The phenotypic switching called epithelial-to-mesenchymal transition is frequently associated with epithelial tumor cell progression from a comparatively benign to an aggressive, invasive malignancy. Coincident with the emergence of such cellular plasticity is an altered response to transforming growth factor-β (TGF-β) as well as epidermal growth factor (EGF) receptor amplification. TGF-β in the tumor microenvironment promotes invasive traits largely through reprogramming gene expression, which paradoxically supports matrix-disruptive as well as stabilizing processes. ras-transformed HaCaT II-4 keratinocytes undergo phenotypic changes typical of epithelial-to-mesenchymal transition, acquire a collagenolytic phenotype, and effectively invade collagen type 1 gels as a consequence of TGF-β1 + EGF stimulation in a three-dimensional physiologically relevant model system that monitors collagen remodeling. Enhanced collagen degradation was coupled to a significant increase in matrix metalloproteinase (MMP)-10 expression and involved a proteolytic axis composed of plasmin, MMP-10, and MMP-1. Neutralization of any one component in this cascade inhibited collagen gel lysis. Similarly, addition of plasminogen activator inhibitor type 1 (SERPINE1) blocked collagen degradation as well as the conversion of both proMMP-10 and proMMP-1 to their catalytically active forms. This study therefore identifies an important mechanism in TGF-β1 + EGF-initiated collagen remodeling by transformed human keratinocytes and proposes a crucial upstream role for plasminogen activator inhibitor type 1–dependent regulation in this event. [Cancer Res 2009;69(9):4081–91]
- epithelial-to-mesenchymal transition
Epithelial tumor progression, from a relatively indolent to a more aggressive phenotype, is frequently accompanied by acquisition of a plastic phenotype reminiscent of a developmental program called epithelial-to-mesenchymal transition (EMT; ref. 1). This process is typified by loss of normal epithelial properties (e.g., cell polarity and junctional complexes) and a gain in mesenchymal traits (expression of vimentin and smooth muscle actin and enhanced cell motility; ref. 2). Although essential during embryonic development, EMT is relatively limited in the adult organism, occurring during wound repair or, more atypically, in advanced pathologies largely in response to specific growth factors associated with tumor progression ( 3– 5). Epidermal growth factor (EGF) receptor amplification and an altered cellular response to transforming growth factor-β (TGF-β), for example, accompany the progression of epithelial tumor cells from a benign phenotype to an aggressive, metastatic carcinoma ( 6– 8). During this pathologic EMT, and despite increased autocrine/paracrine expression of TGF-β, cells become refractory to the normally growth-suppressive effects of TGF-β family members. Mouse models of multistage skin carcinogenesis support the concept that TGF-β functions as a tumor suppressor in the early stages of benign growth; in late stage tumors, however, TGF-β accelerates malignant conversion ( 6). Down-regulation of TGF-β receptors, alterations in TGF-β-dependent Smad signaling components, or a combination of both appear to contribute to this functional switch ( 8, 9).
TGF-β likely promotes tumor-invasive properties through expression of genes that encode stromal remodeling proteins, which paradoxically support matrix-disruptive as well as stabilizing processes. Structural extracellular matrix proteins such as fibronectin and collagen ( 10, 11) are up-regulated by TGF-β in conjunction with their proteolytic regulators, including plasminogen activator inhibitor type 1 (PAI-1; ref. 12) and matrix metalloproteinase (MMP)-1, -3, -9, -10, -11, and -13 ( 13– 15). Similar to TGF-β, EGF-dependent signaling contributes to up-regulation of several MMPs ( 15– 19) and is enhanced through increased receptor levels in various cancers ( 7).
Stringent temporal and spatial controls on MMP activation are essential for maintaining tissue homeostasis. In addition to transcriptional regulation, MMP-dependent activities are also modulated through proteolytic activation ( 20). Proteolytic cascades within the pericellular environment are largely initiated through conversion of matrix plasminogen to the broad-spectrum protease plasmin, which, in turn, directly converts several proMMPs to their active form and triggers a positive feedback mechanism for MMP activation ( 21). Indeed, regulation of plasminogen activation may substantially affect MMP-dependent remodeling processes and thereby cellular invasive traits. The ability of TGF-β and/or EGF stimulation to also up-regulate PAI-1 in several cell types (this study; refs. 12, 22) provides a potential mechanism for upstream negative regulation or titration of the MMP cascade.
The immortalized adult human keratinocyte cell line HaCaT ( 23) harbors genetic changes similar to those that accompany progression of a normal keratinocyte to an invasive squamous cell carcinoma ( 24). Stimulation of activated ras-expressing HaCaT II-4 cells with TGF-β alone or, more effectively, a combination of TGF-β1 and EGF promotes a highly plastic phenotype typified by loss of E-cadherin and de novo synthesis of N-cadherin and vimentin ( 4, 25). The ability of TGF-β1 and/or EGF to elicit EMT-related responses such as these in a more physiologically significant model, however, has not been explored. This article describes the use of a three-dimensional collagen gel system to evaluate proteolytic events associated with TGF-β1 + EGF-stimulated EMT and collagen invasion by HaCaT II-4 keratinocytes. The invasive potential of these keratinocytes was coupled to a plasmin/MMP-10/MMP-1–dependent collagen-remodeling axis, and a role for PAI-1 as a critical upstream regulator of this remodeling process was established.
Materials and Methods
Reagents. Vitrogen (Cohesion Technologies) or PureCol (Inamed; Advanced BioMatrix) provided sources of bovine collagen type 1. Both products yielded comparable results and were used interchangeably. Where indicated, FITC-labeled collagen type 1 (Sigma-Aldrich) or DQ FITC-labeled collagen type 1 (Molecular Probes/Invitrogen) were added to monitor gel degradation. Recombinant human TGF-β1 (R&D Systems) was used at 1 ng/mL and recombinant human EGF (Upstate/Millipore) at 10 ng/mL. Plasminogen, aprotinin, E-64, amiloride, o-phenylenediamine dihydrochloride, hydrogen peroxide, and phosphate-citrate buffer were from Sigma-Aldrich. Recombinant human PAI-1 protein and α2-antiplasmin were from Calbiochem. GM6001 was obtained from Chemicon/Millipore and AG1478 was obtained from Biosource International/Invitrogen. Immunofluorescence antibodies included tubulin (clone DM1A; Sigma), vimentin (LN6 Ab-1; Calbiochem), E-cadherin (clone 36), and N-cadherin (clone 32) from BD Biosciences; MMP-10 from Santa Cruz Biotechnology; and 4′,6-diamidino-2-phenylindole stain, phalloidin-594, and Cell Tracker CMTPX from Molecular Probes/Invitrogen. For Western blotting, antibodies to MMP-1 and MMP-10 or biotinylated antibodies to MMP-1 and MMP-10 were obtained from R&D Systems; antibodies to actin or extracellular signal-regulated kinase 1/2 were from Santa Cruz Biotechnology. Antibodies against human plasminogen and PAI-1 (neutralizing) were from American Diagnostica. Neutralizing antibodies to MMP-1 and MMP-10 were obtained from R&D Systems. A polyclonal antibody to PAI-1 was used for ELISA and immunofluorescence. Unless otherwise indicated, Alexa Fluor 488 (green) or 594 (red) conjugates (Molecular Probes/Invitrogen) were used for immunocytochemistry detection; horseradish peroxidase conjugates (Pierce Biotechnology) were used for Western blot and ELISA analyses.
Cell culture. HaCaT II-4 keratinocytes were maintained in low-glucose DMEM supplemented with 10% fetal bovine serum (Life Technologies/Invitrogen). Cells were harvested with trypsin/EDTA, washed with PBS, and seeded onto collagen in serum-free Advanced DMEM overnight before stimulation with TGF-β1 and/or EGF. Phenol red-free medium (Life Technologies/Invitrogen) was used in fluorescence assays.
Collagen gel-based studies. Collagen type 1 was neutralized according to the manufacturer's instructions using 10× PBS and 0.1 N NaOH and then diluted to 1.8 mg/mL (unless stated otherwise) with DMEM. Gels were polymerized in 48-well tissue culture plates (150 μL), OptiCell Chambers (1.0 mL; USA Scientific), MatTek glass bottom dishes (200 μL; MatTek), or onto cell culture inserts (20 μL, 700 μg/mL) at 37°C for 2 to 3 h. For OptiCell-based invasion assays, 2 × 105 cells were added in 1 mL Advanced DMEM to gels polymerized vertically within the chamber. Cells were viewed on an inverted Olympus IX70 by laying the chamber on its side and images were captured with Image-Pro Plus software. For Transwell invasion assays, 1 × 105 cells were seeded onto thin collagen gels in Advanced DMEM. Invading cells were visualized with 4′,6-diamidino-2-phenylindole and counted in four random fields. For collagen gel dissolution assays, 5 × 104 cells were seeded onto polymerized gels in Advanced DMEM; plasminogen (5-20 μg/mL) was added 24 to 48 h post-growth factor stimulation for up to 24 h. Cells were pretreated with inhibitors or neutralizing antibodies as indicated. In experiments with anti-PAI-1 neutralizing antibody, time (not shown) and/or plasminogen concentration were reduced to capture differences in the state of dissolution with increasing antibody concentration. Cells on intact gels were fixed in 3% paraformaldehyde before viewing on an inverted Olympus IX70 microscope. To quantify collagen degradation, FITC-labeled collagen (25 μg/mL) was incorporated into polymerized gels. Cells were stimulated in phenol red-free DMEM and incubated with plasminogen for 7.5 h and 100 μL conditioned medium was removed for fluorescence spectroscopy using Synergy HT microplate reader equipped with KC4 software (BioTek Instruments).
Microscopy. HaCaT II-4 cells were seeded onto collagen-coated coverslips (50 μg/mL) or onto collagen gels polymerized in MatTek glass-bottomed dishes. Following treatment, cells were fixed in 3% paraformaldehyde, permeabilized, blocked, and incubated with primary and secondary antibodies for 1 h each. Coverslips were mounted using ProLong Gold with 4′,6-diamidino-2-phenylindole and viewed on an Olympus BX61 microscope with Image-Pro Lab software version 3.6.5. or an Olympus IX70 inverted scope with Image-Pro Plus software. For visualization of collagen digestion, cells were seeded onto coverslips coated with collagen type 1 (50 μg/mL) + DQ FITC-labeled collagen type 1 (25 μg/mL).
Protein analysis. Collagen gels were digested with collagenase D; cells were separated from digested collagen by centrifugation and lysed in a 50 mmol/L HEPES containing 150 mmol/L NaCl, 1% Triton X-100, 0.5% deoxycholate, 1% NP-40, 10 mmol/L NaF, 1 mmol/L orthovanadate, and protease inhibitors; and extracts were probed for N-cadherin and E-cadherin. Western blot analysis of MMP-1 and MMP-10 used conditioned medium from cells stimulated with TGF-β1 + EGF followed by incubation with plasminogen ± inhibitors (as indicated). The Human MMP Antibody Array 1.1 from RayBio (RayBiotech) was used to detect changes in MMP protein levels in conditioned medium. For measurement of PAI-1 levels by ELISA, 1.2 × 105 cells seeded on collagen type 1-coated, BSA-blocked wells were maintained under serum-free conditions for 6 h, pretreated with AG1478, as indicated, and then stimulated with TGF-β1 and/or EGF overnight. Cells were fixed with 3% paraformaldehyde, permeabilized, blocked, and incubated with PAI-1 polyclonal antibodies for 1 h followed by a horseradish peroxidase-conjugated secondary antibody. Cell layer PAI-1 was detected by colorimetric assay using an o-phenylenediamine dihydrochloride substrate and measured by spectrophotometer at 492 nm. Results were normalized to cell number by measuring the level of cell-associated crystal violet staining.
Statistical analysis. The Student's t test for two samples, assuming unequal variance, was used to compare conditions within a group. Two-tailed values with P ≤ 0.05 were considered significant.
HaCaT II-4 keratinocytes stimulated with TGF-β1 + EGF undergo EMT and invade collagen gels in a MMP-dependent manner. To recapitulate events associated with cutaneous EMT in a relevant context, p53 mutant, Ha-ras-expressing human keratinocytes (HaCaT II-4 cells; ref. 23) were cultured on a collagen coat ( Fig. 1A ) or onto a more physiologically related three-dimensional collagen gel ( Fig. 1B and C) and simultaneously treated with TGF-β1 and EGF to mimic the increased TGF-β expression/EGF receptor signaling characteristics of late-stage tumors. Under these conditions, EGF stimulation was mitogenic, whereas TGF-β1 maintained its growth-suppressive activity even in the presence of EGF (Supplementary Fig. S1). Whereas HaCaT II-4 colonies cultured on a three-dimensional collagen gel appeared more compact than cells cultured on a collagen coat, dually stimulated cells displayed traits typical of an EMT ( 2) on both substrates including increased scattering ( Fig. 1A and B; tubulin), de novo vimentin, and N-cadherin expression ( Fig. 1A–C) as well as loss of E-cadherin at cell-cell junctions ( Fig. 1A–C). To our knowledge, these observations represent the first evidence that EMT-related events take place in human keratinocytes cultured in a three-dimensional environment.
Because the phenotypic plasticity characteristic of EMT may promote tumor metastasis ( 5), it was important to evaluate the invasive capacities of TGF-β1 + EGF-treated HaCaT II-4 cells. TGF-β1 + EGF enhanced cell invasion and migration into a collagen matrix, as assessed in both OptiCell and Transwell three-dimensional systems ( Fig. 2A and B ), and was effectively attenuated by the broad-spectrum MMP inhibitor GM6001 ( Fig. 2C). Confocal imagery of cells seeded onto thin collagen gels (Supplementary Fig. S2) also supported the observation that TGF-β1 + EGF promoted HaCaT II-4 collagen gel invasion. Enhanced collagen type 1 degradation following TGF-β1 + EGF treatment was confirmed by using collagen matrices prepared from a quenched FITC-labeled collagen type 1 substrate that fluoresces on cleavage ( Fig. 2D). Together, these data indicate that TGF-β1 and EGF play integral roles in modulating HaCaT II-4-based collagenase activity, effectively supporting collagen gel invasion.
TGF-β1 + EGF treatment enhances collagen degradation via a plasmin/MMP dependent mechanism. Physiologic control of pericellular proteolysis occurs primarily through the regulation of plasminogen activation at the cell surface, which, in turn, contributes to downstream extracellular MMP activity ( Fig. 3A ). To explore the mechanisms associated with plasmin-based proteolysis in a cutaneous model, exogenous plasminogen was added to HaCaT II-4 cultures stimulated with TGF-β1 and/or EGF, as HaCaT II-4 cells secrete only low levels of plasminogen ( Fig. 3A). Dissolution of the supporting collagen matrix accompanied TGF-β1 + EGF + plasminogen treatment ( Fig. 3B), a process significantly reduced by the plasmin inhibitors aprotinin and α2-antiplasmin, but not with the cysteine protease inhibitor E-64 ( Fig. 3C and D), which affects cellular cathepsins. Inhibition of MMP activity with GM6001 also blocked plasmin-initiated collagen degradation, confirming a role for MMPs in the remodeling process ( Figs. 2D and 3C and D). Control experiments revealed no evidence of adverse effects arising from treatment with these inhibitors (data not shown). Cells that were detached as a result of collagen gel dissolution, in fact, reattached to tissue culture wells within 24 h (Supplementary Fig. S3).
Plasmin-dependent collagen degradation was quantified through the release of digested FITC-labeled collagen type 1 ( Fig. 3D). Stimulation with either TGF-β1 or EGF independently significantly increased collagen type 1 proteolysis within 7.5 h of plasminogen addition. This by itself, however, was insufficient to trigger a distinguishable loss in the fibrillar network even at later time points ( Fig. 3B). Stimulation with the combination of TGF-β1 + EGF clearly evoked a more substantial proteolytic response ( Fig. 3D) that resulted in dissolution of the polymerized gel within 20 h ( Fig. 3B), suggesting that the collagenolytic activity promoted by combining these two growth factors in the presence of plasminogen surpassed any threshold limitations.
TGF-β1 + EGF-stimulated collagen gel dissolution occurs via a plasmin/MMP-10/MMP-1–dependent axis. Because plasmin-dependent collagen degradation has been linked to MMP-13 up-regulation in mouse keratinocytes ( 26), it was necessary to assess whether MMP-13 might be involved in TGF-β1 + EGF-initiated collagenolysis. Similar to what has been reported previously for other HaCaT variants ( 13, 27), TGF-β1 stimulation significantly increased MMP-13 levels in HaCaT II-4 cells ( Fig. 4A ). Only a modest elevation in MMP-13 was evident, however, following coincubation with TGF-β1 + EGF ( Fig. 4A). MMP-10, in contrast, was substantially increased under these conditions ( Fig. 4A and B).
ProMMP-10 is a plasmin substrate ( 21), and whereas active MMP-10 does not cleave collagen type 1 directly, it does activate the collagenase MMP-1 ( 20). Subsequent to TGF-β1 + EGF stimulation, MMP-10 activation was evident by 4 h post-plasminogen addition and complete by 24 h ( Fig. 5A, top ), whereas the kinetics of MMP-1 activation closely followed the conversion of MMP-10 to a catalytic form ( Fig. 5A, bottom). To confirm the role of MMP-10 in collagen matrix degradation, plasminogen was added to TGF-β1 + EGF-stimulated HaCaT II-4 cultures in the presence of increasing concentrations of neutralizing antibodies to either MMP-1 or MMP-10. MMP-1 inhibition prevented plasminogen-dependent collagen dissolution ( Fig. 5B, top). Importantly, neutralization of MMP-10 activity also blocked plasminogen-initiated collagen degradation ( Fig. 5B, bottom), supporting a plasmin/MMP-10/MMP-1–dependent axis in matrix remodeling. A notable decrease in the level of active MMP-1 was also consistently evident following MMP-10 neutralization ( Fig. 5C). Despite residual levels of active, likely plasmin-generated MMP-1, this activity by itself was insufficient, however, to trigger gel dissolution on MMP-10 inhibition ( Fig. 5B).
PAI-1 functions as an upstream regulator of a MMP-10–initiated collagenolytic phenotype. Similar to MMP-10, PAI-1 expression in HaCaT II-4 cells is increased in response to TGF-β1 stimulation ( 12), whereas the combination of TGF-β1 + EGF synergistically enhanced PAI-1 protein levels ( Fig. 6A ; Supplementary Fig. S4A). Despite the inability of EGF alone to increase PAI-1 levels in this system, enhanced PAI-1 synthesis resulting from TGF-β1 + EGF stimulation, as well as from TGF-β1 alone, was attenuated by inhibition of EGF receptor signaling with AG1478 ( Fig. 6A). Similar results were evident in human dermal fibroblasts (Supplementary Fig. S4B) and kidney epithelial cells ( 22), emphasizing the generality of EGF receptor involvement in TGF-β1–dependent PAI-1 production.
PAI-1, through its inhibition of urokinase-type plasminogen activator, is critical for regulating the generation of pericellular plasmin. It was important, therefore, to assess the effect of PAI-1 on collagen gel dissolution. Blocking urokinase-type plasminogen activator activity with the inhibitor amiloride, or by adding a stable recombinant form of PAI-1 protein (N150H, K154T, Q319L, and M345I; ref. 28), completely inhibited collagen gel dissolution ( Fig. 6B). Addition of PAI-1 protein also effectively blocked conversion of MMP-10 and MMP-1 to their active forms ( Fig. 6C). In contrast, neutralization of endogenous PAI-1 with function-blocking antibodies accelerated both collagenolysis ( Fig. 6B) and activation of MMP-10 and MMP-1 ( Fig. 6C). Collectively, these results indicate that, in the physiologically relevant setting of a complex three-dimensional collagen environment, PAI-1 regulates MMP-10–initiated collagenolytic activity ( Fig. 6D). A key factor in this model is the ability of active MMP-10 to superactivate MMP-1, creating a plasmin/MMP-10/MMP-1 proteolytic axis that enhances collagen type 1 degradation and facilitates collagen gel invasion.
MMPs are integral components of a complex stromal remodeling program designed to modulate matrix integrity, release bioactive fragments, growth factors, and cytokines from matrix constituents, and enhance cell motility ( 29). Amplified MMP expression appears linked to increased tumor aggressiveness, metastasis, and poor patient survival ( 30). Not surprisingly, recently, studies also implicate several MMPs, including MMP-3, -7, -9, and -28, in directly triggering EMT-related processes ( 30). The combination of TGF-β and EGF, which effectively promotes EMT, also up-regulates certain MMPs synergistically including MMP-1, -3, -9, -10, and -14 ( 17, 18), posing some interesting questions regarding potential mechanisms that support amplification of EMT-associated events.
An acute collagenolytic phenotype linked to plasmin-dependent activation of stromelysin-2 (MMP-10) emerged in response to costimulation of HaCaT II-4 keratinocytes with TGF-β1 and EGF and was coincident with collagen gel invasion. MMP-10, which is generally limited to epithelial cells ( 15, 19), has broad substrate specificity, targeting proMMP-1, -7, -8, -9, and -13 as well as collagen types III, IV, and V, gelatin elastin, fibronectin, proteoglycans, and laminin ( 20, 21). Rigorous control over MMP-10 levels and activation are likely critical, therefore, for normal cutaneous homeostasis. MMP-10, in fact, is not evident in intact skin but is expressed during cutaneous injury repair where it localizes to migrating keratinocytes at the wound edge, suggesting that its presence may facilitate invasive behavior ( 31).
Similar to what has been observed in other systems ( 15, 18, 19, 27), MMP-10 and MMP-1 were up-regulated in HaCaT II-4 cells seeded onto collagen gels and stimulated with TGF-β1 and/or EGF. Both proenzymes are plasmin substrates; however, following MMP-10 inhibition, the residual level of active, likely plasmin-generated MMP-1 ( Fig. 5C) was insufficient by itself to trigger collagen gel dissolution ( Fig. 5B). These data likely reflect the established ability of MMP-10 to “superactivate” MMP-1 and enhance its collagenase activity 7- to 10-fold over that observed with plasmin alone ( 19). Given these parameters, neutralization of MMP-10 activity would have quenched this hypercollagenase activity and, as observed, impeded collagen degradation. Similar results regarding MMP-10–dependent superactivation of MMP-1, -8, and -13 in an arthritis-based model have been reported but not linked to plasminogen activation ( 32). Consequently, this article is the first to show a plasmin/MMP-10/MMP-1–dependent collagen remodeling axis and establish its relevance in a keratinocyte-based three-dimensional model.
Clearly, MMP-10 activity can have significant stromal consequences, particularly in a cutaneous environment, irrespective of its level of up-regulation. Amplified MMP-10 expression does, however, accompany the progression of several epithelial cancers, including squamous cell carcinomas of the head and neck and esophagus ( 33, 34). Increased MMP-10 expression also occurs in colorectal carcinoma ( 35), breast cancer ( 36), prostate cancer ( 37), and lymphoma ( 38). It localizes to cells at the invasive front of renal cell carcinomas and signals a lower survival rate compared with patients with MMP-10–negative tumors ( 39). Similarly, tumor MMP-10 levels predict poor survival in non-small cell lung cancer ( 40).
Like MMPs, lysosomal proteinase cathepsins are also associated with tumor cell invasion, particularly the cysteine proteinases cathepsins L and B, which degrade collagen type 1 and activate MMP-1, respectively ( 41). Inhibition of cysteine cathepsins had no effect, however, on collagen gel dissolution in the HaCaT II-4 model, whereas serine proteinase blockade effectively attenuated collagen degradation. These data emphasize a critical role for active plasmin, and not cathepsins, in the initiation of collagen degradation by TGF-β1 + EGF-stimulated cells and are consistent with observations regarding the ability of TGF-β to down-regulate cathepsins ( 42).
Previously, intermediates other than MMP-10, including MMP-13, have been associated with linking keratinocyte-based collagen type 1 dissolution and plasminogen activation ( 26, 43). Studies suggest, however, that contrary to observations in primary human keratinocytes, MMP-13 expression is, in fact, correlated with transformation of human keratinocytes and enhanced in these cells, including HaCaT derivatives, following stimulation with TGF-β1 ( 13, 27). Our data in a three-dimensional system also indicate that TGF-β1 stimulation alone increases the level of MMP-13; however, the combination of TGF-β1 + EGF did not produce this effect. Costimulated cells instead exhibited a robust induction of MMP-10. This disparity may be due, in part, to differences among the ras-HaCaT variants in MMP expression programs ( 44), TGF-β/EGF-related receptor cross-talk promoting EGF-dependent down-regulation of TGF-β1–enhanced MMP-13 levels ( 18, 45), or culture in two-dimensional versus a more complex three-dimensional stromal-equivalent system. The potential for EGF to counteract a TGF-β1–dependent increase in MMP-13 production reinforces the complexity of this process and presents some intriguing questions for future investigation regarding the regulation of these matrix-modifying enzymes in cancer progression.
Consistent with recent observations regarding receptor crosstalk and synergy ( 17, 18), PAI-1 levels increased synergistically following TGF-β1 + EGF treatment of HaCaT II-4 cells on a collagen substrate ( Fig. 6A). During tumor progression, synergistic amplification of PAI-1 would, in effect, inhibit a disproportionate level of stromal degradation and, in doing so, facilitate cell migration by preserving stromal architecture as well as by interacting with the low-density lipoprotein-related receptor ( 46). Up-regulation of both MMP-10 and PAI-1 in conjunction with a TGF-β1 + EGF-stimulated EMT, therefore, promotes an environment conducive to the acquisition of an invasive phenotype. Indeed, like MMP-10, PAI-1 expression is up-regulated in various cancers where its presence is associated with poor patient outcome ( 47, 48). The incidence of stromal PAI-1 is an important factor in determining tumor progression, reflecting its capacity to stabilize the microenvironment and promote tumor vascularization ( 49, 50).
Temporal and spatial balance between extracellular components that reorganize tissue architecture is a significant aspect of tumor progression. This study has identified an important proteolytic axis for regulating collagen type 1 degradation in a three-dimensional environment. The data provided are consistent with a model ( Fig. 6D) in which transformed keratinocytes, in response to TGF-β1 + EGF in the microenvironment, up-regulate proMMP-10, which is converted to its active form in the presence of plasmin and can subsequently superactivate the catalytic activity of MMP-1. This process results in a plasmin/MMP-10/MMP-1–dependent proteolytic axis that effectively enhances collagen type 1 degradation and facilitates collagen gel invasion. PAI-1 plays a crucial role in this paradigm through its ability to counter excessive collagen degradation and maintain stromal integrity for cell migration. Identification of critical components involved in managing the rate and level of collagen type 1 degradation may have far reaching implications for therapeutic targeting of cutaneous pathologies.
Disclosure of Potential Conflicts of Interest
No potential conflicts of interest were disclosed.
Grant support: NIH grant GM57242 (P.J. Higgins) and NIH training grant T32-HL07194 (C.E. Wilkins-Port).
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Note: Supplementary data for this article are available at Cancer Research Online (http://cancerres.aacrjournals.org/).
- Received January 7, 2009.
- Revision received March 3, 2009.
- Accepted March 6, 2009.
- ©2009 American Association for Cancer Research.