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Molecular and Cellular Biology, December 2007, p. 8454-8465, Vol. 27, No. 24
0270-7306/07/$08.00+0     doi:10.1128/MCB.00821-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Identification of Candidate Angiogenic Inhibitors Processed by Matrix Metalloproteinase 2 (MMP-2) in Cell-Based Proteomic Screens: Disruption of Vascular Endothelial Growth Factor (VEGF)/Heparin Affin Regulatory Peptide (Pleiotrophin) and VEGF/Connective Tissue Growth Factor Angiogenic Inhibitory Complexes by MMP-2 Proteolysis{triangledown} ,{dagger}

Richard A. Dean,1 Georgina S. Butler,1 Yamina Hamma-Kourbali,2 Jean Delbé,2 David R. Brigstock,3 José Courty,2 and Christopher M. Overall1*

University of British Columbia, Centre for Blood Research, 4.401 Life Sciences Institute, 2350 Health Sciences Mall, Vancouver, British Columbia, Canada V6T 1Z3,1 Laboratoire de recherche sur la Croissance Cellulaire, la Réparation et la Régénération Tissulaires, CNRS UMR 7149, Université Paris XII, Avenue du Général de Gaulle, 94010 Crétail Cedex, France,2 Children's Research Institute, Room WA2022, 700 Children's Drive, Columbus, Ohio 432053

Received 9 May 2007/ Returned for modification 28 May 2007/ Accepted 5 September 2007


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ABSTRACT
 
Matrix metalloproteinases (MMPs) exert both pro- and antiangiogenic functions by the release of cytokines or proteolytically generated angiogenic inhibitors from extracellular matrix and basement membrane remodeling. In the Mmp2–/– mouse neovascularization is greatly reduced, but the mechanistic aspects of this remain unclear. Using isotope-coded affinity tag labeling of proteins analyzed by multidimensional liquid chromatography and tandem mass spectrometry we explored proteome differences between Mmp2–/– cells and those rescued by MMP-2 transfection. Proteome signatures that are hallmarks of proteolysis revealed cleavage of many known MMP-2 substrates in the cellular context. Proteomic evidence of MMP-2 processing of novel substrates was found. Insulin-like growth factor binding protein 6, follistatin-like 1, and cystatin C protein cleavage by MMP-2 was biochemically confirmed, and the cleavage sites in heparin affin regulatory peptide (HARP; pleiotrophin) and connective tissue growth factor (CTGF) were sequenced by matrix-assisted laser desorption ionization-time of flight mass spectrometry. MMP-2 processing of HARP and CTGF released vascular endothelial growth factor (VEGF) from angiogenic inhibitory complexes. The cleaved HARP N-terminal domain increased HARP-induced cell proliferation, whereas the HARP C-terminal domain was antagonistic and decreased cell proliferation and migration. Hence the unmasking of cytokines, such as VEGF, by metalloproteinase processing of their binding proteins is a new mechanism in the control of cytokine activation and angiogenesis.


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INTRODUCTION
 
Matrix metalloproteinase 2 (MMP-2) (also known as gelatinase A), like many of the 23 MMPs in humans, has long been associated with angiogenesis, tumor progression, and metastasis (20, 50). More recently the critical importance of MMP-2 in breast cancer metastasis was shown by Minn et al. (52), who found that the MMP2 gene was one of the four key genes in the genomic signature associated with the most virulent breast cancer lung metastases. In this model, the development of angiogenesis was highly dependent upon MMP-2 expression (23). With other new functions of MMPs that promote cancer cell growth, invasion, metastasis, and immune cell evasion (20, 50) a more complex role for MMPs than just degradation of extracellular matrix proteins is being revealed (12, 56).

Contrary to their many carcinogenic roles, several MMPs are now being recognized as cancer antitargets due to their normal physiological roles in cell signaling and tissue homeostasis that are protective against cancer (62). MMP-3 overexpression reduces 12-dimethylbenz[a]anthracene (DMBA)-induced skin carcinogenesis (76); Mmp8–/– mice treated with DMBA and the phorbol ester 12-myristate 13-acetate develop more skin papillomas and fibrosarcomas than wild-type mice (4), and Mmp12–/– mice show increased lung carcinogenesis (30). The detailed study of individual MMPs is therefore necessary to understand their specific functions in different physiological and pathological processes, information that is necessary for protease drug target validation (62). Elucidation of each protease substrate repertoire or degradome is crucial to understanding the biological roles of MMPs (44). Genetic models of disease in mice are powerful, yet time-consuming (20, 60, 64), so high-content proteomic approaches using human cell systems offer great promise in understanding the biological functions of human proteases (60, 70).

Angiogenesis is a highly regulated process in embryogenesis, postnatal blood vessel remodeling, and normal physiological events such as wound healing and the female reproductive cycle. Here, the extracellular matrix is thought to both direct and impair endothelial cell invasion, with the primary role of MMPs hypothesized to be the removal of this barrier, thereby allowing endothelial cell migration (14, 56). By binding to glycosylated proteins the extracellular matrix stores many cytokines that direct these processes (73) including epidermal growth factor, fibroblast growth factor, platelet-derived growth factor, transforming growth factor ß, connective tissue growth factor (CTGF), and vascular endothelial growth factor (VEGF) (42, 49, 67). Of these, VEGF is a potent endothelial cell mitogen (41) and permeability-enhancing factor that influences the egress of plasma proteins and cells that both directly and indirectly stimulate angiogenesis (19). Its importance in carcinogenesis and angiogenesis is reflected by the numerous therapeutic strategies to control its activity. Upon tissue remodeling by MMPs these proangiogenic growth factors are released and angiogenic regulatory integrin contacts on endothelial cells are disrupted (71). Indeed, angiogenesis is impaired in mice lacking the MMP-2, -9, or -14 gene (15, 32, 35, 75, 77).

Angiogenesis is also rate limiting for expansive tumor growth, with the "angiogenic switch" now viewed as one of the critical events that must be turned on in carcinogenesis (7). MMP-9 (also known as gelatinase B) is implicated in VEGF mobilization and neovascularization in cancer (6). However, it is not clear if this is through cleavage of the matrix proteins ligating VEGF (6, 67) or by cleavage of VEGF itself to release its bioactive domain from the matrix-binding domain (59). In gelatinase A-deficient mice neovascularization is also reduced in tumors (32) and in corneal angiogenesis assays (35, 39), but neither the angiogenic factors nor the mechanistic aspects of MMP-2 proangiogenic pathways have been elucidated.

In addition to matrix degradation, MMPs have other proangiogenic roles by direct effects on endothelial cells to enhance cell proliferation and up-regulate endothelial expression of MMPs (57), as well as to cleave the ectodomain of vascular endothelial cadherin, thereby breaking cell-cell adhesion (29). Consistent with this, the endogenous tissue inhibitors of metalloproteinases, as well as synthetic MMP inhibitor drugs, are potent inhibitors of angiogenesis (2, 33, 45). Hence, the overarching role of many MMPs is a proangiogenic response. Nonetheless, MMPs also generate angiogenic inhibitors in vitro by proteolytically cleaving fragments from the pericellular matrix (MMP-3, -7, -9, -13, and -20 generate endostatin from collagen XVIII [27]; MMP-9 generates tumstatin from type IV collagen [25]), from other proteases such as plasminogen (MMP-2, -3, -9, and -12 generate angiostatin [16]), or from hormones (MMP-1, -3, -8, and -13 generate angiogenic inhibitor fragments from prolactin [46]). If these proteins are generated in sufficient concentrations in vivo, then a feedback loop might operate to fine-tune angiogenesis. Hence, by exhibiting both pro- and antiangiogenic functions, MMPs are potentially dual regulators of angiogenesis in vivo (69).

Here we report the use of isotope-coded affinity tag (ICAT) labeling and tandem mass spectrometry (MS) to identify novel substrates of MMP-2 in the cellular context. Upon biochemical validation we identified the angiogenic regulatory factors heparin affin regulatory peptide (HARP) (also known as pleiotrophin) and CTGF as novel substrates of MMP-2. Precise proteolytic processing of HARP and CTGF inactivates these growth factors and mobilizes VEGF from HARP-VEGF and CTGF-VEGF complexes. With numerous other novel substrates of MMP-2 being identified such as insulin-like growth factor 6 (IGFBP-6) and follistatin-like 1 protein, which also bind and mask active cytokines to inhibit cell proliferation, our proteomic screen reveals potential new roles for MMP-2 in angiogenesis and carcinogenesis. Rather than merely being involved in extracellular matrix degradation to facilitate angiogenesis, MMP-2 processing releases proangiogenic factors from stable inhibitory complexes to directly stimulate neovascularization and so is a new mechanism for the control of angiogenesis.


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MATERIALS AND METHODS
 
Cell culture. Human umbilical vein endothelial cells, EBM-2 BulletKit medium, and fetal bovine serum (FBS) were purchased from BioWhittaker Clonetics (Emerainville, France). Mouse NIH 3T3 cells were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% FBS. Immortalized Mmp2–/– mouse embryonic fibroblasts were stably transfected with the pGW1HG vector containing cDNA encoding full-length human MMP-2 engineered with an immunoglobulin G leader sequence to aid secretion (54). Empty-vector transfectants served as controls. MMP-2 activation was achieved by addition of concanavalin A (ConA) (63) to the growth medium at the minimal concentration required to achieve MMP-2 activation (selected from dose-response studies). MMP-2 gelatinolytic activity was assayed by zymography using nonreduced conditioned-medium samples electrophoresed on 10% polyacrylamide gels copolymerized with 1 mg/ml gelatin. Following sodium dodecyl sulfate (SDS) removal using Triton X-100, gels were incubated in 50 mM Tris-HCl, 200 mM NaCl, and 5 mM CaCl2, pH 7.4, for 16 h and then stained with Coomassie brilliant blue R250.

For protein analysis, stable transfected cells were grown in DMEM, 5% cosmic calf serum (HyClone), 2 mM L-glutamine, 70 mM xanthine, 1x HT supplement (100 µM sodium hypoxanthine, 16 µM thymidine; Gibco) under 25-µg/ml mycophenolic acid selection until 30% confluent, washed extensively with phosphate-buffered saline (PBS) to remove serum proteins, and grown overnight in serum-free medium. Cells were then carefully washed again with PBS, incubated in phenol red-free, serum-free medium containing 14 µg/ml ConA, and grown for 48 h. Conditioned-medium proteins were then harvested and protease inhibitors (1 mM EDTA and 0.5 mM phenylmethylsulfonyl fluoride) immediately added. Whole cells and cell debris were removed from the conditioned medium by centrifugation (5 min, 500 x g, then 30 min, 8,000 x g) and filtration (0.2-µM cutoff). ConA was removed from the medium using {alpha}-mannose-Sepharose (Sigma) chromatography so as to accurately quantitate the cellular protein content in the conditioned medium and to increase the specific activity of ICAT-labeled cell proteins. Centripreps (3-kDa cutoff; Millipore) were used to concentrate conditioned medium (generally ~120 ml) to a final protein concentration of 2 to 3 mg/ml.

ICAT labeling and tandem MS. Conditioned-medium proteins (100 µg) from MMP-2-transfected cells incubated with 14 µg/ml ConA were labeled with isotopically heavy (13C) cleavable ICAT reagent (Applied Biosystems [ABI], Foster City, CA). Vector control cells were similarly treated with ConA, and the conditioned-medium proteins were labeled with isotopically light (12C) ICAT label. ICAT-labeled conditioned-medium proteins were combined for overnight digestion with sequence grade modified trypsin (Promega) (1:10) at 37°C and then fractionated on a cation exchange column (4 by 15 mm) by elution with 350 mM KCl, pH 3.0, to remove trypsin and unincorporated free label. ICAT-labeled peptides were purified using an avidin affinity column (4 by 15 mm) and eluted with 20% acetonitrile-0.4% trifluoroacetic acid before being dried by centrifugation under vacuum. Samples were then incubated in 95% trifluoroacetic acid (2 h, 37°C) to cleave the biotin tag from the ICAT-labeled peptides, and the samples were simplified for mass spectrometry by multidimensional liquid chromatography fractionation and analyzed using a QStar Pulsar mass spectrometer (ABI) as previously described (18, 74). Tandem MS fragmentation (2 s, m/z 65 to 1,800) was performed on four of the most intense ions as determined from a 1-s survey scan (m/z 300 to 1,500).

Proteins were identified from peptide sequences queried against the National Center for Biotechnology Information nonredundant protein database using MASCOT (Matrix Science). All proteins identified at ≥99% confidence were then manually reconfirmed using the Swiss-Prot sequence database (http://us.expasy.org/). ICAT ratios for identical but isotopically heavy- and light-labeled tryptic peptides were calculated using PROICAT (ABI) software and averaged if multiple peptides for a single parent protein were found.

Proteases and substrate validation. Recombinant full-length human proMMP-2 (zymogen form of MMP-2) was expressed and purified as previously described (11). The concentration of active enzyme after p-aminophenylmercuric acetate (APMA) activation (1 mM, 37°C, 15 min) was determined by active-site titration against a standard preparation of tissue inhibitor of metalloproteinases 2 (11). A monoclonal antibody against human MMP-2 (MAB13489) was purchased from Chemicon. Cathepsin L and the synthetic substrate Z-LR-7-amino-4-methylcoumarin were a kind gift from John Mort (Shriners Hospital for Children, Montreal, Quebec, Canada). Cystatin C and IGFBP-6 were kind gifts from Magnus Abrahamson (University of Lund, Lund, Sweden) and John Fowlkes (University of Arkansas for Medical Sciences), respectively. Follistatin-like 1 protein and HARP were purchased from ProSci (Poway, CA) and R&D Systems, respectively. Recombinant VEGF (VEGF165) was purchased from Pepro Tech EC Ltd. (London, United Kingdom).

Substrate cleavage validation. APMA-activated MMP-2 was incubated with the substrate candidates in 50 mM Tris-HCl, 200 mM NaCl, 5 mM CaCl2, and 0.025% NaN3 for 16 h at 37°C. Reaction products were analyzed by Tris-glycine or Tris-Tricine SDS-polyacrylamide gel electrophoresis (PAGE) and Western blotted or silver stained. The mass of each cleavage product was determined by matrix-assisted laser desorption ionization-time of flight (MALDI-TOF) MS on a Voyager-DE STR biospectrometry workstation (ABI). MS data were deconvoluted to identify the cleavage sites. N-terminal Edman sequencing was used to confirm the neo-N termini of cleavage products.

Analysis of CTGF-induced secreted protein synthesis of fibroblasts. Mmp2–/– embryonic fibroblasts were seeded at 5 x 104 cells per well in 24-well plates in DMEM supplemented with 10% FBS and cultured until 80% confluent. Cells were then serum starved for 24 h, and the medium was replaced with phenol red-free, serum-free medium containing CTGF (final concentration, 10 or 100 ng/ml), MMP-2-proteolyzed CTGF (final concentration, 10 or 100 ng/ml; cleaved to completion using 15 ng/ml MMP-2 for 24 h), or the appropriate concentration of MMP-2 as controls. After 48 h of cell growth the conditioned medium was collected, whole cells were removed by centrifugation (5 min, 500 x g), and protein concentration was analyzed by bicinchoninic acid assay.

Expression of HARP domains. To express analogues of MMP-2 cleavage products of HARP, glutathione S-transferase (GST) fusion proteins with HARP residues 9 to 59 (N-HARP) and 60 to 110 (C-HARP) were constructed (28). cDNAs of the N-HARP and C-HARP proteins were amplified by PCR using the human HARP cDNA as the template, and the products were subcloned in frame with the GST into the pGEX6P1 vector (Pharmacia Amersham Biotech, France) (28). After expression, the GST-HARP fusion proteins were isolated by glutathione agarose affinity and the GST tag was removed using PreScission protease (Pharmacia Amersham Biotech) treatment. N-HARP and C-HARP were further purified by Mono S fast protein liquid chromatography, and the integrity of the truncated proteins was assessed by using 15% Tris-Tricine gels and MALDI-TOF MS.

Thymidine incorporation assay. NIH 3T3 cells were seeded at 3 x 104 cells per well in 48-well plates for 24 h in DMEM supplemented with 10% FBS. Cells were then serum starved for 24 h and incubated with HARP, MMP-2-proteolyzed HARP (digested to N- and C-HARP completely over 24 h), or N-HARP or C-HARP added alone or in combination with HARP for 18 h at 37°C. The cells were then labeled for a further 6 h with 0.5 µCi of [methyl-3H]thymidine (MP Biomedical, Irvine, CA), fixed with 10% trichloroacetic acid, washed with water, and lysed with 0.1 M NaOH, and the total incorporated radioactivity was counted (8).

Endothelial cell migration assay. Endothelial cell migration assays were performed using a 24-well chemotaxis chamber (Transwell; Costar, France). Polycarbonate filters with 8-µm pore size were coated with 10 µg/ml type I collagen (collagen R solution; 0.2%, wt/vol; Serva, Heidelberg, Germany) for 1 h at 22°C and dried under sterile air. HARP, MMP-2-proteolyzed HARP, N-HARP, or C-HARP was diluted to the appropriate concentrations in EBM-2 supplemented with 1% (vol/vol) FBS, and 500 µl of the final dilutions was placed in the lower chamber. Confluent human umbilical vein endothelial cells were trypsinized, and 1 x 106 cells/ml in EBM-2 supplemented with 1% (vol/vol) FBS were seeded (100 µl) in the upper compartment. Transwells were incubated for 4 h at 37°C, and then the cells on the upper surface of the filters were removed by wiping with a cotton tip. The filters were then fixed with methanol, and cells that had migrated to the lower surface were stained with May-Grünwald-Giemsa solution and quantified by counting cells in three random high-power fields in each well (magnification, x100). Results presented are the means of two independent experiments.

MMP-2 mobilization of VEGF from HARP or CTGF complexes. Microtiter plates with 96 wells (Falcon; Becton Dickinson) were coated overnight at 4°C with equimolar HARP (200 ng), CTGF (465 ng), or 0.1% bovine serum albumin (BSA) control diluted in 100 µl PBS. The plates were then washed three times with PBS containing 0.1% (vol/vol) Tween 20, blocked with 1% BSA in PBS for 4 h at room temperature, and washed three times with PBS-Tween 20. To form complexes with VEGF, the coated wells were then incubated overnight at 4°C with 100 ng human VEGF diluted in PBS containing 1% BSA and then washed five times with PBS-Tween 20. The immobilized complexes were incubated for 24 h at 37°C with 1.5 pmol AMPA-activated MMP-2 (100 ng) diluted in 100 µl assay buffer (50 mM Tris-HCl, 200 mM NaCl, 5 mM CaCl2, 0.05% NaN3, pH 7.4). After digestion, the wells were washed five times with PBS-Tween 20 and HARP- or CTGF-bound VEGF was quantified by enzyme-linked immunosorbent assay (ELISA). Anti-human VEGF antibody (100 µl at 1 µg/ml) (MAB293; R&D Systems) diluted in PBS-Tween 20 was detected with goat anti-mouse immunoglobulin G conjugated to alkaline phosphatase (Bio-Rad Laboratories) and developed with 100 µl p-nitrophenylphosphate solution (Sigma), according to standard procedures. Substrate formation corresponding to bound VEGF was determined by measuring absorbance at 405 nm using an ELISA microplate reader (Molecular Devices).

Proteolysis of HARP-VEGF and CTGF-VEGF complexes in solution by MMP-2. HARP-VEGF and CTGF-VEGF complexes were prepared by incubation of 500 ng VEGF with HARP or CTGF at a 1:1 molar ratio at 4°C for 24 h in 50 mM Tris-HCl, 200 mM NaCl, 5 mM CaCl2, and 0.05% NaN3, pH 7.4. APMA-activated MMP-2 (20 ng) was then added to the complexes, and they were incubated at 37°C for 24 h. Reaction products were analyzed by 15% Tris-Tricine SDS-PAGE and silver stained.

Cathepsin L activity assay. Cystatin C was incubated for 16 h at 37°C with APMA-activated MMP-2. Since APMA is a cysteine protease inhibitor, it was removed from the cleaved cystatin C samples prior to addition of cathepsin L using protein desalting spin columns (Pierce). Cathepsin L (0.1 nM) was assayed using 0.2 µM Z-LR-7-amino-4-methylcoumarin in buffer (100 mM sodium phosphate buffer, pH 6.15, 100 mM NaCl, 2 mM EDTA, 1 mM dithiothreitol, 3% dimethyl sulfoxide, and 0.01% Tween 20). Cathepsin L was activated for 20 min on ice in 1 mM dithiothreitol. The increase in fluorescence following the release of amino-4-methylcoumarin by cathepsin L cleavage was measured using a 390-nm excitation and 460-nm emission filter pair. The 50% inhibitory concentrations (IC50) of full-length and cleaved cystatin C and MMP-2 alone with cathepsin L were calculated.


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RESULTS
 
MMP-2 expression and activation. To screen for new MMP-2 substrates, human MMP-2 was stably transfected in Mmp2–/– embryonic fibroblast cells to provide a high signal-to-noise ratio compared to vector transfectants. We used stably transfected cell lines in which the enzyme was expressed at low levels so as not to utilize an experimental system that has unnaturally high enzyme-to-substrate ratios. The clones selected for proteomic analysis exhibited gelatinolytic activity in the conditioned medium by zymography (Fig. 1A) but not by determination of quenched fluorescent synthetic peptide cleavage activity, indicating a very low level of MMP-2 expression.


Figure 1
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  		FIG. 1. Characterization of MMP-2 transfectants. (A) Medium conditioned for 48 h from immortalized Mmp2–/– embryonic fibroblast transfectants expressing pGW1GH vector (vector) or human MMP-2 (MMP-2) with or without 20 µg/ml ConA was analyzed by gelatin zymography. Std, 5 ng recombinant human MMP-2. (B) Titration of ConA (0 to 20 µg/ml) was used to determine the lowest concentration necessary for robust activation of MMP-2 at 48 h. The migration of the zymogen form (proMMP-2) and active forms of MMP-2 is shown. The square indicates the concentration of ConA (14 µg/ml) used to activate MMP-2 in the ICAT experiments. (C) ICAT-labeled MMP-2 tryptic peptides identified at ≥99% confidence in 48-h-conditioned medium from ConA-treated Mmp2–/– cells transfected with MMP-2 or vector alone are shown with their corresponding ICAT ratios. A ratio >1.0 reflects relative accumulation of the protein in conditioned medium.

Gelatin zymography confirmed the absence of MMP-2 activity in vector control Mmp2–/– cells (Fig. 1A). Addition of ConA to proMMP-2-transfected cells induced the cellular activation of MMP-2 (72 kDa) to the 62-kDa active form via the 67-kDa intermediate (Fig. 1A), as first reported (63). The lowest concentration of ConA at which significant proMMP-2 activation occurred was 14 µg/ml, and so this was used in the proteomic screens (Fig. 1B). Since ConA is a 290-amino-acid protein that does not contain a cysteine, we selected the cysteine-reactive ICAT reagent for isotopic labeling rather than the amine-reactive iTRAQ reagent (18). Experiments were conducted under serum-free conditions to increase specific labeling of the secreted proteins. In the ICAT-labeled conditioned-medium proteins, two tryptic peptides of human MMP-2 were identified at ≥99% confidence in the MMP-2 transfectants and not at all in the vector controls both of which had been treated with ConA. The MMP-2 peptides had an average ICAT ratio (MMP-2/vector) of >6.0 (Fig. 1C). These peptides were judged to be singletons, that is, being present only in the MMP-2 transfectants, as the intensities of the ion peaks for the two peptides in the vector cell proteome were within the background noise of the spectra.

ICAT analysis to identify known MMP-2 substrates. We quantified the relative abundance of the ICAT-labeled peptide pairs identified by tandem MS to determine the effects on the cell following expression of MMP-2 and as a screen for substrates of MMP-2. Tam et al. first showed by ICAT that cleaved proteins, ectodomains, and cleavage fragments would increase in quantity in the conditioned medium if proteolytically shed from the cell surface or released from the pericellular matrix by the cell surface membrane type 1 MMP (MT1-MMP) and if not then degraded (74). Similarly, such protease substrate types in cells transfected with MMP-2 will therefore have an ICAT ratio of MMP-2/vector of >1.0. On the other hand, secreted proteins were also reported to decrease in quantity if degraded by the transfected protease or if processed and subsequently cleared by the cell following reduction in protein stability (74); in this case the ICAT ratio of MMP-2/vector will be <1.0. These ratio trends can be used only as a screen for substrates and must be followed by biochemical validation since ConA also induces MT1-MMP. As well, MMP-2 protease activity might also alter the expression of other proteins and the transcription of their genes indirectly or induce the expression and activation of other proteases.

Between 53 and 166 proteins were identified per ICAT experiment using high-stringency peptide confidence levels (≥99%) (see Tables S1 and S2 in the supplemental material for full data sets). ICAT analysis of vector transfectants treated or not with ConA did not show as marked alterations in peptide ICAT ratios as when MMP-2 transfectants were analyzed (see Table S2A in the supplemental material). Tryptic peptides of known MMP-2 substrates that were generated from cleaved substrate fragments that accumulated in the conditioned medium, including several collagen types, osteonectin, amyloid beta A4, fibronectin, and procollagen C-proteinase enhancer protein, were identified with elevated levels from the MMP-2 transfectants compared to those from vector control cells (Table 1). These values were used as a guide to set ICAT ratio cutoffs for identifying candidate MMP-2 substrates. Hence, only secreted or cell membrane protein fragments in the conditioned medium having ICAT ratios ≥1.38 or ≤0.76 were considered to be potential, biologically relevant substrates.


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  		TABLE 1. Tryptic peptides of known MMP-2 substrates identified by ICAT quantitative proteomics in conditioned medium from ConA-treated Mmp2–/– fibroblasts transfected with human MMP-2 or vector alonea

Analysis of the cellular distribution of all proteins identified in the conditioned medium from ConA-treated Mmp2–/– fibroblasts transfected with MMP-2 is shown in Fig. 2A, where proteins from eight different cellular compartments were detected. Each protein was assigned a cellular localization based on information available from Swiss-Prot, Genome Ontology, or Human Protein Reference Database. Cytoplasmic proteins form the largest category (51%), followed by extracellular (24%), nuclear (14%), and plasma membrane (5%) proteins, reflecting the fact that the number of cytoplasmic proteins in the cellular proteome is larger than the number of secreted proteins. Moreover, even small numbers of cells lysing in culture can contribute up to 50% of the proteins in the conditioned medium (37). For proteins with an ICAT ratio (MMP-2/vector) of ≥1.38 only proteins from three cellular compartments were identified: extracellular (50%), cytoplasmic (40%), and plasma membrane (10%) (Fig. 2B). Proteins with an ICAT ratio of ≤0.76 were mainly from the cytoplasm (58%), with secreted proteins making up 15% of those identified (Fig. 2C). Even though intracellular proteins released to the medium in vitro or to the extracellular space in vivo upon cell death might be cleaved by extracellular MMP-2 and so could be biologically relevant, these were not further considered for substrate analysis.


Figure 2
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  		FIG. 2. Distribution of subcellular and extracellular proteins (A) identified by ICAT-labeled tryptic peptides detected in 48-h-conditioned medium from ConA-treated MMP-2 transfectants. For a full list of proteins from which these graphs were plotted, see Table S1 in the supplemental material. An ICAT ratio MMP-2/vector of ≥1.38 (B) indicates relative accumulation of potential substrate protein found as fragments in the conditioned medium, and an ICAT ratio of ≤0.76 (C) indicates relative depletion of proteins from conditioned medium by reduced expression or degradation, as was reflected by the smaller numbers of peptides identified per protein due to MMP-2 activity.

Substrate prediction by changes in ICAT ratio. A selected list of candidate MMP-2 substrates was generated, where an overall change in the MMP-2/vector ICAT ratio (≥1.38 and ≤0.76) was observed (Table 2). The substrate included were known substrates of other MMPs, candidates family members of known substrates, extracellular matrix proteins, molecules with the potential to be important in angiogenesis and cancer such as signaling molecules, and disease markers. To verify that the addition of ConA to the cells was not causing alterations in the levels of known and candidate MMP-2 substrates, we ICAT labeled and proteomically analyzed conditioned medium from vector control cells treated with ConA or vehicle alone (see Table S2A in the supplemental material). This analysis showed that in general ConA did not significantly change the levels of known substrates (fibronectin, decorin, collagens, dystroglycan, procollagen C-proteinase enhancer, amyloid beta A4, and tenascin) or the candidate substrates listed in Table 2 (IGFBP-6, follistatin-like 1 protein, biglycan, and nidogen-1).


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  		TABLE 2. Biologically interesting candidate MMP-2 substrates identified by ICAT quantitative proteomics in conditioned medium from ConA-treated Mmp2–/– fibroblasts transfected with human MMP-2 or vector alonea

Validation of substrate prediction. To discover new MMP-2 substrates, secondary biochemical confirmation was first performed on proteins having family members that are MMP substrates or known substrates of other MMPs. The levels of peptides of IGFBP-6, a glycoprotein known to inhibit growth of insulin-like growth factor II (IGF-II)-dependent cancers (3), were increased in the conditioned medium from ConA-treated MMP-2 transfectants compared to ConA-treated vector cells (ICAT ratio of 2.41) (Table 2). IGFBP-3 and IGFBP-5 are substrates of MMP-2 (21), and by biochemical assays we confirmed the MMP-2 cleavage of IGFBP-6 (Fig. 3A). Hence, IGFBP-6 is a new MMP-2 substrate. Notably, the elevated levels of IGFBP-6 in the conditioned medium were not directly due to ConA alone, as the ICAT ratios of IGFBP-6 were not significantly changed on proteomic analysis of vector transfectants treated with ConA or vehicle (Fig. 3A).


Figure 3
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  		FIG. 3. Confirmation of MMP-2 processing of human IGFBP-6 and CTGF. (A) Analysis of MMP-2 proteolysis of recombinant IGFBP-6 on silver-stained 15% SDS-PAGE gels. Arrows indicate cleaved protein fragments. Molecular weight markers (in thousands) are shown. ICAT ratios for IGFBP-6 identified in conditioned medium from Mmp2–/– cells transfected with MMP-2 or vector alone and with ConA or treated with vehicle are shown, confirming that the increase in proteolytic fragments of this substrate occurs only in the MMP-2-expressing cells and not in the vector-transfected cells or due to ConA treatment alone. (B) MMP-2 cleavage of CTGF analyzed on a 10% SDS-PAGE gel. The schematic depicts the four domains of CTGF: IGFBP, von Willebrand factor type C (vWFC-1), thrombospondin type 1 repeat (TSP-1), and C-terminal cysteine knot (CYS), with the major MMP-2 cleavage site generating the 16,374-Da N-terminal fragment and the 18,745-Da C-terminal fragment indicated. Amino acid numbering commences from the mature protein. Arrows show full-length and cleaved proteins with corresponding MALDI-TOF MS-derived mass and Edman N-terminal sequence analysis. N/D, not detectable.

CTGF, which promotes protein expression and extracellular matrix formation (55) and cell adhesion of fibroblasts and endothelial and epithelial cells (5) and which can inhibit VEGF-induced angiogenesis (31), exhibited elevated ICAT ratios in cells with active MMP-2 (Table 2). CTGF is a known substrate of MMP-1, -3, -7, and -13 (26), as well as MMP-14 (74). We confirmed the proteomic analysis indicating that CTGF is also cleaved by MMP-2 (Fig. 3B). Deconvolution of MALDI-TOF MS data and N-terminal sequencing of the cleavage products confirmed the major site of MMP-2 cleavage to be 152ALA{uparrow}AYR (Fig. 3B). Cleavage at this site in the interdomain linker, between the von Willebrand factor type C and thrombospondin type 1 repeat domains, generates an N-terminal 16,374-Da fragment and a stable 18,745-Da C-terminal fragment.

Follistatin-like 1 protein, also known as transforming growth factor ß-stimulated clone 36, is a secreted glycoprotein that has negative regulatory effects on the growth of human lung cancer cells (72). Seven peptides of follistatin-like 1 were greatly increased in the conditioned-medium fraction of the MMP-2 transfectants (Table 2), indicating accumulation of stable cleavage fragments. In vitro biochemical assays validated follistatin-like 1 as a novel MMP-2 substrate (Fig. 4A). As with IGFBP-6, the elevated levels of follistatin-like 1 in the conditioned medium were not directly due to ConA alone, as the ICAT ratios were not significantly changed on proteomic analysis of vector transfectants treated with ConA or vehicle (Fig. 4A).


Figure 4
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  		FIG. 4. Validation of MMP-2 processing of human follistatin-like 1 protein and HARP in vitro. (A) MMP-2 cleavage of follistatin-like 1 (FSTL1) on silver-stained 15% SDS-PAGE gel. Arrows indicate cleaved protein fragments. Molecular weight markers (in thousands) are shown. ICAT ratios for follistatin-like 1 identified in conditioned medium from Mmp2–/– cells transfected with MMP-2 or vector alone and treated with ConA or vehicle are shown, confirming that the increase in proteolytic fragments occurs only in the MMP-2-expressing cells and not in the vector-transfected cells or due to ConA treatment alone. (B) SDS-PAGE analysis of MMP-2 cleavage of HARP analyzed on a 15% Tris-Tricine gel. The schematic depicts the four domains of HARP, with the two MMP-2 cleavage sites indicated. The corresponding recombinant analogues of MMP-2 cleaved HARP and are shown below. Amino acid numbering commences from the mature protein. Arrows show full-length and cleaved proteins, with corresponding MALDI-TOF MS-derived masses and N-terminal sequences determined by Edman sequencing. N/D, not detectable.

HARP, also known as pleiotrophin, is an inducer of angiogenesis (66) and endothelial cell proliferation (65). We identified elevated levels of HARP peptides in the conditioned medium from ConA-treated Mmp2–/– cells transfected with MMP-2 in comparison to vector alone (ICAT ratio, 3.92). Biochemical cleavage assays revealed that MMP-2 efficiently cleaved HARP into two major fragments (Fig. 4B) with MALDI-TOF MS data and N-terminal sequencing confirming the cleavages to occur between the two thrombospondin-1 repeats at 56PCN{uparrow}WKK and in the C-terminal lysine-rich domain at 115PKP{uparrow}QAE (Fig. 4B).

Cystatin C, an important cysteine protease inhibitor involved in regulating cathepsin activity, was identified by tryptic peptides with ICAT ratios of 1.60 and 1.57. Biochemical assays confirmed that MMP-2 cleaved recombinant cystatin C, and MALDI-TOF MS analysis revealed that cleavage occurred in the sequence 6PPR{uparrow}LVG, which was confirmed by Edman sequencing (see Fig. S1A in the supplemental material). The removal of the first 8 amino acid residues of cystatin C by MMP-2 decreased inhibition of cathepsin L by 55% (see Fig. S1B in the supplemental material) (IC50 for full-length cystatin C = 48 pM; IC50 for cleaved cystatin C = 86 pM). Complete loss of the cathepsin L-inhibitory activity of cystatin C was not observed, consistent with the presence of the two remaining inhibitory loops (10), which remain after cleavage of the N-terminal inhibitory peptide (see Fig. S1C in the supplemental material).

MMP-2 proteolysis of CTGF abrogates CTGF-induced secreted protein synthesis by fibroblasts. CTGF induces extracellular matrix synthesis in fibroblasts (22). Therefore we investigated how proteolysis of CTGF by MMP-2 affected this activity. Cleavage of CTGF by MMP-2 decreased the amount of protein secreted by Mmp2–/– embryonic fibroblast cells in comparison to that induced by full-length CTGF (Fig. 5), revealing that the processing of CTGF at Ala154-Ala155 is an inactivating cleavage.


Figure 5
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  		FIG. 5. MMP-2 processing of CTGF is an inactivating cleavage of protein synthesis stimulation. Mmp2–/– embryonic fibroblasts were cultured with CTGF or MMP-2-processed CTGF (0, 10, and 100 ng/ml) for 48 h. Protein concentration of the 48-h-conditioned medium was analyzed by bicinchoninic acid assay.

MMP-2 cleavage of HARP modulates HARP-induced cell proliferation and migration. We found that 4 nM HARP stimulated NIH 3T3 murine fibroblast proliferation 1.8-fold (Fig. 6A). HARP cleaved by MMP-2 or MMP-2 alone added to the NIH 3T3 cells had no effect on cell proliferation after 18 h (Fig. 6A). Hence, MMP-2 cleavage abrogates the mitogenic activity of HARP. A similar effect was also observed with the migration of human umbilical vein endothelial cells through collagen-coated Transwells, where full-length HARP stimulated migration; however, this induction was significantly reduced after MMP-2 cleavage (Fig. 6A).


Figure 6
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  		FIG. 6. Proteolytic cleavage of HARP by MMP-2 inactivates HARP induction of cell proliferation and migration. Mitogenesis of serum-starved NIH 3T3 cells was determined by measuring [3H]thymidine incorporation over 18 h in the presence of (A) 4 nM recombinant HARP, HARP cleaved by MMP-2, MMP-2 alone, or buffer or (B) 40 nM N-HARP or C-HARP domains alone. Human umbilical vein endothelial cell migration was studied using Transwell chambers. Cells were incubated in the presence of (A) 4 nM recombinant HARP, HARP cleaved by MMP-2, MMP-2 alone, or buffer or (B) 40 nM N-HARP or C-HARP domains alone. After 4 h of incubation at 37°C, the migrated cells were stained using May-Grünwald-Giemsa solution and quantified by counting in three high-power microscopic fields (HPF)/well magnified 100x. Quantification of endothelial cell migration was determined as the mean of three fields/well for two independent experiments. Standard error bars are shown. **, Student's t test values (P < 0.01) comparing 4 nM HARP with all other conditions.

To address whether the HARP domains released by MMP-2 proteolysis retain individual bioactivity, two recombinant domains, HARP residues 9 to 59 (N-HARP) and HARP residues 60 to 110 (C-HARP), were expressed in Escherichia coli. Neither N-HARP (4 nM) nor C-HARP (4 nM) induced proliferation or displayed chemoattractant activities in comparison to those observed with full-length HARP (Fig. 6B).

During proteolysis cleavage fragments progressively accumulate in relation to the full-length substrate. Therefore, we investigated whether the stimulation of mitogenic activity and cell migration by full-length HARP would be modulated in the presence of MMP-2-proteolyzed HARP. Both cell proliferation and migration induced by HARP were significantly reduced upon the addition of MMP-2-cleaved HARP (Fig. 7A and B). This occurs despite the opposite activity observed when the individual cleavage products were added to HARP (Fig. 7C and D). When increasing concentrations of N-HARP (0 to 40 nM) were added to the culture medium containing HARP (4 nM), a significant dose-dependent increase in mitogenic activity was observed (Fig. 7C) whereas C-HARP (0 to 40 nM) reduced mitogenesis when added with HARP (Fig. 7D). When both domains were added together in fivefold excess to full-length HARP, a decrease of mitogenic activity occurred (data not shown), similar to that for the MMP-2-cleaved HARP or the C-HARP domain added in combination with full-length HARP. This indicates that the antagonistic action of the C-HARP domain dominated the weaker actions of N-HARP after cleavage.


Figure 7
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  		FIG. 7. MMP-2-proteolyzed HARP antagonizes HARP-induced cell proliferation and migration. HARP (4 nM) stimulation of mitogenesis of serum-starved NIH 3T3 cells was determined by measuring [3H]thymidine incorporation over 18 h. (A) MMP-2-proteolyzed HARP (4 nM) was added with 4 nM HARP. Concentrations of 0 to 40 nM recombinant N-HARP (C) and C-HARP with or without 4 nM HARP (D) were also studied. Cell migration was quantified using Transwell chambers. (B) Human umbilical vein endothelial cells were incubated in the presence of MMP-2-proteolyzed HARP (4 nM) with 4 nM HARP. After 4 h of incubation at 37°C, the migrated cells were stained using May-Grünwald-Giemsa solution and quantified by counting in three high-power microscopic fields (HPF)/well magnified 100x. Quantification of endothelial cell migration was determined as the mean of three fields/well for two independent experiments. Standard error bars are shown. ** and *, Student's t test P values of <0.01 and <0.05, respectively.

Cleavage of HARP and CTGF mobilizes VEGF. HARP and CTGF bind VEGF and inhibit its angiogenic activity (28, 31), and we showed complex formation by ELISA (Fig. 8B). VEGF might either mask or stabilize the cleavage sites in HARP or CTGF, thus preventing cleavage. Therefore, we investigated whether these proteins, when complexed with VEGF, were still processed by MMP-2. SDS-PAGE analysis showed that identical patterns of cleavage were observed for HARP and CTGF whether incubated alone with MMP-2 or coupled with VEGF (Fig. 8A). VEGF was not cleaved by MMP-2 (Fig. 8A) and was released from HARP or CTGF complexes immobilized to solid matrices upon incubation with MMP-2 (Fig. 8B).


Figure 8
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  		FIG. 8. MMP-2 proteolysis releases intact VEGF from HARP-VEGF and CTGF-VEGF complexes. (A) HARP or CTGF was complexed with VEGF for 24 h at 4°C and then incubated with MMP-2 for 24 h at 37°C. VEGF, HARP, and CTGF were also incubated with MMP-2 for 24 h at 37°C. The resulting cleavage fragments were analyzed by 15% Tris-Tricine SDS-PAGE and silver stained. HARP and CTGF were cleaved by MMP-2 whether incubated alone or when complexed with VEGF and generated identical cleavage fragments, indicating that VEGF did not mask these cleavage sites. VEGF was not processed by MMP-2. VEGF and CTGF comigrate at 38 kDa. Arrows indicate full-length and cleaved protein fragments. (B) VEGF was complexed with HARP or CTGF on a microtiter plate for 24 h prior to a 24-h incubation at 37°C with activated MMP-2. VEGF ELISA was used to quantify the residual VEGF bound to CTGF or HARP after MMP-2 cleavage.


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DISCUSSION
 
Isotope mass tagging quantitative proteomics was used to study the effects of proteolysis on the secretome of MMP-2-transfected cells. We discovered that HARP and CTGF are novel MMP-2 substrates that are cleaved and inactivated upon proteolysis. By cleaving these angiogenic and mitogenic cytokine inhibitors in complex with VEGF, MMP-2 releases VEGF. Notably, we found that MMP-2 did not cleave VEGF and so is one of the few MMPs not to process and inactivate this cytokine (40). Hence, MMP-2 exhibits potential proangiogenic activity by mobilizing intact VEGF from HARP or CTGF cytokine-inhibitory complexes. Mobilized VEGF has been proposed to be important for the "angiogenic switch" although the MMP substrate that upon cleavage released VEGF was not identified (6). Consistent with this, MMP-2 displays poor enzyme kinetics in the generation of endostatin- and prolactin-derived angiogenic inhibitors (27, 46). Hence, from the biochemical validation of the proteomic signature of HARP and CTGF proteolysis in a cell-based system, our mechanistic data point to MMP-2 being a potent pluripotential angiogenic stimulator: it unmasks VEGF by cleavage and inactivation of VEGF-binding inhibitory proteins but without cleaving and inactivating VEGF. Further, the remodeling of the endothelial cell basement membrane by MMP-2 facilitates endothelial cell migration and lumen formation without generating angiogenic inhibitors from the cleaved matrix proteins. Together, these pathways are consistent with the impaired angiogenesis observed in the postnatal Mmp2–/– mouse (32, 35, 58).

HARP, also known as pleiotrophin, is an 18-kDa secreted protein that forms homodimers and, along with midkine (34), forms a two-member family of heparin-binding growth factors. HARP stimulates cellular proliferation in a wide variety of cell types (38), and its role in tumor growth (13) and angiogenesis (17) has long been established. MMP-2 proteolysis of HARP occurred in the flexible linker between the two ß-sheet domains at the same cleavage site as plasmin (68). HARP directly interacts with VEGF via the TSR-1 sequence motif present on both ß-sheets of HARP and inhibits VEGF-induced angiogenesis (28). We showed that proteolytic separation of these two domains releases VEGF from inhibitory complexes in solution or from HARP immobilized on solid matrices.

In addition to releasing VEGF, HARP cleaved by MMP-2 had other direct biological effects: cleavage results in loss of HARP mitogenic and chemotactic activities and also releases HARP fragments with different biological activities. We found that C-HARP antagonizes HARP mitogenic and chemotactic activities but that, lacking the N-domain, it does not stimulate a cellular response. How this occurs is not clear. C-HARP might be a receptor antagonist or block HARP homodimer formation. Indeed, HARP mutants lacking the last 25 amino acids can dissociate HARP homodimers and form nonfunctional heterodimers (9). Since N-HARP had no effect on human umbilical vein endothelial cell migration, this supports the hypothesis that this biological activity resides in the C-terminal ß-sheet domain from residues 60 to 110. We observed a small synergistic induction of the HARP mitogenic effect induced by N-HARP. However, fivefold-higher concentrations of N-HARP or C-HARP over HARP were required to reveal both these different activities, and these may not be readily attainable in vivo. However, when the local levels of intact HARP are depleted by MMP-2 proteolysis, these cleavage fragments might attain high ratios relative to full-length HARP and so modulate HARP activity.

CTGF, another VEGF binding protein that suppresses VEGF angiogenic activity (31), was also cleaved by MMP-2. Like HARP, CTGF-VEGF complexes could be cleaved in solution or when immobilized on solid matrices. CTGF is also disrupted from its VEGF complex by MMP-3 and MMP-7, leading to the recovery of the angiogenic properties of VEGF (26). MMP-3 and -7 cleave CTGF at 154AAY{uparrow}RLE, only 2 residues C-terminal to the MMP-2 cleavage site at Ala154-Ala155 in the interdomain flexible linker. Hence, MMP-2 cleavage of CTGF and release of VEGF should lead to recovery of angiogenic stimulation. We also found that cleavage of CTGF inactivates its own anabolic actions, with loss of the CTGF-induced stimulation of fibroblast secreted protein synthesis. Since CTGF is a well-characterized stimulator of extracellular matrix synthesis (22), its cleavage and inactivation by MMP-2 have a biological outcome in tissues similar to that of MMP-mediated extracellular matrix degradation, but by a different, more potent upstream mechanism.

Among the other new substrates we identified, IGFBP-6 also functions as a cytokine binding protein. IGFBP-6 binds and inhibits the activity of IGF-II cell proliferation by preventing IGF-II interaction with the IGF-I receptor (36). The complete degradation of IGFBP-6 into multiple fragments by MMP-2 should disrupt IGF-II binding and so increase IGF-II bioavailability. Follistatin-like 1 protein, the smallest member of the SPARC (secreted protein acidic and rich in cysteine) family and a distant homologue of follistatin (48), is potently proinflammatory (53), and so its cleavage is consistent with other anti-inflammatory activities of MMP-2 resulting from cleavage of CCL7 (also known as monocyte chemoattractant protein 3) (51). Follistatin-like 1 also inhibits cell growth in vascular smooth muscle cells (43) as well as human lung cancer cell lines (72). Hence, its cleavage by MMP-2 may accelerate cell growth in human lung cancer, thereby suggesting another carcinogenic mechanism of MMP-2, in addition to stimulating angiogenesis, in the four-gene signature of the most highly virulent breast cancer metastases to the lung (52).

Follistatin is an important glycoprotein that tightly binds mature myostatin and inhibits myostatin receptor activation (1). It is also an antagonist binding protein of activin and bone morphogenic protein. In serum follistatin-like 3 performs similar functions. Although potential ligands for follistatin-like 1 have not been identified, a recurring theme for these newly discovered MMP-2 substrates is that they are cytokine binding proteins that function as cytokine inhibitors or antagonists. Upon MMP-2 cleavage, this not only inactivates the substrate but also releases the bound cytokine and so potentiates their functions.

The use of genetic models of disease has led to the identification of several substrates for MMPs in animal models in vivo that point to potential new roles for human MMPs (60, 64). However, high-content proteomic screens of human protease activity in the cellular context have rapidly increased the discovery of MMP substrates (47, 74). We found that perturbing a cellular system by transfection with MMP-2 led to many changes in the levels of proteins in the conditioned medium, even though the protease was expressed at low levels. This raises an important caveat in the interpretation of protease genetic knockout and transgenic-mouse models, in which many downstream effects may contribute to the phenotype observed. One important example of this is the cleavage of cystatin C by MMP-2. We found that this led to a 50% loss of its inhibitory properties (see Fig. S1 in the supplemental material). Hence, this is an example of how altering the expression and activity of a protease, in this case MMP-2, can alter the activity of proteases in other classes in the interconnected protease web (61).

Discovery of new substrates continues to reveal that the MMP substrate degradome is far more extensive than traditionally thought and so highlights the pleiotrophic roles of MMPs in normal cell function and in cancer (18). The new substrates reported here and their potential roles for angiogenic enhancement by MMP-2 other than by degrading the basement membrane are therefore important in the mechanistic understanding of carcinogenesis, angiogenesis, and metastasis. Our present work also suggests possible mechanisms for the reduced angiogenesis in tumors observed for the Mmp2–/– mouse (32) and in the corneal neovascularization models (35). Moreover, at sites of MMP-2 overexpression VEGF mobilization might be due to proteolytic release of VEGF from inhibitory complexes with HARP or CTGF rather than passive release from MMP-cleaved extracellular matrix proteins in tissue degradation (6).

With the other new substrates identified here, including IGFBP-6 and follistatin-like 1 protein, a theme has emerged in the control of extracellular homeostasis and cell function by MMPs (Fig. 9). MMP-2 includes in its substrate degradome many cytokine binding proteins and antagonists that are cleaved and inactivated by processing. By releasing the bound cytokine, MMP cleavage potentiates cytokine activity. This cleavage of HARP and CTGF suggests a new mechanism in the control of angiogenesis and in the regulation of connective-tissue formation. Indeed, the very many previous studies on the roles of MMPs in angiogenesis and matrix turnover might now be reinterpreted, with MMPs not only doing tissue degradation but also performing crucial roles in down-regulating matrix formation by abrogating CTGF signals and releasing angiogenic factors from inhibitory complexes to stimulate angiogenesis.


Figure 9
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  		FIG. 9. Poteolytic unmasking of growth factor activities by MMP cleavage of cytokine binding proteins. With the four new substrates of MMP-2 discovered here (HARP, CTGF, IGFBP-6, and follistatin-like 1) we suggest that a common role for MMPs, and MMP-2 in particular, is to proteolytically liberate active cytokines from inhibitory complexes. Depicted in the schematic are the cleavage and inactivation of HARP/pleiotrophin and CTGF from inhibitory complexes with VEGF. Upon cleavage, VEGF and other cytokines (black squares) are mobilized and free to interact in an autocrine or paracrine manner with cells. These potent growth factor-regulatory activities complement any matrix-degradative roles in facilitating the neovascularization shown here or in other biological processes.


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ACKNOWLEDGMENTS
 
We thank Derek Smith and the University of Victoria Proteomics Centre for help with the ICAT data.

C.M.O. is supported by a Canada Research Chair in Metalloproteinase Proteomics and Systems Biology, with research grants from the Canadian Institutes of Health Research (MT-11633), the National Cancer Institute of Canada (with funds raised by the Canadian Cancer Association), and the Canadian Breast Cancer Research Alliance Special Program Grant on Metastasis, as well as with a Centre Grant from the Michael Smith Research Foundation.


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FOOTNOTES
 
* Corresponding author. Mailing address: University of British Columbia, Centre for Blood Research, 4.401 Life Sciences Institute, 2350 Health Sciences Mall, Vancouver, British Columbia, Canada V6T 1Z3. Phone: (604) 822-2958. Fax: (604) 822-7742. E-mail: chris.overall{at}ubc.ca Back

{triangledown} Published ahead of print on 1 October 2007. Back

{dagger} Supplemental material for this article may be found at http://mcb.asm.org/. Back


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Molecular and Cellular Biology, December 2007, p. 8454-8465, Vol. 27, No. 24
0270-7306/07/$08.00+0     doi:10.1128/MCB.00821-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.



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