Perivascular Cells Regulate Endothelial Membrane Type-1 Matrix Metalloproteinase Activity

Biochemical and Biophysical Research Communications 282, 463– 473 (2001) doi:10.1006/bbrc.2001.4596, available online at http://www.idealibrary.com on

Perivascular Cells Regulate Endothelial Membrane Type-1 Matrix Metalloproteinase Activity Marc A. Lafleur,* Peter A. Forsyth,† Susan J. Atkinson,* Gillian Murphy,* and Dylan R. Edwards* ,1 *School of Biological Sciences, University of East Anglia, Norwich, NR4 7TJ, England; and †Department of Clinical Neurosciences, University of Calgary and Tom Baker Cancer Centre, 1331 29th St. N.W., Calgary, Alberta T2N 4N2, Canada

Received December 11, 2001

Angiogenic stimuli selectively induced expression of membrane type-1 matrix metalloproteinase (MT1MMP) transcripts and protein in human umbilical vein endothelial cells (HUVECs). Pro-MMP-2 activation was blocked by treatment with tissue inhibitor of metalloproteinases-2 (TIMP-2), but not by TIMP-1 or inhibitors of other proteinase classes. Anti-MT1-MMP antibodies abrogated recombinant pro-MMP-2 activation by plasma membranes, indicating that MT1-MMP is the main mediator of pro-MMP-2 activation in HUVECs. Cocultures of HUVECs with smooth muscle cells (SMC) or pericytes (PC) resulted in the suppression of HUVEC pro-MMP-2 activation. Treatment of A10 SMC conditioned media with a neutralising antiTIMP-2 antibody prevented the suppression of HUVEC pro-MMP-2 activation. Inhibition of HUVEC MT1-MMP function by PC and SM3 SMC correlated with elevated TIMP-3 expression. Thus, perivascular supporting cells regulate the functions of proangiogenic MMPs elaborated by endothelial cells via selective expression of TIMPs. This interplay may be important for maintenance of blood vessel architecture and neovascularisation. © 2001 Academic Press Key Words: angiogenesis; MMPs; TIMPs; MT1-MMP; endothelial cells; smooth muscle cells; pericytes.

Angiogenesis is a controlled process whereby new blood vessels sprout from the preexisting microvasculature (1). Although the adult endothelium is usually in a quiescent state, angiogenesis is vital during a variety of normal physiological processes such as ovulation and wound healing. Dysregulation of angiogenesis is associated with the pathogenesis of a number of diseases such as arthritis, diabetes, and tumour growth and metastasis (2). If a solid tumour is to grow 1

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beyond a certain critical size (2–3 mm 3) it must acquire the ability to increase its own blood supply in order to meet its flourishing metabolic needs (3). The acquisition of angiogenic capabilities is a key event in tumour progression, which is controlled by the balance between angiogenic factors and inhibitory molecules (2, 4, 5). Extracellular matrix (ECM) remodeling is an important aspect of angiogenesis because of the structural barriers encountered by invading endothelial cells (EC), and for liberating ECM-bound angiogenic factors such as VEGF (6). The ECs must first dissolve their underlying BM as well as degrade ECM components as they invade into the surrounding perivascular stroma (7). Matrix metalloproteinases (MMPs) are a family of zinc-binding, calcium-dependent endopeptidases that in concert can degrade all of the protein constituents of the ECM (8). The MMP family is subdivided into four main categories based on structural and substrate similarities. These are the collagenases, gelatinases, stromelysins, and the membrane type MMPs (9). All MMPs are synthesised as inactive zymogens and require proteolysis of a propeptide for activation (10). There are currently four known tissue inhibitors of metalloproteinases (TIMPs), which can inhibit the catalytic activity of most MMPs (11). Angiogenesis is dependent, at least in part, on the actions of MMPs since both TIMPs and synthetic metalloproteinase (MP) inhibitors such as BB-94 (Batimastat) display anti-angiogenic properties (12, 13). In vivo, tumours arising from B16F10 melanoma cells overexpressing TIMP-2 show reduced angiogenesis (14). Moreover, endothelial tube formation in type I collagen gels induced by bFGF and VEGF is inhibited by TIMP-2 and TIMP-3, but not TIMP-1 (12). Activation of pro-MMP-2 occurs on the plasma membrane, potentially involving the urokinase plasminogen activator (uPA)-uPA receptor (uPAR)-plasminogen system (15) as well as MT1-, MT2-, MT3-, MT5-, or MT6-MMP

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(16 –22). The activation via MT-MMPs has been proposed to involve formation of a MT-MMP/TIMP-2 complex that acts as a “receptor” for pro-MMP-2, which is then cleaved by an adjacent TIMP free MT-MMP molecule (23). Several observations indicate a link between MMP-2, MT-MMPs and angiogenesis. PEX, a fragment of MMP-2 comprising of the C-terminal haemopexin domain blocks pro-MMP-2 activation on the chick chorioallantoic membrane where it disrupts angiogenesis and tumour growth (24). Tumours generated by malignant cell lines in MMP-2 null mice displayed reduced tumour volumes and decreased levels of angiogenesis (25). Finally, endothelial tubulogenesis within fibrin gels was shown to be dependent on MMP fibrinolytic activity and that Madine-Darby canine kidney (MDCK) epithelial cell tubulogenesis was dependent on MT1-MMP (26). EC are regulated via their interactions with perivascular supporting cells, which include smooth muscle cells (SMC) in the macrovasculature and pericytes (PC) in small capillaries and venules (27). ECs and PC/SMC make direct contact with each other by adhesion molecules and gap junctions through discontinuities in the BM, and also by soluble factors (27). Proliferation of EC is inhibited by contact with PC and SMC (28). In the present study, we investigated EC-PC/SMC interactions in relation to MMP and TIMP activity. Our data indicate a mechanism by which the ECM degrading capabilities of EC is modulated via their interactions with perivascular cells. Specifically, we show that MT1-MMP activates pro-MMP-2 in human umbilical vein endothelial cells (HUVECs) in response to angiogenic stimuli, and that both PC and SMC can regulate MT1-MMP activity via selective expression of TIMP-2 and TIMP-3. This may be important for the maintenance of blood vessel architecture and angiogenesis. MATERIALS AND METHODS Cell culture. HUVECs were obtained from TCS Biologicals (Buckingham, UK) and were grown on type I collagen (SigmaAldrich, Poole, UK) coated flasks (60 ␮g/ml) in large vessel endothelial growth medium supplied by the manufacturer. Cells were grown at 37°C and 5% (v/v) CO 2 and were used between the 1st and 5th passages for all experiments. The A10 SMC line was obtained from ATCC (CRL-1476). A10 cells were maintained in Dulbecco’s modified Eagle medium-F12 (DMEMF12; Gibco-BRL, Paisley, UK) supplemented with 10% (v/v) fetal bovine serum (Gibco-BRL) and grown at 37°C and 5% (v/v) CO 2. SM3 SMC (29) were obtained from Dr. Michael Walsh, (University of Calgary, Calgary, Canada) and maintained in Eagle’s basal medium (Gibco-BRL) supplemented with 10% (v/v) foetal bovine serum (FBS) and grown at 37°C and 5% (v/v) CO 2. PC were isolated from bovine retina capillaries as previously described (30). Briefly, bovine eyes were collected from a slaughterhouse and disinfected with a 20% (v/v) betadine-PBS solution. The eye was then punctured 5 mm posterior from the limbus and cut around the globe. The vitreous body was then removed and the retina gently separated from the eye cup. After several PBS washes, the retina was minced with scalpel blades, washed, and the tissue digested with a 0.2% (w/v) collagenase/BSA (Sigma-Aldrich) solution

for 1 h. The digested tissue was then filtered through a nylon mesh (110 ␮m) and washed with DMEM-F12 (5% FBS). The flow through was centrifuged at 800g and washed twice in DMEM-F12 (5% FBS). The washed pellet was resuspended in DMEM-F12 (10% FBS) and plated into a T25 cm 2 flask. PC were cultured in DMEM- high glucose (4500 mg/L) (Gibco-BRL) supplemented with 10% (v/v) FBS and an antibiotic/antimycotic solution (Gibco-BRL) at 37°C and 5% (v/v) CO 2. PC were subcultured by the use of trypsin-EDTA and used between 1st and 3rd passages. Morphologically the cells are large and irregularly shaped with numerous extending processes as previously described by D’Amore (30). The cells showed strong immunostaining for ␣-smooth muscle actin (data not shown). For stimulation with angiogenic factors, cells were grown to confluency on type I collagen (60 ␮g/ml) coated 24 well dishes. The cells were then washed twice and incubated with large vessel endothelial cell basal medium (TCS Biologicals) and stimulated where indicated with the following factors (or a combination of these factors): vascular endothelial growth factor (VEGF, 25 ng/ml, Chemicon International Inc., Temucula, CA), basic fibroblast growth factor (bFGF, 10 ng/ml, R & D Systems, Abingdon, UK), tumour necrosis factor-␣ (TNF-␣, 10 ng/ml, Chemicon International, Inc.), transforming growth factor-␤1 (TGF-␤1, 1 ng/ml, R&D Systems), epidermal growth factor (EGF, 50 ng/ml, Chemicon International Inc.), transforming growth factor-␣ (TGF-␣, 10 ng/ml, R & D Systems), hepatocyte growth factor/scatter factor (HGF/SF, 300 U/ml, was a kind gift from Alba Warn, University of East Anglia, Norwich, UK), interleukin-1␣ (IL-1␣, 10 ng/ml, R & D Systems), angiogenin (100 ng/ml, R & D Systems), and phorbol 12-myristate 13-acetate (PMA, 10 ⫺7 M Sigma-Aldrich). In some experiments, recombinant TIMP-1 (31) and TIMP-2 (32) (1– 400 ng/ml) were also used to treat cells. In all cases, the conditioned medium from treated cells was collected after 24 h and centrifuged at 2000 rpm for 10 min to remove cells in suspension. 1 M Tris–HCl pH 8.0 was added (for a final concentration of 50 mM) to the conditioned media of these samples. Coculture experiments were carried out in 24-well plates precoated with fibronectin (25 ␮g/ml) (Biomedical Technologies, Stoughton, MA). Combinations of HUVEC:A10 SMC, HUVEC:SM3 SMC, and HUVEC:PC were used, varying the cell ratio from 1:0, 0:1, 1:1, 1:2, 2:1, 1:5, 5:1, 1:10, and 10:1. In each case a total of 1 ⫻ 10 5 cells were plated per well. The cells were left to interact with one another in medium as described above used for the HUVECs at 37°C and 5% (v/v) CO 2 for 24 h. This medium was then removed and the cells were washed twice with serum-free medium. Fresh serum-free medium was added to the cells, including PMA (10 ⫺7 M) in the desired treatments and left at 37°C, 5% (v/v) CO 2 for 24 h. The conditioned media was then collected and centrifuged at 2000 rpm for 10 min. 1 M Tris–HCl pH 8.0 was then added (for a final concentration of 50 mM) to the supernatants of these samples. For the conditioned medium transfer experiments, HUVECs and A10 SMC were grown until confluent in 6-well plates in their respective medium as described above at 37°C and 5% (v/v) CO 2. The conditioned medium was then removed and the cells washed twice with serum-free medium. Fresh serum-free medium (as used for HUVECs) was added to the confluent HUVECs and A10 SMC cells for 24 h. The medium was then removed from the cells, spun at 2000 rpm and transferred to a corresponding 6-well plate with confluent monolayers of HUVECs or A10 SMC. Where indicated, a neutralising sheep anti-human TIMP-2 polyclonal antibody or a non-specific sheep IgG control antibody (as described in (33)) was preincubated with the A10 SMC conditioned medium for 30 min prior to addition to HUVECs. PMA (10⫺7 M) was then added to the cells where appropriate and the cells incubated at 37°C and 5% (v/v) CO 2 for 24 h. The conditioned media was then collected and centrifuged at 2000 rpm for 10 min. 1 M Tris–HCl pH 8.0 was then added (for a final concentration of 50 mM) to the supernatants and gelatin zymography performed on these samples. The sheep anti-human TIMP-2 polyclonal antibody was demonstrated to be inhibitory by its ability to

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block TIMP-2’s inhibitory activity against MMP degradation of soluble 14C type I collagen (data not shown). Zymography and reverse zymography. MMP and TIMP activity in the conditioned medium of cultured cells was analysed by substrate gel electrophoresis. Zymography was performed using a 10% polyacrylamide gel co-polymerised with 1 mg/ml gelatin (SigmaAldrich). Equal amounts of samples were mixed with 1% (w/v) SDS sample buffer under nonreducing conditions and loaded onto the gel. The gels were then washed in 50 mM Tris–HCl, 5 mM CaCl 2 (pH 8.0) and 2.5% (v/v) Triton X-100 overnight and then incubated in 50 mM Tris–HCl, 5 mM CaCl 2 (pH 7.5) for 18 h at 37°C. Gels were stained with Coomassie blue and destained in 10% (v/v) acetic acid and 10% (v/v) isopropanol. Gelatinolytic activity appears as a clear band on a blue background. Reverse zymography was performed in a similar fashion as above, with a solution of gelatinase A (serum-free conditioned medium from baby hamster kidney cells) copolymerised with a 12% polyacrylamide gel. TIMP activity appeared as a blue band on a clear background. RNA analysis. HUVECs were grown to confluency in T75 cm 2 flasks and incubated in serum-free medium and stimulated without, or with PMA (10 ⫺7 M) for 3, 6, 12, and 24 h. Total RNA from HUVECs was harvested at these times using the acid guanidinium isothiocyanate method (34). Reverse transcription (RT) reactions contained 3 ␮g of total RNA, 1 ⫻ PCR buffer (10 mM Tris–HCl pH 9.0, 50 mM KCl, 1.5 mM MgCl 2), 1 mM each deoxynucleotide triphosphates (dATP, dGTP, dCTP, and dTTP), 20 units placental ribonuclease inhibitor (RNAguard, Amersham Pharmacia Biotech UK Limited, Little Chalfont, UK), 100 pmol of random hexamer oligodeoxynucleotides and 200 units of reverse transcriptase (Superscript I, Gibco-BRL). The final reaction volume was 20 ␮l. Each reaction was pre-incubated at 20°C for 10 min and the reverse transcription reaction was performed at 42°C for 50 min. Polymerase chain reactions (PCR) contained 1 ␮l of cDNA from the above RT reactions, 1 ⫻ PCR buffer (10 mM Tris–HCl pH 9.0, 50 mM KCl, 1.5 mM MgCl 2), 80 ␮M of each 4 deoxynucleotide (dATP, dGTP, dCTP, and dTTP) and 20 pmol of each forward and reverse primers. The primer sequences for the forward and the reverse primers for MT1-MMP are GCCCATTGGCCAGTTCTGGCGGG and CCTCGTCCACCTCAATGATGATC, respectively. The forward and reverse primer sequences for GAPDH are CGGAGTCAACGGATTTGGTCGTAT and AGCCTTCTCCATGGTGGTGAAGAC, respectively. Two units of Taq DNA polymerase (Gibco-BRL) was added to each reaction during the first denaturation step (“hot start”). The GAPDH primers (20 pmol) were added at the appropriate cycle number by the “primer dropping method” (35). The PCR cycles for each reaction were as follows: heat-denaturation step at 94°C for 1 min, primer annealing step at 55°C for 30 s and primer extension step at 72°C for 1 min. Thirty PCR cycles were used to amplify MT1-MMP while 23 cycles were used for GAPDH. All PCR reactions were carried out in a PCT-200 Peltier thermal cycler. Aliquots of each PCR reaction were run on a 2% (w/v) agarose gel containing 0.2 ␮g/ml ethidium bromide and equalised in order to give equivalent signals with an internal GAPDH control. Agarose gels were visualised under UV light and photographed using Polaroid film. Western blot analysis. Cell lysates from stimulated HUVECs were collected by lysing the cells with the following lysis buffer: 10 mM Tris–HCl pH 7.6, 10 mM NaCl, 3 mM MgCl 2, 1% (v/v) NP-40, and 100 ␮M PMSF (Sigma-Aldrich). This suspension was then centrifuged at 6000 rpm for 1 min to pellet the nuclei, and the supernatant collected. Total cellular protein was quantified using the BCA protein assay reagent (Pierce, Rockford IL). Samples (10 ␮g of total protein) were added to SDS–PAGE buffer containing 100 mM DTT and boiled for 5 min, run on a 12% SDS–PAGE and transferred to PVDF membrane. The membrane was then probed with a sheep polyclonal anti-human-MT1-MMP antibody (N175, as described in (36)). This primary antibody was then detected using a horseradish

peroxidase conjugated donkey-anti-sheep secondary antibody (Jackson ImmunoResearch Laboratories Inc., Luton UK) and the ECL⫹plus system (Amersham Pharmacia Biotech, UK). Plasma membrane preparation. HUVECs were grown to confluency in 4 T175 cm 2 tissue culture flasks. The cells were washed twice and incubated in serum-free medium and stimulated with PMA (10 ⫺7 M) for 24 h. The cells were then rinsed in ice-cold serum-free medium containing 100 ␮M PMSF, 1 ␮g/ml pepstatin A (SigmaAldrich), and 1 ␮g/ml of E64 (Sigma-Aldrich) and scraped in serumfree DMEM (with protease inhibitors as above), centrifuged and washed twice. The pellets were then resuspended in 5 mM Tris–HCl (pH 7.6), 0.02% (v/v) sodium azide, and protease inhibitors (as above) followed by homogenisation of the cell suspension. The cell suspension was then centrifuged for 10 min at 28,000g and the supernatant centrifuged further for 1 h at 100,000g. Crude membrane pellets were then resuspended in 20 mM Tris–HCl (pH 7.8), 10 mM CaCl 2, 0.05% (v/v) Brij35, 0.02% (w/v) sodium azide, and the protease inhibitors (as above). Plasma membrane incubation with recombinant pro-MMP-2 and anti-MT1-MMP antibodies. Plasma membranes were incubated for 30 min in the following buffer; 10 mM Tris–HCl (pH 7.9), 10 mM CaCl 2, 0.05% (v/v) Brij35, and 0.02% (w/v) sodium azide, with either no antibody, a nonspecific mouse IgG (Sigma-Aldrich; 1 mg/ml), or 2 mouse monoclonal anti-human-MT1-MMP monoclonal antibodies (500 ␮g/ml each of Fuji 113-5B7 and Fuji 114-1F2. Fuji 113-5B7 was raised against the sequence CDGNFDTVAMLRGEM in the hinge/ haemopexin region of MT1-MMP, and Fuji 114-1F2 was raised against the sequence REVPYAYIREGHEK in the MT1-MMP catalytic domain; both antibodies were gifts of Dr. K. Iwata, Fuji Chemicals, Japan. Ten nanograms of recombinant pro-MMP-2 (as described in (37)) was then added to this mixture and incubated overnight at 37°C. Samples were then analysed by gelatin zymography.

RESULTS Activation of Pro-MMP-2 by Angiogenic Agents and PMA in HUVECs HUVECs secrete significant amounts of 72 kDa proMMP-2 which can be activated to 64 and 62 kDa bands (corresponding to the intermediate and fully mature forms respectively) following PMA treatment of cells (38, 39) (Fig. 1a). We also tested nine angiogenic factors (VEGF, bFGF, EGF, TGF-␣, TNF-␣, TGF-␤1, HGF/SF, IL-1␣, and angiogenin) for their relative ability to induce activation of pro-MMP-2 in HUVECs. We found that none of the agents significantly activated pro-MMP-2 alone, but that a cocktail of all nine angiogenic agents listed above was able to activate proMMP-2 in a similar fashion to PMA (Fig. 1a). A combination of TNF-␣, TGF-␤1, and EGF also caused activation of pro-MMP-2 (Fig. 1a) to similar levels as the angiogenic cocktail suggesting that these three agents might be the key factors responsible for inducing activation of pro-MMP-2 in the angiogenic cocktail. However, although PMA and the angiogenic cocktail consistently activated pro-MMP-2, there was batch to batch variability in the HUVECs in their ability to respond to the TNF-␣, TGF-␤1, and EGF combination. Because PMA behaved just as the cocktail of physiological angiogenic agents, PMA was used in further

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FIG. 1. Mechanism of pro-MMP-2 activation in HUVECs. (a) Effect of PMA and angiogenic factors on pro-MMP-2 activation in HUVECs. Confluent HUVECs were cultured in serum-free media and stimulated either with PMA (10 ⫺7 M) or the indicated angiogenic factors for a 24-h period. The angiogenic cocktail consisted of VEGF (25 ng/ml), bFGF (10 ng/ml), TNF-␣ (10 ng/ml), TGF-␤1 (1 ng/ml), EGF (50 ng/ml), TGF-␣ (10 ng/ml), HGF/SF (300 U/ml), IL-1␣ (10 ng/ml), and angiogenin (100 ng/ml). Gelatin zymography was then performed on the conditioned media of these samples. Pro-MMP-2 has a molecular weight of 72 kDa and the intermediate and fully active forms migrate at 64 and 62 kDa, respectively. (b and c) PMA increases MT1-MMP mRNA and protein in HUVECs. (b) HUVECs were cultured in serum-free medium and stimulated with PMA (10 ⫺7 M) at various time points after which total RNA, or cell lysates were harvested. RT-PCR was performed on these samples for MT1-MMP and mRNA levels normalised to GAPDH levels. The PCR product size for MT1-MMP is 530 bp while the size for GAPDH PCR product is 307 bp. (c) Western blot analysis was performed using a sheep polyclonal antibody raised against human MT1-MMP (N175). The 60 kDa band corresponds to the active enzyme, while the smaller 45 kDa band is a processed form of MT1-MMP. (d) Inhibition of recombinant pro-MMP-2 activation by HUVEC membranes by anti-MT1-MMP antibodies. Crude membrane preparations from PMA-stimulated HUVECs were incubated with recombinant proMMP-2 with or without monoclonal anti-MT1-MMP antibodies (Fuji 113-5B7 and Fuji 114-1F2, 0.5 mg/ml of each) or a nonspecific IgG control (1 mg/ml). Gelatin zymography of these samples was then performed.

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experiments due to its reproducibility in activating pro-MMP-2 and to simplify the system.

shown) demonstrating the specificity of these antibodies for MT1-MMP.

PMA-Induced Pro-MMP-2 Activation in HUVECs Is MT1-MMP-Dependent

A10 SMC Suppress PMA-Induced Pro-MMP-2 Activation by HUVECs

To explore the proteolytic requirements for proMMP-2 activation in HUVECs, we treated cells with metallo, serine, and cysteine proteinase inhibitors along with PMA stimulation. Pro-MMP-2 activation in HUVECs was blocked only by MMP inhibitors such as TIMP-2 (see Fig. 3a), BB-94, EDTA, o-phenanthroline (data not shown), but not serine or cysteine protease inhibitors (data not shown). This result suggests that in this system, the mechanism of pro-MMP-2 activation is strictly metalloproteinase-dependent. Five of the six known MT-MMPs have been shown in vitro to activate pro-MMP-2 (16, 19 –22). We therefore performed RT-PCR on total RNA from HUVECs stimulated with PMA for various times to determine which members of the group were expressed. Semiquantitative RT-PCR was carried out using the primerdropping method with internal normalisation to GAPDH levels. It is evident in Fig. 1b that the mRNA levels for MT1-MMP increased as soon as 3 h after PMA stimulation, reaching maximal levels after 24 h. Cell lysates were also collected at these time points and a Western blot probed with a polyclonal anti-MT1MMP antibody. Figure 1c demonstrates that the protein levels for MT1-MMP increase in a time-dependent manner, similarly to the RNA. The angiogenic cocktail also selectively increased production of MT1-MMP protein (data not shown). The mRNA levels for the other members of the MT-MMP family (MT2-, MT3-, MT4-, MT5, and MT6-MMP) were not increased following PMA stimulation (data not shown). These data indicate that MT1-MMP is selectively induced in HUVECs in response to treatment with PMA, consistent with a major role for MT1-MMP in pro-MMP-2 activation. To further explore the functional involvement of MT-MMPs in pro-MMP-2 activation, membrane preparations from PMA-stimulated HUVECs were prepared and incubated with recombinant pro-MMP-2 plus a combination of two anti-MT1MMP monoclonal antibodies or control nonspecific IgG. Membranes from PMA-stimulated HUVECs activated recombinant pro-MMP-2, generating the 64 and 62 kDa forms. The anti-MT1-MMP antibodies (Fuji 1135B7 and Fuji 114-1F2 used in combination) fully suppressed pro-MMP-2 activation, while non-specific control IgG had no inhibitory effect (Fig. 1d), supporting a role for MT1-MMP in the activation process. Both monoclonal antibodies (Fuji 113-5B7 and Fuji 114-1F2) and the polyclonal anti-MT1-MMP antibody (N175) only recognised MT1-MMP by Western blot and not other members of the MT-MMP family (data not

We next asked the question of how interactions between endothelial cells and perivascular supporting cells might influence matrix remodeling capabilities. We monitored production and activation of pro-MMP-2 (and gelatinase-B/MMP-9) by zymography and TIMP activities using reverse zymography in cultures of HUVECs and A10 SMC, grown separately or cocultured in various ratios. Activation of pro-MMP-2 is an effective readout of MT1-MMP function in HUVECs. Unstimulated HUVECs secreted a large amount of pro-MMP-2, but only low levels of TIMP-1, TIMP-2, and TIMP-3 as demonstrated by zymography and reverse zymography (Figs. 2a and 2b). Upon PMA stimulation of HUVECs, there was a weak induction of MMP-9 in some batches of HUVECs, though this was not observed in the batches used for these experiments. The A10 SMC on the other hand secreted very low levels of gelatinases, TIMP-1 and TIMP-3, but higher amounts of TIMP-2 than HUVECs. The A10 SMC were unresponsive to PMA stimulation in terms of gelatinase production/activation or TIMP expression. As the A10 SMC were cocultured at increasing ratios with the HUVECs, the SMC suppressed the PMA induced activation of pro-MMP-2 carried out by the HUVECs. There was complete suppression at 1:1, 1:2, 2:1, 1:5, 1:10 cell ratios of HUVEC:A10 SMC and partial suppression at the 5:1 cell ratio (Fig. 2a). These results indicated that the A10 SMC produce a factor in relatively high amounts that was able to suppress MT1MMP-mediated pro-MMP-2 activation by HUVECs, even at high HUVEC:SMC cell ratios (2:1 and 5:1). We were then interested to see if this inhibitory effect was due to a diffusible molecule, or if cell– cell contact was required. We performed a conditioned medium transfer experiment whereby the conditioned medium from the A10 SMC was transferred to a growing culture of HUVECs (and vice versa) to see if a secreted factor produced by the A10 SMC could suppress the PMA induced activation of pro-MMP-2. As is illustrated in Fig. 2c, the conditioned medium from the A10 SMC was able to suppress the PMA induced activation of pro-MMP-2 secreted by the HUVECs. This suggests that cell-cell contact was not required for this inhibitory effect, but that a diffusible factor was responsible for the inhibition of pro-MMP-2 activation. Suppression of Pro-MMP-2 Activation in HUVECs by A10 SMC is TIMP-2-Dependent The TIMP-2 concentration in A10 SMC conditioned medium was six fold higher than the amount of

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FIG. 2. Suppression of MT-MMP-mediated pro-MMP-2 activation in HUVECs by coculture with SMC. (a and b) Coculture of HUVECs and A10 SMC. Serum-free conditioned medium from HUVECs and A10 SMC cultured alone, or cocultured at various cell ratios was analysed for secreted gelatinases by (a) gelatin zymography and for TIMPs by (b) reverse zymography. Cells were cultured in the absence or presence of PMA (10 ⫺7 M). The arrows in (b) indicate the gelatinolytic inhibitory activity corresponding to human TIMP-1 (28 kDa) and TIMP-2 (21 kDa). The A10 SMC secrete a protein with gelatinolytic inhibitory activity larger than the 28 kDa human TIMP-1 protein. This protein may be rat TIMP-1, or some other unidentified TIMP. The 24 kDa bands and the minor 23 kDa bands also with MMP inhibitory activity comigrate with unglycosylated TIMP-3. TIMP standards were prepared from conditioned medium of baby hamster kidney (BHK) cells stably transfected with either the TIMP-1 or TIMP-2 genes. Conditioned media from untransfected BHK cells was used as a standard for MMP-2. (c) Inhibition of pro-MMP-2 activation in HUVECs is due to a diffusible molecule produced by A10 SMC. Serum-free conditioned medium from a 24-h culture of HUVECs was transferred to confluent HUVECs and A10 SMC in the presence of PMA (10 ⫺7 M) and cultured for an additional 24 h. Likewise, conditioned medium from A10 SMC was transferred to confluent HUVECs and A10 SMC and cultured for a further 24 h in the presence of PMA (10 ⫺7 M). The MMP-9 standard was prepared from conditioned media of BHK cells stably transfected with the human MMP-9 gene.

TIMP-2 found in HUVEC conditioned medium, as determined by calibration of a reverse zymogram against a dilution series of pure recombinant TIMP-2 preparation (data not shown). This suggested that TIMP-2 secreted by the A10 SMC might be responsible for inhibiting PMA induced pro-MMP-2 activation. We therefore added rTIMP-1 and rTIMP-2 to HUVECs in

culture to examine effects on phorbol ester-induced activation of pro-MMP-2. Figure 3 (a and b) illustrates that TIMP-1 was unable to block pro-MMP-2 activation produced by the HUVECs up to a concentration of 400 ng/ml. TIMP-2 on the other hand was fully able to subdue the activation of pro-MMP-2 produced by the HUVECs, at a concentration as low as 100 ng/ml,

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FIG. 3. Suppression of MT-MMP-mediated pro-MMP-2 activation by A10 SMC is TIMP-2-dependent. (a and b) Effects of rTIMP-1 and rTIMP-2 on pro-MMP-2 activation in HUVECs. HUVECs were cultured for 24 h in serum-free media in the absence or presence of PMA (10 ⫺7 M) and either rTIMP-1 or rTIMP-2 (1– 400 ng/ml). Gelatin zymography was performed on these samples as illustrated in (a) and reverse zymography as demonstrated in (b). The TIMP-3 standard was prepared from the ECM of BHK cells stably transfected with the human TIMP-3 gene. (c) Serum-free conditioned media from a 24-h culture of A10 SMC was transferred to confluent HUVECs in the absence or presence of PMA (10 ⫺7 M) and in the absence or presence of an anti-TIMP-2 neutralising antibody (100 ␮g/ml) or nonspecific IgG (100 ␮g/ml).

which represents a level similar to that produced by the A10 SMC (see Fig. 3b). This suggests that TIMP-2 produced by the A10 SMC may be responsible for inhibition of the activation of pro-MMP-2 by the HUVECs. To confirm the identity of the activation inhibitory activity of the A10 SMC conditioned medium, we used a function-blocking polyclonal anti-TIMP-2 antibody. A10 SMC conditioned media was collected and added to confluent monolayers of HUVECs at which point the cells were treated with anti-TIMP-2 antibody, or control nonspecific IgG. As demonstrated in Fig. 3c, A10 SMC conditioned medium blocked PMA-induced proMMP-2 activation by HUVECs, and this inhibition was

unaffected by the presence of nonspecific IgG. However, in the presence of the anti-TIMP-2 neutralising antibody, the inhibitory effect of A10 SMC medium on pro-MMP-2 activation was eliminated. Thus, the high level of TIMP-2 produced by the A10 SMC was responsible for the suppression of activation of HUVEC proMMP-2 in this coculture system. SMC and PC Express High Levels of TIMP-3 We also observed suppression of pro-MMP-2 activation with conditioned media from SM3 SMC (data not shown) and low passage cultures of bovine retinal PC (Fig. 4). Bovine retinal PC also expressed pro-MMP-9

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FIG. 4. High levels of TIMP-3 secreted by PC inhibit pro-MMP-2 activation in HUVECs. Serum-free conditioned medium from HUVECs and PC cultured alone, or cocultured at various cell ratios was analysed for gelatinolytic activity by (a) gelatin zymography and for TIMPs by (b) reverse zymography. Cells were cultured in the absence or presence of PMA (10 ⫺7 M). The arrows in (b) indicate the gelatinolytic inhibitory activity corresponding to TIMP-2 (21 kDa), soluble glycosylated TIMP-3 (27 kDa) and soluble unglycosylated TIMP-3 (24 kDa).

constitutively, which was further increased and slightly activated following PMA treatment (Fig. 4a). These cells also secreted large amounts of pro-MMP-2, which was not activated upon PMA stimulation. The PC expressed relatively low levels TIMP-2, but instead we observed large amount of soluble TIMP-3 (Fig. 4b), and also extracellular matrix-associated TIMP-3 (data not shown). A set of TIMP activities at 27–29 kDa likely includes both TIMP-1 and the glycosylated form of TIMP-3. Coculture of HUVECs with PC showed inhibition of pro-MMP-2 activation at cell ratios of 1:1, 1:2, 2:1, 1:5, and 1:10 of HUVEC:PC (Fig. 4a). No inhibition was observed at the 5:1 and 10:1 cell ratio. Essentially equivalent data were obtained with the SM3 SMC (i.e., elevated TIMP-3 expression and suppression of pro-MMP-2 activation), the only difference being that the SM3 SMC did not express MMP-2 (data not shown). Thus, with both the SM3 SMC and the PC, inhibition of MT1-MMP-mediated pro-MMP-2 activation by HUVECs was evident, though this correlated with elevated TIMP-3 production rather than TIMP-2.

We have also studied primary human SMC cocultured with HUVECs and again observed inhibition of proMMP-2 activation; the primary SMC secreted high levels of TIMP-1, -2, and -3 (data not shown). DISCUSSION Angiogenesis involves the proliferation and migration of EC and their assembly into tubular structures. Subsequently, these neo-vessels mature by recruitment of perivascular supporting cells that modulate EC behaviour by fostering ECM deposition and inhibition of proliferation (40, 41). There is compelling evidence for the involvement of MMPs in angiogenesis from knockout mice (25, 42) and from in vitro and in vivo models of tumour vascularisation that have examined the effects of MMPs and TIMPs and synthetic MP inhibitors (43, 44). The contributions of MMPs may be made at several points, such as MMP-9-mediated ECM degradation leading to increased bioavailability of factors such as VEGF during endochondral bone forma-

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tion, (45) or activation of the angiogenic switch during pancreatic carcinogenesis (6). The cell surface location of MT-MMPs provides an ideal mechanism for focal control of proteolysis that would allow penetration of ECM structures by an invading vessel sprout, while preserving matrix integrity distally (26). The MTMMPs are themselves very effective, broad-spectrum ECM degrading proteinases (36, 46, 47) and pericellular proteolysis would be further assisted by their immediate downstream effector, MMP-2. Our data show that in HUVECs, pro-MMP-2 activation induced by PMA or an angiogenic cocktail consisting of nine angiogenic agents is metalloproteinasedependent, without a requirement for input from other extracellular proteinase classes. Moreover, although transcripts for all six known MT-MMPs are detected in HUVECs, the selective induction of MT1-MMP following PMA stimulation and the blocking effect of monoclonal anti-MT1-MMP antibodies on pro-MMP-2 activation in HUVEC membrane preparations argue that MT1-MMP itself is the main enzyme involved in this activation mechanism. Our data linking MT1-MMP function and pro-MMP-2 activation in HUVECs are supported by other studies that have demonstrated a correlation between increased MT1-MMP transcript levels and pro-MMP-2 activation (39). Also, anti-MT1MMP antibodies have been shown to inhibit proMMP-2 activation by human dermal microvascular EC membranes (48). However, additional contributions to pro-MMP-2 activation could come from both the uPA/ uPAR/plasmin cascade and thrombin through further processing of the MT-MMP-cleaved 64 kDa intermediate MMP-2 species to the fully active form (49, 50) (Lafleur et al. in preparation). Thus in vivo, several mechanisms might conspire to render MMP-2 fully active on the surface of EC of the neovasculature. The specific requirement for MT1-MMP for PMA-induced pro-MMP-2 activation in HUVECs is intriguing, given that MT1-, MT2-, MT3- MT5-, and MT6-MMP have been shown to activate pro-MMP-2 when overexpressed in vitro (16 –22). For MT1-MMP to confer invasive behaviour on RPMI1791 melanoma cells, the enzyme has to be directed to specific cell-surface sites termed “invadopodia” (51). This trafficking depends on the presence of the transmembrane region and cytoplasmic tail of MT1-MMP, which may interact with specific intracellular proteins. It is possible that following PMA stimulation, newly synthesised MT1-MMP becomes associated preferentially with specific membrane micro-domains, leading to optimal activation of pro-MMP-2 and focussed matrix proteolysis. Thus cell type-specific involvement of specific MT-MMPs in proMMP-2 activation may reflect a combination of both expression level and correct sub-cellular trafficking. This is the first study to explore the relative MP proteolytic balance in EC and cocultured perivascular cells, and our data highlight some key points. We have

shown that HUVECs respond to various angiogenic agents by up-regulating cell surface MT1-MMP activity, and the consequence of this response, namely increased pro-MMP-2 activation, can be suppressed by coculture with either SMC or PC. The SMC and PC secrete significant amounts of TIMP-2 or TIMP-3 protein, which can inhibit MT1-MMP and block the activation of pro-MMP-2 produced by HUVECs. In the case of the A10 SMC, the factor responsible for suppression of MT1-MMP action was shown specifically to be TIMP-2 from the use of neutralising antibodies. With the bovine retinal PC and SM3 SMC the dominant TIMP species was TIMP-3, which is also an efficient inhibitor of MT1-MMP (52). HUVECs express low levels of TIMPs, and respond to stimulation by upregulating expression of TIMP-1, which does not block MT-MMP activity. SMC and PC express abundant gelatinolytic activities (either MMP-2 or MMP-9), but show no evidence of pro-MMP-2 activation following PMA stimulation, which may be attributed to their high endogenous TIMP-2/TIMP-3 expression. Recruitment of PC to developing blood vessels is an essential aspect of vascular maturation (53). Plateletderived growth factor (PDGF)-B-deficient mice die late during gestation due to hemorrhage resulting from a lack of PC surrounding their capillaries (54). This picture is reinforced from the phenotypes of mice in which genes for the EC specific receptor tyrosine kinases Tie1 and Tie2 and the Tie-2 ligand, angiopoietin-1, have been disrupted (41, 55, 56). All of these are lethal mutations resulting from deficiencies at various stages in the maturation of the vasculature, with inadequate encapsulation of immature vessels by peri-endothelial support cells. In humans, mutation of Tie2 leads to the development of venous malformations with thin-walled veins and sparse coverage by SMC, instead of the normal situation where vessels of increasing diameter are surrounded by increasing numbers of layers of SMC (57). Thus the proliferation, differentiation, migration, and ECM remodeling capabilities of EC and SMC/PC are normally tightly coupled to generate mature vessels (28, 40, 58). Our data indicate that one way that mural perivascular cells may contribute to the maturation of blood vessels and the stabilisation of their architecture is via the creation of a TIMP-2 or TIMP3-rich environment, thereby containing the MMPmediated ECM remodeling capacity of EC. It will be of interest to explore further the roles of TIMP-2 and TIMP-3 in angiogenesis and the maintenance of blood vessel architecture in vivo using knockout mouse models. ACKNOWLEDGMENTS The authors wish to thank Mary Tretiak for technical assistance and Dr. K. Iwata at Fuji Chemicals for the anti-MT1-MMP antibodies. This work was supported by a grant from the Medical Research

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Council of Canada to DRE, a grant from the Alberta Cancer Board Research Initiative Program to DRE and PAF, and a grant from the MRC UK to GM. MAL was supported by a Natural Sciences and Engineering Research Council of Canada studentship, with additional support from the Alberta Heritage Foundation for Medical Research (AHFMR). DRE acknowledges support from AHFMR and the Norfolk and Norwich Big C Appeal.

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