FRINK Linked Data Fragments server

Matches in SemOpenAlex for { <https://semopenalex.org/work/W2168296320> ?p ?o ?g. }

• W2168296320 endingPage "30524" @default.
• W2168296320 startingPage "30516" @default.
• W2168296320 abstract "Angiogenesis depends on proper collagen biosynthesis and cross-linking, and type I collagen is an ideal angiogenic scaffold, although its mechanism is unknown. We examined angiogenesis using an assay wherein confluent monolayers of human umbilical vein endothelial cells were overlain with collagen in a serum-free defined medium. Small spaces formed in the cell layer by 2 h, and cells formed net-like arrays by 6–8 h and capillary-like lumens by 24 h. Blocking of α2β1, but not α1 or αvβ3 integrin function halted morphogenesis. We found that a triple-helical, homotrimeric peptide mimetic of a putative α2β1 binding site: α1(I)496–507 GARGERGFP*GER (where single-letter amino acid nomenclature is used, P* = hydroxyproline) inhibited tube formation, whereas a peptide carrying another putative site: α1(I)127–138 GLP*GERGRP*GAP* or control peptides did not. A chemical inhibitor of p38 mitogen-activated protein kinase (p38 MAPK), SB202190, blocked tube formation, and p38 MAPK activity was increased in collagen-treated cultures, whereas targeting MAPK kinase (MEK), focal adhesion kinase (FAK), or phosphatidylinositol 3-kinase (PI3K) had little effect. Collagen-treated cells had fewer focal adhesions and 3- to 5-fold less activated FAK. Thus capillary morphogenesis requires endothelial α2β1 integrin engagement of a single type I collagen integrin-binding site, possibly signaling via p38 MAPK and focal adhesion disassembly/FAK inactivation. Angiogenesis depends on proper collagen biosynthesis and cross-linking, and type I collagen is an ideal angiogenic scaffold, although its mechanism is unknown. We examined angiogenesis using an assay wherein confluent monolayers of human umbilical vein endothelial cells were overlain with collagen in a serum-free defined medium. Small spaces formed in the cell layer by 2 h, and cells formed net-like arrays by 6–8 h and capillary-like lumens by 24 h. Blocking of α2β1, but not α1 or αvβ3 integrin function halted morphogenesis. We found that a triple-helical, homotrimeric peptide mimetic of a putative α2β1 binding site: α1(I)496–507 GARGERGFP*GER (where single-letter amino acid nomenclature is used, P* = hydroxyproline) inhibited tube formation, whereas a peptide carrying another putative site: α1(I)127–138 GLP*GERGRP*GAP* or control peptides did not. A chemical inhibitor of p38 mitogen-activated protein kinase (p38 MAPK), SB202190, blocked tube formation, and p38 MAPK activity was increased in collagen-treated cultures, whereas targeting MAPK kinase (MEK), focal adhesion kinase (FAK), or phosphatidylinositol 3-kinase (PI3K) had little effect. Collagen-treated cells had fewer focal adhesions and 3- to 5-fold less activated FAK. Thus capillary morphogenesis requires endothelial α2β1 integrin engagement of a single type I collagen integrin-binding site, possibly signaling via p38 MAPK and focal adhesion disassembly/FAK inactivation. In adult mammals, angiogenesis, the growth of new capillaries from the existing vasculature, is the exclusive mechanism by which new vessels are formed, and is involved in normal homeostasis as well as in various diseases, including growth and metastasis of solid tumors, rheumatoid arthritis, and diabetic retinopathy (1Folkman J. Nat. Med. 1995; 1: 27-31Crossref PubMed Scopus (7189) Google Scholar). It is a complex process by which endothelial cells degrade their matrix, migrate, proliferate, and differentiate into new vessels (2Strömblad S. Cheresh D.A. Trends Cell Biol. 1996; 6: 462-467Abstract Full Text PDF PubMed Scopus (168) Google Scholar). Angiogenesis depends upon the interaction of endothelial cells with extracellular matrix proteins via cell adhesion molecules, and the activities of growth factors and cytokines (3Eliceiri B.P. Cheresh D.A. J. Clin. Invest. 1999; 103: 1227-1230Crossref PubMed Scopus (616) Google Scholar). Because type I collagen is a ubiquitous component of many tissues that undergo angiogenesis during embryonic development, it may play a role in promoting blood vessel development and contribute to pathological angiogenesis. In fact, type I collagen is among the most ideal scaffolds for the induction of angiogenesis in vitro. Thus, bovine aortic endothelial cells synthesize type I collagen (4Cotta-Pereira G. Sage H. Bornstein P. Ross R. Schwartz S. J. Cell Physiol. 1980; 102: 183-191Crossref PubMed Scopus (72) Google Scholar, 5Levene C.I. Bartlet C. Heale G. Atherosclerosis. 1984; 52: 59-71Abstract Full Text PDF PubMed Scopus (10) Google Scholar), and its expression precedes angiogenesis and is limited to the vicinity of cells forming capillary tubes in endothelial cell monolayers (6Iruela-Arispe M.L. Hasselaar P. Sage H. Lab. Invest. 1991; 64: 174-186PubMed Google Scholar, 7Fouser L. Iruela-Arispe L. Bornstein P. Sage E.H. J. Biol. Chem. 1991; 266: 18345-18351Abstract Full Text PDF PubMed Google Scholar). Furthermore, endothelial cells grown between or dispersed within collagen gels form branching networks of tubes (8Nicosia R.F. Ottinetti A. Lab. Invest. 1990; 63: 115-122PubMed Google Scholar, 9Montesano R. Orci L. Vassalli P. J. Cell Biol. 1983; 97: 1648-1652Crossref PubMed Scopus (507) Google Scholar, 10Ilan N. Mahooti S. Madri J.A. J. Cell Sci. 1998; 111: 3621-3631Crossref PubMed Google Scholar), and in HUVEC 1The abbreviations used are: HUVECs, human umbilical vein endothelial cells; EBM, endothelial cell basal medium; ERK, extracellular regulated protein kinase; FAK, focal adhesion kinase; MAPK, mitogen-activated protein kinase; MEK, MAPK kinase; MAPKAP, MAPK-activated protein; PI3K, phosphatidyl inositol 3-kinase; SFDM, serum-free defined medium; SPR, surface plasmon resonance; SSP, single stranded peptide; THP, triple helical peptide; VEGF, vascular endothelial growth factor; PMA, phorbol 12-myristate 13-acetate; FGF-2, fibroblast growth factor-2; BSA, bovine serum albumin; PBS, phosphate-buffered saline; TBS, Tris-buffered saline.1The abbreviations used are: HUVECs, human umbilical vein endothelial cells; EBM, endothelial cell basal medium; ERK, extracellular regulated protein kinase; FAK, focal adhesion kinase; MAPK, mitogen-activated protein kinase; MEK, MAPK kinase; MAPKAP, MAPK-activated protein; PI3K, phosphatidyl inositol 3-kinase; SFDM, serum-free defined medium; SPR, surface plasmon resonance; SSP, single stranded peptide; THP, triple helical peptide; VEGF, vascular endothelial growth factor; PMA, phorbol 12-myristate 13-acetate; FGF-2, fibroblast growth factor-2; BSA, bovine serum albumin; PBS, phosphate-buffered saline; TBS, Tris-buffered saline. monolayers, angiogenesis rapidly proceeds in the presence of type I collagen and sulfated glycosaminoglycans (11Jackson R.L. Busch S.J. Cardin A.D. Physiol. Rev. 1991; 71: 481-539Crossref PubMed Scopus (958) Google Scholar, 12Jackson C.J. Giles I. Knop A. Nethery A. Schrieber L. Exp. Cell Res. 1994; 215: 294-302Crossref PubMed Scopus (21) Google Scholar, 13Sweeney S.M. Guy C.A. Fields G.B. San Antonio J.D. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 7275-7280Crossref PubMed Scopus (88) Google Scholar). In vivo, angiogenesis is disrupted in the chick embryo by inhibiting collagen triple helix formation or fibrillogenesis using α,α-dipyridyl or β-aminopropionitrile, respectively (14Ingber D.E. Folkman J. Lab. Invest. 1988; 59: 44-51PubMed Google Scholar). Therefore, a definitive role for collagen exists, but the mechanisms by which it exerts its pro-angiogenic effect remain elusive. Roles for several intracellular signaling pathways are proposed for collagen-induced angiogenesis. Thus, in long term collagen gel cultures, the ERK1/2 and p38 MAPK pathways are implicated in mediating angiogenesis of the endothelial cell line MSS31, although their relative contributions to specific phases of angiogenesis are unknown (15Tanaka K. Abe M. Sato Y. Jpn. J. Cancer Res. 1999; 90: 647-654Crossref PubMed Scopus (120) Google Scholar). It is of interest that in mice null for p38α MAPK, primary angiogenesis proceeds normally, but remodeling of the capillary plexus and placental angiogenesis are defective (16Mudgett J.S. Ding J. Guh-Siesel L. Chartrain N.A. Yang L. Gopal S. Shen M.M. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 10454-10459Crossref PubMed Scopus (325) Google Scholar), whereas lack of Mekk3, the p38 MAPK upstream activator, causes deficiency in both primary as well as placental angiogenesis (17Yang J. Boerm M. McCarty M. Bucana C. Fidler I.J. Zhuang Y. Su B. Nat. Genet. 2000; 24: 309-313Crossref PubMed Scopus (192) Google Scholar). Additionally, links between the collagen-associated integrins and p38 MAPK have been established, albeit in a non-angiogenic context (18Ivaska J. Reunanen H. Westermarck J. Koivisto L. Kahari V.M. Heino J. J. Cell Biol. 1999; 147: 401-416Crossref PubMed Scopus (191) Google Scholar, 19Klekotka P.A. Santoro S.A. Zutter M.M. J. Biol. Chem. 2001; 276: 9503-9511Abstract Full Text Full Text PDF PubMed Scopus (82) Google Scholar). In three-dimensional collagen gels, PMA treatment induces HUVEC survival and tube formation by activating MAPK, PI3K, and Akt/protein kinase B pathways, which are postulated to act by up-regulating VEGF receptor expression and cell-cell adhesion molecules such as vascular endothelial cadherin (10Ilan N. Mahooti S. Madri J.A. J. Cell Sci. 1998; 111: 3621-3631Crossref PubMed Google Scholar). Within other matrix scaffolds, FGF-2-dependent angiogenesis correlates with a sustained, αVβ3 integrin-dependent wave of ERK activity, necessary for endothelial cell migration (20Eliceiri B.P. Klemke R. Stromblad S. Cheresh D.A. J. Cell Biol. 1998; 140: 1255-1263Crossref PubMed Scopus (362) Google Scholar), whereas VEGF promotes angiogenesis, but requires src kinase and ERK activities (21Eliceiri B.P. Paul R. Schwartzberg P.L. Hood J.D. Leng J. Cheresh D.A. Mol. Cell. 1999; 4: 915-924Abstract Full Text Full Text PDF PubMed Scopus (663) Google Scholar). In vivo, type I collagen implants induce angiogenesis and require matrix metalloproteinase activity (22Seandel M. Noack-Kunnmann K. Zhu D. Aimes R.T. Quigley J.P. Blood. 2001; 97: 2323-2332Crossref PubMed Scopus (132) Google Scholar); in the case of αVβ3 integrin-mediated angiogenesis, matrix metalloproteinase 2 association is also necessary and proposed to facilitate cell migration (23Silletti S. Kessler T. Goldberg J. Boger D.L. Cheresh D.A. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 119-124PubMed Google Scholar). Thus, we seek to identify the endothelial integrin receptors used, the determinants in the type I collagen fibril to which they bind, and the intracellular signaling pathways employed during tube formation in type I collagen. Here we developed an in vitro model representing the capillary tube formation phase of angiogenesis, using a novel SFDM containing only BSA, FGF-2, and VEGF. We found that α2β1 integrins and a single integrin binding site on type I collagen are essential for tube formation and identified potential roles for p38 MAPK and FAK. Reagents—Cell culture reagents were from Invitrogen. Endothelial cell basal media (EBM) was from Clonetics. Unless specified, all other materials were from Fisher Scientific. Signaling pathway inhibitors SB202190, SB203580, LY294002, PD98059, Ro318220, and calphostin C were from Calbiochem, and geldanamycin was from Sigma. Monoclonal antibodies to α1 (MAB1973Z, clone FB12), α2β1 (MAB1998, clone BHA2.1), αVβ3 (MAB1976Z, clone LM609), β1 (MAB1959, clone P5D2), purified integrin receptors α1β1, αVβ3, control mouse and fluorescein-tagged IgG were from Chemicon. Antibodies to phospho-FAK (Y397) and p38 MAPK (T180/Y182) were from Upstate Biotechnology and Cell Signaling, respectively. Endothelial cell growth supplement was isolated from bovine hypothalami as described previously (24Maciag T. Cerundolo J. Ilsley S. Kelley P.R. Forand R. Proc. Natl. Acad. Sci. U. S. A. 1979; 76: 5674-5678Crossref PubMed Scopus (577) Google Scholar). CM5 chips were from Biacore. Recombinant α2 I domain was a generous gift from Dr. Santoro (University of Washington, St. Louis, MO). Type I Collagen Isolation—Collagen was isolated from rat tail tendons as previously described and solubilized in 10 mm acetic acid overnight with agitation at 4 °C prior to use (25San Antonio J.D. Lander A.D. Wright T.C. Karnovsky M.J. J. Cell. Physiol. 1992; 150: 8-16Crossref PubMed Scopus (46) Google Scholar). HUVEC Isolation—Umbilical cords were from the Obstetrics Department, Thomas Jefferson University. Cells were isolated as described and used through passage six (26Gimbrone M.A. Collen B.S. Progress in Hematology and Thrombosis. Vol. III. W. B. Saunders, Philadelphia1976: 1-28Google Scholar). In Vitro Angiogenesis Assays—Our assay was adapted from that of Montesano et al. (9Montesano R. Orci L. Vassalli P. J. Cell Biol. 1983; 97: 1648-1652Crossref PubMed Scopus (507) Google Scholar). 6- and 12-well cell culture dishes (Falcon) were coated with 100 μg/ml rat-tail type I collagen in sterile 10 mm acetic acid (10.5 μg/cm2) overnight at 4 °C. Plates were rinsed twice in warm, sterile PBS+, and cells were plated at 105 cells/cm2 in complete media. Alternatively, suspended cells were rinsed twice with SFDM lacking growth factors and plated at 105 cells/cm2 in SFDM, incubated overnight at 37 °C, 5% CO2 in a humidified atmosphere. The next day wells were rinsed with PBS+ and given 80–100 μl/cm2 of a collagen gel. Gels were prepared by mixing seven parts of a 1.4 mg/ml type I collagen stock in sterile 10 mm acetic acid with one part of 10× BME or Medium 199 and two parts of an 11.8 mg/ml sodium bicarbonate stock on ice. Cultures were incubated as above until gels had solidified for ∼15 min then given 0.6 ml of media with test agents to yield a final volume of 1.0 ml. In gel removal experiments, we used a sterile scalpel to cut along the circumference of the gel then pipetted the medium under the gel to dislodge it. Finally, gels and media were aspirated, and media were replaced. Geldanamycin was dissolved in Medium 199, and all other inhibitors were dissolved in Me2SO. Inhibitors were diluted in SFDM and added after gel formation. Transmission Electron Microscopy of Cultures—Methods were as detailed previously (27Iozzo R.V. J. Cell Biol. 1984; 99: 403-417Crossref PubMed Scopus (103) Google Scholar). At 24 h, embedded control monolayers and collagen-treated cultures were cut on an Ultracut S ultramicrotome, with the plane of sectioning perpendicular to the monolayer. Analysis was performed on a JEM-100CX II, and images were imported into Adobe Photoshop 6.0. Quantitation of Angiogenesis—Endothelial tube morphogenesis was examined by phase contrast microscopy and assessed quantitatively. Photographs taken using a FUJI X HC-300Z digital camera were adjusted to the same numerical values using the levels function of Adobe Photoshop 5.5. Typical quantitation methods count the area occupied by or numbers of tube-like structures, making the uncertain assumption that all structures counted are bona fide tubes. We used a more objective measurement of HUVEC network formation, i.e. the formation of tube-like structures and their intervening spaces, as the total empty area in a given culture. Area measurements were performed using Image 1.62 (National Institutes of Health). All assays were repeated at least three times with at least two wells per treatment group. In some experiments the progression of morphogenesis was also quantitated by measuring the width of cellular processes at the midpoint, as a measure of the stage of tube formation, i.e. well-formed tubes have narrow diameters, whereas poorly or partially formed tubes do not. p values were generated using the Student's t test. Fluorescent Staining of Angiogenic Cultures—100 μg of collagen I was dried onto permanox 2-well chamber slides overnight and stored at 4 °C prior to use. Slides were rehydrated with warm PBS for 5 min, before plating 105 HUVEC/cm2 in SFDM; gels were prepared as outlined above. After 30 min or 1 h of collagen gel exposure, both the monolayer cultures, and the treated cultures (following gel removal) were rinsed in PBS, fixed in 4% paraformaldehyde for 10 min, rinsed in TBS, permeabilized for 5 min with TBS/0.1% triton X 100, and blocked with TBS/1% BSA for 30 min. Cells were incubated with 1:100 anti-phospho-FAK in TBS for 1 h. In some cases, cells were dual-stained with 1:200 anti-human vinculin (hVIN1, Sigma) or 1:50 anti-talin (Research Diagnostics, Inc.). Following incubation with primary antibodies, cells were rinsed, incubated with the appropriate secondary antibodies, and rinsed again; coverslips were glycerol-mounted. Photographs were taken using Kodak 1600 color slide film and an Olympus BX50 upright microscope. Analysis of the Phosphorylation State of Focal Adhesion Kinase—Two 12-well plates were set up for angiogenesis assays per time point; collagen gels were prepared as outlined above. One plate received collagen while the other SFDM; following polymerization, plates were extracted using 40 μl (10 μl/cm2) of 1× SDS loading buffer at various times after collagen addition. Gels were removed as outlined above, and cells were rinsed twice with cold PBS prior to extraction. Equivalent microgram quantities of total lysate were loaded and run on 12% pre-cast polyacrylamide gels (ICN Biomedicals). Proteins were then transferred onto activated polyvinylidene difluoride membranes, blocked with 5% nonfat dry milk, and incubated with primary antibody, 1:500 phospho-FAK, 4 h at room temperature, and 1:250 fluorescein isothiocyanate-conjugated secondary antibody, 1 h at 25 °C. Blots were visualized using a Molecular Dynamics Fluorimager. Relative levels of phosphorylation were determined using the integrate volume and local average background subtraction functions of ImageQuant software. Analysis of p38 MAPK Activity—We determined relative p38 MAPK activity in cultures using a non-radioactive assay (Cell Signaling). Two 12-well plates were set up for angiogenesis assays as outlined above; one plate received collagen while the other SFDM. At various times following the addition of the collagen, gels and media were removed, cells rinsed twice with PBS, and extracted with 40 μl of lysis buffer containing 2 μm phenylmethylsulfonyl fluoride per well. Extracts were clarified at 14,000 × g/10 min/4 °C. Supernatants were immunoprecipitated overnight at 4 °C using an immobilized anti-phospho-p38 MAPK antibody (T180/Y182). On the following day, pellets were rinsed in lysis and kinase buffers then given 2 μg of recombinant ATF-2 and 10 μm ATP at 30 °C/30 min. Reactions were quenched using SDS loading buffer, run on 10% PAGE gels (ICN), transferred to polyvinylidene difluoride membranes, blocked in 5% nonfat milk in TBS, then incubated with 1:1000 phospho-ATF-2 antibody overnight at 4 °C. Bands were visualized using an horseradish peroxidase-linked secondary antibody and ECL. Films were scanned into a computer, and band intensities were analyzed using the gel analysis macros of NIH Image 1.62. Toxicity of Chemical Inhibitors—HUVECs were plated at 105 cells/cm2 on collagen-coated 96-well plates overnight in SFDM. The next day, cultures were given SFDM containing various chemical inhibitors. Plates were incubated at 37 °C for 4, 8, 12, and 24 h, media were withdrawn, and PBS containing 1 mm calcein and 2 μm ethidium homodimer (Molecular Probes) was added; plates were incubated for 1 h at 25 °C in the dark. Following labeling, plates were read using the Molecular Dynamics Fluorimager, and fluorescent signals were analyzed using Fluorsep and ImageQuant. Flow Cytometry Analysis of Integrin Expression—HUVEC passages 0–7 were trypsinized, allowed to recover in suspension for 2 h, rinsed twice in PBS/1% BSA, and resuspended at 5 × 106 cells/ml. 5 × 105 cells were placed in a non-tissue culture-treated 96-well plate with 5–40 μg/ml anti-α2β1, α1, or αVβ3 antibodies at 24 °C for 1 h. Plates were rinsed twice in PBS and incubated with a 1:100 dilution of a fluorescein isothiocyanate-labeled secondary antibody for 30 min at 24 °C. Cells were rinsed twice with PBS, resuspended in PBS, and mixed 1:1 with 4% paraformaldehyde. Samples were read on a Beckman Coulter XL/MCL analyzer using a 488-nm argon ion laser and system II software. Triple Helical Peptide Synthesis—Chemicals, including 9-fluorenylmethoxycarbonyl-amino acid derivatives, of analytical reagent grade or better were from Novabiochem (San Diego, CA) or Fisher. Amino acids, except for Gly, were l-isomers. The monoalkyl chains decanoic acid (CH3-(CH2)8-CO2H, designated C10) and palmitic acid (CH3-(CH2)14-CO2H, designated C16) were purchased from Aldrich. THPs and single-stranded peptides (SSP) were synthesized and purified as described (28Lauer-Fields J.L. Broder T. Sritharan T. Chung L. Nagase H. Fields G.B. Biochemistry. 2001; 50: 5795-5803Crossref Scopus (82) Google Scholar, 29Yu Y.-C. Berndt P. Tirrell M. Fields G.B. J. Am. Chem. Soc. 1996; 118: 12515-12520Crossref Scopus (247) Google Scholar, 30Yu Y.-C. Tirrell M. Fields G.B. J. Am. Chem. Soc. 1998; 120: 9979-9987Crossref Scopus (160) Google Scholar). Three THPs and one SSP were constructed: α1(I)256–270 (C10-(GPP*)4GEP*GAP*GNKGDTGEP*(GPP*)4-NH2) 2Where single-letter amino acid nomenclature is used, P* = hydroxyproline. THP (α1(I)256 –270 THP); α1(I)127–138 (C10-(GPP*)4GLP*GERGRP*GAP*(GPP*)4-NH2) THP (α1(I)127–138 THP); α1(I)496 –507 (C10-(GPP*)4GARGERGFP*GER(GPP*)4-NH2) THP (α1(I)496 –507 THP); and α1(I)499 –510 (C16GAP*GFP*GERGEK-NH2) SSP (α1(I)499 –510 SSP). Matrix-assisted laser desorption ionization-mass spectrometry analysis was performed on a Hewlett-Packard G2025A LD-TOF mass spectrometer using either a sinapinic acid or 2,5-dihydroxybenzoic acid/2-hydroxy-5-methoxybenzoic acid (9:1, v/v) matrix. Single chain mass values were as follows: α1(I)256 –270 THP, [M+H]+ 3721.8 Da (theoretical, 3722.0 Da); α1(I)127–138 THP, [M+H]+ 3498.8 Da (theoretical, 3502.9 Da); α1(I)496 –507 THP, [M+H]+ 3595.5 Da (theoretical, 3595.5 Da); and α1(I)499 –510 SSP, [M+H]+ 1469.9 Da (theoretical, 1470.7 Da). Circular dichroism spectra were recorded over the range λ = 190 –250 nm on a JASCO J-600 spectropolarimeter using a 10-mm path-length quartz cell. We obtained thermal transition curves by recording the molar ellipticity ([Θ]) at λ = 225 nm while continuously increasing the temperature from 5 to 80 °C at 0.2 °C/min, using a JASCO PTC-348WI temperature control unit. For samples exhibiting sigmoidal melting curves, the reflection point in the transition region (first derivative) is defined as the melting temperature (T m). T m values were 44.5, 56.0, and 52.5 °C for α1(I)256 –270 THP, α1(I)127–138 THP, and α1(I)496 –507 THP, respectively. Surface Plasmon Resonance Measurements of Integrin-THP Interactions—Binding of THP and SSP peptides to α1β1, αVβ3, and α2 integrins was determined at 25 °C using SPR from measurements of the accompanying increase in refractive index through time using a Biacore™ 2000 Biosensor (Biacore, Inc., Piscataway, NJ). Initially, the α1β1, αVβ3, and α2 integrins were captured on individual channels of a CM5 sensor chip by amine coupling. Solutions containing THPs and SSPs in elution buffer (10 mm HEPES, 150 mm NaCl, 1 mm MgCl2, pH 7.4) at the required concentrations were injected over each surface, and the response was measured as a function of time. The surface was regenerated after each injection by a single 10-s injection of 1 m NaCl. After subtraction of the contribution of bulk refractive index changes and nonspecific interactions with the CM5 chip surface, typically less than 5%, the individual association (k a) and dissociation (k d) rate constants were obtained by global fitting of data to a 1:1 Langmuir binding model using BIAevaluation™ (Biacore, Inc.). These values were then used to calculate the dissociation constant (K D). The values of average squared residual (c 2) obtained were not significantly improved by fitting data to models that assumed bivalent or heterogeneous interactions. Conditions were chosen so that the contribution of mass transport to the observed values of K D was negligible. To examine mechanisms of type I collagen-induced angiogenesis, we first modified the classic method for the induction of angiogenesis by a type I collagen gel deposited apically on a confluent HUVEC monolayer plated on a collagen gel (9Montesano R. Orci L. Vassalli P. J. Cell Biol. 1983; 97: 1648-1652Crossref PubMed Scopus (507) Google Scholar). Thus, we simplified the system by replacing the basal collagen gel with a collagen film to prevent cellular invasion and developed a serum-free medium (SFDM) that optimally supported angiogenesis. Development of a Novel Serum-free Defined Medium—Typically, endothelial cells are grown and used in angiogenesis assays in 10% serum with either endothelial cell growth factor or endothelial cell growth supplement, a heterogeneous mixture derived from bovine hypothalamic extracts (24Maciag T. Cerundolo J. Ilsley S. Kelley P.R. Forand R. Proc. Natl. Acad. Sci. U. S. A. 1979; 76: 5674-5678Crossref PubMed Scopus (577) Google Scholar); in such media angiogenesis occurs over 24–72 h. To develop a SFDM substitute, we tested the effectiveness of three base media: EBM, Medium 199, and Dulbecco's modified Eagle's medium: Ham's F-12 (1:1) in supporting tube formation. We found that EBM, a commercially available serum-free media for endothelial cells, although more complex and more expensive, was no more effective than Medium 199. The Dulbecco's modified Eagle's medium/Ham's F-12 mixture, often used because it is a richer nutrient medium, was also no better than Medium 199. Next, we found that insulin, transferrin, and selenium, often included in serum-free formulations to enhance cell survival and growth, had no benefit. Finally, we evaluated the effect of FGF-2 and VEGF and found that VEGF alone at either 5 or 10 ng/ml was less effective than 5 ng/ml FGF-2 alone in supporting tube formation (10Ilan N. Mahooti S. Madri J.A. J. Cell Sci. 1998; 111: 3621-3631Crossref PubMed Google Scholar); however, a combination of 5 ng/ml of both growth factors was optimal in agreement with previous studies (31Pepper M.S. Ferrara N. Orci L. Montesano R. Biochem. Biophys. Res. Commun. 1992; 189: 824-831Crossref PubMed Scopus (752) Google Scholar). Thus, our final SFDM was composed of Medium 199, 1% BSA (w/v), and 5 ng/ml FGF-2 and VEGF. Notably, use of this formulation resulted in the formation of widespread tube-like structures typically within 8–12 h. Time Course of Angiogenesis in SFDM—Observation of live cultures revealed immediate changes within the cell layer following polymerization of the collagen gel, when cells appear flattened as compared with non-collagen-treated cells. Within 2 h small spaces appeared in the cell layer, enlarging to form a net-like structure by 6–8 h as cells condensed and elongated forming capillary-like tubes that persisted to 24 h (Fig. 1). At this time, intracellular vacuoles were apparent by phase contrast microscopy, and it is proposed that such structures are precursors to lumen formation (32Folkman J. Haudenschild C. Nature. 1980; 288: 551-556Crossref PubMed Scopus (857) Google Scholar, 33Davis G.E. Camarillo C.W. Exp. Cell Res. 1996; 224: 39-51Crossref PubMed Scopus (303) Google Scholar). Ultrastructural analysis of cell layers at 24 h revealed structures reminiscent of capillaries enclosing lumen-like cavities formed by several HUVEC (Fig. 2). The lumen-like cavities occasionally contained filopodia, debris, and cellular protrusions yet always excluded the fibrillar collagen gel (Fig. 2, B–D). Thus our system resulted in the rapid formation of capillary-like tubes and therefore replicated the end point or capillary morphogenesis phase of angiogenesis.Fig. 2Capillary tube-like structures form in endothelial cell layers in response to an apical collagen gel. Control (A) and collagen-treated (B–D) cultures were fixed at 24 h and processed for transmission electron microscopy; sections were perpendicular to the plane of the cultures. Cultures stimulated with an apical collagen gel showed lumen-like structures (Lu) (B–D), and lumen-like structures with filopodia (arrow). Arrow-heads indicate electron dense intercellular junctions in closely apposed cells. Some lumen-like structures contained cellular protrusions (cp) and debris (d) (D). In some micrographs the fibrillar collagen gel (cg) can be seen (C and D). Upper panel, ×3600; middle panel, ×2900; lower panel, ×1900.View Large Image Figure ViewerDownload Hi-res image Download (PPT) Temporal Requirement for Collagen—To determine whether the collagen gel initiates angiogenesis, or is required throughout the process, collagen gels were added to cultures, then removed and replaced with SFDM from 2–12 h following polymerization and were examined by microscopy an additional 24 h (not shown). We found that, when apical gels were removed at times less than 8 h after their addition, cellular reorganization was arrested and cells returned to a near-confluent monolayer as previously reported (34Deroanne C.F. Colige A.C. Nusgens B.V. Lapiere C.M. Exp. Cell Res. 1996; 224: 215-223Crossref PubMed Scopus (32) Google Scholar). However, in cultures where collagen gels were present for at least 8 h and capillary like networks had formed, some remnants of the net-like structure remained following gel removal and persisted even at 24 h. Thus collagen must be constantly present during HUVEC tube formation. However, once morphogenesis is complete, parts of the capillary-like network remain quite stable morphologically, even in the absence of collagen. Requirement for the α2β1 Integrin—We next examined the role of various integrins, the cell surface receptors likely to mediate HUVEC-collagen inte" @default.
• W2168296320 created "2016-06-24" @default.
• W2168296320 creator A5027441705 @default.
• W2168296320 creator A5030553564 @default.
• W2168296320 creator A5050632500 @default.
• W2168296320 creator A5054779535 @default.
• W2168296320 creator A5059131009 @default.
• W2168296320 creator A5059836393 @default.
• W2168296320 creator A5068707393 @default.
• W2168296320 creator A5087639379 @default.
• W2168296320 date "2003-08-01" @default.
• W2168296320 modified "2023-10-18" @default.
• W2168296320 title "Angiogenesis in Collagen I Requires α2β1 Ligation of a GFP*GER Sequence and Possibly p38 MAPK Activation and Focal Adhesion Disassembly" @default.
• W2168296320 cites W1547774078 @default.
• W2168296320 cites W1574213882 @default.
• W2168296320 cites W1582963778 @default.
• W2168296320 cites W1586440254 @default.
• W2168296320 cites W1602538488 @default.
• W2168296320 cites W1950519844 @default.
• W2168296320 cites W1964257347 @default.
• W2168296320 cites W1966931906 @default.
• W2168296320 cites W1976772770 @default.
• W2168296320 cites W1977013517 @default.
• W2168296320 cites W1978094951 @default.
• W2168296320 cites W1980969801 @default.
• W2168296320 cites W1982405831 @default.
• W2168296320 cites W1992437868 @default.
• W2168296320 cites W1993928278 @default.
• W2168296320 cites W1995857537 @default.
• W2168296320 cites W1996537616 @default.
• W2168296320 cites W1998004834 @default.
• W2168296320 cites W2000000541 @default.
• W2168296320 cites W2002782430 @default.
• W2168296320 cites W2004487950 @default.
• W2168296320 cites W2010308831 @default.
• W2168296320 cites W2011083738 @default.
• W2168296320 cites W2014128175 @default.
• W2168296320 cites W2022415270 @default.
• W2168296320 cites W2022914311 @default.
• W2168296320 cites W2024538453 @default.
• W2168296320 cites W2026614330 @default.
• W2168296320 cites W2032945902 @default.
• W2168296320 cites W2038963002 @default.
• W2168296320 cites W2039317622 @default.
• W2168296320 cites W2042204791 @default.
• W2168296320 cites W2047158623 @default.
• W2168296320 cites W2047408781 @default.
• W2168296320 cites W2050128446 @default.
• W2168296320 cites W2051554687 @default.
• W2168296320 cites W2051914085 @default.
• W2168296320 cites W2055628427 @default.
• W2168296320 cites W2059225346 @default.
• W2168296320 cites W2060409716 @default.
• W2168296320 cites W2065295487 @default.
• W2168296320 cites W2065621361 @default.
• W2168296320 cites W2066711736 @default.
• W2168296320 cites W2067844964 @default.
• W2168296320 cites W2072410510 @default.
• W2168296320 cites W2074801137 @default.
• W2168296320 cites W2079070267 @default.
• W2168296320 cites W2093060454 @default.
• W2168296320 cites W2100363932 @default.
• W2168296320 cites W2106333878 @default.
• W2168296320 cites W2109594092 @default.
• W2168296320 cites W2111618967 @default.
• W2168296320 cites W2115341926 @default.
• W2168296320 cites W2127670194 @default.
• W2168296320 cites W2135112491 @default.
• W2168296320 cites W2146787008 @default.
• W2168296320 cites W2154387825 @default.
• W2168296320 cites W2161615417 @default.
• W2168296320 cites W2165336459 @default.
• W2168296320 cites W4375905804 @default.
• W2168296320 cites W2087003171 @default.
• W2168296320 doi "https://doi.org/10.1074/jbc.m304237200" @default.
• W2168296320 hasPubMedId "https://pubmed.ncbi.nlm.nih.gov/12788934" @default.
• W2168296320 hasPublicationYear "2003" @default.
• W2168296320 type Work @default.
• W2168296320 sameAs 2168296320 @default.
• W2168296320 citedByCount "129" @default.
• W2168296320 countsByYear W21682963202012 @default.
• W2168296320 countsByYear W21682963202013 @default.
• W2168296320 countsByYear W21682963202014 @default.
• W2168296320 countsByYear W21682963202015 @default.
• W2168296320 countsByYear W21682963202016 @default.
• W2168296320 countsByYear W21682963202017 @default.
• W2168296320 countsByYear W21682963202018 @default.
• W2168296320 countsByYear W21682963202019 @default.
• W2168296320 countsByYear W21682963202020 @default.
• W2168296320 countsByYear W21682963202021 @default.
• W2168296320 countsByYear W21682963202022 @default.
• W2168296320 crossrefType "journal-article" @default.
• W2168296320 hasAuthorship W2168296320A5027441705 @default.
• W2168296320 hasAuthorship W2168296320A5030553564 @default.
• W2168296320 hasAuthorship W2168296320A5050632500 @default.
• W2168296320 hasAuthorship W2168296320A5054779535 @default.
• W2168296320 hasAuthorship W2168296320A5059131009 @default.
• W2168296320 hasAuthorship W2168296320A5059836393 @default.

• next