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VASCULAR BIOLOGY
School of Kinesiology and Health Sciences, York University, Toronto, Ontario, Canada M3J 1P3
Submitted 3 October 2007 ; accepted in final form 12 June 2008
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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vascular endothelia growth factor; phosphatidylinositol 3-kinase; Rho-GTPase; cytoskeleton; endothelium; angiogenesis
We, and others (24, 50, 59), have shown that the MMPs, specifically MMP-2 and MT1-MMP, are upregulated in response to reorganization of the actin cytoskeleton. The Rho-GTPases are known regulators of the actin cytoskeleton (39) and contribute to the transformation of extracellular stimuli into angiogenic responses (9). Angiogenic factors such as vascular endothelial growth factor (VEGF) and thrombin recruit downstream pathways, including the mitogen-activated protein kinases (MAPKs), phosphatidylinositol 3-kinase (PI3K), and various transcription factors through Rho-GTPase activation (28, 32–34, 54).
RhoA induces stress fiber formation and is required for VEGF-induced increases in endothelial cell permeability, migration, and stabilization of capillary tubes (1, 12, 38). Shear stress increases RhoA activity (46, 57) and stress fiber formation (7) and results in increased focal adhesion and junctional complex formation (45).
Rac1 controls lamellipodia and is linked with angiogenic signaling cascades. It is responsible for endothelial branching morphogenesis and capillary assembly in Matrigel overlay assays (11) through its contributions to endothelial migration (3) and MMP-2 activation (63). Downstream targets of Rac1 include the MAPKs and the AP-1 family of transcription factors, which in turn regulate MMP production (34, 54).
Cdc42 activation occurs with VEGF activation (31) and triggers the formation of filopodia and regulates cell polarization through microtubule organization. Activation of Cdc42 induces lumen and vacuole formation in endothelial cells and confers an invasive phenotype to T-lymphocytes (36, 48). Nobes and Hall (39) proposed that a hierarchy of cross talk occurs in which Cdc42 lies upstream of Rac1 and RhoA and that Cdc42 activates Rac1 while suppressing RhoA activity.
We hypothesized that manipulation of Rho-GTPase activity would result in changes in MMP-2 and MT1-MMP expression and activity in microvascular endothelial cells. We show that RhoA suppresses MMP-2 expression while limiting the amount of cell surface MT1-MMP and, thereby, MMP-2 activation. Conversely, Cdc42 activation promotes angiogenesis and increases the amount of cell surface MT1-MMP, thereby increasing MMP-2 activation. Furthermore, we demonstrate that these pathways contribute to VEGF-dependent activation of MMP-2.
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Three-dimensional collagen culture. SMEC were embedded in a three-dimensional type I collagen matrix (2.5 mg/ml Vitrogen) as previously described (8). SMEC were cultured in the presence or absence of 10 µM LY-294002 for 24 h at 37°C.
Gelatin zymography. Cells were lysed using 120 mM Tris·HCl (pH 8.7), 0.1% Triton X-100, and 5% glycerol supplemented with protease inhibitors (Sigma, catalog no. P8340) and 100 mM sodium orthovandate (lysis buffer). Whole cell protein extracts (10 µg) were analyzed by gelatin zymography as previously described (20). Gels were visualized and imaged using the Fluorchem gel doc system and analyzed using Alphaease (Alpha Innotech) software. Images were inverted for publication for ease of visualization. Total MMP-2 protein was calculated as the sum of the latent (72 kDa) and active (62 kDa) bands and was expressed as fold increase verses control. Active MMP-2 (62 kDa) was expressed as fold increase versus control.
Northern blot analysis. Total RNA was isolated and analyzed by Northern blot analysis using conventional techniques as described previously (20). Films were scanned and densitometry was performed using Alphaease (AlphaInnotech) software. Loading was normalized to the 28S rRNA band.
Transient transfection and promoter assays. Cells were plated on type I collagen-coated, 12-well dishes at a density of 50,000 cells/well, and 18 h later transient transfections were performed using LipofectAMINE 2000 (Invitrogen) reagent as per manufacturer's instructions. Cells were transfected with 0.5 µg of full-length MMP-2 promoter (21) and 0.05 µg of pRenilla (Promega). Twenty four hours posttransfection, cell were treated with either H1152 or GGTI-298 for 24 h. Forty-eight hours after transfection, cells were lysed with 1x passive lysis reagent (Promega), and reporter assays were performed using the Dual-Glo Luciferase Assay system (Promega). Firefly luciferase values were normalized to Renilla luciferase values. Normalized MMP-2 luciferase values from conditions that were treated with H1152 or GGTI-298 were then compared relative with those control untreated conditions.
Western blot analysis. Whole cell extracts were analyzed by Western blot analysis as described previously (24). Primary antibodies were the following: MT1-MMP (1:1,000, Novus), AKT (1:1,000), and Phospho-AKT (1:1,000, Cell Signaling). Films were scanned and densitometry was performed using Alphaease (AlphaInnotech) software. Phospho-AKT values were normalized to total-AKT values to account for variability in loading.
Immunofluorescence staining. Cells were plated on type I collagen-coated glass coverslips, incubated overnight at 37°C, and then treated with 10 µM H1152 for varying time points. Cells were fixed with 3.75% paraformaldehyde and then blocked and permeabilized in PBS + 5% normal goat serum + 0.05% Triton-X 100. Cells were incubated with primary phospho-c-Jun NH2-terminal kinase (JNK) antibody (1:300, Upstate), followed by secondary goat anti-rabbit Alexa-568 (1:400, Molecular Probes). Actin was visualized with fluorescein isothiocyanate (FITC)-phalloidin (4 µM, Sigma), and nuclei were counterstained with 4',6'-diaminodino-2-phenylindole (DAPI, 1:1,500, Molecular Probes). Cells were visualized by fluorescence microscopy (Zeiss Axiovert 200M). Images were captured using a cooled digital CCD Camera (Quantix 57) and imaging software (Metamorph, Universal Imaging).
TAT-Rho-GTPase fusion protein creation and transduction. Rho-GTPase DNA plasmids (RhoAN19, RhoAQL, Rac1N17, Rac1QL, Cdc42N17, and Cdc42QL were gifts from Dr. Ken Yamada) were subcloned into the psecTAG-TAT vector (amino acids 47–57 of the HIV-Tat, a gift from Dr. Tibor Barka) (5). Cos7 cells (provided Dr. Imogen Coe, York University) were transfected with 6 µl of TransPassD1 (New England Biolabs) and 5 µg TAG-TAT-Rho-GTPase or TAT-TAT plasmid DNA in OptiMEM for 3 h, as per manufacturer's instructions. OptiMEM was replaced with cDMEM and cells were maintained at 37°C overnight after which media was replaced with fresh OptiMEM for 3 h. The Cos7-conditioned OptiMEM, containing the secreted TAT or TAT-Rho-GTPase proteins, was collected and spun down, and the supernatant was stored at –20°C. SMECs were incubated with equal volumes of the OptiMEM containing mutant Rho-GTPase protein after which the cells were lysed as described above. Because the TAT peptide is known to elicit changes in endothelial cell cytoskeleton organization (58) and signaling (4), experimental conditions utilizing the TAT-Rho-GTPase fusion proteins were compared with TAT peptide-treated controls. In some experiments, cyclohexamide (20 µg/ml, Sigma) was added during the OptiMEM incubation to eliminate the effects of mRNA translation on MMP-2 protein levels.
Cell surface immunofluorescence staining. Cells were plated on type I collagen-coated glass coverslips and incubated overnight at 37°C and then treated with 10 µM H1152 or TAT-Cdc42QL-containing media for 2 h. Cells were kept on ice and washed with physiological saline solution (PSS, in mM: 130 NaCl, 1.18 KH2PO4, 1.17 MgSO4·7H2O, 14.9 NaHCO3, pH = 7.3, 5.5 dextrose, 0.026 CaNa2EDTA, and 1.6 CaCl2), blocked for 1 h with 5% BSA in PSS, and incubated with MT1-MMP antibody (1:300, Chemicon, Ab815) in PSS for 1.5 h. After three washes with PSS, cells were fixed with 3.75% paraformaldehyde and then permeabilized in PSS + 0.05% Triton-X 100. Cells were then incubated with secondary goat anti-rabbit Alexa-568 (1:400 dilution, Molecular Probes). Actin was stained with FITC-phalloidin (4 µM, Sigma). Cells were visualized by confocal microscopy (Olympus Fluoview 300 with Argon laser, 488 nm, and HeNe laser, 543 nm, pinhole aperture = 2) using Fluoview software (Olympus). Z-sections were captured (0.1 µm, 10 slices), and complete Z-stacks were used for actin staining images. For cell surface MT1-MMP images, only the first three slices of the Z-section were included in the final Z-stack.
Surface biotinylation. SMEC were cultured on type I collagen for 24 h, pretreated with 50 µM LY-294002, and then stimulated with 25 ng/ml VEGF or 10 µM H1152 for 60 min or treated with TAT-Cdc42QL-containing media for 2 h. Cells were washed with ice-cold PBS and incubated with 1 mg/ml sulfo-NHS-biotin in PBS for 30 min on ice. The reaction was terminated by washing the cells with 100 mM glycine for 20 min. Cells were lysed as described above, and 75 µg of protein from total cell lysates were incubated overnight at 4°C with Streptavidin-Agarose beads (Pierce) with gentle rocking. Beads were collected by centrifugation and the pellet was washed several times with PBS containing 0.1% NP-40. Fifty microliters of 1x loading buffer were added, the samples were boiled, and the proteins were separated by 10% SDS-PAGE followed by MT1-MMP Western blot analysis as previously described above. Blots were then stripped and reprobed for β1-integrin (Cell Signaling) as a loading control.
Capillary sprout formation assay. Capillary segments were isolated from the rat epididymal fat pad and cultured within a three-dimensional type I collagen matrix (Vitrogen) as previously described (20). Fragments were cultured overnight at 37°C after which time culture media was replaced with Opti-MEM (Invitrogen) containing constitutively active or dominant negative TAT-Rho-GTPase proteins or a TAT peptide control for 3 h. Opti-MEM was then replaced with cDMEM, and the fragments were cultured for an additional 48 h. Phase-contrast images were taken, and sprout length was measured using Metamorph imaging software (Universal Imaging).
RhoGTPase activity assays. SMEC were lysed with IP lysis buffer (50 mM Tris, pH 7.5, 10 mM MgCl2, 0.3 M NaCl, and 2% NP-40), and 100 µg of protein were incubated with Rhotekin-GST or PAK-GST protein beads (Cytoskeleton) for 2 h at 4°C with gentle rocking for RhoA or Rac1 and Cdc42 activity measurements respectively. Beads were collected by centrifugation and the pellet was washed 3x with wash buffer (25 mM Tris, pH 7.5, 30 mM MgCl2, 40 mM NaCl). Fifty microliters of 1x reducing loading buffer were added, the samples were boiled, and the proteins were separated by SDS-PAGE through a 15% acrylamide gel followed by Western blot analysis for RhoA, Rac1, or Cdc42 (1:1,000, Cell Signaling).
Statistics. Data were normalized to control and are presented as fold change ± SE versus control. Student's t-test or one-way ANOVA followed by Tukey post hoc tests, were applied to determine statistical significance (P < 0.05).
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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JNK regulates MMP-2 and MT1-MMP mRNA expression following ROCK inhibition. From our previous findings that JNK regulates MMP-2 and MT1-MMP mRNA expression in endothelial cells (24), we tested whether JNK contributes to the increase in mRNA expression following ROCK inhibition. Inhibition of ROCK with H1152 caused depolymerization of actin stress fibers and an increase in cortical actin (Fig. 2B, arrows). H1152 treatment caused nuclear translocation of phospho-JNK (Fig. 2B, arrowheads). Correlating with this response, we observed that the H1152-induced increase in MMP-2 mRNA expression was attenuated partially by JNK inhibition, whereas the increase in MT1-MMP was blocked completely by pretreatment with SP-600125 (Fig. 2C).
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To test whether PI3K modulates endothelial cell sprouting, we isolated capillary segments and cultured them within a three-dimensional type I collagen matrix in the absence or presence of LY-294009 (Fig. 4C). The typical formation of endothelial cell sprouts from the preexisting capillaries was significant in those treated with the PI3K inhibitor (Fig. 4D). Correspondingly, the increase in total MMP-2 protein observed in endothelial cells following 24 h in three-dimensional culture was attenuated partially by PI3K inhibition (3.3 ± 0.4-fold vs. two-dimensional control, P < 0.05, and 2.3 ± 0.4-fold vs. two-dimensional control, P > 0.05, respectively). Notably, MMP-2 protein activation was significantly reduced by pretreatment with LY-294009 compared with control three-dimensional cultures (Fig. 4E).
Overexpression of Cdc42 increases activation of MMP-2. To further define the roles of individual Rho-GTPase members in MMP-2 activation, we utilized Rho-GTPase fusion proteins of both the dominant negative and constitutively active forms of RhoA, Rac1, and Cdc42, coupled to the 11 amino acid cell entry domain of the HIV-Tat peptide (TAT). Western blot analysis verified the function of the TAT-Rho-GTPase fusion proteins. Activation of Cdc42 and Rac1 by treatment with TAT-Cdc42QL and TAT-Rac1QL, respectively, are shown as representative experiments (Fig. 5A).
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Transduction of endothelial cells with TAT-Cdc42QL for 4 h resulted in a significant increase in MMP-2 protein activation (Fig. 5C) without a concurrent increase in total MMP-2 protein. This increase in active MMP-2 was not attenuated in Cdc42QL-transduced cells treated with the translational inhibitor cycloheximide (1.66 ± 0.1-fold above TAT-only in untreated vs. 1.50 ± 0.2-fold above TAT-only in cyclohexmide-treated cells; n = 6–7). There was a trend toward increased MMP-2 activation in SMEC treated with TAT-Rac1QL, but no change in MMP-2 activation was seen in cells treated with TAT-RhoAQL. Increased levels of P-AKT also were observed following transduction of cells with Cdc42QL (Fig. 5D), similar to that observed in response to inhibition of the RhoA effector ROCK (Fig. 4A).
Given the effect of Cdc42QL on MMP-2 activation, we examined MT1-MMP localization. Treatment of endothelial cells with TAT-Cdc42QL caused cell retraction, formation of filopodia-like protrusions, and an increase in cortical actin (Fig. 6A). Cell surface MT1-MMP localization also increased as evidenced by immunostaining (Fig. 6A) and cell surface biotinylation (Fig. 6B).
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The level of RhoA activity has been linked previously to MMP-2 activity as Matsumoto et al. (35) showed that in lysophosphatidic acid-stimulated cells, low levels of RhoA activity increased MMP-2 activity but increased RhoA activity-inhibited MMP-2 activity. Similarly, confluent nonmigratory human umbilical vein endothelial cells exhibit high levels of RhoA activity, and RhoA inhibition increases MT1-MMP activity (16). However, mechanisms linking theses effects were not established by these studies. Zucker et al. (64) showed that actin depolymerization increases the number of cell surface receptors for tissue inhibitor of metalloproteinase-2 (TIMP-2) (i.e., MT1-MMP) without altering the binding affinity for TIMP-2. MT1-MMP is internalized by both clathrin-dependent (26, 43) and clathrin-independent (through caveolae) (16, 43) endocytosis, both of which rely on actin scaffolding. Disruption of the actin cytoskeleton may affect normal internalization of MT1-MMP (17), quickly increasing MT1-MMP molecules on the cell surface. These observations suggest that high levels of RhoA and stress fiber formation contribute to a stabilized, nonproteolytic endothelial phenotype.
Increasing Cdc42 activity caused the same effect on MMP-2 activation as inhibition of RhoA/ROCK activity. Overexpression of Cdc42 induced an increase in MT1-MMP on the cell surface and increased angiogenesis in three-dimensional type I collagen. Previous studies indicated a significant permissive role for Cdc42 in tumor cell migration and invasion (19, 30, 55). Our results extend these findings by providing a mechanism by which these events may be mediated, namely Cdc42-dependent MMP-2 activation via increased cell surface MT1-MMP. It is possible that in addition to inhibition of MT1-MMP internalization, activation of Cdc42 may increase MT1-MMP cell surface localization by increasing exocytosis of MT1-MMP from intracellular stores to the plasma membrane. For example, Cdc42 activation plays a positive role in exocytosis of von Willebrand factor in endothelial cells (29, 51) and the second phase of insulin secretion in pancreatic β-cells (53).
The Rho family of GTPases signal to one another in the process of coordinating the appropriate intracellular responses to extracellular signals. We demonstrated that Cdc42 activation suppressed basal RhoA activity. This is in agreement with the hierarchy of Rho-GTPase signaling proposed by Nobes et al. (39). We observed that Cdc42 activation led to decreased RhoA activity, stress fiber depolymerization, increased MT1-MMP cell surface localization, and greater MMP-2 activation. Concurrently, Cdc42 activation can increase Rac1 activation, which may contribute to increase endothelial sprouting.
Coordination of Cdc42 and RhoA signaling was recently demonstrated in podosome formation in primary endothelial cells (49). Cdc42 activation resulted in podosome formation, which was accompanied by localized stress fiber depolymerization and correlated with decreased RhoA activity (49). Conversely, Cdc42-dependent Rac1 activation has been reported wherein Cdc42 activates IRSp53, an intermediary protein that allows for Rac1 to bind to the WAVE-Arp 2/3 complex, and is required for Rac1-dependent lamellipodia formation (18).
Several studies indicate Cdc42-dependent activation of PI3K and AKT (10, 60), which is consistent with our observation that PI3K is required for MMP-2 activation following RhoA inhibition. When considered with the report that increased RhoA activity inhibited AKT-dependent phosphorylation of endothelial nitric oxide synthase in endothelial cells (52) and that Cdc42 activation decreases RhoA activity, we postulate that activation of Cdc42 alleviates the RhoA-induced suppression of PI3K/P-AKT signaling. The activation of these pathways then increases cell surface MT1-MMP and induces MMP-2 activation.
The mechanism by which P-AKT regulates MT1-MMP on the cell surface remains to be elucidated. Galvez et al. (14) suggested that clustering of MT1-MMP molecules is more important than internalization in regulating MT1-MMP activation of MMP-2. They proposed that MT1-MMP clustering is dependent on cortical actin polymerization, which is regulated by PI3K. Our data are consistent with this hypothesis as ROCK inhibition increased P-AKT levels at time points corresponding to increased MMP-2 activation, and inhibition of PI3K abolishes the increases in MMP-2 activation without altering the amount of MT1-MMP on the cell surface.
Our results provide evidence that VEGF induction of MMP-2 activity utilizes a combination of increased Cdc42 activity and decreased RhoA activity. The signal cascade leading to MMP-2 activation proposed here is in line with events following VEGF stimulation of endothelial cells. Cdc42 activation has been reported in response to VEGF stimulation (61) and is activated during endothelial cell haptotaxis on type I collagen (47). Interestingly, activation of Cdc42 can induce VEGF promoter activity via a c-Jun-dependent mechanism (44), and we have shown previously that c-Jun is necessary for MMP-2 mRNA expression following VEGF stimulation (24). Together, these results suggest that VEGF can induce both rapid and sustained production activation of MMP-2 through upregulation of Cdc42 activity.
In summary, inhibition of RhoA or activation of Cdc42 in microvascular endothelial cells induced a shift from stress fibers to cortical actin and increased MT1-MMP localization to the cell surface. Longer-term consequences of RhoA inhibition or Cdc42 activation included increased MMP-2 and MT1-MMP mRNA expression and greater capillary sprouting. These findings further elucidate the process of endothelial cell activation and initiation of angiogenesis.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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