Complex Regulation of the Pericellular Proteolytic Microenvironment during Tumor Progression and Wound Repair: Functional Interactions between the Serine Protease and Matrix Metalloproteinase Cascades

Wilkins-Port, Cynthia E.; Higgins, Stephen P.; Higgins, Craig E.; Kobori-Hotchkiss, Issey; Higgins, Paul J.

doi:https://doi.org/10.1155/2012/454368

Biochemistry Research International

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Volume 2012 | Article ID 454368 | https://doi.org/10.1155/2012/454368

Complex Regulation of the Pericellular Proteolytic Microenvironment during Tumor Progression and Wound Repair: Functional Interactions between the Serine Protease and Matrix Metalloproteinase Cascades

Cynthia E. Wilkins-Port,¹Stephen P. Higgins,¹Craig E. Higgins,¹Issey Kobori-Hotchkiss,¹and Paul J. Higgins¹

Academic Editor: Maria L. Urso

Received31 Aug 2011

Accepted21 Nov 2011

Published20 Feb 2012

Abstract

Spatial and temporal regulation of the pericellular proteolytic environment by local growth factors, such as EGF and TGF-β, initiates a wide repertoire of cellular responses coupled to a plasmin/matrix metalloproteinase (MMP) dependent stromal-remodeling axis. Cell motility and invasion, tumor metastasis, wound healing, and organ fibrosis, for example, represent diverse events controlled by expression of a subset of genes that encode various classes of tissue remodeling proteins. These include members of the serine protease and MMP families that functionally constitute a complex system of interacting protease cascades and titrated by their respective inhibitors. Several structural components of the extracellular matrix are upregulated by TGF-β as are matrix-active proteases (e.g., urokinase (uPA), plasmin, MMP-1, -3, -9, -10, -11, -13, -14). Stringent controls on serine protease/MMP expression and their topographic activity are essential for maintaining tissue homeostasis. Targeting individual elements in this highly interactive network may lead to novel therapeutic approaches for the treatment of cancer, fibrotic diseases, and chronic wounds.

1. Introduction

Epithelial transdifferentiation or cellular “plasticity” refers to a specialized morphogenetic switch typified by loss of normal epithelial properties, a gain in the expression of genes generally restricted to the mesenchymal lineage and conversion of sessile, nonmotile cells to a migratory phenotype [1, 2]. While essential during development and organogenesis (i.e., embryonic patterning) (also termed Type 1 transition), this process is relatively limited in the adult organism, occurring during wound healing and regenerative repair [3–5] or, more atypically, in tissue fibrosis (Type 2) and tumor metastasis (Type 3) [6–9]. Whether a true epithelial-mesenchymal-myofibroblast transition, or a more intermediate state of transdifferentiation, contributes to the pathophysiology of human fibrotic disease, however, is the subject of considerable debate [10–16].

The temporal and spatial regulation of cellular plasticity, as well as the subsequent restitution of an epitheloid phenotype, is likely a collective response to specific growth factors (individually or in combination) and informational cues from the extracellular environment [2, 8, 17]. The nature of the initiating stimulus as well as the underlying pathology and associated genetic reprogramming also impact temporal control versus persistence of the plastic restructuring. Epidermal growth factor receptor (EGFR) amplification and alterations in the transforming growth factor-β (TGF-β) signal transduction network, for example, frequently accompany epithelial tumor progression from a benign or noninvasive lesion to an aggressive, metastatic carcinoma [18–20]. During this transition, and despite increased autocrine/paracrine expression of TGF-β, cells often become refractory to the normally growth-suppressive effects of TGF-β family members due, in part, to downregulation of TGF-β receptors and/or anomalies in TGF-β-initiated signaling pathways [20, 21]. Similarly, in tissue fibrotic disorders, differentiation, proliferation, and subsequent interstitial accumulation of “fibroblastoid” elements in the kidney and lung (two of the most well-studied organ systems in the context of fibroproliferative disease) appear to result from a programmed, but persistent, response to several profibrotic cytokines, the most prominent of which is TGF-β [11].

2. The Serine Protease-Matrix Metalloproteinase Cascade in Tissue Remodeling

TGF-β promotes cellular motile and invasive properties, as well as the emergence of the plastic cohort, through expression of a subset of genes that encode various classes of stromal remodeling proteins [22, 23]. These include members of the serine protease and matrix metalloproteinase (MMP) families and their respective inhibitors which, paradoxically, support matrix disruptive as well as stabilizing processes. Several structural components of the extracellular matrix [24, 25] are, in fact, upregulated by TGF-β as are matrix-active proteases (e.g., urokinase (uPA), plasmin, MMP-1, -3, -9, -10, -11, and -13) and protease inhibitors [26–29]. Stringent controls on serine protease/MMP transcription, duration of expression, and topographic activity are essential for maintaining tissue homeostasis in the intact organism as well as in organotypic systems [30]. Proteolytic networks within the pericellular microenvironment, moreover, are frequently activated by the conversion of plasminogen to plasmin, a broad-spectrum protease. Plasmin, in turn, targets stromal elements directly while also activating several MMPs triggering a complex cascade leading to matrix degradation [31]. Upstream plasmin generation substantially impacts MMP-dependent stromal remodeling and, thereby, cellular invasive traits. Such in vivo pathologies can be elegantly modeled in vitro upon addition of EGF + TGF-β1 to malignant human epithelial cells to mimic the frequently observed TGF-β1 elevation in the tumor microenvironment and amplified EGFR signaling characteristic of late-stage cancers [32]. Combined EGF + TGF-β1 costimulation resulted in the synergistic upregulation of a defined set of proinvasive genes, the most prominent of which encodes plasminogen activator inhibitor-type-1 (PAI-1 or SERPINE1, the clade E member 1 of the family of serine protease inhibitors). PAI-1 is a potent and fast-acting inhibitor of uPA-dependent plasmin production [22, 23]. This finding is of considerable translational relevance since increased PAI-1 levels often occur in concert with epithelial cell plasticity, paralleling the requirement for enhanced cell motility [33]. The ability of TGF-β and/or EGF to increase PAI-1 expression in several cell types [26, 34] provides a potential mechanism for upstream titration of the MMP cascade via controlled generation of pericellular plasmin consequently modulating, both in time and space, extracellular matrix proteolysis and stromal remodeling. Indeed, elevated PAI-1 levels commonly accompany the development of such diverse pathologies as tumor progression, inflammation, hypertrophic scarring, atherosclerosis, thrombosis, myocardial infarction, diabetes, and the obesity-associated metabolic syndrome [11, 31, 35–40].

3. Focal Proteolysis: Regulation of Cell Migration and Signaling

The contribution of PAI-1 as a promoting element in various disease states is thought to occur through multiple avenues involving proteolytic control, an essential aspect in the maintenance of a “stromal scaffold” that impacts cell survival, growth and transdifferentiation, cellular motile processes, and signal transduction. Focal proteolysis within the pericellular microenvironment is controlled primarily through mechanisms that regulate plasminogen activation at the cell surface that, in turn, affect MMP activation downstream with subsequent engagement of a complex tissue remodeling program [41] (Figure 1).

Importantly, focalized proteolysis promotes the discrete release of several physiologically significant bioactive fragments and growth factors from the stromal compartment that influence cell proliferation and cell migration. MMP-dependent generation of degradation products of extracellular matrix structural elements, for example, affects both angiogenic and antiangiogenic activities with an impact on endothelial motile characteristics under in vivo-relevant conditions [42]. MMP-2 and MMP-9 cleave collagen IV, exposing cryptic epitopes that stimulate angiogenesis [43, 44] while matrikines such as arrestin, canstatin, tumstatin and metastatin, also generated from collagen IV, are antiangiogenic [41, 42]. Proteolytically derived fragments of collagen XVIII (endostatin and neostatin), collagen VIII (vastatin), collagen XV (restin), and perlecan (endorepellin) similarly exhibit anti-angiogenic properties [41, 42]. These fragments often compete with intact extracellular matrix molecules for binding to various cell surface receptors as one mechanistic basis for their effects [41]. A particularly important event involves the MMP-dependent release of laminin-332 fragments that promote epithelial cell migration. Indeed, the recombinant domain III of the laminin-332 γ2 chain (processed from laminin-332 by MT1-MMP, a membrane type 1 MMP, and MMP-2) binds to EGFR and initiates signaling events which culminate in enhanced cell motility [45–47]. Similarly, MMP-based proteolysis of fibronectin yields fragments that affect migration (MSF) [41, 42, 48], angiogenesis (anastellin) [49, 50], cell proliferation, and differentiation [41]. Stromal PAI-1 is itself a substrate for several extracellular proteases including elastase, MMP-3 and plasmin resulting in the generation of rather specific PAI-1 cleavage products [32, 33, 51–53]. Such “cleaved” PAI-1 is unable to bind its target plasminogen activators uPA and tissue-type PA (tPA) to inhibit plasmin-based proteolysis but retains the ability to bind to the low-density lipoprotein receptor-related protein-1 (LRP1) where it effectively augments cell migration through a uPA/tPA complex-independent interaction [54]. The mechanistic basis for this response is not clear but appears to involve LRP1 function as a key signaling mediator in several intracellular pathways as a consequence of, in part, interactions with multiple adaptor and scaffolding proteins [55]. LRP1 ligand binding and/or complex formation with additional surface receptors also activates specific mitogen-activated protein (MAP) and src kinases [56–60] stimulating cell proliferation [58, 61–63] and migration [54, 56, 64] with the motile outcome dependent on Rho family GTPases [64]. Alternatively, PAI-1 can also initiate signaling events that impact cell migration through engagement of LRP1 and the related very low-density lipoprotein receptor [65]. Indeed, different conformations of PAI-1 (active, latent as well as plasmin- or MMP-cleaved) all interact with LRP1 to enhance cell migration into physiological scaffolds or stromal equivalents [66]. The three forms of PAI-1 appear to increase LRP1-dependent cell motility via specific engagement of the Jak/Stat1 signaling pathway [54, 67, 68]. These data are consistent with recent findings that the migratory response to TGF-β1 + EGF in transformed human epidermal keratinocytes is PAI-1-dependent [22, 23] and that stabilized (i.e., long half-life) recombinant PAI-1 alone, in the absence of added growth factors, stimulates motility comparable to that attained by growth factor supplementation (Figure 2). The receptor-associated protein (RAP), an LRP1 antagonist which binds LRP1 and blocks interactions with all known ligands including PAI-1, inhibits the migration-promoting effects of PAI-1 in chemotactic and wound healing assays [54] and effectively inhibited EGF-stimulated motility, which is known to be dependent on LRP1/PAI-1 (Figure 2). 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 [33]. Collectively, these data illustrate that serine protease/MMP-initiated proteolytic processing of the extracellular environment impacts multiple aspects with regard to the regulation of cell motility.

(a)

(b)

Figure 2

HaCaT II-4 cultures were grown to confluency, monolayers washed 2X with PBS and incubated in serum-free DMEM for 24 hours. Cultures were scrape-wounded with a P1000 pipette tip using a constant-pressure press, washed 3X to remove liberated cells, and returned to serum-free medium without (control) or with the following additives: EGF (10 ng/mL), PAI-1 (40 mM), RAP (5 μg/mL), or the combination RAP (5 μg/mL) + EGF (10 ng/mL). Initial wound sizes were measured with a calibrated ocular grid at multiple marked sites; 24 hours later, the injury sites were remeasured at the identical marked regions used to calibrate the initial denuded area. The extent of wound site closure (motility index = grid distance migrated) was plotted on the -axis for each condition; shown are data from 2 representative experiments. PAI-1 stimulated cell migration to the same extent as EGF. Ctl: control unstimulated cultures.

4. Tumor Microenvironment and Cutaneous Wound Repair: Sites of Interacting Proteolytic Cascades

Amplified MMP expression correlates with tumor aggressiveness, metastasis, and poor prognosis [69]. Not surprisingly, therefore, recent studies implicate several MMPs, including MMP-3, -7, -9, and -28 in triggering plasticity-related processes [69]. The combination of TGF-β + EGF effectively promotes epithelial-to-mesenchymal transition and upregulates MMPs-1, -3, -9, -10, and 14 [22, 23, 70, 71], coupling cellular invasive potential to a plasmin/MMP-10/MMP-1-dependent collagen-remodeling axis. As a proof-of-concept, an acute collagenolytic phenotype, linked to plasmin-dependent activation of stromelysin-2 (MMP-10), accompanies costimulation of malignant (HaCaT II-4) keratinocytes with TGF-β1 and EGF coincident with collagen invasion [32] (Figure 3). MMP-10, which is generally limited to epithelial cells [29, 72], targets proMMPs-1, -7, -8, -9, and -13, as well as collagens type III, IV, and V, gelatin elastin, fibronectin, proteoglycans, and laminin [30, 31]. MMP-10 induction in response to TGF-β + EGF suggests that precise control over its levels and activation are likely critical for cutaneous homeostasis. MMP-10, in fact, is not evident in intact skin but expressed during cutaneous injury repair localizing to migrating keratinocytes at the wound edge, suggesting a role in invasive behavior [73].

Figure 3

Model of TGF-β1 + EGF-enhanced plasmin-dependent collagen matrix remodeling and development of an invasive phenotype. In the presence of plasmin, increased MMP-10 levels promote MMP activation creating a proteolytic axis that accelerates collagen degradation through “superactivation” of MMP-1. STAT3 may act as a positive switch in this process, via promotion of EGF-stimulated proMMP-10 expression [23]. Upregulation of PAI-1 in response to TGF-β1 + EGF may subsequently shift this proteolytic balance, enabling PAI-1 to “titrate” the extent and locale of collagen matrix remodeling to facilitate stromal invasion. Indeed, PAI-1 induction occurs early in this transition and required for stimulated migration and collagen invasion since PAI-1 knockdown (with siRNA constructs) effectively inhibited both events [22].

Similar to other systems [29, 71, 72, 74], MMP-10 and MMP-1 are upregulated in TGF-β1 and/or EGF-stimulated human malignant keratinocytes maintained on collagen substrates [32]. The proenzyme forms of MMP-1 and MMP-10 are plasmin substrates. Following MMP-10 inhibition, however, the residual level of active plasmin-generated MMP-1 appears insufficient to initiate collagen dissolution [32]. This reflects the established ability of MMP-10 to “superactivate” or enhance MMP-1-dependent proteolysis, as well as that of MMP-8 and -13, and significantly enhances collagenolytic activity over that observed with plasmin alone [72, 75]. Although plasmin is clearly an important primary activator of this complex cascade, the cathepsins, like MMPs, also associate with tumor cell invasion. This is particularly true for the cysteine proteinases cathepsin L and cathepsin B, which degrade type-1 collagen and mobilize several MMPs (including MMP-1), respectively [76]. Inhibition of cysteine cathepsins had no effect, however, on collagen dissolution in the HaCaT II-4 model while serine proteinase blockade effectively attenuated collagen degradation [32]. These data reinforce the critical role for active plasmin, and not cathepsins, in the initiation of collagen degradation by TGF-β1 + EGF and are consistent with observations regarding the downregulation of several cathepsins by TGF-β [77].

The coupling of keratinocyte-based type-1 collagen degradation with plasminogen activation implicates intermediates other than MMP-10, including MMP-13 [78, 79]. The available evidence indicates the existence of a functional overlap between MMP-13 and uPA in the context of cutaneous wound repair, at least in the murine system [80]. Contrary to what has been observed in primary human keratinocytes, MMP-13 expression is, in fact, linked to transformation of human keratinocytes (i.e., HaCaT cells and various derivatives) [27, 74] and actually enhanced following addition of TGF-β1 to HaCaT II-4 cells [32, 74]. Use of a physiological 3-D reconstruct model of the stromal microenvironment, however, led to the conclusion that the combination of TGF-β1 + EGF does not significantly upregulate MMP-13 protein in HaCaT II-4 cells but, instead, leads to a robust induction of MMP-10 [32]. This disparity may be due in part to differences among the ras-HaCaT variants in MMP expression programs [81], TGF-β/EGF receptor crosstalk [11, 71, 82], or culture in 2D versus a more complex 3D stromal-equivalent system [32].

Models of cutaneous injury repair have shed considerable light on the complex roles of growth factors and cascading protease systems in the tissue response to trauma. Generally, both TGF-β1 and EGF levels increase substantially following acute injury, partially due to their release from platelet α granules, but also through increased cellular expression, particularly at the wound edge [83]. These growth factors appear critical to the initial stages of cutaneous tissue regeneration through promotion of keratinocyte migration, as well as proliferation [84–87]. TGF-β1 and EGF upregulate MMP-10 in keratinocytes [72, 88] and, during cutaneous wound repair, MMP-10 is specifically localized to cells in the migrating tongue where it appears to enhance migration [87, 88]. Similarly, uPA and MMP-3, -9, and -13 all localize to leading edge epidermal cells [80]. Overexpression of constitutively active MMP-10 in the epidermis, moreover, has deleterious effects on the coordinated migration of keratinocytes into the wound bed; an effect attributed to excessive laminin-5 (laminin-332) processing [87]. Unconstrained MMP-10 activity leads to excessive collagenolysis [32] which impacts negatively on cell migration and, ultimately, the restoration of tissue integrity. Notably, PAI-1 expression also increases in keratinocytes at the wound margin and is deposited into the migration tracks of these cells, suggesting that this SERPIN, as well, plays an integral role in regulating directional migration and wound closure [89–92]. Coordinate upregulation of proteolytic enzymes such as the MMPs, together with the upstream inhibitor of plasmin generation (i.e., PAI-1) by individual growth factors provides an exquisite mechanism for fine control of focal proteolysis to facilitate optimal cell motility in complex environments.

5. Conclusions

Increases in epithelial MMP-10 expression, and its subsequent activation by catalytic-levels of plasmin, mobilize an MMP cascade creating a proteolytic axis that accelerates collagen degradation through “superactivation” of MMP-1 while enhancing stromal proteolysis largely by MMP-7, -8, -9, and -13. Upregulation of the serine protease inhibitor PAI-1, in tumor cells, in mesenchymal cells within the tumor microenvironment as well as by “wound-stimulated” epithelial cells, may subsequently shift this proteolytic balance to optimize creation of a migratory “scaffold.” The available data clearly implicate PAI-1 as a major upstream modulator of a uPA→plasmin-generating system that exerts fine control over the MMP-dependent pericellular proteolytic cascade. In this context, PAI-1 may “titrate” the extent and locale of collagen matrix remodeling to facilitate cellular invasion within the stromal compartment as part of the metastatic and tissue repair programs. Further clarification of the complexity of controls and the extent of interdependency of individual cascading “arms” in this highly interactive network of matrix proteases and protease inhibitors will be necessary for the rationale design of focused therapeutic approaches for the treatment of cancer, fibrotic disorders and chronic wounds. Assessments of MMP inhibitors in clinical trials is already ongoing. Indeed, the emergence of uPA and PAI-1 as significant level-of-evidence-1 prognostic markers of overall survival in breast cancer is well established [93]. The development of small molecule inhibitors of PAI-1 (i.e., tiplaxtinin or PAI-039) that effectively attenuate aortic remodeling in the context of vascular injury [94] suggest that targeting this SERPIN may have translational implications for treatment of chronic fibrotic and, perhaps, malignant disease.

Acknowledgment

This work was supported by NIH Grant GM57242 and a Grant from the Butler Family Foundation for Mesothelioma Research to P. J. H.

References

J. P. Thiery, “Epithelial-mesenchymal transitions in development and pathologies,” Current Opinion in Cell Biology, vol. 15, no. 6, pp. 740–746, 2003.
View at: Publisher Site | Google Scholar
B. Boyer, A. M. Vallés, and N. Edme, “Induction and regulation of epithelial-mesenchymal transitions,” Biochemical Pharmacology, vol. 60, no. 8, pp. 1091–1099, 2000.
View at: Publisher Site | Google Scholar
C. Yan, W. A. Grimm, W. L. Garner et al., “Epithelial to mesenchymal transition in human skin wound healing is induced by tumor necrosis factor-α through bone morphogenic protein-2,” American Journal of Pathology, vol. 176, no. 5, pp. 2247–2258, 2010.
View at: Publisher Site | Google Scholar
P. Savagner, D. F. Kusewitt, E. A. Carver et al., “Developmental transcription factor slug is required for effective re-epithelialization by adult keratinocytes,” Journal of Cellular Physiology, vol. 202, no. 3, pp. 858–866, 2005.
View at: Publisher Site | Google Scholar
R. Kalluri, “EMT: when epithelial cells decide to become mesenchymal-like cells,” Journal of Clinical Investigation, vol. 119, no. 6, pp. 1417–1419, 2009.
View at: Publisher Site | Google Scholar
M. L. Ackland, D. F. Newgreen, M. Fridman et al., “Epidermal growth factor-induced epithelio-mesenchymal transition in human breast carcinoma cells,” Laboratory Investigation, vol. 83, no. 3, pp. 435–448, 2003.
View at: Google Scholar
J. Zavadil, M. Bitzer, D. Liang et al., “Genetic programs of epithelial cell plasticity directed by transforming growth factor-β,” Proceedings of the National Academy of Sciences of the United States of America, vol. 98, no. 12, pp. 6686–6691, 2001.
View at: Publisher Site | Google Scholar
J. P. Their, “Epithelial-mesenchymal transitions in tumor progression,” Nature Reviews Cancer, vol. 2, no. 6, pp. 442–454, 2002.
View at: Google Scholar
R. M. Carew, B. Wang, and P. Kantharidis, “The role of EMT in renal fibrosis,” Cell and Tissue Research, vol. 347, no. 2, pp. 103–116, 2012.
View at: Google Scholar
S. E. Quaggin and A. Kapus, “Scar wars: mapping the fate of epithelial-mesenchymal-myofibroblast transition,” Kidney International, vol. 80, no. 1, pp. 41–50, 2011.
View at: Publisher Site | Google Scholar
R. Samarakoon, J. M. Overstreet, S. P. Higgins et al., “TGF-b1 $?$ SMAD/p53/USF2 $?$ PAI-1 transcriptional axis in ureteral obstruction-induced renal fibrosis,” Cell and Tissue Research, vol. 347, no. 2, pp. 117–128, 2012.
View at: Google Scholar
I. Grgic, J. S. Duffield, and B. D. Humphreys, “The origin of interstitial myofibroblasts in chronic kidney disease,” Pediatric Nephrology, vol. 27, no. 2, pp. 183–193, 2012.
View at: Google Scholar
W. Kriz, B. Kaissling, and M. Le Hir, “Epithelial-mesenchymal transition (EMT) in kidney fibrosis: fact or fantasy?” Journal of Clinical Investigation, vol. 121, no. 2, pp. 468–474, 2011.
View at: Publisher Site | Google Scholar
A. Hertig, S. N. Flier, and R. Kalluri, “Contribution of epithelial plasticity to renal transplantation-associated fibrosis,” Transplantation Proceedings, vol. 42, no. 9, pp. S7–S12, 2010.
View at: Publisher Site | Google Scholar
M. Fragiadaki and R. M. Mason, “Epithelial-mesenchymal transition in renal fibrosis—evidence for and against,” International Journal of Experimental Pathology, vol. 92, no. 3, pp. 143–150, 2011.
View at: Publisher Site | Google Scholar
R. Kalluri and R. A. Weinberg, “The basics of epithelial-mesenchymal transition,” Journal of Clinical Investigation, vol. 119, no. 6, pp. 1420–1428, 2009.
View at: Publisher Site | Google Scholar
J. P. Thiery, H. Acloque, R. Y. J. Huang, and M. A. Nieto, “Epithelial-mesenchymal transitions in development and disease,” Cell, vol. 139, no. 5, pp. 871–890, 2009.
View at: Publisher Site | Google Scholar
W. Cui, D. J. Fowlis, S. Bryson et al., “TGFβ1 inhibits the formation of benign skin tumors, but enhances progression to invasive spindle carcinomas in transgenic mice,” Cell, vol. 86, no. 4, pp. 531–542, 1996.
View at: Publisher Site | Google Scholar
O. Rho, L. M. Beltran, I. B. Gimenez-Conti, and J. DiGiovanni, “Altered expression of the epidermal growth factor receptor and transforming growth factor-α during multistage skin carcinogenesis in SENCAR mice,” Molecular Carcinogenesis, vol. 11, no. 1, pp. 19–28, 1994.
View at: Publisher Site | Google Scholar
R. Derynck, R. J. Akhurst, and A. Balmain, “TGF-β signaling in tumor suppression and cancer progression,” Nature Genetics, vol. 29, no. 2, pp. 117–129, 2001.
View at: Publisher Site | Google Scholar
G. Han, S. L. Lu, A. G. Li et al., “Distinct mechanisms of TGF-β1-mediated epithelial-to- mesenchymal transition and metastasis during skin carcinogenesis,” Journal of Clinical Investigation, vol. 115, no. 7, pp. 1714–1723, 2005.
View at: Publisher Site | 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-β1+EGF-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
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, vol. 2009, article 963209, pp. 1–12, 2009.
View at: Google Scholar
A. B. Roberts, U. I. Heine, K. C. Flanders, and M. B. Sporn, “Transforming growth factor-β. Major role in regulation of extracellular matrix,” Annals of the New York Academy of Sciences, vol. 580, pp. 225–232, 1990.
View at: Publisher Site | Google Scholar
T. M. Vollberg, M. D. George, and A. M. Jetten, “Induction of extracellular matrix gene expression in normal human keratinocytes by transforming growth factor β is altered by cellular differentiation,” Experimental Cell Research, vol. 193, no. 1, pp. 93–100, 1991.
View at: Google Scholar
S. Akiyoshi, M. Ishii, N. Nemoto, M. Kawabata, H. Aburatani, and K. Miyazono, “Targets of transcriptional regulation by transforming growth factor-β: expression profile analysis using oligonucleotide arrays,” Japanese Journal of Cancer Research, vol. 92, no. 3, pp. 257–268, 2001.
View at: Google Scholar
N. Johansson, J. Westermarck, S. Leppä et al., “Collagenase 3 (matrix metalloproteinase 13) gene expression by HaCaT keratinocytes is enhanced by tumor necrosis factor α and transforming growth factor β1,” Cell Growth and Differentiation, vol. 8, no. 2, pp. 243–250, 1997.
View at: Google Scholar
H. S. Kim, T. Shang, Z. Chen, S. C. Pflugfelder, and D. Q. Li, “TGF-β1 stimulates production of gelatinase (MMP-9), collagenases (MMP-1, -13) and stromelysins (MMP-3, -10, -11) by human corneal epithelial cells,” Experimental Eye Research, vol. 79, no. 2, pp. 263–274, 2004.
View at: Publisher Site | Google Scholar
M. Madlener, C. Mauch, W. Conca, M. Brauchle, W. C. Parks, and S. Werner, “Regulation of the expression of stromelysin-2 by growth factors in keratinocytes: implications for normal and impaired wound healing,” Biochemical Journal, vol. 320, no. 2, pp. 659–664, 1996.
View at: Google Scholar
S. Chakraborti, M. Mandal, S. Das, A. Mandal, and T. Chakraborti, “Regulation of matrix metalloproteinases. An overview,” Molecular and Cellular Biochemistry, vol. 253, no. 1-2, pp. 269–285, 2003.
View at: Publisher Site | Google Scholar
H. R. Lijnen, “Matrix metalloproteinases and cellular fibrinolytic activity,” Biochemistry, vol. 67, no. 1, pp. 92–98, 2002.
View at: Google Scholar
C. E. Wilkins-Port, Q. Ye, J. E. Mazurkiewicz, and P. J. Higgins, “TGF-β1+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
R.-P. Czekay, C. E. Wilkins-Port, S. P. Higgins et al., “PAI-1: an integrator of cell signaling and migration,” International Journal of Cell Biology, vol. 2011, article 562481, pp. 1–9, 2011.
View at: Publisher Site | Google Scholar
S. M. Kutz, C. E. Higgins, R. Samarakoon et al., “TGF-β1-induced PAI-1 expression is e box/USF-dependent and requires EGFR signaling,” Experimental Cell Research, vol. 312, no. 7, pp. 1093–1105, 2006.
View at: Publisher Site | Google Scholar
P. A. Andreasen, L. Kjøller, L. Christensen, and M. J. Duffy, “The urokinase-type plasminogen activator system in cancer metastasis: a review,” International Journal of Cancer, vol. 72, no. 1, pp. 1–22, 1997.
View at: Publisher Site | Google Scholar
R. D. Balsara, Z. Xu, and V. A. Ploplis, “Targeting plasminogen activator inhibitor-1: role in cell signaling and the biology of domain-specific knock-in mice,” Current Drug Targets, vol. 8, no. 9, pp. 982–995, 2007.
View at: Publisher Site | Google Scholar
M. K. Durand, J. S. Bødker, 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
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
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
A. K. Ghosh and D. E. Vaughan, “PAI-1 in tissue fibrosis,” Journal of Cellular Physiology, vol. 227, no. 2, pp. 493–507, 2012.
View at: Publisher Site | Google Scholar
J. D. Mott and Z. Werb, “Regulation of matrix biology by matrix metalloproteinases,” Current Opinion in Cell Biology, vol. 16, no. 5, pp. 558–564, 2004.
View at: Publisher Site | Google Scholar
A. G. Arroyo and M. L. Iruela-Arispe, “Extracellular matrix, inflammation, and the angiogenic response,” Cardiovascular Research, vol. 86, no. 2, pp. 226–235, 2010.
View at: Publisher Site | Google Scholar
M. Hangai, N. Kitaya, J. Xu et al., “Matrix metalloproteinase-9-dependent exposure of a cryptic migratory control site in collagen is required before retinal angiogenesis,” American Journal of Pathology, vol. 161, no. 4, pp. 1429–1437, 2002.
View at: Google Scholar
J. Xu, D. Rodriguez, E. Petitclerc et al., “Proteolytic exposure of a cryptic site within collagen type IV is required for angiogenesis and tumor growth in vivo,” Journal of Cell Biology, vol. 154, no. 5, pp. 1069–1079, 2001.
View at: Publisher Site | Google Scholar
C. Gilles, M. Polette, C. Coraux et al., “Contribution of MT1-MMP and of human laminin-5 γ2 chain degradation to mammary epithelial cell migration,” Journal of Cell Science, vol. 114, no. 16, pp. 2967–2976, 2001.
View at: Google Scholar
N. Koshikawa, G. Giannelli, V. Cirulli, K. Miyazaki, and V. Quaranta, “Role of cell surface metalloprotease MT1-MMP in epithelial cell migration over laminin-5,” Journal of Cell Biology, vol. 148, no. 3, pp. 615–624, 2000.
View at: Publisher Site | Google Scholar
S. Schenk, E. Hintermann, M. Bilban et al., “Binding to EGF receptor of a laminin-5 EGF-like fragment liberated during MMP-dependent mammary gland involution,” Journal of Cell Biology, vol. 161, no. 1, pp. 197–209, 2003.
View at: Publisher Site | Google Scholar
S. L. Schor, I. R. Ellis, S. J. Jones et al., “Migration-stimulating factor: a genetically truncated onco-fetal fibronectin isoform expressed by carcinoma and tumor-associated stromal cells,” Cancer Research, vol. 63, no. 24, pp. 8827–8836, 2003.
View at: Google Scholar
R. Pasqualini, S. Bourdoulous, E. Koivunen, V. L. Woods, and E. Ruoslahti, “A polymeric form of fibronectin has antimetastatic effects against multiple tumor types,” Nature Medicine, vol. 2, no. 11, pp. 1197–1203, 1996.
View at: Publisher Site | Google Scholar
M. Yi and E. Ruoslahti, “A fibronectin fragment inhibits tumor growth, angiogenesis, and metastasis,” Proceedings of the National Academy of Sciences of the United States of America, vol. 98, no. 2, pp. 620–624, 2001.
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
H. R. Lijnen, B. Arza, B. van Hoef, D. Collen, and P. J. Declerck, “Inactivation of plasminogen activator inhibitor-1 by specific proteolysis with stromelysin-1 (MMP-3),” Journal of Biological Chemistry, vol. 275, no. 48, pp. 37645–37650, 2000.
View at: Publisher Site | 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
J. Herz and D. K. Strickland, “LRP: a multifunctional scavenger and signaling receptor,” Journal of Clinical Investigation, vol. 108, no. 6, pp. 779–784, 2001.
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 α2-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
S. C. Muratoglu, I. Mikhailenko, C. Newton, M. Migliorini, and D. K. Strickland, “The LDL receptor-related protein 1 (LRP1) forms a signaling complex with platelet-derived growth factor receptor-β 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
Y. Shi, E. Mantuano, G. Inoue, W. M. Campana, and S. L. Gonias, “Ligand binding to LRP1 transactivates trk receptors by a src family kinase-Dependent pathway,” Science Signaling, vol. 2, no. 68, p. ra18, 2009.
View at: Publisher Site | Google Scholar
L. Zhou, Y. Takayama, P. Boucher, M. D. Tallquist, and J. Herz, “LRP1 regulates architecture of the vascular wall by controlling PDGFRβ-dependent phosphatidylinositol 3-kinase activation,” PLoS ONE, vol. 4, no. 9, Article ID e6922, 2009.
View at: Publisher Site | Google Scholar
P. Boucher, M. Gotthardt, W. P. Li, R. G. W. Anderson, and J. Herz, “LRP: role in vascular wall integrity and protection from atherosclerosis,” Science, vol. 300, no. 5617, pp. 329–332, 2003.
View at: Publisher Site | Google Scholar
P. Boucker, W.-P. Li, R. L. Matz et al., “LRP1 functions as an atheroprotective integrator of TGFβ and PDGF signals in the vascular wall: implications for Marfan syndrome,” PLoS ONE, vol. 2, no. 5, article e448, 2007.
View at: Publisher Site | Google Scholar
S. S. Huang, T. Y. Ling, W. F. Tseng et al., “Cellular growth inhibition by IGFBP-3 and TGF-β1 requires LRP-1,” FASEB Journal, vol. 17, no. 14, pp. 2068–2081, 2003.
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. 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
S. X. Hou, Z. Zheng, X. Chen, and N. Perrimon, “The JAK/STAT pathway in model organisms: emerging roles in cell movement,” Developmental Cell, vol. 3, no. 6, pp. 765–778, 2002.
View at: Publisher Site | Google Scholar
L. S. Orlichenko and D. C. Radisky, “Matrix metalloproteinases stimulate epithelial-mesenchymal transition during tumor development,” Clinical and Experimental Metastasis, vol. 25, no. 6, pp. 593–600, 2008.
View at: Publisher Site | Google Scholar
Y. C. Tian, Y. C. Chen, C. T. Chang et al., “Epidermal growth factor and transforming growth factor-β1 enhance HK-2 cell migration through a synergistic increase of matrix metalloproteinase and sustained activation of ERK signaling pathway,” Experimental Cell Research, vol. 313, no. 11, pp. 2367–2377, 2007.
View at: Publisher Site | Google Scholar
S. Uttamsingh, X. Bao, K. T. Nguyen et al., “Synergistic effect between EGF and TGF-β1 in inducing oncogenic properties of intestinal epithelial cells,” Oncogene, vol. 27, no. 18, pp. 2626–2634, 2008.
View at: Publisher Site | Google Scholar
L. J. Windsor, H. Grenett, B. Birkedal-Hansen, M. K. Bodden, J. A. Engler, and H. Birkedal-Hansen, “Cell type-specific regulation of SL-1 and SL-2 genes. Induction of the SL- 2 gene but not the SL-1 gene by human keratinocytes in response to cytokines and phorbolesters,” Journal of Biological Chemistry, vol. 268, no. 23, pp. 17341–17347, 1993.
View at: Google Scholar
M. Krampert, W. Bloch, T. Sasaki et al., “Activities of the matrix metalloproteinase stromelysin-2 (MMP-10) in matrix degradation and keratinocyte organization in wounded skin,” Molecular Biology of the Cell, vol. 15, no. 12, pp. 5242–5254, 2004.
View at: Publisher Site | Google Scholar
N. Johansson, R. Ala-Aho, V. J. Uitto et al., “Expression of collagenase-3 (MMP-13) and collagenase-1 (MMP-1) by transformed keratinocytes is dependent on the activity of p38 mitogen-activated protein kinase,” Journal of Cell Science, vol. 113, no. 2, pp. 227–235, 2000.
View at: Google Scholar
H. E. Barksby, J. M. Milner, A. M. Patterson et al., “Matrix metalloproteinase 10 promotion of collagenolysis via procollagenase activation: implications for cartilage degradation in arthritis,” Arthritis and Rheumatism, vol. 54, no. 10, pp. 3244–3253, 2006.
View at: Publisher Site | Google Scholar
F. Song, K. Wisithphrom, J. Zhou, and L. J. Windsor, “Matrix metalloproteinase dependent and independent collagen degradation,” Frontiers in Bioscience, vol. 11, supplement 3, pp. 3100–3120, 2006.
View at: Publisher Site | Google Scholar
A. Gerber, A. Wille, T. Welte et al., “Interleukin-6 and transforming growth factor-ß1 control expression of cathepsins B and L in human lung epithelial cells,” Journal of Interferon and Cytokine Research, vol. 21, no. 1, pp. 11–19, 2001.
View at: Publisher Site | Google Scholar
S. Netzel-Arnett, D. J. Mitola, S. S. Yamada et al., “Collagen dissolution by keratinocytes requires cell surface plasminogen activation and matrix metalloproteinase activity,” Journal of Biological Chemistry, vol. 277, no. 47, pp. 45154–45161, 2002.
View at: Publisher Site | Google Scholar
H. Birkedal-Hansen, H. Y. Lin, B. Birkedal-Hansen, L. J. Windsor, and M. C. Pierson, “Degradation of collagen fibrils by live cells: role of expression and activation of procollagenase,” Matrix, supplement 1, pp. 368–374, 1992.
View at: Google Scholar
A. Juncker-Jensen and L. R. Lund, “Phenotypic overlap between MMP-13 and the plasminogen activation system during wound healing in mice,” PLoS ONE, vol. 6, no. 2, article e16954, 2011.
View at: Publisher Site | Google Scholar
L. C. Meade-Tollin, P. Boukamp, N. E. Fusenig, C. P. R. Bowen, T. C. Tsang, and G. T. Bowden, “Differential expression of matrix metalloproteinases in activated c-ras(Ha)-transfected immortalized human keratinocytes,” British Journal of Cancer, vol. 77, no. 5, pp. 724–730, 1998.
View at: Google Scholar
C. K. Joo, H. S. Kim, J. Y. Park, Y. Seomun, M. J. Son, and J. T. Kim, “Ligand release-independent transactivation of epidermal growth factor receptor by transforming growth factor-β involves multiple signaling pathways,” Oncogene, vol. 27, no. 5, pp. 614–628, 2008.
View at: Publisher Site | Google Scholar
S. Barrientos, O. Stojadinovic, M. S. Golinko, H. Brem, and M. Tomic-Canic, “Growth factors and cytokines in wound healing,” Wound Repair and Regeneration, vol. 16, no. 5, pp. 585–601, 2008.
View at: Publisher Site | Google Scholar
C. K. Jiang, T. Magnaldo, M. Ohtsuki, I. M. Freedberg, F. Bernerd, and M. Blumenberg, “Epidermal growth factor and transforming growth factor α specifically induce the activation- and hyperproliferation-associated keratins 6 and 16,” Proceedings of the National Academy of Sciences of the United States of America, vol. 90, no. 4, pp. 6786–6790, 1993.
View at: Google Scholar
Y. Li, J. Fan, M. Chen, W. Li, and D. T. Woodley, “Transforming growth factor-alpha: a major human serum factor that promotes human keratinocyte migration,” Journal of Investigative Dermatology, vol. 126, no. 9, pp. 2096–2105, 2006.
View at: Publisher Site | Google Scholar
G. Schultz, D. S. Rotatori, and W. Clark, “EGF and TGF-α in wound healing and repair,” Journal of Cellular Biochemistry, vol. 45, no. 4, pp. 346–352, 1991.
View at: Google Scholar
M. Krampert, W. Bloch, T. Sasaki et al., “Activities of the matrix metalloproteinase stromelysin-2 (MMP-10) in matrix degradation and keratinocyte organization in wounded skin,” Molecular Biology of the Cell, vol. 15, no. 12, pp. 5242–5254, 2004.
View at: Publisher Site | Google Scholar
M. Madlener, C. Mauch, W. Conca, M. Brauchle, W. C. Parks, and S. Werner, “Regulation of the expression of stromelysin-2 by growth factors in keratinocytes: implications for normal and impaired wound healing,” Biochemical Journal, vol. 320, no. 2, pp. 659–664, 1996.
View at: Google Scholar
F. Li, J. Goncalves, K. Faughnan et al., “Targeted inhibition of wound-induced PAI-1 expression alters migration and differentiation in human epidermal keratinocytes,” Experimental Cell Research, vol. 258, no. 2, pp. 245–253, 2000.
View at: Publisher Site | Google Scholar
K. M. Providence, S. M. Kutz, L. Staiano-Coico, and P. J. Higgins, “PAI-1 gene expression is regionally induced in wounded epithelial cell monolayers and required for injury repair,” Journal of Cellular Physiology, vol. 182, no. 2, pp. 269–280, 2000.
View at: Publisher Site | Google Scholar
K. M. Providence and P. J. Higgins, “PAI-1 expression is required for epithelial cell migration in two distinct phases of in vitro wound repair,” Journal of Cellular Physiology, vol. 200, no. 2, pp. 297–308, 2004.
View at: Publisher Site | Google Scholar
K. M. Providence, S. P. Higgins, A. Mullen et al., “SERPINE1 (PAI-1) is deposited into keratinocyte migration "trails" and required for optimal monolayer wound repair,” Archives of Dermatological Research, vol. 300, no. 6, pp. 303–310, 2008.
View at: Publisher Site | Google Scholar
M. Schmitt, K. Mengele, R. Napieralski et al., “Clinical utility of level-of-evidence-1 disease forecast cancer biomarkers uPA and its inhibitor PAI-1,” Expert Review of Molecular Diagnostics, vol. 10, no. 8, pp. 1051–1067, 2010.
View at: Publisher Site | Google Scholar
A. D. Weisberg, F. Albornoz, J. P. Griffin et al., “Pharmacological inhibition and genetic deficiency of plasminogen activator inhibitor-1 attenuates angiotensin II/salt-induced aortic remodeling,” Arteriosclerosis, Thrombosis, and Vascular Biology, vol. 25, no. 2, pp. 365–371, 2005.
View at: Publisher Site | Google Scholar

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Copyright © 2012 Cynthia E. Wilkins-Port et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

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