PAI-1: An Integrator of Cell Signaling and Migration
Cellular migration, over simple surfaces or through complex stromal barriers, requires coordination between detachment/re-adhesion cycles, involving structural components of the extracellular matrix and their surface-binding elements (integrins), and the precise regulation of the pericellular proteolytic microenvironment. It is now apparent that several proteases and protease inhibitors, most notably urokinase plasminogen activator (uPA) and plasminogen activator inhibitor type-1 (PAI-1), also interact with several cell surface receptors transducing intracellular signals that significantly affect both motile and proliferative programs. These events appear distinct from the original function of uPA/PAI-1 as modulators of the plasmin-based proteolytic cascade. The multifaceted interactions of PAI-1 with specific matrix components (i.e., vitronectin), the low-density lipoprotein receptor-related protein-1 (LRP1), and the uPA/uPA receptor complex have dramatic consequences on the migratory phenotype and may underlie the pathophysiologic sequalae of PAI-1 deficiency and overexpression. This paper focuses on the increasingly intricate role of PAI-1 as a major mechanistic determinant of the cellular migratory phenotype.
The switch between a sessile and migratory cellular phenotype is triggered, in part, by the activation of signaling pathways that regulate the expression of the involved genes, (e.g., [1, 2]). While the actual genomic response varies as a consequence of cell type, the acquisition of a core “plasticity” signature (at both the mRNA and proteomic levels) represents the transition to a motile phenotype whether over simple planar surfaces or through complex matrix barriers in normal as well as transformed keratinocytes, (e.g., [2–7]). Global transcriptome profiling of both wounded keratinocyte cultures and epithelial tumor cells has highlighted the requirement for precise spatial/temporal control of pericellular proteolysis and matrix remodeling in the integration of the cellular motile/tissue repair responses [2, 5]. Indeed, among the transcriptional outputs (i.e., genes with altered expression) that typify the migratory or invasive phenotype, urokinase plasminogen activator (uPA) and its major negative regulator plasminogen activator inhibitor type-1 (PAI-1) are among the most highly induced transcripts, (e.g., [4, 5, 8]) (Figure 1). PAI-1 belongs to the serine protease inhibitor (SERPIN) protein family that also includes PAI-2 and PAI-3 (protein C inhibitor), protease nexin-1, and neuroserpin (reviewed in ). uPA and PAI-1 (also known as SERPINE1) are both the targets and modifiers of pathways that impact proliferative/migratory events (Figure 2) and coordinately titrate the overall pericellular proteolytic balance directly (via generation of plasmin) as well as indirectly by activating several members of the matrix metalloproteinase (MMP) family (reviewed in [4, 7]). Motile epithelial cells focalize both uPA, following interaction with its cell surface receptor uPAR, and PAI-1, upon binding of this SERPIN to uPA/uPAR or vitronectin (VN), to the leading edge where they modulate the interrelated events of matrix remodeling and migration, (e.g., [10–12]). Focal proteolysis reorganizes extracellular matrix (ECM) architecture, affecting cell-ECM interactions with integrin receptors and releasing bioactive fragments of matrix molecules as well as activating growth factors that stimulate the migratory behavior (Figure 3) (reviewed in ). These findings have important implications. While uPA and uPAR are widely implicated in tumor invasion, deficiencies in PAI-1 levels also correlate with significantly reduced epithelial cell migration and tumor progression [1, 4, 7, 13]. A critical balance between uPA and PAI-1 appears required, therefore, to create a microenvironment compatible with efficient cell motility. High stromal PAI-1 levels, in fact, correlate with a poor prognosis in various cancers [14–16] and typify diseases in which fibrosis and/or cellular infiltration are common pathologic features (e.g., scarring anomalies, renal fibrosis, atherosclerosis) [17–21]. Collectively, these findings suggest that PAI-1-dependent preservation of the surrounding matrix may facilitate cell locomotion in vivo, perhaps by fine-tuning the proteolytic activity to optimize tissue penetration. This paper focuses on the most recent developments in this field and on the complex proteolytic as well as nonproteolytic functions of PAI-1 in the cellular motile program.
2. PAI-1-Regulated Cell Migration: Receptor Interactions
Stromal PAI-1 is itself a substrate for several extracellular proteases including elastase, MMP-3, and plasmin [22–24]. “Cleaved” PAI-1 is unable to interact with its target plasminogen activators uPA and tissue-type PA (tPA) to inhibit plasmin-based proteolysis but retains its ability to bind the low-density lipoprotein receptor-related protein-1 (LRP1) and augment cell migration, through a u/tPA complex-independent interaction (Figure 4, left) . LRP1, in addition to its function as a major endocytic receptor for multiple ligands, is also a key signaling mediator in several pathways due, in part, to its ability to support interactions with multiple adaptor and scaffolding proteins . LRP1 ligand binding and/or its complex formation with cell surface partners including integrins [27–29], growth factor receptors [30–32], and proteoglycans  activates mitogen-activated protein (MAP) and nonreceptor src kinases [34–37], impacting cell proliferation [30, 31, 38, 39] and migration [25, 34, 40] with the motile response involving activation of Rho family GTPases . Alternatively, PAI-1 can also function as a signaling molecule that directly affects cell migration through engagement of LRP1 and the very low-density lipoprotein receptor . Indeed, the different conformations of PAI-1 (active, latent, cleaved) interact with LRP1 to stimulate cellular migration into 3D collagen gels through a LRP1-dependent mechanism . All three forms of PAI-1 increase LRP1-dependent cell motility with the activation of the Jak/Stat1 pathway [25, 43, 44] (Figure 4, left). While active PAI-1 is routinely cleared from the extracellular environment in a complex with uPA/uPAR/LRP1, latent and cleaved species of PAI-1, with a preserved motile function, remain embedded in the matrix likely serving as a reservoir to maintain cell movement.
One prerequisite for efficient cellular migration is a sustainable, flexible state of cell adhesion. PAI-1 significantly impacts adhesion through interaction with LRP1 and VN. PAI-1 mutants that vary in their capacity to bind uPA, VN, or LRP1 can attenuate smooth muscle cell adhesive forces through deregulation of integrin activity . This mechanism, targeting only active, matrix-engaged integrins, results in cell detachment from VN, fibronectin (FN), and collagen matrices , allowing for readhesion to alternative matrix structural elements, thus promoting migration. It appears that even low concentrations of PAI-1 lead to substantial and rapid changes in the actin cytoskeleton and the loss of focal adhesions  with likely consequences on the motile phenotype.
PAI-1 also regulates levels of cell surface integrins by triggering their internalization in an LRP1-dependent manner [27, 45, 46] resulting in cell detachment from various substrates [27, 45] (Figure 4, middle). Integrin internalization by LRP1, however, is not a requirement during PAI-1-initiated cell release . This mechanism appears to differ from that which modulates PAI-1-stimulated migration directly via LRP1, as uPA and uPAR are required for deadhesion but not for the migratory response [25, 27, 43, 46]. Although LRP1-mediated integrin endocytosis seems not to be necessary for efficient cell detachment, integrin endocytosis would allow for their subcellular redistribution (i.e., to the leading edge) in support of cell locomotion and stromal invasion. While the interaction between PAI-1 and uPA/uPAR/integrin complexes would ultimately enhance the integrin/uPAR “attachment-detachment-reattachment” cycle , thereby, increasing cell motility, it is apparent that PAI-1 can utilize multiple avenues to impact LRP1-dependent cell migration (Figure 4, left and middle). Further complicating this process is the potential for PAI-1 to modulate syndecan-dependent keratinocyte migration, as evident during wound healing. Keratinocytes at the wound margin begin to synthesize and deposit unprocessed laminin-332, supporting syndecan-1 binding through the LG4/5 domain (Figure 4, right). PAI-1, which is also expressed by cells at the wound edge, stabilizes this interaction by preventing plasmin-initiated proteolytic processing of laminin-332  and syndecan-1 shedding [49, 50]. The presence of VN at the wound edge can augment this event through its ability to focalize PAI-1 and extend the half-life of active PAI-1 (discussed below) as well as engage syndecan-1 . PAI-1, through its ability to reduce pericellular levels of active plasmin, promotes syndecan-1-dependent migration on unprocessed laminin-332 by preventing cleavage of the syndecan-binding site LG4/5. Additionally, PAI-1 inhibition of plasmin activation facilitates migration on unprocessed laminin-332 by reducing the shedding of syndecan-1 from the cell surface. As the proteolytic environment matures, PAI-1 and VN are endocytosed and degraded [52, 53]. Syndecan-1 binding is lost due to proteolytic processing of laminin-332, as well as syndecan-1 ectodomain shedding; α3β1 binding to processed laminin-332 begins to slow keratinocyte migration and initiate hemidesmosome formation [48, 54] (see Figure 4, right).
3. PAI-1-Regulated Cell Migration: Interactions with Vitronectin
PAI-1/VN interactions impact several mechanisms associated with cell migration. Whereas PAI-1 had been recognized earlier as a highly significant prognostic indicator for malignant disease outcome , the importance of stromal VN as inducer of cell motility came in focus only more recently [56–58]. In part, it does so by stabilizing PAI-1 in an active conformation, extending its half-life and amplifying the inhibition of focal proteolysis modulating the extent, locale, and duration of matrix remodeling, thereby preserving a stromal architecture permissive for cell motility [59, 60]. This is particularly important following cutaneous injury where restoration of barrier function and tissue integrity is dependent upon keratinocyte movement. PAI-1 and VN are both released from the α granules of platelets during hemostasis, where their combined presence would presumably promote the formation of a fibrin clot and subsequently contribute to provisional matrix remodeling [61, 62]. PAI-1 upregulation in keratinocytes at the wound margin [1, 12] highlights the potential involvement of this SERPIN in initiating tissue repair. VN expression, however, is limited under normal physiological conditions [63–66] but similarly enhanced under circumstances requiring stromal remodeling (i.e., wound repair [67–69] or tumor progression [70–74]) suggesting a continuing, albeit dynamic, molecular interaction with PAI-1 of potential physiologic significance. This dynamic might reflect the fact that the binding of PAI-1 to VN alters the motogenic properties of PAI-1, rendering PAI-1/VN complexes nonmotogenic, whereas all non-VN-bound PAI-1s (cleaved, latent, or active) exhibit strong motogenic properties .
The interaction between PAI-1 and VN also affects cell motility through mechanisms that directly modulate cell surface receptor binding (Figure 5). VN promotes cellular locomotion via RGD-dependent interactions with αvβ3 and αvβ5 integrins [75–78], as well as through binding to uPAR [79, 80]. The recognition site for PAI-1 on VN, however, approximates those for both integrin and uPAR docking , and, as a result, the interaction of PAI-1 with VN regulates the ability of these receptors to engage VN [47, 79–82] (Figure 5). PAI-1, in addition to regulating cell-to-substrate attachment, also affects cellular release from VN by two distinct mechanisms. The affinity of PAI-1 for VN is significantly higher than that of uPAR for VN. Consequently, PAI-1 can competitively displace uPAR from VN, initiating detachment of cells that rely mainly on uPAR for cell adhesion to VN [79, 80, 82]. However, PAI-1 is unable to promote its binding to VN by competitive displacement of preengaged integrins from VN. In the presence of uPA/uPAR/αv-integrin complexes; moreover, PAI-1 binding to complexed uPA will initiate integrin deactivation, promoting their detachment from VN and endocytic clearance [27, 45]. These receptors are subsequently recycled back to the cell surface to reengage matrix molecules and promote cell migration  (Figure 4, middle). In contrast to the effects of PAI-1 on cell attachment, the deadhesive effect of PAI-1 is strictly uPA-dependent and VN-independent since PAI-1 can also initiate cell release from FN, collagen-I, and laminin-332 matrices .
In addition, PAI-1/VN binding blocks PAI-1 interaction with LRP1, thus preventing the LRP1-dependent migration signaling  (Figure 5). The question remains how PAI-1 will react to the presence of the other two binding partners, VN and uPA. Recent observations would suggest that the stoichiometry between these three molecules will determine the result of their interactions . Migration of human vascular smooth muscle cells on 2D and through 3D collagen gels, in the presence of VN, was significantly reduced in low PAI-1, whereas high PAI-1 concentrations strongly promoted cell migration.
Cell migration requires the temporal/spatial regulation of a series of complex proteolytic events coupled with the activation of critical surface receptors (uPAR, integrins, LRP1) and initiation of downstream signaling, by several elements intimately involved in pericellular proteolysis. PAI-1, through its varied interactions with VN and cellular receptors, is centrally positioned to coordinate the duration and locale of both intracellular (signal initiation) and extracellular (detachment/readhesion cycles, receptor binding) events that manage the intricate process of cell movement in both physiologic and pathologic contexts. Clearly, the binding of PAI-1 with its several targets including VN, uPA, uPA/uPAR, and LRP1 has the potential to affect the motile program on multiple levels providing opportunities to therapeutically manipulate this pathway in pathophysiologic settings.
This work was supported by grants from NIH (GM57242), the NYSDOH Empire State Stem Cell Trust Fund (C024312), the Friedman Family Cancer Research Endowment, and the Kevin J. Butler Foundation for Mesothelioma Research.
G. Fitsialos, A. A. Chassot, L. Turchi et al., “Transcriptional signature of epidermal keratinocytes subjected to in vitro scratch wounding reveals selective roles for ERK1/2, p38, and phosphatidylinositol 3-kinase signaling pathways,” Journal of Biological Chemistry, vol. 282, no. 20, pp. 15090–15102, 2007.View at: Publisher Site | Google Scholar
J. Freytag, C. E. Wilkins-Port, C. E. Higgins et al., “PAI-1 regulates the invasive phenotype in human cutaneous squamous cell carcinoma,” Journal of Oncology, no. 2, pp. 1–12, 2009.View at: Google Scholar
C. E. Wilkins-Port, Q. Ye, J. E. Mazurkiewicz, and P. J. Higgins, “TGF-1 + EGF-initiated invasive potential in transformed human keratinocytes is coupled to a plasmin/mmp-10/mmp-1-dependent collagen remodeling axis: role for PAI-1,” Cancer Research, vol. 69, no. 9, pp. 4081–4091, 2009.View at: Publisher Site | Google Scholar
C. E. Wilkins-Port, J. Freytag, S. P. Higgins, and P. J. Higgins, “PAI-1: a multifunctional SERPIN with complex roles in cell signaling and migration,” Cell Communication Insights, vol. 2010, no. 3, pp. 1–10, 2010.View at: Google Scholar
J. Freytag, C. E. Wilkins-Port, C. E. Higgins, S. P. Higgins, R. Samarakoon, and P. J. Higgins, “PAI-1 mediates the TGF-1EGF-induced scatter response in transformed human keratinocytes,” Journal of Investigative Dermatology, vol. 130, no. 9, pp. 2179–2190, 2010.View at: Publisher Site | Google Scholar
K. M. Providence, L. A. White, J. Tang, J. Gonclaves, L. Staiano-Coico, and P. J. Higgins, “Epithelial monolayer wounding stimulates binding of USF-1 to an E-box motif in the plasminogen activator inhibitor type 1 gene,” Journal of Cell Science, vol. 115, no. 19, pp. 3767–3777, 2002.View at: Publisher Site | Google Scholar
L. S. Gutierrez, A. Schulman, T. Brito-Robinson, F. Noria, V. A. Ploplis, and F. J. Castellino, “Tumor development is retarded in mice lacking the gene for urokinase-type plasminogen activator or its inhibitor, plasminogen activator inhibitor-1,” Cancer Research, vol. 60, no. 20, pp. 5839–5847, 2000.View at: Google Scholar
B. Hundsdorfer, H. F. Zeilhofer, K. P. Bock, P. Dettmar, M. Schmitt, and H. H. Horch, “The prognostic importance of urinase type plasminogen activators (uPA) and plasminogen activator inhibitors (PAI-1) in the primary resection of oral squamous cell carcinoma,” Mund-, Kiefer- und Gesichtschirurgie, vol. 8, no. 3, pp. 173–179, 2004.View at: Google Scholar
M. K. V. Durand, J. S. Bödker, A. Christensen et al., “Plasminogen activator inhibitor-1 and tumour growth, invasion, and metastasis,” Thrombosis and Haemostasis, vol. 91, no. 3, pp. 438–449, 2004.View at: Google Scholar
Q. Zhang, Y. Wu, C. H. Chau, D. K. Ann, C. N. Bertolami, and A. D. Le, “Crosstalk of hypoxia-mediated signaling pathways in upregulating plasminogen activator inhibitor-1 expression in keloid fibroblasts,” Journal of Cellular Physiology, vol. 199, no. 1, pp. 89–97, 2004.View at: Publisher Site | Google Scholar
R. Samarakoon, S. P. Higgins, C. E. Higgins, and P. J. Higgins, “TGF-1-induced plasminogen activator inhibitor-1 expression in vascular smooth muscle cells requires pp60c-src/EGFRY845 and Rho/ROCK signaling,” Journal of Molecular and Cellular Cardiology, vol. 44, no. 3, pp. 527–538, 2008.View at: Publisher Site | Google Scholar
A. M. Audenaert, I. Knockaert, D. Collen, and P. J. Declerck, “Conversion of plasminogen activator inhibitor-1 from inhibitor to substrate by point mutations in the reactive-site loop,” Journal of Biological Chemistry, vol. 269, no. 30, pp. 19559–19564, 1994.View at: Google Scholar
D. A. Lawrence, S. T. Olson, S. Palaniappan, and D. Ginsburg, “Serpin reactive center loop mobility is required for inhibitor function but not for enzyme recognition,” Journal of Biological Chemistry, vol. 269, no. 44, pp. 27657–27662, 1994.View at: Google Scholar
B. Degryse, J. G. Neels, R. P. Czekay, K. Aertgeerts, Y. I. Kamikubo, and D. J. Loskutoff, “The low density lipoprotein receptor-related protein is a motogenic receptor for plasminogen activator inhibitor-1,” Journal of Biological Chemistry, vol. 279, no. 21, pp. 22595–22604, 2004.View at: Publisher Site | Google Scholar
S. C. Muratoglu, I. Mikhailenko, C. Newton, M. Migliorini, and D. K. Strickland, “Low density lipoprotein receptor-related protein 1 (LRP1) forms a signaling complex with platelet-derived growth factor receptor- in endosomes and regulates activation of the MAPK pathway,” Journal of Biological Chemistry, vol. 285, no. 19, pp. 14308–14317, 2010.View at: Publisher Site | Google Scholar
L. C. Wilsie and R. A. Orlando, “The low density lipoprotein receptor-related protein complexes with cell surface heparan sulfate proteoglycans to regulate proteoglycan-mediated lipoprotein catabolism,” Journal of Biological Chemistry, vol. 278, no. 18, pp. 15758–15764, 2003.View at: Publisher Site | Google Scholar
E. Mantuano, G. Inoue, X. Li et al., “The hemopexin domain of matrix metalloproteinase-9 activates cell signaling and promotes migration of Schwann cells by binding to low-density lipoprotein receptor-related protein,” Journal of Neuroscience, vol. 28, no. 45, pp. 11571–11582, 2008.View at: Publisher Site | Google Scholar
E. Mantuano, G. Mukandala, X. Li, W. M. Campana, and S. L. Gonias, “Molecular dissection of the human 2-macroglobulin subunit reveals domains with antagonistic activities in cell signaling,” Journal of Biological Chemistry, vol. 283, no. 29, pp. 19904–19911, 2008.View at: Publisher Site | Google Scholar
E. Mantuano, M. Jo, S. L. Gonias, and W. M. Campana, “Low density lipoprotein receptor-related protein (LRP1) regulates Rac1 and RhoA reciprocally to control Schwann cell adhesion and migration,” Journal of Biological Chemistry, vol. 285, no. 19, pp. 14259–14266, 2010.View at: Publisher Site | Google Scholar
C. W. Heegaard, A. C. W. Simonsen, K. Oka et al., “Very low density lipoprotein receptor binds and mediates endocytosis of urokinase-type plasminogen activator-type-1 plasminogen activator inhibitor complex,” Journal of Biological Chemistry, vol. 270, no. 35, pp. 20855–20861, 1995.View at: Publisher Site | Google Scholar
N. Garg, N. Goyal, T. L. Strawn et al., “Plasminogen activator inhibitor-1 and vitronectin expression level and stoichiometry regulate vascular smooth muscle cell migration through physiological collagen matrices,” Journal of Thrombosis and Haemostasis, vol. 8, no. 8, pp. 1847–1854, 2010.View at: Publisher Site | Google Scholar
Y. Kamikubo, J. G. Neels, and B. Degryse, “Vitronectin inhibits plasminogen activator inhibitor-1-induced signalling and chemotaxis by blocking plasminogen activator inhibitor-1 binding to the low-density lipoprotein receptor-related protein,” International Journal of Biochemistry and Cell Biology, vol. 41, no. 3, pp. 578–585, 2009.View at: Publisher Site | Google Scholar
B. S. Pedroja, L. E. Kang, A. O. Imas, P. Carmeliet, and A. M. Bernstein, “Plasminogen activator inhibitor-1 regulates integrin v3 expression and autocrine transforming growth factor signaling,” Journal of Biological Chemistry, vol. 284, no. 31, pp. 20708–20717, 2009.View at: Publisher Site | Google Scholar
S. Stefansson and D. A. Lawrence, “Old dogs and new tricks: proteases, inhibitors, and cell migration,” Science Stke, vol. 2003, no. 189, p. pe24, 2003.View at: Google Scholar
L. J. Marshall, L. S. P. Ramdin, T. Brooks, P. C. DPhil, and J. K. Shute, “Plasminogen activator inhibitor-1 supports IL-8-mediated neutrophil transendothelial migration by inhibition of the constitutive shedding of endothelial IL-8/heparan sulfate/syndecan-1 complexes,” Journal of Immunology, vol. 171, no. 4, pp. 2057–2065, 2003.View at: Google Scholar
C. E. Wilkins-Port, R. D. Sanderson, E. Tominna-Sebald, and P. J. McKeown-Longo, “Vitronectin's basic domain is a syndecan ligand which functions in trans to regulate vitronectin turnover,” Cell Communication and Adhesion, vol. 10, no. 2, pp. 85–103, 2003.View at: Google Scholar
R. P. Czekay, T. A. Kuemmel, R. A. Orlando, and M. G. Farquhar, “Direct binding of occupied urokinase receptor (uPAR) to LDL receptor-related protein is required for endocytosis of uPAR and regulation of cell surface urokinase activity,” Molecular Biology of the Cell, vol. 12, no. 5, pp. 1467–1479, 2001.View at: Google Scholar
J. Grondahl-Hansen, I. J. Christensen, P. Briand et al., “Plasminogen activator inhibitor type 1 in cytosolic tumor extracts predicts prognosis in low-risk breast cancer patients,” Clinical Cancer Research, vol. 3, no. 2, pp. 233–239, 1997.View at: Google Scholar
J. Mimuro and D. J. Loskutoff, “Binding of type 1 plasminogen activator inhibitor to the extracellular matrix of cultured bovine endothelial cells,” Journal of Biological Chemistry, vol. 264, no. 9, pp. 5058–5063, 1989.View at: Google Scholar
J. Mimuro and D. J. Loskutoff, “Purification of a protein from bovine plasma that binds to type 1 plasminogen activator inhibitor and prevents its interaction with extracellular matrix. Evidence that the protein is vitronectin,” Journal of Biological Chemistry, vol. 264, no. 2, pp. 936–939, 1989.View at: Google Scholar
S. A. Hill, S. G. Shaughnessy, P. Joshua, J. Ribau, R. C. Austin, and T. J. Podor, “Differential mechanisms targeting type 1 plasminogen activator inhibitor and vitronectin into the storage granules of a human megakaryocytic cell line,” Blood, vol. 87, no. 12, pp. 5061–5073, 1996.View at: Google Scholar
D. Seiffert and R. R. Schleef, “Two functionally distinct pools of vitronectin (Vn) in the blood circulation: identification of a heparin-binding competent population of Vn within platelet -granules,” Blood, vol. 88, no. 2, pp. 552–560, 1996.View at: Google Scholar
D. Seiffert, M. Keeton, Y. Eguchi, M. Sawdey, and D. J. Loskutoff, “Detection of vitronectin mRNA in tissues and cells of the mouse,” Proceedings of the National Academy of Sciences of the United States of America, vol. 88, no. 21, pp. 9402–9406, 1991.View at: Google Scholar
D. Seiffert, M. L. Iruela-Arispe, E. H. Sage, and D. J. Loskutoff, “Distribution of vitronectin mRNA during murine development,” Developmental Dynamics, vol. 203, no. 1, pp. 71–79, 1995.View at: Google Scholar
D. Seiffert, “Constitutive and regulated expression of vitronectin,” Histology and Histopathology, vol. 12, no. 3, pp. 787–797, 1997.View at: Google Scholar
B. R. Tomasini and D. F. Mosher, “Vitronectin,” Progress in Hemostasis and Thrombosis, vol. 10, pp. 269–305, 1991.View at: Google Scholar
Y. C. Jang, R. Tsou, N. S. Gibran, and F. F. Isik, “Vitronectin deficiency is associated with increased wound fibrinolysis and decreased microvascular angiogenesis in mice,” Surgery, vol. 127, no. 6, pp. 696–704, 2000.View at: Google Scholar
L. Taliana, M. D. M. Evans, S. Ang, and J. W. McAvoy, “Vitronectin is present in epithelial cells of the intact lens and promotes epithelial mesenchymal transition in lens epithelial explants,” Molecular Vision, vol. 12, pp. 1233–1242, 2006.View at: Google Scholar
C. L. Gladson and D. A. Cheresh, “Glioblastoma expression of vitronectin and the v3 integrin. Adhesion mechanism for transformed glial cells,” Journal of Clinical Investigation, vol. 88, no. 6, pp. 1924–1932, 1991.View at: Google Scholar
C. L. Gladson, J. N. Wilcox, L. Sanders, G. Y. Gillespie, and D. A. Cheresh, “Cerebral microenvironment influences expression of the vitronectin gene in astrocytic tumors,” Journal of Cell Science, vol. 108, no. 3, pp. 947–956, 1995.View at: Google Scholar
D. A. Cheresh and R. C. Spiro, “Biosynthetic and functional properties of an Arg-Gly-Asp-directed receptor involved in human melanoma cell attachment to vitronectin, fibrinogen, and von Willebrand factor,” Journal of Biological Chemistry, vol. 262, no. 36, pp. 17703–17711, 1987.View at: Google Scholar
D. A. Cheresh, “Human endothelial cells synthesize and express an Arg-Gly-Asp-directed adhesion receptor involved in attachment to fibrinogen and von Willebrand factor,” Proceedings of the National Academy of Sciences of the United States of America, vol. 84, no. 18, pp. 6471–6475, 1987.View at: Google Scholar
R. Pytela, M. D. Pierschbacher, and E. Ruoslahti, “A 125/115-kDa cell surface receptor specific for vitronectin interacts with the arginine-glycine-aspartic acid adhesion sequence derived from fibronectin,” Proceedings of the National Academy of Sciences of the United States of America, vol. 82, no. 17, pp. 5766–5770, 1985.View at: Google Scholar
J. W. Smith, D. J. Vestal, S. V. Irwin, T. A. Burke, and D. A. Cheresh, “Purification and functional characterization of integrin (v)5. An adhesion receptor for vitronectin,” Journal of Biological Chemistry, vol. 265, no. 19, pp. 11008–11013, 1990.View at: Google Scholar
G. Deng, S. A. Curriden, G. Hu, R.-P. Czekay, and D. J. Loskutoff, “Plasminogen activator inhibitor-1 regulates cell adhesion by binding to the somatomedin B domain of vitronectin,” Journal of Cellular Physiology, vol. 189, pp. 23–33, 2001.View at: Google Scholar
D. A. Waltz, L. R. Natkin, R. M. Fujita, Y. Wei, and H. A. Chapman, “Plasmin and plasminogen activator inhibitor type 1 promote cellular motility by regulating the interaction between the urokinase receptor and vitronectin,” Journal of Clinical Investigation, vol. 100, no. 1, pp. 58–67, 1997.View at: Google Scholar
G. Deng, S. A. Curriden, S. Wang, S. Rosenberg, and D. J. Loskutoff, “Is plasminogen activator inhibitor-1 the molecular switch that governs urokinase receptor-mediated cell adhesion and release?” Journal of Cell Biology, vol. 134, no. 6, pp. 1563–1571, 1996.View at: Google Scholar