D.H. Shaw S. Immunol. Today. 1993; 14: 111-115Abstract Full Text PDF PubMed Scopus (382) Google Scholar, 19Hoogewerf A.J. Kuschert G.S. Proudfoot A.E. Borlat F. Clark-Lewis I. Power C.A. Wells T.N. Biochemistry. 1997; 36: 13570-13578Crossref PubMed Scopus (437) Google Scholar). Thus, it has been proposed that chemokines like macrophage inflammatory protein (MIP)-1β or interleukin-8 (IL-8) would be tethered to circulating leukocytes complexed to membrane-bound proteoglycans in endothelial cells (20Tanaka Y. Adams D.H. Hubscher S. Hirano H. Siebenlist U. Shaw S. Nature. 1993; 361: 79-82Crossref PubMed Scopus (844) Google Scholar, 21Middleton J. Neil S. Wintle J. Clark-Lewis I. Moore H. Lam C. Auer M. Hub E. Rot A. Cell. 1997; 91: 385-395Abstract Full Text Full Text PDF PubMed Scopus (616) Google Scholar). Although retention of SDF-1α in heparin affinity columns indicates that the chemokine has the capacity to complex with GAG (22Bleul C.C. Fuhlbrigge R.C. Casasnovas J.M. Aiuti A. Springer T.A. J. Exp. Med. 1996; 184: 1101-1109Crossref PubMed Scopus (1275) Google Scholar), it is not known whether SDF-1α is capable to interact with GAG under physiological conditions. Moreover, the nature of GAG is not known, and the structural determinants of the protein that eventually would account for such interactions remain unidentified. The aim of this work was to investigate the capacity of SDF-1α to form complexes with isolated or cell-bound GAG and to characterize the GAG family accounting for these interactions. On the other hand, we wanted to identify structural determinants of SDF-1α involved in the physical contact with GAG. Our findings demonstrate that SDF-1α interacts selectively and with relatively high affinity with HS in vitro. HS is also responsible for the binding of SDF-1α to CXCR4-negative epithelial or endothelial cells. Finally, we identified a cluster of basic residues in the first β-strand of the β-sheet of SDF-1α which is necessary for interaction with HS both in vitro and in intact cells. CHO-K1, CHO-pgsB 618, CHO-pgsD 677, Jurkat, CEM, CEMx174, ECV-304, and HeLa were obtained from the American Type Cell Collection. HS, dermatan sulfate, chondroitin sulfate A, and chondroitin sulfate B were obtained from Sigma (catalogue numbers H5393, C3788, C9819, and C0320, respectively). Heparinase (EC 4.2.2.7), heparitinase I (EC 4.2.2.8), and chondroitinase ABC (EC 4.2.2.4) were purchased from Seikagaku Corp. Heparin was from Sanofi Recherche. An upgraded BIAcore system, F1 sensorchips, amine coupling kit, and HBS (10 mm Hepes, 150 mm NaCl, 3.4 mmEDTA, 0.005% surfactant P20, pH 7.4) were obtained from BiAcore AB. Biotin-LC-hydrazide was from Pierce, and streptavidin was obtained from Sigma. SDF-1α 2-67, 3-67, and, 4-67, regulated on activation normal T cell expressed and secreted (RANTES), and MIP-1α and MIP-1β were a gift from Dr. Ian Clark-Lewis (British Columbia University, Vancouver, British Columbia, Canada). Wild type SDF-1α (SDF-1α) and SDF-1 3/6 (substitution of Lys24, His25, and Lys27 by Ser24, Ser25, and Ser27) were synthesized by the Merrifield solid-phase method on a fully automated peptide synthesizer (Pioneer, Perspective Biosystems, and Perkin-Elmer) using fluorenylmethyloxycarbonyl (Fmoc) chemistry. All amino acids were double coupled with O-(7-azabenzotriazol-1-yl)-1,1,3,3-tetramethyluroniumhexafluorophosphate/DIEA activation, and piperidine Fmoc deprotection was optimized by elongating the standard deprotection time. After completion of the synthesis, the polypeptide was released from the resin and precipitated in cold diethyl ether. The precipitate was dissolved in aqueous 0.08% trifluoroacetic acid and lyophilized. The crude polypeptide was dissolved in 6 m guanidine hydrochloride, 0.1 mTris acetate, pH 8.5, and 16% 2-mercaptoethanol, stirred at 37 °C for 2 h, and then acidified to pH 4. The reduced chemokine was purified on a preparative medium pressure liquid chromatography column (313 × 26 mm) packed with C18 100-Å and 20-μm Nucleoprep packing (Macherey-Nagel GmbH & Co, Düren, Germany) using a 20–80% linear gradient of acetonitrile in 0.08% aqueous trifluoroacetic acid over 120 min at a 25 ml/min flow rate. After lyophilization, the purified, reduced chemokine was solubilized in 6m guanidine hydrochloride, 0.1 m Tris acetate, pH 8.5, and rapidly diluted into 0.1 m Tris acetate buffer, pH 8.5. Final concentration of the chemokine was 0.4 mg/ml in 1m guanidine hydrochloride, 0.1 m Tris acetate, pH 8.5. The solution was stirred overnight to allow chemokine folding and gentle air oxidation of the four cysteines. The folded chemokine was purified using the medium pressure liquid chromatography purification procedure described above. Final purity of SDF-1α and SDF-1 3/6 was superior to 95% as judged by high pressure liquid chromatography. The average molecular weights determined by ion spray mass spectrometry were 7830.9 ± 0.5 (theoretic molecular weight, 7831.3) for SDF-1α and 7699.0 ± 0.5 (theoretic molecular weight, 7699.0) for SDF-1 3/6. SDF 1–34 was synthesized using the same procedure. Cysteine in position 11 was replaced by alanine. Cyclization of the reduced half-chemokine was done by H2O2at pH 7 (23Kudryavtseva E.V. Sidorova M.V. Ovchinnikov M.V. Bespalova Z.D. Bushuev V.N. J. Pept. Res. 1997; 49: 52-58Crossref PubMed Scopus (31) Google Scholar), and the cyclic peptide was purified under the same conditions as described above. Ion spray mass spectroscopy showed a molecular weight of 3910.8 ± 1 (theoretic molecular weight, 3910.6). The concentration of each chemokine or derivative was determined by amino acid analysis on a 6300 Beckman amino acid analyzer after hydrolysis for 20 h in 6 n HCl, 0.2% phenol in the presence of a known amount of norleucine as internal standard. All chemicals for the synthesis were purchased from Perspective Biosystems and Perkin-Elmer, France. The anti-SDF-1 monoclonal antibody (mAb) K15C (IgG2a κ) was generated by immunizing BALB/c mice with the SDF-1-derived peptide KPVSLSYRSPSRFFC conjugated via cysteine 15 to bovine serum albumin. Fusions were carried out as described previously (24Tanchou V. Delaunay T. Bodeus M. Roques B. Darlix J.L. Benarous R. J. Gen. Virol. 1995; 76: 2457-2466Crossref PubMed Scopus (12) Google Scholar). mAb was purified from bulk culture by affinity chromatography on a protein A-Sepharose column. Anti-SDF-1 K15C specificity was evaluated by an enzyme-linked immunosorbent assay. Briefly, SDF-1α coated on 96-well plates was incubated for 2 h at 37 °C with K15C mAb (0.05 μg/ml) previously preincubated overnight at 4 °C with various concentrations (10−12 to 10−5m) of SDF-1α or irrelevant chemokines (MIP-1α, MIP-1β, and RANTES) diluted in PBS, 0.1% Tween 20, 0.5% bovine serum albumin. K15C·SDF-1α complexes were revealed with a 5000-fold diluted horseradish peroxidase-labeled goat anti-mouse immunoglobulins G (H + L) (Diagnostic Pasteur, Marnes la Coquettes, France) and ortho-phenylenediamine (Sigma, 2% in 0.05 mphosphate/citrate buffer, pH 5.0) and 0.015% H2O2. The reaction was stopped with 50 μl of 4 m H2SO4, and optical density at 492 nm was measured in a spectrophotometer. K15C mAb specificity was finally investigated by Western blot analysis. Fifty ng of SDF-1α, SDF-1β, or SDF-1α proteins with progressive deletions at the amino terminus (SDF 2–67, SDF 3–67, and SDF 4–67) were separated by electrophoresis on 20% SDS-polyacrylamide gels (PAGE), blotted onto nitrocellulose membranes, and probed with the K15C mAb (1 μg/ml). The binding of the antibody was revealed using an enhanced chemiluminescence assay (ECL kit, Amersham Pharmacia Biotech, Les Ulis, France). The ability of K15C mAb to block CXCR4 endocytosis induced by SDF-1α was also investigated. Jurkat cells (5 × 105cells/sample) were incubated with 10 nm SDF-1α or 10 nm SDF-1α previously neutralized with either 30 μg/ml K15C mAb or an equal amount of an irrelevant, isotype-matched, mouse immunoglobulin. Down-regulation of CXCR4 expression was evaluated as described previously (11Amara A. Legall S. Schwartz O. Salamero J. Montes M. Loetscher P. Baggiolini M. Virelizier J.L. Arenzana-Seisdedos F. J. Exp. Med. 1997; 186: 139-146Crossref PubMed Scopus (518) Google Scholar). The anti-CXCR4 6H8 (IgG1κ) mAb used in this work was obtained by immunizing BALB/c mice with a peptide spanning the first 28 amino-terminal amino acids of CXCR4 conjugated via cysteine 28 to ovalbumin (25Mondor I. Moulard M. Ugolini S. Klasse P.J. Hoxie J. Amara A. Delaunay T. Wyatt R. Sodroski J. Sattentau Q.J. Virology. 1998; 248: 394-405Crossref PubMed Scopus (69) Google Scholar). 6H8 was screened by flow cytometry analysis of CEM cells and was shown to bind to human peripheral blood lymphocytes and human cell lines (Jurkat, U937, THP-1, HeLa, and SupT1) that constitutively express CXCR4. 6H8 reacts specifically with CXCR4-transfected CHO-K1 cells but not with CHO-K1 cells transfected with CCR5, CCR3, CCR1, or CCR2b (data not shown). The specificity of CXCR4 detection by 6H8 mAb was compared with that obtained using the well characterized 12G5 anti-CXCR4-mAb. Both antibodies exhibit identical patterns of recognition and specificity for a large panel of human or animal cells expressing CXCR4 constitutively or upon transfection. Adherent cells were plated 2 days before binding experiments. Cells were detached with 2 mm EDTA in PBS and washed twice with ice-cold binding buffer (RPMI 1640, 20 mmHepes, 1% bovine serum albumin). 4 × 105 cells were resuspended in the presence of the indicated concentration of chemokines in a total volume of 200 μl and incubated for 90 min at 4 °C under agitation. Unbound chemokine was removed by washing with binding buffer, and cell-bound SDF-1α was detected by incubation with the anti-SDF-1 mAb K15C (15 μg/ml, diluted in PBS, 1% bovine serum albumin). After staining with phycoerythrin-conjugated anti-mouse immunoglobulins (Southern Biotechnology), cells were fixed in 1% formaldehyde buffer and analyzed in a FACSscan (Becton Dickinson, CA). To remove cell-surface GAG, CHO-K1 and HeLa cells (106cells) were washed twice and incubated for 90 min at 37 °C with 1 milliunit/ml GAG-degrading enzymes. For trypsin treatment, cells were washed once with PBS containing 2 mm EDTA and incubated for 4 min at 37 °C with 0.25% trypsin in PBS. After enzyme treatment, cells were washed four times with 5 ml of binding buffer, detached with PBS/EDTA, and then assayed for SDF-1α binding as described above. Fractionated heparin (9 kDa) resuspended in PBS at 1 mm was reacted for 24 h at room temperature with 10 mm biotin/LC-hydrazine. The mixture was then extensively dialyzed against water to remove unreacted biotin and freeze-dried. Two flow cells of an F1 sensorchip were activated with 50 μl of a mixture of 0.2 m EDC, 0.05 m NHS before injection of 50 μl of streptavidin (0.2 mg/ml in 10 mm acetate buffer, pH 4.2). Remaining activated groups were blocked with 50 μl of 1m ethanolamine, pH 8.5. Typically, this procedure allowed coupling of approximately 2000–2500 resonance units (RU) of streptavidin. Biotinylated heparin (5–10 μg/ml in HBS containing 0.3m NaCl) was then injected on one of the two streptavidin surfaces (the other one being a negative control). Approximately 50 RU of material was immobilized. Both flow cells were then conditioned with several injections of 1.5 m NaCl. The conversion of RU to surface concentration of proteins was performed using a conversion factor of 1000 RU = 1 ng/mm2. Test samples were diluted in HBS maintained at 25 °C and injected over the heparin surface at a flow rate of 50 μl/min. This high flow rate was necessary to eliminate mass transport effect due to the relatively high association rate of the proteins being studied. In a typical analysis, six different SDF-1α concentrations (usually ranging between 0 and 200 nm) were injected onto the heparin surface for 3 min (to study the association phase and equilibrium). Thereafter, the formed complexes were washed at 50 μl/min with HBS to study the dissociation phase. The sensorchip surface was regenerated with a 2-min pulse of 1.5 m NaCl in HBS. In some experiments, soluble heparin was included in the buffer during the dissociation phase to minimize possible rebinding effects. Kinetic constants were derived from the sensorgrams by fitting the data to different interaction models, using BIAevaluation software, essentially as described (26Sadir R. Forest E. Lortat-Jacob H. J. Biol. Chem. 1998; 273: 10919-10925Abstract Full Text Full Text PDF PubMed Scopus (91) Google Scholar). Affinities (dissociation equilibrium constants: Kd) were calculated from the ratio of dissociation and association rate constants (Kd =koff/kon). CEM cells (5 × 106 cells/ml) were incubated in PBS with 0.25 nm iodinated SDF-1α (New England Nuclear, specific activity, 2200 Ci/mmol) and various concentrations of unlabeled SDF-1α or SDF-1 3/6 for 1 h at 4 °C in a final volume of 300 μl. Incubations were terminated by centrifugation at 4 °C. The cell pellets were washed twice in ice-cold PBS. Nonspecific binding was determined in the presence of 1 μm unlabeled SDF-1α. Cell pellet-associated radioactivity was counted using a LKB-Wallac microcomputer controlled 1272 CliniGamma counter. The binding data were analyzed using a GraphPrad Prism 2.0 software. CEMx174 cells were resuspended in chemotaxis medium (RPMI 1640 containing 1% pasteurized human plasma protein (Swiss Red Cross Laboratory) and buffered with 20 mm Hepes, pH 7.3). Cell migration was performed in 48-well chemotaxis chambers (Neuro Probe Inc., Cabin John, MD) as described previously (27Loetscher P. Seitz M. Baggiolini M. Moser B. J. Exp. Med. 1996; 184: 569-777Crossref PubMed Scopus (394) Google Scholar). Chemokines diluted in chemotaxis medium were added to the lower and 105 CEMx174 cells in the same medium to the upper wells. Polycarbonate membranes with 5-μm pores (PC 5 μm PVPF, Costar, Cambridge, MA) were used to measure cell migration for 2 h at 37 °C. Membranes were removed, and the upper side was washed with PBS, fixed, and stained. Cell migration was assessed at 1000× magnification in five randomly selected fields. For calcium measurements, CEMx174 cells were washed twice with PBS and resuspended (2 × 106/ml) in 20 mm Hepes, 136 mm NaCl, 4.6 mm KCl, 1 mmCaCl2, pH 7.4. Aliquots of 1 ml were loaded with 0.8 μm Fura2/AM (Fluka AG, Buchs, Switzerland) for 20 min at 37 °C. The cells were sedimented and resuspended in fresh medium (1.25 × 106/ml). Calcium mobilization was determined as described previously (8Oberlin E. Amara A. Bachelerie F. Bessia C. Virelizier J.L. Arenzana-Seisdedos F. Schwartz O. Heard J.M. Clark-Lewis I. Legler D.F. Loetscher M. Baggiolini M. Moser B. Nature. 1996; 382: 833-835Crossref PubMed Scopus (1478) Google Scholar). Single-cell Ca2+ measurements were performed in HeLa cells loaded with Fura-2 (3 μm) as described previously (11Amara A. Legall S. Schwartz O. Salamero J. Montes M. Loetscher P. Baggiolini M. Virelizier J.L. Arenzana-Seisdedos F. J. Exp. Med. 1997; 186: 139-146Crossref PubMed Scopus (518) Google Scholar). NMR experiments were acquired at 35 °C on a Varian Unity 500 spectrometer operating at 499.84 MHz for1H and equipped with a triple resonance z gradient 5-mm probe. Data were processed on a Sun workstation using the VNMR 5.3 program. All two-dimensional proton NMR experiments were acquired in the phase-sensitive mode using the hypercomplex scheme (28States D.J. Haberkorn R.A. Ruben D.J. J. Magn. Reson. 1982; 48: 286-292Google Scholar). A two-dimensional nuclear Overhauser experiment with pure absorption phase was performed in four quadrants. The SDF-1 3/6 sample was prepared by dissolving 6 mg of freeze-dried powder in 350 μl of 20 mm acetate buffer in 90% H2O/10% D2O, pH 5. The final concentration was 2.1 mm. The TOCSY and NOESY experiments were collected with mixing times of 80 and 200 ms, respectively, and with 512 t1increments of 32 scans each and 2048 points in the t2 dimension. In both dimensions, the data were apodized with a shifted sine-bell function before Fourier transformation and zero-filled to a final matrix of 4096 × 2048 points. The capacity of SDF-1α to bind cell membrane molecules other than CXCR4 was investigated using a novel mAb. The K15C mAb was obtained by immunizing mice with a linear peptide derived from the amino terminus of SDF-1α. In enzyme-linked immunosorbent assay experiments, the antibody showed specific reactivity with immobilized SDF-1α (Fig.1 a) as competition occurred when the antibodies were preincubated with free SDF-1α but not with the non-related chemokines RANTES, MIP-1α, or MIP-1β. Apart from SDF-1α or SDF-1β (Fig. 1 b) which differ exclusively by four amino acids at the carboxyl terminus domain, the antibody failed to recognize any other known CC or CXC chemokine (data not shown). To characterize further the specificity of epitope recognition by K15C mAb, we performed Western blot experiments using SDF-1α derivatives with progressive deletion of amino acids at the amino terminus of SDF-1α. SDF-1α derivatives lacking the two first residues (Lys1 and Pro2) (29Crump M.P. Gong J.H. Loetscher P. Rajarathnam K. Amara A. Arenzana-Seisdedos F. Virelizier J.L. Baggiolini M. Sykes B.D. Clark-Lewis I. EMBO J. 1997; 16: 6996-7007Crossref PubMed Scopus (631) Google Scholar) were not recognized by the antibody. This proves that Lys1 and Pro2 are essential residues in the epitope recognized by K15C mAb (Fig.1 b). Importantly, incubation of K15C mAb fully prevented the biological activity of SDF-1α as assessed by its capacity to block SDF-1α-mediated CXCR4 endocytosis in Jurkat cells (Fig.1 c) or chemotaxis of human T lymphocytes (data not shown). CXCR4-negative CHO-K1 cells (Fig.2 b) were incubated with 10–1000 nm SDF-1α and labeled with the K15C mAb. The cytofluorographic analysis showed that SDF-1α bound to the cell surface in a concentration-dependent manner (Fig.2 a). The recognition of the SDF-1α·K15C complex by the secondary antibody was specific since no fluorescence was observed when SDF-1α was omitted or replaced by MIP-1β (1000 nm) (Fig. 2 a). The capacity of SDF-1α to bind to cells in a CXCR4-independent manner is not restricted to a particular cell type. Indeed, SDF-1α also bound to CXCR4-negative, endothelial cells (ECV304 cell line, Fig.2 b) with a comparable efficiency as to CHO-K1. Similarly, cell membrane-bound SDF-1α was found when the chemokine was incubated with HeLa cells that constitutively express CXCR4 (Fig. 2 b). SDF-1α binding detected by FACS analysis on HeLa cells was not due to interaction of the chemokine with CXCR4 because the K15C mAb exclusively recognizes the critical amino-terminal residues that are engaged in binding to CXCR4 and are required for SDF-1α-induced signal transduction (29Crump M.P. Gong J.H. Loetscher P. Rajarathnam K. Amara A. Arenzana-Seisdedos F. Virelizier J.L. Baggiolini M. Sykes B.D. Clark-Lewis I. EMBO J. 1997; 16: 6996-7007Crossref PubMed Scopus (631) Google Scholar). Accordingly, we failed to detect SDF-1α bound to CXCR4-positive, CEM T lymphoblastoid cells (Fig.2 b). CEM cells express high levels of functional CXCR4 receptors, and saturable binding of SDF-1α in these cells has been shown (29Crump M.P. Gong J.H. Loetscher P. Rajarathnam K. Amara A. Arenzana-Seisdedos F. Virelizier J.L. Baggiolini M. Sykes B.D. Clark-Lewis I. EMBO J. 1997; 16: 6996-7007Crossref PubMed Scopus (631) Google Scholar). This suggests that in CEM cells, CXCR4 accounts for most of the cellular binding of SDF-1α. Collectively, our results indicate that other structures apart from CXCR4 have the capacity to attach SDF-1α to the cell surface. These interactions apparently do not involve the amino terminus of SDF-1α that is recognized by the K15C mAb and is masked after interaction with CXCR4. To ascertain wheth
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