Helsinki University Biomedical Dissertations. No. 11

Membrane-type-1 matrix metalloproteinase in pericellular

proteolysis and cell migration

Kaisa I. Lehti

Departments of Pathology and Virology, Haartman Institute and

Helsinki University Hospital, Biomedicum Helsinki

and Division of Biochemistry

Department of Biosciences University of Helsinki

Finland

Academic Dissertation

To be presented, with the permission of the Faculty of Science, University of Helsinki, for public criticism, in the Lecture Hall 2 at Biomedicum Helsinki,

Haartmaninkatu 8, Helsinki, on April 26th, 2002, at 12 o ’clock noon

Helsinki 2002

Supervised by: Professor Jorma Keski-Oja, M.D. Departments of Pathology and Virology Biomedicum and Haartman Institute University of Helsinki Helsinki, Finland

Reviewed by: Docent Erkki Koivunen, Ph.D. Division of Biochemistry Department of Biosciences University of Helsinki Helsinki, Finland

and Professor Leif C. Andersson, M.D. Department of Pathology, Haartman Institute University of Helsinki Helsinki, Finland

Opponent: Professor Karl Tryggvason, M.D. Division of Matrix Biology Department of Medical Biochemistry and Biophysics Karolinska Institutet Stockholm, Sweden

ISSN: 1457-8433 ISBN: 952-10-0510-6 (Printed version) ISBN: 952-10-0511-4 (PDF version) Yliopistopaino Helsinki 2002

Contents

Original publications………………………………………………… 7 Abbreviations…………………………………………………………. 8 Abstract…………………………..……………………………………. 9 Introduction………………………..………………………………… 10

Extracellular matrix……………………………………………….…. 10 Interstitial matrix…………………………………………….… 10 Basement membrane…………………………………………… 12

Role of extracellular matrix on cellular communication…………… 14

Extracellular matrix remodeling……………………………..……… 15 Matrix metalloproteinases………………………..…………….. 15

Collagenases………………………..……………...…… 16 Gelatinases……………………………..……….………. 17 Stromelysins and other MMPs…………………...…….. 19

Membrane-type matrix metalloproteinases…………………..… 19 Structural features………………………..………..……. 20 Activity and substrate specificity………………………. 21

Regulation of MT-MMP activity………………………….…… 23 Gene expression………………………..………………. 23 Zymogen activation………………………..…………… 25 Metalloproteinase inhibitors……………………….....… 27

MT1-MMP and TIMP-2 in proMMP-2 activation……...………29

Cell migration………………………..…….…………………..………. 31 Pericellular proteolysis………………………..………………... 32

Interplay between proteolysis, adhesion, and signaling... 32 Proteolytic cascades……………………….…………….33

Role of MT1-MMP in cell migration…………………………... 34

Biological roles of MT1-MMP ….…………………………………….34 Angiogenesis……………………………………………..…….. 35 Bone development…………………………….…………….….. 36 Cancer…………………………………….……………………..38

Aims of the present study……………………………….…………..41 Materials and methods……………………………………………... 42

Growth factors, chemicals, and enzymes…………………..…... 42 Cell culture and treatments……………………………….…….. 42 Antibodies…………………………………………………..….. 43 Northern hybridization and RNAse protection………………….43 Expression constructs ………………………………………….. 43

Transfection of cells ………………………………..………….. 44 SDS-PAGE and immunoblotting …………………………..….. 44 Gelatin zymography and reverse zymography…………….…… 44 Pulse-chase analysis and immunoprecipitation……………….... 45 Purification and N-terminal sequencing of MT1-MMP……….. 45 Cell surface biotinylation ……………………………………… 46 Cell surface cross-linking ……………………………………… 46 In vitro analyses of MT1-MMP cleavage……………………… 46 GST-fusion protein purification……………………………..… 46 Immunofluorescence and internalization assays………….…… 47 In vitro invasion assay………………………………………….. 47

Results………………………………………………………………… 48 Differential regulation of MT1-MMP gene expression and

proMMP-2 activation (I) ……………………………... …...48 Calcium ionophores inhibit proMT1-MMP activation,

binding of TIMP-2 and proMMP-2, and proMMP-2 activation (I, II) …………………………………………….49

Inactivating autocatalytic MT1-MMP cleavage produces 43-kDa cell surface and 20-kDa soluble fragments (I, II, IV) …49

Oligomerization through hemopexin and cytoplasmic domains regulates the activity and turnover of MT1-MMP (III) ……… ………………………………………….50

MT1-MMP targeting through its cytoplasmic domain is critical for cell invasion through Matrigel (IV) ……… ……50

Endocytosis in clathrin-coated pits regulates cell surface expression of MT1-MMP (V) ……… ……………………..51

Discussion…..……… …………………………………………………52

Correlation between MT1-MMP expression and proMMP-2 activation (I) ……… ……………………………………….52

Role of proMT1-MMP processing/activation in proMMP-2 activation (II) ……… ………………………………………53

Autocatalytic MT1-MMP inactivation (I, II, IV) ……… ………54 Identification of oligomeric MTI-MMP complexes (III) …… …55 ProMMP-2 activation and autocatalytic MT1-MMP

inactivation in the oligomeric complexes (II, III, IV).... …...57 MT1-MMP targeting and trafficking during cell migration

(IV, V)…………………………………………………… ...60

Perspective…………………………………………………………….63

Acknowledgements……… …………………………………………..64

References…………………………………………………………..…65

5

Original publications

This thesis is based on the following original articles, which are referred to by their Roman numerals in the text.

I. Lohi, J., Lehti, K., Westermarck, J., Kähäri, V.-M., and Keski-Oja, J.: Regulation of membrane-type matrix metalloproteinase-1 expression by growth factors and phorbol 12-myristate 13-acetate. Eur. J. Biochem. 239: 239-247, 1996.

II. Lehti, K., Lohi, J., Valtanen, H., and Keski-Oja, J.: Proteolytic processing of membrane-type-1 matrix metalloproteinase is associated with gelatinase A activation at the cell surface. Biochem. J. 334: 345-353, 1998.

III. Lehti, K., Lohi, J., Juntunen, M.M., Pei, D., and Keski-Oja, J.: Oligomerization through hemopexin and cytoplasmic domains regulates the activity and turnover of membrane-type 1 matrix metalloproteinase (MT1-MMP). J. Biol. Chem. 277: 8440-8448, 2002.

IV. Lehti, K., Valtanen, H., Wickström S., Lohi, J., and Keski-Oja, J.: Regulation of membrane-type-1 matrix metalloproteinase (MT1-MMP) activity by its cytoplasmic domain. J. Biol. Chem. 275: 15006-15013, 2000.

V. Jiang, A., Lehti, K., Wang, X., Weiss, S. J., Keski-Oja, J., and Pei, D.: Regulation of membrane-type matrix metalloproteinase 1 activity by dynamin-mediated endocytosis. Proc. Natl. Acad. Sci. U. S. A. 98: 13693-13698, 2001.

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Abbreviations

aa amino acid ADAM adamalysin-related metalloproteinase APMA p-aminophenylmercuric acetate bp base pair FGF fibroblast growth factor BM basement membrane cDNA complementary deoxyribonucleic acid CHX cycloheximide ConA concanavalin A ECM extracellular matrix EDTA ethylenediamine tetra-acetic acid EGF epidermal growth factor GAPDH glyceraldehyde-3-phosphate dehydrogenase GPI glycosyl-phosphatidyl inositol IGF-BP insulin-like growth factor binding protein IL-1 interleukin-1 kDa kilodalton MMP matrix metalloproteinase MT-MMP membrane-type matrix metalloproteinase PAGE polyacrylamide gel electrophoresis PAI plasminogen activator inhibitor PCR polymerase chain reaction PG proteoglycan PMA phorbol 12-myristate 13-acetate RECK revision-inducing cysteine-rich protein with kazal motifs SPARC secreted protein, acidic and rich in cysteine TGF-β transforming growth factor-β TIMP tissue inhibitor of metalloproteinases TNF-α tumor necrosis factor-α TTSP type II transmembrane serine proteinase uPA urokinase type plasminogen activator uPAR uPA receptor wt wild-type

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Abstract

Regulation of matrix metalloproteinase (MMP) activity occurs at the levels of gene expression, secretion, compartmentalization, zymogen activation, and inhibition by tissue inhibitors of metalloproteinases (TIMPs). Membrane-type (MT)-MMPs are a membrane-anchored subset of MMPs. They are powerful ECM-degrading proteases, which may also cleave cell surface proteins and serve as receptors and activators for secreted proteinases. The severe connective tissue disorders of MT1-MMP knockout mice, and the efficacy of MT-MMPs to promote migratory and invasive phenotype of normal and neoplastic cells, emphasize the importance of these pericellular proteolytic pathways.

We found that proteolytic processing and oligomerization are essential mechanisms to control MT1-MMP activity. Fibroblastic cells constitutively expressed MT1-MMP mRNA. MT1-MMP was synthesized as a 63-kDa zymogen, which was rapidly processed to the 60-kDa activated enzyme with N-terminal Tyr112. The MT1-MMP mRNA expression was increased two to four-fold by phorbol myristyl acetate or concanavalin A. However, neither the relative levels of mRNAs for MT1-MMP or TIMP-2, nor the levels of activated enzyme correlated with MT1-MMP-mediated proMMP-2 activation. In human HT-1080 fibrosarcoma cells, proMMP-2 activation correlated with the accelerated turnover of mature MT1-MMP through an autocatalytic inactivating cleavage to membrane-bound 43-kDa and soluble ~20-kDa forms. When crosslinked, MT1-MMP was detected as homo-oligomeric cell-surface complexes composed of mature 60-kDa and inactivated 43-kDa species. Competitive inhibition of MT1-MMP oligomerization through the cytoplasmic and hemopexin domains interfered with both proMMP-2 activation and autocatalytic MT1-MMP inactivation.

We characterized cytoplasmic domain as a critical element to regulate the cellular distribution of MT1-MMP. The enzyme was clustered at the leading edge of migrating human Bowes melanoma cells. Cytoplasmic truncations abolished clustering and MT1-MMP-induced cell invasion through reconstituted basement membrane matrixes. On the other hand, in various cell types MT1-MMP was mainly localized to intracellular compartments due to endocytosis in clathrin-coated pits. Inhibition of internalization by cytoplasmic deletion or by co-transfection with dominant negative mutant of dynamin enhanced both the cell surface localization of MT1-MMP and proMMP-2 activation.

In conclusion, MT1-MMP can be uniquely regulated at the levels of membrane localization and traffic, oligomerization, and autoproteolytic inactivation. Such cellular regulation of MT1-MMP in cooperation with adhesion, cytoskeletal rearrangement, and signaling pathways provides cells with mechanisms for precisely targeted pericellular proteolysis during cell migration and invasion.

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Introduction

Cell migration is a recurring phenomenon in development, vascular remodeling, wound healing, inflammatory responses, and cancer. During these processes many types of cells traverse basement membrane barriers and move across interstitial, basement membrane, or temporary matrixes. Invasive cell phenotype can be activated by altered interactions of cellular receptors with extracellular matrix (ECM) components and soluble factors. Limited proteolysis by cell surface proteinases such as membrane-type matrix metalloproteinases (MT-MMPs) allows cells to move through modified matrix. This movement involves continuous ECM attachments and detachments by adhesion receptors. At the cell surface proteinases may remodel not only ECM components, but also pericellular growth factors or their binding proteins and cell surface proteins including receptors and other proteolytic enzymes. Therefore, when targeted to the leading edge of migrating cells, proteolysis may not only clear the way for movement, but it also provides cells with directional information by modifying the environment in front of the cell.

Extracellular matrix ECM consists of collagens, glycoproteins, proteoglycans, and glycosaminoglycans (Aumailley and Gayraud, 1998; Zagris, 2001). It is a highly organized fibrillar meshwork, which serves as substratum for cell adhesion and migration. ECM also constitutes barriers that maintain tissue integrity, impede cell migration, and regulate molecular diffusion and transfer of stimuli. In addition, ECM forms a dynamic cellular microenvironment, which plays an important role in the determination of cell phenotype (Boudreau and Bissell, 1998; Streuli, 1999). It mediates information to and from cells directly through its components and by storing and modulating the function of growth factors/cytokines and other regulatory factors including processing enzymes and their inhibitors.

Two structurally and functionally distinct ECM categories are the specialized ECMs of interstitial connective tissues and basement membranes. Blood fibrin clot is a temporary form of connective tissue during tissue repair, the formation of which is initiated by thrombin-induced cleavage of fibrinogen to fibrin.

Interstitial matrix Interstitial matrix surrounds and is formed by connective tissue cells such as fibroblasts, osteoblasts, chondrocytes, and macrophages. It consists of a meshwork of protein fibers embedded in amorphous glycosaminoglycan/ proteoclycan substance (Aumailley and Gayraud, 1998). The composition and

9

molecular architecture of matrices differs substantially in tissues like bone, cartilage, tendons, ligaments, dermis, vessel walls, and the stroma of parenchymal organs.

Collagens are the most abundant structural components of interstitial ECM in all tissues (Prockop and Kivirikko, 1995; Aumailley and Gayraud, 1998). Fibrillar collagens (types I, II, III, V, and XI) are the most important molecules in conferring mechanical strength. They are synthesized and secreted as procollagens with large globular N- and C-terminal propeptides, which are proteolytically processed to mature collagen. These triple-helical molecules, composed of three α-chains with series of Gly-X-Y triplet sequences, are highly resistant to proteolysis. In tissues, individual 2 nm thick collagen molecules assemble into 20-200 nM diameter fibrils. Covalent crosslinking stabilizes these fibrils and they can associate laterally to form fibers. Type I collagen is the major component of collagen fibrils in a variety of tissues including bone, skin, tendon, and other fibrous tissues. It is a heterotrimer of two α1(I) and one α2(I) chains. Type II collagen, the main component of cartilage, is a homotrimer α1(II)3. Type III [α1(III)3] and V [α1(V), α2(V), α3(V)] collagens form heterotypic fibrils with type I collagen in soft connective tissues. In bone, type I collagen is accompanied by type V and XI [α1(XI), α2(XI), α3(XI)] collagens. Type XI collagen also forms heterotypic fibrils with type II collagen.

Non-fibrillar collagens form a rather heterogeneous group (Prockop and Kivirikko, 1995; Aumailley and Gayraud, 1998). Fibril-associated collagens with interrupted triple-helices (FACIT-collagens, type IX, XII, XIV, XVI, and XIX) are found in association with collagen fibrils. Type VI collagen forms microfibrils in most stromal connective tissues. The network-forming collagens are found in hypertrophic cartilage (type X), subendothelial matrices (VIII), and basement membranes (type IV). Long-chain collagen (type VII) is a component of anchoring fibrils, which stabilize the attachment of basement membranes and epithelia to underlying stroma. In addition, collagens with a transmembrane domain (type XIII and XVII), multiplexin (multiple triple-helix domain and interruptions) collagens (type XV and XVIII), and other proteins containing triple-helical domains have been characterized (Prockop and Kivirikko, 1995).

Elastic fibers are responsible for the elastic properties of tissues like lung, dermis, and large blood vessels (Rosenbloom et al., 1993). Elastin, the main component of elastic fibers, is a hydrophobic and extensively crosslinked protein, which is resistant to harsh physical treatments and to most proteinases. The elastic fibers interact with microfibrils consisting of fibrillins, other glycoproteins, and proteoglycans (Debelle and Tamburro, 1999; Saharinen et al. 1999).

Various glycoproteins are present in the interstitial ECMs. Fibronectin and vitronectin are structural ECM glycoproteins also present in plasma, which mediate cell attachment to the ECM (Tryggvason et al., 1987). Fibronectins are high-molecular weight (235-270 kDa) glycoproteins that form disulfide-linked dimers and fibrillar structures. Extensive alternative splicing of a single gene generates different forms of fibronectins. They are composed of three types of

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repeats, and other functional domains. Fibronectins interact with cell surface integrin receptors principally through RGD-sequence. They also bind to different matrix components including collagens, fibrin, and proteoglycans. During development, adhesion of embryonic cells to fibronectin is essential for their migration through fibronectin rich matrixes (George et al., 1993). Upon cell transformation, decreased synthesis and enhanced proteolytic degradation down regulates fibronectins at the cell surface and ECM (Tryggvason et al., 1987; Vartio et al., 1983). Many other glycoproteins associate with various structural ECM elements. Chondronectin, for example, is a cell-associated glycoprotein in cartilages, which promotes the attachment of chondrocytes to type II collagen. Thrombospondins, tenascins, and SPARC are capable of mediating both adhesive and anti-adhesive interactions and can induce disassembly of focal contact structures (Murphy-Ullrich, 2001).

Proteoglycans (PGs) and glycosaminoglycans (GAGs) constitute the amorphous substance of interstitial ECM. PGs consist of a core polypeptide chain, which serine and threonine residues have O-linked GAG chains of heparan, keratan, dermatan, or chondroitin sulfate (Iozzo, 1998). The metabolism of PGs is faster than that of collagens, with half-lives in tissues of a few days to several weeks. PGs form structural frameworks and act on matrix organization. As hydrophilic molecules they are important for retaining water and maintaining tissue volume. In addition, PGs play a role in the modulation of variety of biological processes including cell growth, adhesion, and invasion (Schwartz, 2000). Many ECM associated growth factors, such as FGF and VEGF family members bind to heparan sulphate PGs (Taipale and Keski-Oja, 1997). Small leucine-rich PGs, such as decorin and fibromodulin bind to ECM components and participate in the regulation of collagen fibrillogenesis and organization of the matrix (Iozzo, 1999). They also bind TGF-β and may thus modulate the biological effects of this growth factor (Yamaguchi et al., 1990). Aggregan and versican, the main chondroitin sulphate PGs of cartilage and noncartilagenous tissues, respectively, interact with glycoproteins and hyaluronic acid to create extensive networks (Wight et al., 1992). Cell surface associated PGs such as glypicans, syndecans, and CD44 modulate cell adhesion.

Basement membrane Basement membranes (BM) are specialized extracellular matrix sheets that separate epithelial and endothelial cell layers from underlying cells of collagenous stroma (Tryggvason et al., 1987; Yurchenco and O'Rear, 1994; Timpl, 1996). They are produced and assembled in co-operation by cells situated on both sides of the BM. The main BM components include type IV collagen, laminin, entactin/nidogen, and heparan and chondroitin sulfate proteoglycans (PGs). PGs are present in all BM structures where they may function in charge-dependent molecular sieving and immobilizing growth factors like FGF-2 and VEGF, which can bind to perlecan, the main BM heparan sulfate PG (Handler et al., 1997; Iozzo

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and San Antonio, 2001). Some other components, such as SPARC (secreted protein, acidic and rich in cysteine), fibulin, and multiplexin collagen types XV and XVIII have also been found in association with basement membranes. Fibronectin is abundant in fetal BMs.

Type IV collagens are trimeric proteins (~540 kDa) composed of three parallel α(IV) chains, which form a partially triple helical structure with numerous interruptions (Timpl, 1996). Six different chains have been cloned (α1(IV) to α6(IV)). The α1(IV) and α2(IV) chains are found in most basement membranes, whereas the expression of α3-6(IV) chains is more restricted (Hudson et al., 1993; Yurchenco and O'Rear, 1994). The most common trimer is α1(IV)2α2(IV). Type IV collagen trimers can assemble into a three-dimensional network through three types of interactions (Yurchenco, 1994; Timpl, 1996). Four molecules can bind to each other at the N-terminal cysteine-rich domain (7S) to form tetramers, which are joined to a network structure by a dimeric interaction between C-terminal noncollagenous (NC1) domains. Additional lateral associations of collagen molecules lead to the formation of irregular three-dimensional networks, which are stabilized by disulfide bonds and covalent cross-links.

Laminins are a group of large heterotrimeric glycoproteins (~400-900 kDa) consisting of three distinct polypeptide chains: α, β, and γ chains (Tryggvason, 1993). At least twelve different heterotrimeric laminin isoforms assembled from five α, three β, and three γ chains have been characterized with different tissue distributions and functions (Tryggvason, 1993; Colognato and Yurchenco, 2000). Like type IV collagen, laminins also self-assemble through calcium dependent interactions involving the terminal domain of three chains to form a polymer network in vitro (Yurchenco and O'Rear, 1994; Timpl, 1996).

The networks of type IV collagen and laminin are connected through various interactions. For example, nidogen is a sulfated glycoprotein (150 kDa), which bridges these molecules together (Colognato and Yurchenco, 2000). It has a C-terminal binding site to EGF repeat of laminin γ chains that is located near the center of laminin trimer, and N-terminal binding site to type IV collagen. In addition, perlecan contains binding sites for both type IV collagen and laminin (Kallunki and Tryggvason, 1992). The NC1 domains of type XV and XVIII collagens bind to perlecan and laminin/nidogen complexes (Sasaki et al. 1998; 2000)

Much of the information required for the assembly of complicated ECM structures including collagen fibrils and type IV collagen and laminin networks appear to be intrinsic to the molecules. Indeed, these components can be induced to self-assemble in vitro in cell-free systems to structures resembling those found in vivo (Yurchenco, 1994). In contrast, cell surface integrins and cytoskeletal components can control fibronectin fibrillogenesis (Magnusson and Mosher, 1998), and in tissues also basement membrane assembly may be regulated through cellular interactions.

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Role of extracellular matrix on cellular communication During embryonic development cells actively synthesize, degrade, and remodel ECM. A dynamic reciprocal interplay of the cells and the ECM environment corresponds to the flow of information that regulates cell survival, proliferation, differentiation, and migration during developmental processes (Zagris, 2001). The major players mediating and modulating this information include growth factors, ECM components, growth factor and adhesion receptors, intracellular signal transduction cascades, and extracellular modifying enzymes. In most adult tissues, ECM remodeling and cell proliferation are slow, the capacity for cell motility is repressed, and stable cell adhesion is essential for the maintenance and function of the tissues. For example, epithelial cells receive signals for quiescence through interactions with intact basement membrane and neighboring cells (Radisky et al., 2001). In the absence of specific ECM interactions cells fail to correctly respond to growth factors and maintain or achieve an appropriate phenotype (Streuli, 1999). Consequently, they function inappropriately or die. Indeed, changes in the ECM, adhesive interactions, and remodeling machinery take place early in malignant transformation and metastasis (Yap, 1998; Bissell and Radisky, 2001).

Cells attach and respond to the surrounding ECM through large diversity of adhesion receptors including integrins, which are heterodimeric transmembrane glycoproteins, composed of noncovalently associated α- and β-chains. The combination of α- and β-chain provides each integrin with a unique range of specificities for a variety of ECM components and cell surface counter receptors (Ruoslahti, 1996). Clustering of integrins by ligand specific interactions induces cytoskeletal accumulation of multiprotein complexes composed of interacting adaptor proteins and signaling molecules. Intracellular events can then feed back on the expression and activity of the integrins and other gene products of the cell. The cellular responses depend on the composition and architecture of the ECM network as well as on the repertoire of cell receptors.

Another way how ECM can affect cell behavior is by storing and mobilizing growth factors and cytokines. These signal to cells through receptor tyrosine (or serine/threonine) kinases and receptors coupled with G-proteins or signaling complexes with tyrosine or serine/threonine kinases and phosphatases. Both the remodeling of ECM components and the release or activation of ECM bound growth factors by proteolysis modulate transduced signals, which control cell phenotype. Some of these changes participate in the modulation of proteolysis by feedback signaling (Taipale and Keski-Oja, 1997). Interestingly, a distinct subfamily of discoidin domain tyrosine kinase receptors (DDR1 and DDR2), which signal in response to collagens rather than growth factors, have also been identified (Alves et al., 1995; Vogel et al., 1997). These receptors can also establish a feedback loop for cell phenotype through induced proteolysis (Olaso et al., 2001).

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Extracellular matrix remodeling Coordinated degradation of ECM is involved in various physiological tissue remodeling processes such as tissue morphogenesis and growth, angiogenesis, trophoblast implantation, bone remodeling, wound healing, and involution of postpartum uterus or postlactation mammary gland (Vu et al., 2000; Sternlicht and Werb, 2001; Zagris, 2001). Both excessive and deficient proteolysis are associated with a number of pathological conditions like arthritis, periodontitis, chronic wounds, and scleroderma (Birkedal-Hansen, 1995; Sternlicht and Werb, 2001). During tumor invasion, neoplastic cells utilize proteolytic and invasive mechanisms in a controlled but abnormally regulated fashion to allow cell attachment, localized degradation of the ECM, and cell migration through the digested barrier (Stetler-Stevenson and Yu, 2001).

The remodeling of the differentiated ECMs of various organs and the subepithelial or subendothelial basement membranes is dependent on the concerted action of several proteinases from different proteinase families. Proteases are classified into exo- or endopeptidases according to the terminal or internal cleavage site on the target proteins (Woessner, 1998). Endopeptidases are divided into the major classes of serine, cysteine, aspartic, and metalloproteinases based on amino acid sequences and cofactors determining their catalytic activity and mechanism. Matrix metalloproteinases (MMPs) form one of the four subfamilies that belong to metzincins, which in turn is one of numerous metalloproteinase superfamilies (Woessner, 1998).

Matrix metalloproteinases The MMP family consists currently of 26 distinct but structurally related vertebrate enzymes, and 21 characterized human homologues with partially overlapping substrate specificities (Nagase and Woessner, 1999; Pei, 1999b,c; Velasco et al., 1999; Park et al. 2000; Lohi et al. 2001). They are zinc-dependent neutral endopeptidases, whose activation requires the removal of the N-terminal prodomain (Fig.1). MMPs function in the degradation and remodeling of different ECM proteins and proteoglycans, but recent studies also suggest various other roles for these enzymes (Tables 1 and 2). For example, MMPs can cleave and thus modulate the assembly and activity of membrane or ECM-bound cytokine precursors, chemokines, growth factors, hormone receptors, growth factor binding proteins, and proteinase inhibitors (McCawley and Matrisian, 2001; Sternlicht and Werb, 2001). MMP activity is regulated by diverse mechanisms at the levels of gene transcription, mRNA stability, enzyme secretion and binding, zymogen activation, and inhibition by endogenous inhibitors to achieve precise proteolysis during normal tissue remodeling (Nagase and Woessner, 1999). Dysregulated MMP activity is characteristic to a number of pathological conditions such as chronic wounds, arthritis, periodontitis, cardiovascular disease, and cancer.

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Collagenases:MMP-1, -8,and -13

Stromelysins and others: 20, MMP-3, -10, -12, -19, and -

Gelatinases:MMP-2 and -9

Prod

omai

n

Cata

lytic

Hing

e

Pexi

n-lik

e

MMP-11 and -28

MMP-7 and -26

Sign

al p

eptid

e

Prototype

Cys Zn++

Cys Zn++

Cys Zn++

Cys Zn++

Cys Zn++

Zn++Cys

Fibronectin type II repeats

Membrane-bound MMPs:MMP-14, -15, -16, and -24

Cys Zn++

Cys Zn++TM

MMP-23MMP-17 and -25

Zn++ C.A. Ig-like GPI

Figure 1. Domain structure of MMPs. TM, transmembrane domain; GPI, glycosyl phospatidyl inositol-anchor; C.A., Cysteine array

MMPs can be divided into subgroups based on structural and functional criteria (Fig. 1). They are either soluble proteins or membrane proteins anchored to cellular membranes by type I or type II transmembrane domains or by a glycosyl-phosphatidyl inositol (GPI)-anchor. The soluble secreted MMPs have been further classified to subgroups of collagenases, which can degrade fibrillar collagens, gelatinases, having high activity against gelatin and type IV collagen, and stromelysins, matrilysins and other MMPs, which degrade variety of ECM components. The membrane-anchored MMPs form the group of membrane-type matrix metalloproteinases.

Collagenases Collagenases-1, -2, and -3 (MMP-1, MMP-8, and MMP-13, respectively) are the main secreted neutral proteinases, which can initiate the degradation of native helix of fibrillar collagens (Jeffrey, 1998). The hemopexin domains of these MMPs are essential for specific binding and cleavage of this substrate (Allan et al., 1991; Clark and Cawston, 1989; Knäuper et al., 1996a, 1997) All three collagenases can cleave a specific site (Gly775-Ile/Leu776) at each α-chain of the trimeric collagen molecule. The resulting N-terminal ¾ and C-terminal ¼ fragments are spontaneously denatured at 37°C to gelatin, which can be further

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degraded by other proteinases. Three collagenases have overlapping activities on the fibrillar collagen types I, II, and III, but their substrate preferences are different. MMP-1 displays highest catalytic efficiency against type III relative to type I collagen, whereas MMP-8 has reversed preference, and MMP-13 preferentially cleaves collagen type II. MMP-13 also displays higher gelatinase activity and has broader substrate specificity than MMP-1 and –8. MT-MMPs (Ohuchi et al., 1997) and MMP-2 (Aimes and Quigley, 1995; Patterson et al., 2001) are also able to degrade fibrillar collagens to the ¾ and ¼ fragments.

MMP-1 was the first MMP discovered based on its activity in metamorphosing tadpole tail (Gross and Lapiere 1962). It was also the first MMP purified to homogeneity (Stricklin et al., 1977) and cloned as a cDNA (Goldberg et al., 1986). Fibroblasts of various origins, chondrocytes, osteoblasts, endothelial cells, keratinocytes, hepatocytes, macrophages and monocytes, eosinophils, and tumor cells secrete MMP-1 in vitro (Jeffrey, 1998). MMP-8 is synthesized by polymorphonuclear leukocytes during their maturation in bone marrow, stored in intracellular granules, and released in response to external stimuli (Hasty et al., 1990). In addition, chondrocytes, rheumatoid synovial fibroblasts, gingival fibroblasts, bronchial epithelial cells, and melanoma cells express MMP-8 (Cole et al., 1996; Hanemaaijer et al., 1997; Abe et al., 2001; Prikk et al., 2001). MMP-13 is expressed during fetal bone development, postnatal bone remodeling, and gingival wound repair (Johansson et al., 1997b; Ravanti et al., 1999). In addition, MMP-13 expression has been associated with pathological conditions such as severe chronic inflammation in osteoarthritic cartilage, rheumatoid synovium, and chronic wounds, as well as malignant tumor invasion (Airola et al., 1997; Johansson et al., 1997a; Vaalamo et al., 1997; Balbin et al., 1999)

Gelatinases MMP-2 (gelatinase A, 72-kDa gelatinase) and MMP-9 (gelatinase B, 92-kDa gelatinase) differ from the other MMPs by containing three head-to-tail repeats homologous to the type II repeat of the collagen-binding domain of fibronectin (Collier et al., 1988). These domains are required for gelatinases to bind and cleave collagen (Keski-Oja and Todaro, 1980; Vartio et al., 1981; Vartio et al., 1982b; Keski-Oja and Vaheri, 1982; Murphy et al., 1994; Steffensen et al., 1995) and elastin (Shipley et al. 1996). The hemopexin domain does not affect MMP-2 binding to collagen (Allan et al., 1995), but similarly to collagenases it is critical for the initial cleavage of the triple helical type I collagen (Patterson et al., 2001). The hinge domain of MMP-9 contains an additional type V collagen-like insert (Wilhelm et al., 1989). Unlike other MMPs, the MMP-2 and -9 proenzymes can bind TIMP-2 and -1, respectively (Olson et al., 1997; O'Connell et al., 1994) A wide range of normal and transformed cells of fibroblastic, endothelial, and epithelial origin constitutively express MMP-2 (Vartio and Vaheri, 1981; Collier et al., 1988; Huhtala et al., 1991; Salo et al., 1983; Salo et al., 1991; Tryggvason et al., 1990). During development it is widely expressed by stromal cells (Reponen et al., 1992). Expression of MMP-9 is more restricted and is often

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Table 1. Secretory matrix metalloproteinases

Potential substrates Enzyme name(s) Matrix components Others

Collagenases MMP-1/collagenase-1 (fibroblast collagenase, interstitial collagenase)

Col I, II, III, VII, VIII, X, XI; gelatin; entactin; aggrecan; tenascin; MBP; perlecan; IGFBP-2, 3;

ProMMP-1, 2; casein, α2M; α1PI; α2AC; proTNFα

MMP-8/ collagenase-2 (neutrophil collagenase)

Col I, II, III, gelatin; entactin; aggrecan; tenascin

ProMMP-8; α2M; α1PI

MMP-13/ collagenase-3 Col I, II, III, IV, IX, X, XIV; gelatin; entactin; aggrecan; tenascin; osteonectin; fibrinogen/fibrin

ProMMP-9; 13; α2M; α2AC; PAI;

Gelatinases (type IV collagenases)

MMP-2/ gelatinase A (72-kDa gelatinase)

Gelatin; elastin; fibronectin; Col I, IV, V, VII, X, XI; laminin; aggrecan; vitronectin; decorin; MBP; IGFBP-3/5

ProMMP-1, 2,13; plasminogen; casein; α2M; α1PI; α2AC; proTNFα; proTGFβ2; proIL1β; MCP3; FGFr1

MMP-9/ gelatinase B (92-kDa gelatinase)

Gelatins; Col IV, V, VII, XI, XIV, XVII; elastin; fibrillin; fibronectin; aggrecan; fibrinogen/fibrin; MBP

Plasminogen; casein; α2M; α1PI; proTNFα; proTGFβ2; proIL1β

Stromelysins, matrilysins and others

MMP-3/ stromelysin-1

Fibronectin; laminin; gelatin; Col III, IV, V, VII, IX, X, XI; elastin; decorin; nidogen; perlecan; aggrecan; tenascin; fibrin/fibrinogen; fibrillin; entactin; vitronectin; IGFBP-3

ProMMP-1, 3,7,8,9,13; plasminogen; casein; α2M; α1PI; α2AC; proTNFα; E-cadherin; proIL-1β; proHB-EGF

MMP-10/ stromelysin-2 Fibronectin; laminin; gelatin; Col III, IV, V, II, IX, X, XI; decorin; elastin; nidogen; fibrin/fibrinogen; fibrillin; entactin; tenascin; vitronectin; aggrecan;

ProMMP-1, 8, 10

MMP-11/ stromelysin-3 Laminin; fibronectin; aggrecan; IGFBP-1

Α2M; α1PI

MMP-7/ matrilysin-1 (PUMP-1)

Fibronectin; laminin; Col IV; gelatin; aggrecan; decorin; nidogen; elastin; fibrillin; laminin; MBP: osteonectin; tenascin; vitronectin

ProMMP-2, 7; casein; α1PI; pro α-defensin; FasL; β4 integrin; E-cadherin; plasminogen; proTNFα

MMP-26/ matrilysin-2 (endometase)

Col IV; gelatin; fibronectin; fibrin/fibrinogen

ProMMP-9; casein; α1PI

MMP-12/ macrophage metalloelastase

Elastin; fibronectin; fibrinogen/fibrin; laminin

Plasminogen; casein

MMP-19/ (RASI) Col IV; gelatin; fibronectin; tenascin; aggrecan; COMP

MMP-20/ enamelysin Amelogenin; aggrecan; COMP MMP-23/ CA-MMP Gelatin MMP-28/ epilysin ND Casein

Modified from (McCawley and Matrisian, 2001; Sternlicht and Werb, 2001). Abbreviations: Col, collagen; COMP, cartilage oligomeric matrix protein; IGFBP, insulin-like growth factor binding protein; Ln, laminin; MBP, myelin basic protein; PAI, plasminogen activator inhibitor; α2M, α2 macroglobulin; α1PI, α1 proteinase inhibitor; α2AC, α2 antichymotrypsin; ND, not determined

17

low in normal tissues, but it can be induced when tissue remodeling occurs during development, wound healing and cancer invasion. MMP-9 is secreted by alveolar macrophages, polymorphonuclear leukocytes, osteoclasts, keratinocytes, and invading trophoblasts, and by several transformed cell lines, but not by fibroblastic cells (Hibbs et al., 1985; Saarialho-Kere et al., 1993; Vartio et al., 1982a; Reponen et al., 1995; Reponen et al., 1994; Salo et al., 1991). In vitro both gelatinases can degrade a variety of proteins, but the in vivo substrates are largely unknown (Sternlicht and Werb, 2001). Based on their ability to degrade type IV collagen and laminin, the main components of BMs and the correlation between the levels of MMP-2 expression and activity, and invasive potential of certain cancer cells, gelatinases, especially MMP-2 have been suggested to degrade BM components in vivo (Tryggvason et al., 1987). Both gelatinases can also efficiently degrade partially denatured collagens of all genetic types following the initial cleavage by collagenases (Overall et al., 1989). MMP-2 but not MMP-9 selectively cleaves γ2-chain of laminin 5 and promotes cell migration in a mammary epithelial cell model (Giannelli et al., 1997). MMP-9 has been suggested to affect angiogenesis by releasing ECM bound VEGF (Vu et al., 1998). The α1-proteinase inhibitor (α1-PI) is an in vivo substrate for MMP-9 in skin blisters (Liu et al., 2000)

Stromelysins, matrilysins, and other MMPs Stromelysins include stromelysin-1, -2 and -3 (MMP-3, MMP-10, and MMP-11, respectively). Matrilysins-1 and -2 (MMP-7 and MMP-26, respectively) and metalloelastase (MMP-12) are other MMPs with broad substrate specificities. Matrilysins are the smallest MMPs (~28-kDa) that lack the C-terminal hemopexin-like domains present in other MMPs. The domain structures of stromelysins resemble those of collagenases. However, they are unable to cleave native fibrillar collagens. Stromelysin-3 is inactive against many ECM components; instead it can cleave proteinase inhibitors, α2-magroglobulin (α2M) and α1-PI (Pei et al., 1994), and insulin-like growth factor binding protein (IGF-BPs; Manes et al., 1997). It also differs from most secreted MMPs by having recognition sequence for proprotein convertases between the pro- and catalytic domains. Some recently characterized MMPs including CA-MMP (MMP-23), and epilysin (MMP-28) also contain insertions with similar basic sequences (Pei 1999a; Velasco et al., 1999; Lohi et al., 2000).

Membrane-type matrix metalloproteinases The finding that plasma membranes from various tumor cells contained proMMP-2 activator sensitive to MMP inhibitors first suggested the existence of membrane-bound MMPs (Brown et al., 1993; Strongin et al., 1993b). Sato et al. (Sato et al., 1994) cloned the cDNA for MT1-MMP (MMP-14), encoding a 63-kDa type I transmembrane protein that could activate proMMP-2 and promote cell

18

invasion. To date, six additional cDNAs for membrane-bound MMPs have been cloned. Their protein products have been named MT-MMPs -2, -3, -4, -5, and –6, and CA-MMP (MMP-14, -15, -16, -17, -24, -25, and –23, respectively; Will and Hinzmann, 1995; Takino et al., 1995; Puente et al., 1996; Llano et al., 1999; Pei, 1999a, b, c; Velasco et al., 1999). MT-MMPs have recently gained considerable attention, partly because of the marked abnormalities associated with MT1-MMP knockout mice in contrast to the subtle phenotypes of mice deficient of secretory MMPs (Holmbeck et al., 1999; Shapiro, 1998; Zhou et al., 2000). MT1-MMP is also more active in ECM degradation and promoting cell invasiveness in experimental models than its soluble form or the secretory MMPs, highlighting the importance of the cell surface localization and cellular regulation of these enzymes (Hiraoka et al., 1998; Holmbeck et al., 1999; Hotary et al., 2000).

MMP-23 differs from all other characterized MMPs by having unique cysteine-rich, proline-rich, and IL-1 type II receptor-like domains instead of the C-terminal hemopexin-like domain (Pei, 1999a; Velasco et al., 1999). It has also an atypical N-terminal prodomain that lacks the conserved “cysteine switch” sequence, but contains a potential membrane spanning region. This binds proMMP-23 to the cellular membranes as a type II transmembrane protein. A single proteolytic cleavage between the pro- and catalytic domains by furin both activates the proenzyme and releases the activated soluble enzyme from the cell membrane (Pei et al., 2000).

Structural features MT1-MMP has a similar domain structure with most MMPs (Fig. 1; Sato et al., 1994). An N-terminal hydrophobic signal sequence is followed by the propeptide domain. The subsequent catalytic domain contains two zinc ion (catalytic and structural) and two calcium ion-binding sites (Fernandez-Catalan et al., 1998). A proline-rich hinge region separates the hemopexin-like domain from the catalytic domain. This hemopexin domain consists of four blades, which are stabilized around central calcium ion by an intramolecular disulfide bond between cysteine residues at both ends of this domain (Li et al., 1995; Libson et al., 1995; Morgunova et al., 1999).

Three insertions not found in most secreted MMPs determine many distinct features of MT1-MMP: 1) an 11-aa insertion between the propeptide and the catalytic domain that contains an RRKR sequence, a recognition motif for cleavage by subtilisin-like Golgi associated proteinases (present also in MMP-11, MMP-23 and MMP-28); 2) an 8-aa insertion in the N-terminal part of the catalytic domain (MT-loop); and 3) a C-terminal insertion containing a stretch of 24-aa hydrophobic sequence (a transmembrane domain) followed by a 20-aa cytoplasmic domain (Sato et al., 1994; Cao et al., 1995). The crystal structure of MT1-MMP catalytic domain in complex with TIMP-2 has been characterized (Fernandez-Catalan et al., 1998). The catalytic domain consists of a five-stranded β sheet and three α helixes characteristic for MMP fold (Fig. 2). The structure

19

differs from classical MMP fold mostly by containing two large insertions remote from the active site cleft: the long flexible loop between βII and βIII strands called as “MT-MMP loop” and the elongated loop between βV strand and αB helix (Fernandez-Catalan et al., 1998).

Based on their membrane anchoring mechanisms the six MT-MMPs can be divided to type I transmembrane proteins and GPI-anchored proteins. (Will and Hinzmann, 1995; Takino et al., 1995; Puente et al., 1996; Llano et al., 1999; Pei, 1999b; Pei, 1999c; Velasco et al., 2000). MT2-, MT3-, and MT5-MMPs (MMP-15, -16, and –24) are type I transmembrane proteins homologous to MT1-MMP and contain the three insertions found in MT1-MMP. The sequence identity of the catalytic domains of these MT-MMPs is over 65%. In contrast, MT4-, and MT6-MMPs (MMP-17 and MMP–25) lack the cytoplasmic tail and the hydrophobic region at the C-terminus acts as a signal for GPI-anchorage (Itoh et al., 1999; Kojima et al., 2000). MT4-, and MT6-MMPs contain the potential furin cleavage sites, but lack the 8-aa insertion present in the catalytic domains of other MT-MMPs.

Figure 2. Ribbon structure of MT1-MMP catalytic domain (dark grey)/ TIMP-2 (light gray) complex (Fernandez-Catalan et al., 1998). The α-helixes (HA-HC) and β-sheets (SI-SVI) of MT1-MMP catalytic domain as well as zinc and calcium ions have been indicated.

Activity and substrate specificity The substrate specificities of the MT-MMPs have been studied with recombinant catalytic domains and transmembrane deletion mutants and with wild-type enzymes expressed in bacterial and mammalian expression systems (Table 2).

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Table 2. Membrane-type matrix metalloproteinases

Potential substrates Enzyme name(s) Matrix components Others

MMP-14/ MT1-MMP

Col I, II, III; gelatin; fibronectin; Ln-1, -5; vitronectin; aggrecan; tenascin; nidogen; perlecan; fibrinogen/fibrin; fibrillin

ProMMP-2, -13; α1PI; α2M; CD44; CXCL12; proTNFα; tissue transglutaminase; αv-integrin

MMP-15/ MT2-MMP

Col I; gelatin; fibronectin; Ln-1; vitronectin; aggrecan; tenascin; nidogen; perlecan; fibrinogen/fibrin

ProMMP-2; cell surface tissue transglutaminase;

MMP-16/ MT3-MMP

Col III; gelatin; casein; fibronectin; Ln-1; vitronectin; aggrecan

ProMMP-2,

MMP-17/ MT4-MMP

Gelatin; fibrillin; fibronectin ProTNFα

MMP-24/ MT5-MMP

Heparan and chondroitin sulphate proteoglycans; gelatin; fibronectin

ProMMP-2

MMP-25/ MT6-MMP

Col IV; gelatin; fibrinogen/fibrin; fibronectin; vitronectin

α1PI

Modified from (McCawley and Matrisian, 2001; Sternlicht and Werb, 2001). Abbreviations: Col, collagen; Ln, laminin; α2M, α2 macroglobulin; α1PI, α1 proteinase inhibitor; ND, not determined

These studies have demonstrated that MT1-, MT2-, MT3-, and MT5-MMP are all potent matrix-degrading proteases. The catalytic domains of MT1- and MT2-MMP degrade gelatin, fibronectin, tenascin, nidogen, aggrecan, perlecan, and laminin and process a proTNF-α fusion protein to release mature TNF-α (Pei and Weiss 1996; d'Ortho et al., 1997). MT1-MMP retaining the hemopexin domain is able to specifically cleave native type I and type III collagens into the 3/4-1/4 fragments typical of the collagenases (Ohuchi et al., 1997), whereas the catalytic domain alone does not have this activity. MT1-MMP is also an efficient fibrinolytic proteinase (Hiraoka et al., 1998). By quantitative analyses of the activities of soluble mutants, MT3-MMP was found to be five fold more efficient in cleaving type III collagen than MT1-MMP, whereas MT3-MMP was inefficient in cleaving fibrillar type I collagen (Shimada et al., 1999). Soluble MT3-MMP also digests aggrecan, gelatin, fibronectin, vitronectin, laminin-1, α1-PI, and α2M. Proteoglycans are the preferred substrates for mouse MT5-MMP (Wang et al., 1999a). At the surface of cultured cells MT1-MMP can also cleave several receptors, such as CD44, cell surface tissue transglutaminase, and integrin αV, α3, and α5 subunits (Belkin et al., 2001; Deryugina et al., 2001b; Kajita et al., 2001; Ratnikov et al., 2001). Recently, receptor of complement component 1q (gC1qR) was found to be susceptible to MT1-MMP proteolysis in vitro and in cell cultures (Rozanov et al., 2001b). Soluble forms of MT1-, MT2-, MT3-, and MT5-MMP are efficient in initiating the proMMP-2 activation by cleavage of its prodomain.

The catalytic activity of MT4- and MT6-MMPs differs from those of the other MT-MMPs. They are ineffective in MMP-2 activation (Itoh et al., 1999; Kojima et al., 2000). They are also ineffective against many ECM components, but can cleave gelatin, fibrinogen, and fibrin. MT4-MMP has also TNF-α-

21

converting activity (English et al., 2000; English et al., 2001b; Wang et al., 1999b).

As is the case for the other MMPs, the in vivo substrates of the MT-MMPs are largely unknown. Previously, MT1-MMP was considered mainly as proMMP-2 activator, and it can also activate proMMP-13 (Knäuper et al., 1996b). However, the characterization of the severe phenotype of MT1-MMP deficient mice, in contrast to the subtle phenotype of MMP-2 deficient mice suggests important MMP-2 independent roles for MT1-MMP in connective tissue metabolism and angiogenesis (Holmbeck et al., 1999; Zhou et al., 2000; Itoh et al., 1997). In addition, overexpression of MT1-MMP in MDCK cells, which do not express MMP-2, renders invasion-incompetent cells able to invade fibrin gels and type I collagen matrix (Hiraoka et al., 1998; Hotary et al., 2000). In this experimental system the overexpression of soluble MMPs has no effect on cell invasion, highlighting the significance of the direct ECM degrading activity of MT1-MMP during invasive processes. On the other hand, MMP-2 activation is impaired in tissues of MT1-MMP and TIMP-2 deficient mice, indicating that the function of MT1-MMP and TIMP-2 is important also for MMP-2 activation in vivo (Caterina et al., 2000; Wang et al., 2000; Zhou et al., 2000).

Regulation of MT-MMP activity

Gene expression Regulatory mechanisms of MMP gene transcription exist for cell-type and stimulus specificity (Birkedal-Hansen et al., 1993; Westermarck and Kähäri, 1999). The expression of many MMPs is confined to only a few cell types or tissues, suggesting the presence of cell or tissue-specific promoters, enhancers and/or silencers (Frisch and Morisaki, 1990; Harendza et al., 1995). Most MMP genes are not expressed in normal adult tissues or by unstimulated cells in vitro, but their expression is markedly induced by growth factors, pro-inflammatory cytokines, and phorbol esters (Birkedal-Hansen et al., 1993; Westermarck and Kähäri, 1999). The induction is mediated by the TPA-responsive element (TRE) and the polyomavirus enhancer element (PEA3) present in the promoter regions of these MMPs (Birkedal-Hansen et al., 1993; Westermarck and Kähäri, 1999). In contrast, MT1-MMP is constitutively expressed in vitro by different cell types including fibroblasts, endothelial cells, and smooth muscle cells, and its expression is only modestly enhanced by PMA, TNF-α, and the lectin concanavalin A (ConA), not affected by TGF-β, and decreased by dexamethasone (Lohi and Keski-Oja, 1995; Migita et al., 1996). In addition, the mRNAs for all MT-MMPs are detectable by Northern blot analysis in normal human tissues (Table 3). MMP-2 is also constitutively expressed in most fibroblastic cells, and

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Table 3. Expression of membrane-type matrix metalloproteinases in human tissues

MT-MMP# 1 2 3 4 5 6

Brain - - ++ +++ +++ - Lung +++ + ++ + + Liver -/+ ++ ++ - - - Heart + ++ - - - - Placenta +++ ++ ++ - - - Pancreas + ++ - - + - Kidney ++ ++ - - ++ - Ovary +++ - ++ - - Intestine +++ ++ - - Prostate ++ - - - - Spleen + - - - + Testis + ++ ++ - - Colon + ++ ++ - - Muscle + ++ - - - - Leukocytes - - ++ - +++

Expression levels: +++, high; ++, considerable; +, detectable by Northern blotting; -, very low or not detectable (Will and Hinzmann, 1995; Takino et al. 1995; Puente et al. 1996; Llano et al. 1999; Pei 1999; Velasco et al. 2000). The expression levels of each MT-MMP are relative only to the expression levels of the same MT-MMP in different tissues.

its expression is enhanced by TGF-β but not by phorbol esters (Collier et al., 1988). The MT1-MMP and MMP-2 genes lack a conserved TATA sequence and AP-1 binding sites (Huhtala et al., 1990; Lohi et al., 2000). In addition, both MT1-MMP and MMP-2 promoters have Sp1 binding sites that are important for basal promoter activity. Coexpression of MT1-MMP with MMP-2 and TIMP-2 during mouse embryogenesis suggests similar regulation of gene transcription in vivo (Kinoh et al., 1996; Apte et al., 1997). The expression of both MT1-MMP and MMP-2 is down regulated after birth, whereas the expression of TIMP-2 remains high favoring tissue stability (Kinoh et al., 1996; Apte et al., 1997).

MT1-MMP expression is regulated by cytoskeleton-ECM interactions. It is induced in fibroblasts and endothelial cells by culture in three dimensional collagen matrixes, by mechanical stretching, and by treatment of the cells with the cytoskeleton disrupting agent cytochalasin D (Ailenberg and Silverman, 1996; Gilles et al., 1997; Tomasek et al., 1997; Tyagi et al., 1998; Haas et al., 1998; Haas et al., 1999). Therefore, signaling pathways initiated through collagen binding β1 integrins may regulate MT1-MMP expression (Haas and Madri, 1999). Interestingly, the induction of MT1-MMP occurs when the cells are cultured on or within a type I collagen gel, but does not occur when cells are grown on a thin coating of type I collagen (Azzam and Thompson, 1992; Haas et al., 1998; Tomasek et al., 1997). MT1-MMP expression levels are also higher in fibroblasts cultured on a floating or stress-relaxed collagen lattice compared with the

23

expression levels in actin stress-fiber containing fibroblasts cultured in stabilized lattices (Tomasek et al., 1997). Therefore, the induction could involve, in addition to ligand-specific integrin interaction, alterations in mechanical force and in the assembly of actin cytoskeleton.

In endothelial cells cultured in collagen matrix, increased binding of Egr-1 transcription factor to MT1-MMP promoter correlates with enhanced MT1-MMP transcription (Haas et al., 1999). At the MT1-MMP promoter the consensus binding sites for Sp1 and Egr-1 are overlapping, whereas MMP-2 promoter does not contain Egr-1 binding sites, thus allowing cells to specifically upregulate MT1-MMP. Interestingly, the induction of Egr-1 occurs in response to potential angiogenesis initiators such as wound formation, mechanical stress, and fluid shear stress (Khachigian et al., 1996; Silverman and Collins, 1999). In murine melanoma B16F10 cells osteopontin, an ECM protein that interacts with αVβ3

integrin and induces cell migration and invasion, enhances MT1-MMP expression via an NF-κB mediated pathway (Philip et al., 2001). Cytokines appear to regulate MT1-MMP expression during inflammation. Although TNF-α alone has little effect on MT1-MMP gene expression in cultured fibroblasts (Han et al., 2001), activation of NF-κB signaling synergistically by TNF-α and type I collagen induces MT1-MMP expression in skin fibroblasts (Han et al., 2001). TNF-α, IL-1α, and IL-1β also up regulate MT1-MMP gene expression in vascular endothelial cells (Rajavashisth et al., 1999). In human monocytes MT1-MMP is induced by lipopolysaccharide through prostaglandin-cAMP dependent pathway (Shankavaram et al., 2001).

Zymogen activation All MMPs are synthesized as latent zymogens containing an about 10-kDa amino terminal globular prodomain that protrudes into the active site of the enzyme and maintains its latency (Becker et al., 1995; Morgunova et al., 1999). A cysteine-zinc bond links the unpaired cysteine residue in the prodomain to the catalytic zinc ion preventing the access of a water molecule that is necessary for catalysis (Bode et al., 1999). The sequence surrounding the cysteine in the prodomain, (K)PRCGV/NPD(V), and the zinc binding sequence in the catalytic domain, HEXGHXXGXXH, are the two best-conserved sequences in MMPs (Nagase and Woessner, 1999). The activation of the soluble proMMPs can be accomplished in vitro by detergents, chaotropic agents, oxidants, mercurial compounds, sulfhydryl reagents, and acidic pH, which expose the prodomain cysteine by conformational change, and cause the disruption of Cys-zinc bond (“cysteine switch”; Springman et al., 1990; Van Wart and Birkedal-Hansen, 1990). In addition, cleavage of the prodomain by various serine, cysteine, and metalloproteinases leads to unfolding and exposure of the remaining part of propeptide and disruption of the cysteine-zinc bond. The initial cleavages occurring usually in the flexible, exposed αI-αII loop is followed by a series of (autocatalytic) cleavages leading to the generation of the mature enzyme.

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In most cases the proteolytic activation of proMMP occurs extracellularly. Because several soluble enzymes can activate most secreted MMPs, their availability may be as important as the type of the activating enzyme(s) in vivo (Mignatti and Rifkin, 1993, 2000). The lack of activation of many MMPs (MMP-3, MMP-9, MMP-12, and MMP-13) in macrophages of uPA deficient mice suggests an important role of the cell surface plasminogen activator/plasmin system on the activation of some secreted MMPs in vivo (Carmeliet et al., 1997).

Characterization of constitutive intracellular activation of prostromelysin-3 (MMP-11) by the Golgi-associated endopeptidase furin described a novel mechanism for the activation of MMPs (Pei and Weiss, 1995). A basic sequence RXRXKR present in the prodomain was found to signal the cleavage as shown by site-directed mutagenesis, deletion mutants, and switching of this region to MMP-1, where it also caused the intracellular activation of this MMP. All MT-MMPs contain similar basic recognition sequences (RXK/RR) at the C-terminal ends of the prodomains, and also these MMPs appear to be activated during transport to the cell surface. Indeed, co-expressed furin intracellularly activates truncated MT1-MMP lacking the transmembrane and cytoplasmic domains, and an irreversible inhibitor of furin, the Pittsburgh mutant of α1-PI prevents the activation (Pei and Weiss, 1996; Sato et al., 1996). The N-terminus of furin-activated enzyme is identical to the endogenous MT1-MMP purified from HT-1080 cells (Strongin et al., 1995). Soluble proMT1-MMP can also be activated by plasmin. However, the initial cleavage by plasmin occurs after RRK sequence leading to, after further processing, only partially the same N-terminus with wild type activated MT1-MMP (Okumura et al., 1997). In addition, extracellularly added serine proteinase inhibitors appear to have no effect on MT1-MMP activity. Furthermore, treatment of human fibroblastic and fibrosarcoma cells as well as several melanoma cells with the synthetic furin inhibitor Dec-RVKR-CH2Cl inhibits the processing of full-length MT1-MMP and MMP-2 activation (Kurschat et al., 1999; Maquoi et al., 1998; Sato et al., 1999). Yana and Weiss (Yana and Weiss, 2000) described furin-dependent and independent pathways for MT1-MMP processing, which were affected by the membrane tethering of the enzyme. They also identified two sets of basic motifs as potential substrates for proprotein convertases in the MT1-MMP prodomain. MT3-MMP is also activated by furin in the trans-Golgi networks of MDCK cells (Kang et al., 2002). However, the potentially cell-type specific regulation of MT-MMP activation by different furin-like proprotein convertases during yet uncharacterized trafficking events from endoplasmic reticulum to the cell surface is still incompletely understood. Furthermore, Cao et al. (Cao et al., 1996; Cao et al., 1998; Cao et al., 2000) have questioned the requirement of this proteolytic removal of the prodomain. In fact, they have suggested that the prodomain is required for the binding of TIMP-2 to the catalytic domain and for the catalytic activity of MT1-MMP (Cao et al., 1998; Cao et al., 2000). A possible explanation for this requirement of the prodomain may be its chaperone function that is essential for folding and trafficking of the enzyme (Cao et al., 2000). This function is dependent on the conserved Y42GYL45 sequence within the propeptide domain (Pavlaki et al., 2001). The level of

25

catalytic MMP activity obtained by conformational changes without removal of the prodomain is currently unclear (Bannikov et al. 2002).

Metalloproteinase inhibitors Endogenous inhibitors like tissue inhibitors of metalloproteinases and serum α2-macroglobulin (α2M) can inhibit the activated MMPs in vivo. α2M is an abundant serum inhibitor of many types of proteinases (Sottrup-Jensen and Birkedal-Hansen, 1989). In addition, thromposbondins can inhibit MMP-2 and -9 activation and induce their clearance through scavenger receptor-mediated endocytosis (Bein and Simons, 2000). New insight for the regulation of cell surface proteolysis was brought by the characterization of a GPI-anchored membrane glycoprotein RECK (revision-inducing cysteine-rich protein with kazal motifs), which was shown to down regulate MT1-MMP, MMP-2 and MMP-9 function and tumor angiogenesis and metastasis (Takahashi et al., 1998; Oh et al., 2001).

Specific inhibitors, TIMPs are considered to be the key MMP regulators locally in tissues. The TIMP family consists of four human TIMPs: TIMP-1, -2, -3, and –4, which share 30-50% sequence identity (Table 4; Docherty et al., 1985; Stetler-Stevenson et al., 1989; Silbiger et al., 1994; Greene et al., 1996). They are composed of N- and C-terminal domains, which both are stabilized by three disulfide bonds between six conserved cysteine residues (Bode et al., 1999; Fernandez-Catalan et al., 1998; Tuuttila et al., 1998). Removal of the C-terminal domain has minor effects on the MMP inhibition by the N-terminal domains of TIMPs (Murphy et al., 1991; Huang et al., 1997). This larger N-terminal domain folds into a β-barrel similar to an oligonucleotide/oligosacharide-binding fold of certain DNA-binding proteins (Fig. 2; Tuuttila et al., 1998; Bode et al., 1999). An N-terminal extended stretch segment important for MMP inhibition binds to the barrel core by two disulfide bridges. The smaller C-terminal domain contains a parallel stranded β-hairpin and a β-loop-β motif. α-helixes from the N- and C-terminal domains form an all-helical center of the protein, and the last acidic C-terminal residues form a flexible tail on the TIMP surface (Tuuttila et al., 1998).

All TIMPs can inhibit most MMPs by tight non-covalent binding to their active site in a 1:1 stoichiometric ratio (Howard et al., 1991; Gomez et al., 1997). MT-MMPs are an exception as TIMP-1 is a poor inhibitor of MT1-, MT2-, MT3-, and MT-5-MMP (Will et al. 1996; Shimada et al. 1999). In contrast, MT4- and MT6-MMP are also effectively inhibited by TIMP-1 (Kolkenbrock et al. 1999; English et al., 2001b). Unlike other TIMPs, TIMP-3 is a good inhibitor of the adamalysin-related metalloproteinases (ADAMs; Amour et al., 1998; Hashimoto et al., 2001). TIMPs also have differential abilities to form complexes with progelatinases through interactions between their C-terminal domains and the C-terminal hemopexin domains of progelatinases (Table 4; Goldberg et al., 1989; Olson et al., 1997; Bigg et al., 1997; Butler et al., 1999).

TIMPs have been thought to be mainly regulated at the level of gene expression. Various cultured cells constitutively express TIMP-2, whereas several

26

growth factors/cytokines and chemicals upregulate the expression of TIMP-1 (Table 4; Gomez et al., 1997). TIMP-3 expression is induced in cultured cells by stimulation with serum, EGF, and TGF-β. Unlike other TIMPs, TIMP-3 is insoluble and binds to ECM both in vitro and in vivo. Generally, TIMPs are very stable, although they may be inactivated by limited proteolysis by serine proteinases like trypsin and elastase (Itoh and Nagase, 1995; Okada et al., 1988). Recently, Maquoi et al. (2000) demonstrated that internalization through an MT1-MMP dependent mechanism and subsequent degradation downregulate TIMP-2 in tumor cell cultures.

Due to their ability to inhibit proteolytic degradation required for cell invasion, TIMPs have been indicated as negative regulators of processes like angiogenesis and cancer invasion (Gomez et al., 1997). Indeed, TIMP-1 and TIMP-2 inhibit endothelial and vascular smooth muscle cell migration (Cheng et al., 1998; Fernandez et al., 1999), TIMP-2 reduces melanoma cell invasion, tumor growth and angiogenesis in vivo (Valente et al., 1998). Adenovirus-mediated gene delivery of TIMP-3 inhibits melanoma cell invasion (Ahonen et al., 1998) and overexpression of TIMP-4 inhibits invasion of breast cancer cells in vitro and tumor growth and metastasis in vivo (Wang et al., 1997). On the other hand, TIMP-1, -2, and -3 stimulate the growth of several cell types (Henriet et al., 1999). TIMP-2 can suppress EGF-mediated mitogenic signaling (Hoegy et al., 2001), and TIMP-3 can induce apoptosis of normal and malignant cells (Ahonen et al., 1998; Baker et. al., 1998). Recent data also propose the involvement of TIMPs in invasive processes like placental invasion (Henriet et al., 1999).

Table 4. Tissue inhibitors of metalloproteinases

Inhibitor Size

Glyco-sylation

Solubility ProMMP binding

MT1-MMP Inhibition/ proMMP-2 activation

Gene expression upregulated by

TIMP-1 28 kDa (184 aa)

Yes Soluble ProMMP-9 No/ no TGF-β; FGF-2; EGF; TNF-α PDGF; IL-1, -6; PMA; retinoic acid; progesterone; oncostatin M

TIMP-2 21 kDa

(194 aa)

No Soluble Pro-MMP-2 Yes/ yes cAMP; LPS; retinoic acid; progesterone

TIMP-3 21-24 kDa

(188 aa)

Yes ECM bound

Pro-MMP-2, -9

Yes/ no TGF-β; PMA

TIMP-4 22 kDa

(195 aa)

No Soluble ProMMP-2 Yes/ no ?

Modified from (Henriet et al., 1999)

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MT1-MMP and TIMP-2 in proMMP-2 activation The activation of proMMP-2 differs from that of most secreted MMPs, as this enzyme cannot be effectively activated by serine proteases (Okada et al., 1990; Strongin et al., 1995). ProMMP-2 binds to cell membranes, where its processing/activation can be induced by phorbol esters, ConA, β1 integrin clustering antibodies, and by cultivation of cells on type I collagen (Brown et al., 1990; Overall and Sodek, 1990; Brown et al., 1993; Strongin et al., 1993a; Young et al., 1995; Yu et al., 1995). MMP inhibitors like 1,10-phenanthroline, EDTA, and TIMP-2 inhibit this processing (Brown et al., 1993; Strongin et al., 1993a).

In the first report of MT1-MMP, Sato et al. (1994) found that transfection of MT1-MMP to COS cells induces proteolytic processing of exogenous proMMP-2 to the active 64-kDa and 62-kDa forms, resembling those seen in PMA or ConA treated HT-1080 cells. Strongin et al. (1995) then characterized a receptor complex in HT-1080 cells, where the N-terminal inhibitory domain of TIMP-2 binds to the catalytic center of activated MT1-MMP, while the C-terminal domain of TIMP-2 interacts with proMMP-2 (Strongin et al., 1995). Proteolytic activity of adjacent TIMP-free MT1-MMP is required for the processing of the bound proMMP-2 to the 64-kDa intermediate form containing N-terminal Leu67 (Knäuper, 1998). Studies with catalytically inactive mutants of MMP-2 suggest that the activated 62-kDa enzyme with N-terminal Tyr110 forms by an autocatalytic MMP-2 cleavage (Crabbe et al., 1994; Atkinson et al., 1995). The stability and concentration dependent activation of the intermediate MMP-2 forms in cell culture systems also supports final cell-surface activation through an intermolecular cleavage (Overall et al., 2000). While similar MT1-MMP dependent two-step activation of MMP-2 occurs also with recombinant proteins in solution in the absence of TIMP-2 (Will et al., 1996), the membrane-anchorage of MT1-MMP is important for this activation in cell culture (Cao et al., 1995).

TIMP-2 has a dual role in the proposed activation mechanism: at low concentrations it induces proMMP-2 activation by promoting the binding of proMMP-2 to MT1-MMP, whereas at high concentrations it prevents the activation by inhibiting all proteolytic MT1-MMP activity (Strongin et al. 1995). In HT-1080 cells proMMP-2 activation is enhanced when recombinant MT1-MMP and TIMP-2 are expressed in molar ratio from 3:1 to 3:2, while an excess of TIMP-2 inhibits the activation (Butler et al., 1998). This enhancing effect requires the TIMP-2 C-terminal domain. Similarly, in a cell free system increasing TIMP-2 concentrations up to molar ratio 0.24 of TIMP-2 to MT1-MMP enhances proMMP-2 activation by agarose-bound MT1-MMP, while further TIMP-2 decreases activation (Kinoshita et al., 1998). The characterization of TIMP-2 knockout mice and the cells derived from them confirmed the dual role of TIMP-2 on proMMP-2 activation (Caterina et al., 2000; Wang et al., 2000). However, studies with modified noninhibitory TIMP-2 and MMP-2 hemopexin domain, as well as characterization of the fibroblasts from TIMP-2 deficient mice, have all indicated that the first cleavage of proMMP-2 can also occur independently of cell surface binding through TIMP-2, whereas the final autocatalytic activation totally

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depends on this binding (Higashi and Miyazaki, 1999; Overall et al., 2000). In contrast, MT2-MMP can mediate proMMP-2 activation by a TIMP-2 independent mechanism (Morrison et al. 2001)

By site-directed mutation analysis the TIMP-2 binding site in proMMP-2 has been localized to the junction of hemopexin blades III and IV (Overall et al., 1999). These blades are localized peripherally away from the catalytic domain in the crystal structure of proMMP-2 (Morgunova et al., 1999). The proMMP-2-binding site in TIMP-2 has been localized to the 9-aa anionic tail of the C-terminal domain (Tuuttila et al., 1998): Deletion of these residues decreases the association rate (Willenbrock et al., 1993) and after TIMP-2 binding this C-terminal tail loses immunoreactivity, indicating that the tail is buried in the complex (Overall et al., 2000).

The major interaction sites for the catalytic domain of active MT1-MMPs are in the N-terminal domain of TIMP-2 (Fig. 2; Fernandez-Catalan et al., 1998), while the C-terminal domain may form stabilizing interactions with the MT1-MMP hemopexin domain. The edge of TIMP-2, which binds to the MT1-MMP active site cleft, is composed of several segments. N-terminal segment Cys1 to Pro5 binds to the active site of MT1-MMP at the S1, S1', S2', and S3' sites, where Cys1 appears to act as fourth ligand for the catalytic zinc and makes a hydrogen bond to the catalytic glutamic acid (Glu240) thus replacing an attacking water molecule (Bode et al., 1999). In addition, βC-strand connector loop (Ala65-Cys72), the disulfide bridge between Cys1 and Cys72, βE-βF loop (Lys100-Cys101), and βG-βH loop participate in interactions to cover the catalytic cleft (Fernandez-Catalan et al., 1998). In the MT1-MMP catalytic domain the "MT-MMP loop" forms a surface pocket, with which the long βA-βB-hairpin loop of TIMP-2 interacts (Fernandez-Catalan et al., 1998). The Tyr36 at the end of this AB loop make an important binding contribution, because mutations of this single residue substantially reduce the rate of TIMP-2 binding to MT1-MMP, but not to other MMPs (Butler et al., 1999; Williamson et al., 2001). The MT-MMP loop is unique to MT1-, MT2-, MT3-, and MT5-MMP, which are all potent proMMP-2 activators. Accordingly, deletion and point mutations in this loop of MT1-MMP affect the kinetics of proMMP-2 activation and TIMP-2 association rate, but have no substantial effect on the MT1-MMP activity against peptide substrates and macromolecules such as fibrinogen or on the inhibition by TIMP-2 (English et al., 2001a).

Specific structural features of proMMP-2 are also likely to affect its interactions and activation. The prodomain of MMP-2 has a three-helix fold, where the helical part is similar to the corresponding part of proMMP-3, but the connecting loops are different (Becker et al., 1995; Morgunova et al., 1999). During the activation, the connecting loops of proMMP-2 are sensitive for proteolytic cleavage. MT1-MMP cleaves the Asn66-Leu67 bond within the loop between αI and αII helixes, which contains a unique disulfide bond. In addition, the catalytic domain of MMP-2 contains three type II fibronectin repeats, of which the third interacts with the prodomain and the first with the blades I and II in the

29

hemopexin domain (Morgunova et al., 1999). The activation of proMMP-2 mutant lacking the fibronectin repeats is slower than that of the full-length enzyme. Interestingly, the soluble mutant MT1-MMP and the corresponding mutant with additional deletion of the MT-loop activate this mutant proMMP-2 lacking the fibronectin repeats with a similar efficiency (English et al., 2001a). This suggests that direct interactions between the MT-loop and the fibronectin repeats in proMMP-2 might also modulate the activation process.

The major differences between TIMPs-1, -2, -3, and -4 reside in the exposed loop regions and in the flexible C-terminus, which are likely to have important roles in the determination of the MT-MMP inhibition specificity as well as in the discrimination between proMMP-2 and -9. TIMP-1 has a short AB loop not capable of extending into the pocket formed by the MT-MMP loop, which has been suggested to partly explain the lack of MT1-MMP inhibition by TIMP-1 (Butler et al., 1999). However, TIMP-3 binds MT1-MMP although it also has a quite short AB loop. TIMP-4 does not support proMMP2 activation in spite of being able to bind to the C-terminal hemopexin domain of MMP-2 (Bigg et al., 1997) and efficiently inhibit MT1-MMP (Bigg et al., 2001).

Cell migration Cell migration is a recurring phenomenon in all morphogenic processes during the embryonic development (Zagris, 2001). In the adults, physiological migration processes also include trophoblast invasion and invasive migration of neutrophils and macrophages (Mignatti and Rifkin, 2000). In many other adult cells the capacity for cell movement can be activated by wounding or trauma or by malignant transformation.

Cell migration is accomplished by propulsive forces transferred from ECM to cytoskeleton through repeated cycles of adhesion, detachment and proteolysis (Lauffenburger and Horwitz, 1996; Mitchison and Cramer, 1996). Binding of ligands to specific cell surface integrins modulates the integrin activity and induces integrin clustering to focal contacts at the edges of cells. The cytoplasmic tails of integrins interact with cytoskeletal multiprotein complexes that link the focal contacts to actin filaments and intracellular signaling pathways. These interactions regulate the activity of Rho family of small GTPases such as Cdc42, Rac, and Rho, which are critical for the assembly of actin cytoskeletal components to form membrane ruffles, filopodia, and lamellipodia (Mitchison and Cramer, 1996; Sander and Collard, 1999; Evers et al., 2000). Cdc42 is a key regulator of filopodia, which are thin cylindrical projections with bundle of long actin filaments stabilized by actin binding proteins. Rac regulates the organization of lamellipodia that are broad, thin protrusive sheets at the leading edge of migrating cells. Rho is more important for the formation of actin stress fibers and for focal contact assembly. Forward movement of the cell results from adhesive traction by focal contacts and actin filament contraction. As the cell moves forward the focal contacts move in a retrograde fashion, but the magnitude of

30

traction is greater than the reward pull through the adhesion complexes. New protrusions and focal contacts form continuously at the leading edge (Lauffenburger and Horwitz, 1996). At the cell rear focal adhesions detach from matrix and filament contraction pulls the rear forward. Here the magnitude of traction is less than contraction.

The direction of cell migration is affected by chemoattractants and by the generation of preferred adhesive interactions. Chemoattractants include cytokines and growth factors such as TGF-β, EGF, FGF, and VEGF, and matrix fragments, which are recognized by adhesion receptors. Proteolytic enzymes and their inhibitors may modulate cell migration at least by: 1) clearing the restricting ECM, 2) remodeling ECM components or cell adhesion molecules to modify affinity and dynamics of cell-ECM interaction, for example, cryptic Arg-Gly-Asp (RGD) sequences exposed from fibrillar collagen by proteolysis may modify the substrate to become suitable for migration (Davis, 1992), 3) mobilizing and activating or inactivating growth factors, cytokines, and chemokines (Imai et al., 1997; Fowlkes et al., 1994; McQuibban et al., 2001; Whitelock et al., 1996), 4) generating regulatory fragments such as endostatin, (Sasaki et al., 1998; Wickström et al., 2001), angiostatin (Cornelius et al., 1998), and sites in laminin-5 α3 chain that may direct the path in front of invading cells (Giannelli et al., 1997; Koshikawa et al., 2000).

Pericellular proteolysis In tissues, considerable molar excess of proteinase inhibitors derived either from plasma or secreted by the tissue cells protects the surrounding ECM from degradation. Focal proteolytic activity can be achieved when proteinases are compartmentalized and activated in the immediate pericellular environment where proteinase inhibitors have limited access (Sternlicht and Werb, 2001). Indeed, localizing proteases and their activators to the plasma membrane and special membrane domains via a transmembrane domain or binding to cell surface receptors has emerged as an important mechanism for the generation of biologically active proteinases that mediate variety of cellular functions (Werb, 1997; Sternlicht and Werb, 2001).

The cell surface enzymes characterized in pericellular proteolysis include MMPs, ADAMs, and serine proteases including plasmin and uPA/uPA receptor system and type II transmembrane serine proteinases (TTSPs) (Werb, 1997; Blobel, 1997; Chapman, 1997; Hooper et al., 2001). All these proteinases have multidomain structures that provide them with capacity to interact with multiple partners in many cases on both sides of plasma membrane and in intracellular compartments. Therefore, they may be involved in multiple cellular pathways.

Interplay between proteolysis, adhesion, and signaling During cell invasive and morphogenic processes, the cell surface association of

31

proteolytic activity has several advantages. First, the proteinase-inhibitor balance can be finely turned to favor pericellular proteolysis, while the surrounding tissue is protected by inhibitors (see above). As a result, sufficient matrix can be degraded to allow forward movement while an attachment surface is left for propulsion (Hiraoka et al., 1998; Hotary et al., 2000; Werb, 1997). Secondly, cell surface-associated proteinases may specifically interact with cell surface signaling receptors or their pericellular ligands, as well as with the cytoplasmic signaling complexes, and in this manner function as key regulators of signaling events. Thirdly, proteolytic enzymes may interact and cooperate with adhesion receptors to control continuous cell adhesion and detachment events required for cell movement. The ADAMs that are thought to be involved in fertilization, development, inflammation, and angiogenesis (Black et al., 1997; Blobel, 1997) have been implicated in the proteolysis of ECM components and cell surface proteins including Notch and TNF-α, in RGD-dependent and -independent interactions with integrins, and signaling via interactions of their cytoplasmic domains (Black and White, 1998). The uPA/uPA receptor system, on the other hand, is an example of complex coordinated regulation of protease and adhesion systems to promote cell migration. While uPAR is the major binding site for uPA, the plasminogen activator generating active plasmin (Chapman, 1997), uPAR may also serve as a high affinity receptor for vitronectin (Wei et al., 1994). Based on the observation that macrophage adhesion is affected by uPA binding to uPAR independently of the proteolytic activity of uPA, uPAR has been suggested to transduce signals to cytoskeleton that regulate cell adhesion (Gyetko et al., 1994). Later it was found that uPAR could form complexes with β1-integrins and caveolin (Wei et al., 1996) while integrin-dependent signaling events regulate the expression of uPAR in T-cells (Bianchi et al., 1996). Recent data also suggest that cell adhesion and migration may be modulated by interactions of MMPs with β1- and β3-integrins (Brooks et al., 1996; Deryugina et al., 2001a; Dumin et al., 2001) and other cell surface components, such as CD44 (Kajita et al., 2001), tissue transglutaminase (Belkin et al., 2001), and melanoma chondroitin sulphate proteoglycan (Iida et al., 2001). Like uPAR, MT1-MMP has also been localized to caveolin-rich lipid rafts, caveolaes (Annabi et al., 2001a; Puyraimond et al., 2001), where distinct interactions and signaling events may occur.

Proteolytic cascades Increasing evidence points to a cross talk between different proteolytic

systems at the cell surface. The uPA/uPA receptor pathway has an important role in the activation of many soluble MMPs (Carmeliet et al., 1997; Mazzieri et al., 1997). Once activated by furin-like serine proteinases, MT-MMPs co-operating with TIMPs may also contribute to the activation of many other proteinases (Knäuper, 1998). Furthermore, various secreted serine proteinases including thrombin and plasmin may modulate MT1-MMP/TIMP/MMP-2 activation cascades (Schwartz et al., 1998; Lafleur et al., 2001a; Lafleur et al., 2001b; Monea et al. 2002). A member of TTSP family, MT-SP1 can activate pro-uPA on the cell

32

surface (Takeuchi et al., 2000). One potentially important feature of these cell surface cascades is that they may allow amplification of the pericellular degradative processes, if one cell surface proteinase can activate many soluble enzymes. MT-MMPs, uPA/uPA receptor as well as TTSPs have all been localized to the leading edges of migrating cells, which with the substantial similarities between their regulation and interactions supports their cooperative and/or compensating function at the cell surface. The set of proteolytic cascades involved in invasive and degradative processes may depend on, for example, the proteinase expression pattern of a given cell type or the nature of the activating signals.

Role of MT1-MMP in cell migration MT1-MMP promotes invasive phenotype of normal and neoplastic cells of fibroblastic, endothelial, and epithelial origin. Laminin-5 cleavage by MT1-MMP and MMP-2 may be one mechanism that triggers migration of cells on epithelial basement membranes (Giannelli et al., 1997; Koshikawa et al., 2000; Seftor et al., 2001). Interestingly, the small GTPase Rac1, which regulates the dynamics of actin cytoskeleton during cell migration, induces MT1-MMP expression and clustering, proMMP-2 activation, and HT-1080 cell invasion on fibrillar collagen (Zhuge and Xu, 2001; Itoh et al., 2001). During cell migration the coordinated interplay between MT1-MMP mediated proteolysis and cell adhesion appears to involve complex regulatory mechanisms. While integrin-dependent signaling events may modulate MT1-MMP expression in response to type I collagen (Haas and Madri, 1999), MT1-MMP mediated processing of integrin pro-αV chain stimulates migration of MCF-7 breast carcinoma cells on vitronectin. This processing also enhances tyrosine phosphorylation of FAK, without affecting RGD-ligand binding by the αVβ3 integrin (Deryugina et al., 2001a; Ratnikov et al., 2001). In addition, MT1-MMP may induce tumor cell migration by cleaving CD44, the main cell surface receptor for hyaluronan (Kajita et al., 2001). Furthermore, MT1-MMP has been reported to induce activation of intracellular signaling cascades involving extracellular signal-regulated protein kinase (ERK) by a mechanism that is dependent on both the catalytic activity and the cytoplasmic domain, and correlates with increased cell invasion through Matrigel (Gingras et al., 2001).

Biological roles of MT1-MMP During mouse embryogenesis, MT1-MMP is expressed at high levels in developing blood vessels, bones, and kidney as well as in osteocartilaginous and musculotendinous tissues (Kinoh et al., 1996; Apte et al., 1997). Different distribution of mRNAs for the six MT-MMPs in human tissues (Table 3) indicate the differential tissue and cell type specific regulation of their gene expression. The high level of MT3-, MT4-, and MT5-MMP expression in brain suggests specific roles in the central nervous system, while expression of MT4- and MT6-

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MMP in leukocytes suggests roles in the inflammatory response. In the tissues of TIMP-2 and MT1-MMP-deficient mice proMMP-2

activation is impaired indicating a functional link between these two enzymes and the TIMP-2 mediated receptor system in vivo (Caterina et al., 2000; Wang et al., 2000; Zhou et al., 2000). However, MMP-2 and TIMP-2 -deficient mice display subtle phenotypes, whereas MT1-MMP deficiency causes severe defects in skeletal and extraskeletal connective tissue metabolism and cartilage angiogenesis (Holmbeck et al., 1999; Zhou et al., 2000) suggesting that the severe phenotype in MT1-MMP-deficient mice is not primarily a consequence of impaired proMMP-2 activation.

Angiogenesis Angiogenesis, i.e. formation of new blood vessels from the existing ones is essential for tissue development and repair, and tumor growth. Angiogenesis may be initiated when the balance turns to the dominance of positive angiogenesis factors by the induction of positive factors such as VEGF and FGF-2, and/or loss of negative regulation (“angiogenic switch”; Hanahan and Folkman, 1996; Pepper, 2001). In most mature tissues, on the other hand, predominating negative regulation through for example, proteinase inhibitors, thrombospondins, and interactions with various ECM components or their proteolytic fragments are important for maintaining endothelial quiescence. The activation of angiogenesis involves coordinated alterations in cell-cell and cell-ECM interactions and cell phenotype accompanied by alterations of dynamic signal transduction cascades downstream of growth factor and adhesion receptors (Hanahan and Folkman, 1996; Stromblad and Cheresh, 1996). These alterations lead to increased vascular permeability, extravascular fibrin deposition, and the induction and activation of MMP and plasminogen activator (PA)/plasmin cascades (Carmeliet and Collen, 1998; Pepper, 2001). This is followed by localized proteolytic degradation of the sub-endothelial basement membrane and the proliferation and migration of activated endothelial cells into fibrin rich matrix to form a capillary bud and then a capillary tube. During the maturation of newly formed blood vessels the regulatory balance has to be restored to favor basement membrane assembly and endothelial cell differentiation.

In MT1-MMP deficient mice angiogenesis of cartilage is defective (Holmbeck et al., 1999; Zhou et al., 2000), and in corneal angiogenesis assay these mice do not have angiogenic response to implanted FGF-2 (Zhou et al., 2000). In addition, angiogenic stimuli selectively upregulate MT1-MMP expression in endothelial cells in vitro suggesting a role for MT1-MMP during these processes (Haas and Madri, 1999). By virtue of the capacities of MT1-MMP to activate proMMP-2 and directly degrade various substrates, it may be involved in several distinct ECM remodeling events during angiogenesis. Substantial evidence supports the involvement of MMP-2 activity in angiogenesis. For example, an increase in MMP-2 activity is associated with angiogenic phenotype

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of cells in rat Swarm chondosarcoma model, while antisense MMP-2 oligonucleotides inhibit angiogenesis and tumor growth in this model (Fang et al., 2000). On the other hand, endogenous angiogenesis inhibitors like thrombospondins and endostatin (a proteolytic fragment of collagen XVIII) have been reported to repress MMP-2 activity or activation (Bein and Simons 2000; Kim et al. 2000). In addition, αvβ3 binds the intermediate form of MMP-2 through the C-terminal hemopexin-like domain (PEX) and facilitates its maturation on the cell surface (Brooks et al., 1996; Deryugina et al., 2001a). Accordingly, PEX inhibits angiogenesis when used as a recombinant protein or when delivered by a viral vector both in vitro and in vivo models (Brooks et al., 1996; Brooks et al., 1998; Pfeifer et al., 2000). Finally, in MMP-2 deficient mice, tumor induced angiogenesis is markedly reduced in a dorsal air sac assay with B16-BL6 melanoma cells (Itoh et al. 1998).

Until recently, the PA/plasmin system has been considered to be responsible for the major fibrinolytic activity of cells. While experiments with PA and plasminogen deficient mice have shown the essential role of these proteinases for the fibrinolysis by several types of cells including keratinocytes, smooth muscle cells, and inflammatory cells, these mice do not exhibit clear angiogenic phenotypes (Carmeliet et al., 1994; Romer et al., 1996; Bugge et al., 1996; Carmeliet et al., 1997; Carmeliet and Collen, 1998). Therefore, endothelial cells have been suggested to invade fibrin rich matrixes in plasmin independent manner. Indeed, Hiraoka et al. (Hiraoka et al., 1998) found that in an ex vivo model tissues from PA and plasminogen deficient mice efficiently neovascularize to surrounding fibrin gel. MMP inhibitors, but not serine, cysteine, or aspartate proteinase inhibitors prevent the neovessel formation in vivo and in vitro and the invasion of isolated endothelial cells to fibrin gel. Of the recombinant MMPs (MMP-1, -2, -3, and MT1-MMP) soluble truncated MT1-MMP has the highest fibrinolytic activity, whereas fibrinolysis by membrane anchored MT1-MMP is critical for the MDCK cell invasion to fibrin gel (Hiraoka et al., 1998). These results suggest that endothelial cells can use membrane-bound MMPs such as MT1-MMP to invade fibrin rich matrix. Interestingly, MMP-9 can also act as a fibrinolytic enzyme and prevent fibrin accumulation in glomeruli (Lelongt et al., 2001).

Bone development Bone development proceeds either by intramembranous or endochondral ossification. Axial skeleton develops through endochondral ossification (Olsen et al., 2000). Mesenchymal cells first aggregate to form bone anlagen, where the peripheral cells begin to flatten to form perichondrium. The cartilage elongates and expands by proliferation of chondrocytes and cartilage matrix deposition. In the central region, chondrocytes differentiate to hypertrophic chondrocytes with enlarged cytoplasms, which then exit cell cycle and synthesize bone matrix. These hypertrophic chondrocytes secrete angiogenic factors, which induce vascular

35

invasion and invasion of chondroclasts, osteoblasts and osteoclasts from the perichondrium. These invading cells express receptors for angiogenic factors and MMP-9 and MT1-MMP (Vu et al., 1998; Colnot and Helms, 2001). In the ossification center, MMP-13 is induced and the hypertrophic cartilage matrix is degraded. The hypertrophic chondrocytes then die, osteoblasts replace the disappearing cartilage with trabecular bone, and bone marrow is formed. In the

perichondrium osteoblasts form compact bone around the middle portion (diaphysis) of the cartilage, so that the primary ossification center stays inside a bone tube. Secondary ossification centers form at one or both ends (epiphyses) of the bone. As a result a plate of cartilage (growth plate) is left between epiphysis and diaphysis. In the growth plate, series of chondrocyte proliferation, hypertrophy, and apoptosis lead to longitudinal growth of long bones, coordinately with growth of the epiphysis and radial growth of the diaphysis. By contrast, some craniofacial bones develop by intramembranous ossification, where osteoblasts directly deposit bone matrix to replace the cartilage matrix, and the bone remodeling is associated with bone vasculation. In both cases, dynamic bone remodeling continues through ECM deposition and resorption by the co-operative action of osteoblasts and osteoclasts.

Mice deficient of either MT1-MMP or MMP-9 show bone malformation phenotypes. In MMP-9 deficient mice, the defects in epiphyseal ossification lead to subtle shortening of the bones (Vu et al., 1998), whereas in MT1-MMP deficient mice the craniofacial, axial, and appendicular skeletons are severely affected (Holmbeck et al., 1999; Zhou et al., 2000). Severe connective tissue disorders in MT1-MMP deficient mice include dwarfism, skeletal dysplasia, osteopenia, soft tissues fibrosis, arthritis, and early death. Defective vascular invasion of cartilage leads to enlargement of hypertrophic zones of growth plates and impaired formation of ossification centers in long bones. Normal endothelial cells, osteoclasts and osteoblasts express MT1-MMP both in vitro and in vivo (Sato et al., 1997; Haas and Madri, 1999). Therefore, MT1-MMP may have important roles in the invasion of any of these cells to hypertrophic cartilage. In migrating osteoclasts, MT1-MMP is focused to the lamellipodia (Sato et al., 1997). MMP activity is required for the migration of osteoclasts in developing marrow cavity (Blavier and Delaisse, 1995), but the distinct roles of MT1-MMP and MMP-9 in this migration are not well understood.

Bone resorption and osteoclastic activity are increased in MT1-MMP deficient mice indicating that MT1-MMP is not critical for bone degradation by osteoclasts, an activity that has been primarily associated for acidic proteinases of cathepsin family (Hall and Chambers, 1996). Rather, the osteopenia correlates with the increased osteoclastic bone resorption and impaired function of osteoblasts. Chondrocyte proliferation is also reduced in the growth plates (Zhou et al., 2000). On the other hand, skin fibroblasts and stromal marrow cells from MT1-MMP deficient mice cannot degrade type I collagen matrix suggesting an indispensable role for MT1-MMP activity in stromal collagen remodeling (Holmbeck et al., 1999). The bone resorption was suggested to be a compensatory

36

response for abundant periskeletal fibrosis resulting from the inability of soft tissue cells to remodel surrounding collagenous matrix (Holmbeck et al., 1999). Interestingly, the growth of MMP-2 knockout mice is also delayed (Itoh et al. 1997). In humans, inactivating mutations of MMP-2 cause a multisendric osteolysis and arthritis syndrome with similarities to the phenotype of MT1-MMP knockout mice (Martignetti et al. 2001). Therefore, the osteolytic phenotypes could result from imbalance between the breakdown and synthesis of stromal and/or bone ECM near bone/soft tissue interfaces, which would cause impaired function of osteoblasts and osteoclasts. Finally, impaired MT1-MMP dependent removal of collagenous uncalcified cartilage primordia interfere with intramembranous bone formation and suture closure.

In MMP-9 deficient mice, the hypertrophic chondrocytes develop normally, but their apoptosis as well as epiphyseal vasculation and ossification are delayed, resulting in lengthening of the growth plate (Vu et al., 1998). Interestingly, the inhibition of VEGF signaling by injection of soluble VEGFR1 receptor (Flt-1-IgG fusion protein) in mice inhibits vascular invasion and recruitment of chondroclasts, osteoblasts, and osteoclasts into hypertrophic cartilage and results in a bone phenotype similar with the MMP-9 knockout (Engsig et al., 2000; Gerber et al., 1999). Furthermore, in a transgenic RIP Tag mice model of SV40 T antigen (Tag) induced pancreatic beta cell carcinogenesis, angiogenesis appears to depend on the mobilization of VEGF from the ECM stores by MMP-9. In this model, the “angiogenic switch” is not associated with transcriptional upregulation of VEGF, its receptor or FGF-2, whereas in RIPTag X MMP-9-deficient double transgenic mice angiogenesis and tumor growth are inhibited (Bergers et al., 2000). MMP-2 deficiency does not affect angiogenesis in this model. Therefore, MMP-9 may be important for recruitment of endothelial cells and other cell types invading to ossification centers by mobilizing VEGF from hypertrophic ECM (Sternlicht and Werb, 2001).

Cancer Cancer invasion and metastasis depend on a series of proteolytic steps. During initial invasion into the stromal tissue cancer cells traverse the basement membrane by a process depending on proteinases that can degrade and remodel basement membrane components (McCawley and Matrisian, 2001; Stetler-Stevenson and Yu, 2001). Continued cancer cell invasion is facilitated by the degradation of the surrounding stromal matrix, where tumor tissue substitutes the normal host tissue. Formation of solid tumor also requires angiogenesis to provide blood supply into the growing tumor tissue (Folkman, 1995). During metastasis a tumor cell dissociates from the primary tumor, invades the surrounding ECM, including interstitial stroma and basement membranes, enters the vascular or lymphatic space (intravasation), adheres to the endothelium at distant sites, egresses from the vasculature (extravasation) into the surrounding tissue, and proliferates to form a secondary tumor (metastasis) (Mignatti and Rifkin, 1993).

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Numerous studies have described the overexpression of various proteinases including serine, metallo-, cysteine, and aspartic proteinases by invasive and metastatic tumor cells or stromal cells (Sloane, 1990; Rochefort et al., 1990; Pyke et al. 1992, 1993; Johnsen et al. 1998; Stetler-Stevenson and Yu, 2001; Sternlicht and Werb). The candidates most often implicated in invasion and metastasis are uPA and various MMPs (Johnsen et al. 1998; Stetler-Stevenson and Yu, 2001). However, invasive growth involves concerted action of different proteinases and proteinase cascades, and the set of proteinases used may vary in cancers originating from different tissues. Accordingly, MMP and serine proteinase inhibitors negatively regulate tumor invasiveness in many in vitro and in vivo models (Mignatti et al., 1986; Khokha et al., 1989; Koop et al., 1994; Martin et al., 1996; Sternlicht and Werb, 2001;). However, TIMP-1 is often overexpressed in colon cancer, and enhanced expression of TIMP-2 in the stroma of breast carcinomas correlates with tumor recurrence (Lu et al., 1991; Visscher et al., 1994). In addition, reduction of MMP-9 in mice leads to decreased angiostatin synthesis and consequent increased tumor growth and vascularization, while high MMP-9 expression causes a reduction in tumor vascularization (Pozzi et al., 2002; Pozzi et al., 2000). These results suggest that even in cancer the extracellular proteolysis has to be tightly regulated to allow cancer and endothelial cells to grow and invade. Indeed, many of the proteolytic mechanisms involved in cancer cell invasion are apparently similar to those used during cell migration in physiological processes. On the other hand, recent studies with transgenic mice lacking or overexpressing a single MMP gene also support a role for MMPs, e.g. MMP-3 and MMP-7, already during cancer initiation (Sternlicht and Werb, 2001).

Cancer cells recruit macrophages and stromal cells surrounding the invading front of metastazing tumor to secrete proteinases like uPA and proMMP-2 (Pyke et al., 1992; Pyke et al., 1993). If the cancer cells express receptors and activators like uPAR and MT1-MMP, the proteolytic potential produced by stromal cells can be focused on the cancer cell surface. Indeed, several forms of cancer, including those of gastric, lung, breast, ovarian, colon, cervical, urothelial, thyroid, hepatocellular, bladder, and pancreatic origin, show high MT1-MMP expression, either by cancer cells themselves or by stromal cells (Nomura et al., 1995; Okada et al., 1995; Polette et al., 1996; Tokuraku et al., 1995; Yamamoto et al., 1996). MT1-MMP expression levels also correlate with MMP-2 activation and the malignancy and invasiveness of the tumor (Yamamoto et al., 1996), and MT1-MMP expression enhances metastatic activity in an experimental metastasis assay (Tsunezuka et al., 1996). In transgenic mice MT1-MMP overexpression induces mammary gland abnormalities and adenocarcinoma (Ha et al., 2001). Cancer cells from different tissues may use distinct MT-MMPs. Indeed, MT3-MMP is expressed in several cultured cancer cell lines (Takino et al., 1995), MT4-MMP is expressed in various breast carcinomas and breast cancer cell lines (Puente et al., 1996), and high levels of MT5-MMP are detectable in brain tumors (Llano et al., 1999).

Recently, new insights into the profiles of gene expression during tumor

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invasion have come from gene expression microarrays. Both MT1-MMP and MMP-2 are highly expressed in invasive pancreatic cancer tissues (Iacobuzio-Donahue et al., 2002). Further characterization of such tissues have indicated that MT1-MMP is overexpressed especially in the neoplastic epithelium, whereas MMP-2 mRNA expression is induced within the stromal response. The expression of MMP-1, -2, -9, and MT1-MMP and also of laminin-5 γ2 chain is significantly increased in aggressive compared with poorly aggressive melanoma cells (Seftor et al. 2001). MT1-MMP expression is also induced in metastatic colorectal cancer as compared to primary cancers and normal colorectal epithelium (Saha et al. 2001).

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Aims of the present study

The activities of MMPs are regulated at multiple levels including gene expression, secretion and binding to the cell surface or ECM, proteolytic activation, and inhibition by TIMPs to achieve targeted ECM remodeling during multiple physiologic processes. Dysregulated MMP activity, in contrast, is associated with various pathological conditions. The discovery of membrane-bound MMPs, which may both remodel ECM and cell surface proteins, and also participate in pericellular cascades to activate other MMPs revealed a new field for cellular regulation of MMP activity. In the current work the aim was to find answers to the question: How is MT1-MMP activity regulated during cell migration and invasion? We first characterized the regulation at the level of gene expression, and I then concentrated on characterizing posttranslational mechanisms of the regulation. The specific questions raised during the work include the following:

1) How do cell growth modulating factors regulate MT1-MMP and TIMP-2 at

the levels of mRNA and protein expression, and how do their expression correlate with proMMP-2 activation?

2) Does proteolytic processing of MT1-MMP play a role in the regulation of its activity?

3) What kind of protein-protein interactions regulate the functions and distribution of MT1-MMP, and do the MT1-MMP molecules interact with each other to form oligomeric complexes?

4) What is the role of the cytoplasmic domain in the regulation of MT1-MMP by membrane trafficking and in the targeting of MT1-MMP activity at the cell surface?

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Materials and methods

Growth factors, chemicals, and enzymes.

Phorbol 12-myristate 13-acetate (PMA), the calcium ionophore A23187 (calcimycin), ionomycin, dexamethasone, cycloheximide, actinomycin D, ConA, human recombinant EGF, thapsigargin, and human type I collagen were purchased from Sigma Chemical Co. (St. Louis, MO). Human recombinant IL-1β and TNF-α were obtained from Boehringer Mannheim GmbH (Mannheim, Germany). Human recombinant bFGF (FGF-2) was kindly provided by Dr. Olli Saksela, Helsinki, Finland. Human platelet derived TGF-β was from R&D Systems (Minneapolis, MN). MMP-2 and TIMP-2 were from Chemicon Co. (Temecula, CA). Cyclic peptide inhibitor CTTHWGFTLC, which inhibits specifically gelatinases (Koivunen et al., 1999), was obtained from Dr. E. Koivunen Helsinki, Finland. The MMP inhibitor with broad inhibitory activity, BB-3103, was from British Biotech Pharmaceuticals Ltd (Oxford, U.K.).

Cell culture and treatments

The following table indicates the cell lines used and their ATCC (American Type Culture Collection, Rockville, MD) identification numbers. All cells were grown in Eagle's minimal essential medium (MEM) containing 10% heat-inactivated fetal calf serum (GIBCO), 100 IU/ml penicillin, and 50 µg/ml streptomycin. The cultures were incubated at 37°C in a humidified 5% CO2 atmosphere until confluent. They were then rinsed twice with serum-free medium and incubated under serum-free conditions for 6 h. For analyses, cells were treated with PMA (4-40 nM), BB-3103 (1-10 µM), ionomycin (500 nM), thapsigargin (500 nM), TNF-α (20 ng/ml), IL-1β (2 U/ml), Con A (50 µg/ml), EGF (30 ng/ml), FGF-2 (30 ng/ml), TGF-β (10 ng/ml), A23187 (500 nM), cycloheximide (10 µg/ml) and dexamethasone (1 µM) under serum free conditions for 24 h or as indicated.

Name Description Used in Bowes human melanoma cells IV CCL-137 human embryonic lung fibroblasts, ATCC I, II, III CCL-127 CHO Chinese hamster ovary epithelial cells, V ATCC CCL-61 COS-7 African green monkey kidney epithelial I, V cells, SV40 transformed, ATCC CRL-1651 HEK293 Human embryonic kidney cells V HT-1080 Human fibrosarcoma cells, ATCC CCL-121 I, II, III, V MDCK Madin-Darby canine kidney epithelial cells V VA-13 SV-40 transformed human lung fibroblasts, III ATCC CCL-75.1

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Antibodies

The antibodies used in the experiments have been listed in the following table. Polyclonal antibodies to MT1-MMP were raised in rabbits and affinity purified as described in detail in the original articles (I, II). The immunogens used were: a bacterially generated recombinant fusion protein of Schistosoma japonicum glutathione S-transferase and amino acid residues 260 to 484 of MT1-MMP (Ab-1), a 26 amino acid long synthetic peptide corresponding to the C-terminal cytoplasmic domain of MT1-MMP (Ab-2) and a 24 amino acid long synthetic peptide corresponding to the amino acids 155-177 of MT1-MMP (Ab-3). Name Immunogen Use Reference Rabbit: Ab-1 GST/MT1-MMP(260-484) IB, IP, IF (I,II, IV) I fusion protein Ab-2 MT1-MMP peptide 557-582 IB, IP (I,II, III,IV) I Ab-3 MT1-MMP peptide 155-177 IB, IP, IF (III, IV, V) IV Ab815 MT1-MMP (hinge) IB, IP, IF (III) Chemicon TIMP-2 IB Chemicon GST IB, (III) Upstate Biotechnology Goat: Ab6613 GST IB, IF (III) Abcam Mouse: Ab92 GST IB, IF (III) Abcam

Northern hybridization and RNAse protection

RNA isolation and Northern blotting: RNA was purified by pelleting a guanidine thiocyanate-N-laurylsarcosyl cell lysate through a CsCl cushion. Total RNA (15 µg) was fractioned by formaldehyde-agarose gel electrophoresis and transferred to a Zeta probe membrane (Bio-Rad, Richmond, CA) and immobilized by UV crosslinking. The membranes were hybridized to MT1-MMP and TIMP-2 cDNA probes labeled with [32P]dCTP (Amersham-Pharmacia Biotech, Upsala, Sweden) using random priming kit (Amersham-Pharmacia Biotech). The signal intensities were quantitated by phosphoimager analysis (Fuji Photo Film Co., Tokyo, Japan). Ribonuclease protection analyses were carried out using Direct Protect kit (Ambion Inc., Austin, TX) and [32P]UTP (Amersham) labeled cRNA probe complementary to a 251 bp fragment of MT1-MMP cDNA (bases 852-1102).

Expression constructs

Amplification of MT1-MMP cDNA by PCR and isolation of cDNA clones has been described in detail in the original article I. An expression plasmid MTpc3SE containing bases 1-2369 of MT-MMP-1 cDNA under transcriptional control by the CMV promoter was constructed on pcDNA3 vector (Invitrogen, San Diego

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CA). All the listed deletion and mutation constructs were generated from MTpc3SE by PCR mediated mutagenesis as described in the respective articles.

Name (Tag/) MT1-MMP amino acids Reference (Used in)

MTpc3SE/wt MT1-MMP 1-582 I, IV (I, II, III, IV, V)

E240A MT1-MMP 1-582, Valtanen et al 2000,IV (IV)

inactivating E240A point mutation

∆∆∆∆577 MT1-MMP 1-577 IV, (IV)

∆∆∆∆573 MT1-MMP 1-573 IV, (IV)

∆∆∆∆567 MT1-MMP 1-567 IV, (IV)

∆∆∆∆TM MT1-MMP 1-538 IV (II, IV,)

∆∆∆∆Cyt MT1-MMP 1-564 Pei et al. 1996 (III, V)

∆∆∆∆Cat GST/MT1-MMP 296-582 III (III)

∆∆∆∆Cat/Pex GST/MT1-MMP 438-582 III (III)

TM/Cyt HA/MT1-MMP 525-582 III (III)

Transfection of cells

Cells were cultured on a six-well plate to reach 50-80 % confluence and transfected using 2 µg of plasmid DNA and 5 µl of FuGENE 6 -transfection reagent for each transfection according to the manufacturer’s instructions (Boehringer Mannheim, Mannheim, Germany). Stable transfectants were selected with 0.5-1.2 mg/ml G418 (Gibco-BRL) in complete medium. Expression of the recombinant MT1-MMPs by the transfected cell clones was analyzed by immunoblotting and Northern blotting.

SDS-PAGE and immunoblotting

SDS-PAGE of was carried out using 4-15% gradient or 7.5% and 10% standard polyacrylamide gels (Bio-Rad Co., Richmond, VA). The proteins were then transferred to nitrocellulose membranes, which were saturated with 5% milk in PBS/Triton X-100 (0.5%) and incubated with primary antibodies in 50 mM Tris-HCl buffer, pH 8.5 containing 500 mM NaCl, 0.1 % Tween 20 and 0.1 % bovine serum albumin. After several washes with the same buffer the bound antibodies were detected using biotinylated antimouse or antirabbit antibodies and peroxidase-conjugated streptavidin (Dakopatts, Copenhagen, Denmark) and enhanced chemiluminescence Western blotting detection system (Amersham).

Gelatin zymography and reverse zymography

To analyze the gelatinolytic proteins, aliquots of cell conditioned medium, cell membrane extracts or immunoprecipitated MT1-MMP immune complexes were

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analyzed by gelatin zymography essentially as described (Lohi et al. 1995). Samples were dissolved in non-reducing Laemmli sample buffer and separated by SDS-PAGE using10% polyacrylamide gels containing 1 mg/ml of gelatin. After electrophoresis the gels were washed twice with 50 mM Tris-HCl, pH 7.6, containing 5 mM CaCl2, 1 µM ZnCl2, 2.5% Triton X-100 (v/v) for 15 min to remove SDS, followed by a brief rinsing in washing buffer without Triton X-100. The gels were then incubated at 37°C for 12-24 h in 50 mM Tris-HCl buffer containing 5 mM CaCl2, 1 µM ZnCl2, 1% Triton X-100, 0.02% NaN3, pH 7.6. The gels were then stained with Coomassie Brilliant Blue R250 followed by destaining with 10% acetic acid, 5% methanol. For reverse zymography 25 ng/ml MMP-2 (Chemicon) was added to the 12% or 15% polyacrylamide gels containing 1 mg/ml gelatin. Washes and the digestion were performed as described for gelatin zymography.

Pulse-chase analysis and immunoprecipitation

For pulse-chase studies cells were incubated in methionine-free MEM followed by metabolic labeling for 10 min with 250 µCi/ml [35S]-methionine (<1000 Ci/mmol). The cells were then chased with MEM (over 500-fold excess of unlabeled methionine). At the indicated time points the cells were washed and lysed with Triton lysis buffer (50 mM Tris-HCl buffer, pH 8.0, containing 150 mM NaCl, 1% Triton X-100, 0.02% sodium azide, 10 mM EDTA, 10 µg/ml aprotinin, 1 µg/ml pepstatin A and 1 µg/ml aminoethylbenzene sulphonylfluoride; Calbiochem, San Diego, CA). For immunoprecipitation analysis, cell lysates were preabsorbed by incubation with preimmune serum and protein A-Sepharose beads at 4°C for 2-20 h. After centrifugation the supernatants were incubated with MT1-MMP antibodies (2 µg/ml) on ice for 1 h, followed by incubation with protein A-Sepharose at 4°C for 1 h. The beads were then washed three times with Triton X-100 lysis buffer and once with PBS. Bound proteins were eluted by reducing Laemmli sample buffer and incubation at 95°C for 5 min. The immune complexes were resolved by SDS-PAGE and detected by fluorography.

Purification and N-terminal sequencing of MT1-MMP

The 60-kDa form of wild-type MT1-MMP was immune affinity purified from stable MT1-MMP transfected HT-1080 cell clone using MT1-MMP antibodies as described for immunoprecipitation analyses. The 43-kDa form of soluble mutant MT1-MMP with six additional C-terminal histidines was purified from ∆TM transfected HT-1080 cells using TALON metal affinity resin (Clontech, Palo Alto, CA) according to the manufacturer’s protocol. Proteins were dissolved with Laemmli sample buffer, resolved by SDS-PAGE and electrotransferred to polyvinylidene difluoride membrane (Millipore Corporation, Bedford, MA). After staining with Coomassie blue R-250 the bands were excised from the membrane and subjected to amino acid sequencing (Ariad Pharmaceuticals, Cambridge, MA).

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Cell surface biotinylation

Cells were rinsed at 4°C with PBS and incubated in PBS containing 0.5 mg/ml of Sulfo-NHS-biotin (Pierce Chemical, Rockford, USA) on ice for 1 h. The reaction was terminated by washing with 10 mM Tris-HCl buffer (pH 7.4) containing 10 mM glycine, 0.4 M sucrose, and 1.5 mM CaCl2, and 5 mM MgCl2 for 10 min. The cells were lysed and subjected to immunoprecipitation with MT1-MMP antibodies. The immune complexes were resolved by SDS-PAGE and analyzed with peroxidase-conjugated streptavidin (DAKO A/S, Denmark) and enhanced chemiluminescence Western blotting detection system (Pierce Chemical). To analyze the processing time courses of cell surface biotinylated MT1-MMP proteins the cells were surface biotinylated as above, shifted to 37oC for increasing time periods (0 to 180 min) to allow processing, then lysed, and subjected to immunoprecipitation.

Cell surface cross-linking

For crosslinking experiments metabolically (50 µCi/ml 35S-methionine) or surface (0.2 mg/ml sulfo-NHS-biotin) labeled cells were incubated with nonpermeant thiol-cleavable crosslinker DTSSP (1 mM) for 1 h on ice, washed and lysed. Aliquots of the lysates were subjected to immunoprecipitation with Ab-2 against the cytoplasmic domain of MT1-MMP followed by SDS-PAGE under nonreducing or reducing (50-100 mM DTT) conditions and detection by fluorography or horseradish-peroxidase conjugated streptavidin. Alternatively, the cell surface proteins were crosslinked with noncleavable BS3 crosslinker (0.5 mg/ml). The cells were then washed and lysed, and aliquots of cell lysates were fractioned by SDS-PAGE under reducing conditions and analyzed by immunoblotting as indicated.

In vitro analyses of MT1-MMP cleavage

Recombinant soluble forms of wt and inactive MT1-MMP (∆TM and E240A∆TM, respectively) were expressed in baculovirus/insect cell system and purified by cation exchange and gel filtration chromatography (Valtanen et al., 2000). ∆TM and E240A∆TM (10 ng) were incubated alone or with proteinase inhibitors as indicated in 50 mM Tris-HCl buffer, pH 7.5, containing 0.15 M NaCl, 10 mM CaCl2, 0.05 % Brij 35 and 0.02 % NaN3 at 37°C for 3 hours. The reactions were terminated with reducing Laemmli sample buffer, and the MT1-MMP fragments were separated by SDS-PAGE and characterized by immunoblotting.

GST-fusion protein purification

For GST pull-down analyses the noncatalytic GST/MT1-MMP fusion proteins were captured from the cell lysates by incubation with glutathione-Sepharose for 1.5 h at 4°C and washed. The bound proteins were eluted with Laemmli sample

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buffer, fractionated by SDS-PAGE under nonreducing or reducing conditions, and analyzed by immunoblotting with anti-MT1-MMP or anti-GST antibodies.

Immunofluorescence and internalization assays

Cells were cultured on glass coverslips, washed with PBS, and fixed with 3% paraformaldehyde for 10 min. For intracellular staining the cells were permeabilized with 0.1 % Triton X-100 in PBS for 20 min. Subsequently, the coverslips were treated with 5% bovine serum albumin in PBS for 30 min. The cells were then labeled with the indicated primary antibodies followed by Rhodamine- or fluorescein isothiocyanate (FITC)-conjugated secondary antibodies (Jackson ImmunoResearch Laboratories, West Grove, PA). Next, the coverslips were washed and mounted on glass slides using Vectashield (Vector Laboratories, Burlingame, CA). The fluorescence images were obtained by using epifluorescence microscope or a Bio-Rad MRC-1024 confocal microscope. For internalization assays, the cells were cultured on glass coverslips and washed twice with cold PBS containing 1% bovine serum albumin. Cells were then incubated with anti-MT1-MMP antibodies (Ab-3) on ice for 1 h, washed and shifted to 37oC for the indicated time periods. Subsequently, the cells were fixed, permeabilized, and stained with FITC-conjugated goat anti-rabbit IgG. For co-localization experiments, cells were labeled with anti-clathrin, dynamin, or EEA1 primary antibodies followed by Rhodamine- or FITC- conjugated secondary antibodies. For the staining of migrating cells, a gel containing serum-free MEM and 0.5 % low melting point agar was poured above the cells. Chemoattractant gradient was generated to the solid agar gel by adding 1 ng FGF-2 or 5 µl fetal calf serum to one side of the coverslips (Alanko et al., 1994). After 24 h the gel was removed, and the cells were fixed and labeled with Ab-1 and FITC–conjugated anti-rabbit antibodies as described above.

In vitro invasion assay

The Boyden-chamber cell invasion assay was carried out according to manufacturer’s instructions (Becton Dickinson, Bedford, MA). Bowes control cells or stably transfected Bowes cells expressing wild-type or mutated MT1-MMP proteins (1x105 cells/ml) were suspended in MEM containing 0.1% BSA and seeded onto uncoated or Matrigel-coated Transwell filters (8 µm pore size) in invasion chambers. Proteinase inhibitors aprotinin (100 µg/ml), BB-3103 (10 µM), or CTTHWGFTLC (500 nM) were added to the suspension of control cells or cells expressing wt MT1-MMP, and the cells were seeded onto the filters after 0.5 h incubation. Fetal calf serum (5%) was used as a chemoattractant in the lower chambers. After 18 h incubation, noninvading cells were removed from the top surface of the filter, and the filters were stained with Diff-Quick (Dade AG, Dudingen, Switzerland). Cells migrated onto the lower surface of the filter were counted under a microscope at x40 or x200 magnification. Each assay was carried out in triplicate, and five fields were counted for each sample.

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Results

Differential regulation of MT1-MMP gene expression and proMMP-2 activation (I) To analyze the function of MT1-MMP and its role on proMMP-2 activation, we cloned from a human embryonic lung cDNA library a cDNA for MT1-MMP and studied MT1-MMP expression in human HT-1080 fibrosarcoma cells and CCL-137 embryonic lung fibroblasts. Northern hybridization analysis indicated that both cell lines constitutively express endogenous MT1-MMP mRNA. Treatment of both cells with PMA caused a 2-3-fold increase in mRNA levels, whereas ConA increased MT1-MMP mRNA levels about 4-fold only in CCL-137 fibroblasts, and TNF-α treatment resulted in a slight increase in MT1-MMP mRNA levels only in HT-1080 cells. EGF, bFGF, TGF-β, and IL-1β had no significant effects on MT1-MMP expression. The increase in MT1-MMP mRNA expression by PMA required protein synthesis, as shown by cycloheximide inhibition. The effect of PMA was also inhibited by treatment of the cells with dexamethasone. Analysis of MT1-MMP mRNA stability using actinomycin D treatment indicated that its half-life was very long and not affected by PMA. These data indicate that PMA affects MT1-MMP expression at the transcriptional level, since no further increase in mRNA stability could account for the observed increase of mRNA levels.

MT1-MMP overexpression induces proMMP-2 activation (Sato et al., 1994). However, only HT-1080 cells had significant proMMP-2 processing activity after treatment with PMA or ConA, while fibroblasts were virtually negative. Hence, proMMP-2 activation did not directly correlate with MT1-MMP mRNA expression. To further characterize the correlation between MT1-MMP expression and proMMP-2 activation, we transfected HT-1080 cells with MT1-MMP and selected stable transfectants producing about 10-fold excess of MT1-MMP as compared with the wild-type HT-1080 cells. Consistent with the primary report (Sato et al., 1994), these cells were able to activate proMMP-2. PMA treatment further enhanced this activation. These results indicate that whereas overexpression of MT1-MMP in some cells results in MMP-2 activation, the contribution of other factors is also important.

To examine the role of potential changes in TIMP-2 expression in the activation of proMMP-2, the RNA blots from the experiments described above were hybridized with a TIMP-2 cDNA probe. TIMP-2 mRNA was expressed at similar levels in both HT-1080 and CCL-137 cells, and none of the chemicals or growth factors tested, including PMA, ConA, the calcium ionophore A23187, TNF-α, EGF, bFGF, TGF-β, and IL-1β, caused significant changes.

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Calcium ionophores inhibit proMT1-MMP activation, binding of TIMP-2 and MMP-2, and proMMP-2 activation (I, II) We next hypothesized that posttranslational processing of MT1-MMP may regulate its cell surface activity. To study the molecular forms of MT1-MMP expressed in HT-1080 and CCL-137 cells we raised polyclonal rabbit antibodies against various intracellular and extracellular regions of MT1-MMP. By using Ab-2 against the C-terminal cytoplasmic domain we found that MT1-MMP zymogen was synthesized as a 63-kDa protein in HT-1080 and CCL-137 cells. This zymogen was rapidly N-terminally processed to an activated 60-kDa form with N-terminal Tyr112. Results of immunoblotting analysis revealed that CCL-137 cells contained mainly activated 60-kDa MT1-MMP. In these cells, PMA and ConA substantially increased the levels of activated MT1-MMP without considerable proMMP-2 activation. Therefore, neither proMT1-MMP processing nor the accumulation of active enzyme was sufficient to promote proMMP-2 activation in these fibroblasts.

Previous results of our group have indicated that the activation of proMMP-2 by HT-1080 cells treated with either PMA, TNF-α, or Con A can be prevented by simultaneous treatment of the cells with calcium ionophores or thapsigargin (Lohi and Keski-Oja, 1995). This effect is not accompanied by any significant changes in MT1-MMP mRNA levels, suggesting an inhibition of some other step in the activation process. To understand this inhibition mechanism, we studied the effects of calcium ionophores on proMMP-2 activation and MT1-MMP processing in transfected MT1-MMP overexpressing HT-1080 cells. Treatment with the calcium ionophore ionomycin also prevented proMMP-2 activation in these cells, while MT1-MMP mRNA expression remained constant. Indeed, ionomycin treatment inhibited the processing of proMT1-MMP to the activated 60-kDa form. Thus, intracellular calcium levels controlled proMMP-2 activation by modulating proMT1-MMP activation.

Inactivating autocatalytic MT1-MMP cleavage produces 43-kDa cell-surface and 20-kDa soluble fragments (I, II, IV) The enhanced MT1-MMP mRNA expression by PMA, ConA, or transfection did not correlate with the relative level of the activated 60-kDa MT1-MMP in HT-1080 cells. Instead, the proMMP-2 activation was associated with accelerated turnover of MT1-MMP through further N-terminal cleavage to a 43-kDa cell surface form. This form was inactive in gelatin zymography unlike the 60-kDa MT1-MMP. Portion of soluble C-terminally truncated MT1-MMP mutant was also processed to ~43-kDa form. PMA treatment increased the levels of this soluble form in a manner similar to that of the wild-type MT1-MMP. The cleavage that resulted in the generation of this form occurred at the Ala255–Ile256 bond in the C-terminal part of the catalytic domain as determined by N-terminal

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sequencing. Overexpression of MT1-MMP induced its cleavage to the 43-kDa cell surface form also in human Bowes melanoma cells. During this cleavage a soluble, ~20-kDa N-terminal fragment containing the catalytic center was released to the culture medium. A synthetic MMP inhibitor, but not a gelatinase specific inhibitor, prevented the cleavage suggesting that it was a result of an autocatalytic event. Another concomitant cleavage resulted in the release of an active soluble ~56-kDa form of wt MT1-MMP that was not recognized by antibodies against the cytoplasmic domain. Furthermore, purified soluble MT1-MMP was also autoproteolytically processed to the 43-kDa and 20-kDa forms in vitro. This processing was concentration dependent (data not shown). These results suggest that autocatalytic intermolecular inactivating cleavage is a putative mechanism for negative regulation of MT1-MMP activity.

Oligomerization through hemopexin and cytoplasmic domains regulates the activity and turnover of MT1-MMP (III) The formation of multimeric complexes by MT1-MMP may facilitate its autocatalytic inactivation or proMMP-2 activation on the cell surface. To test this hypothesis, we expressed various glutathione-S-transferase (GST)/MT1-MMP fusion proteins in HT-1080 cells and SV-40 transformed human lung fibroblasts and analyzed their effects on MT1-MMP activity and potential homophilic interactions. We found that crosslinked MT1-MMP was expressed on the cell surface as oligomeric 200-240-kDa complexes containing both the active 60-kDa and the autocatalytically processed 43-kDa forms. Overexpression of a GST/MT1-MMP fusion protein containing the transmembrane and cytoplasmic domains of MT1-MMP inhibited the PMA-induced autocatalytic cleavage of endogenous MT1-MMP to the 43-kDa species, but not proMMP-2 activation. On the other hand, a similar fusion protein with the hemopexin, transmembrane and cytoplasmic domains inhibited proMMP-2 activation in a dominant negative fashion. These results suggest that both the autocatalytic cleavage of MT1-MMP and proMMP-2 activation may be regulated by oligomerization through cytoplasmic and hemopexin domains. Indeed, either domain, when attached to cell membrane by a transmembrane domain, formed stable homophilic complexes. Co-purification of MT1-MMP with these fusion proteins also correlated with their cell surface co-localization.

MT1-MMP targeting through its cytoplasmic domain is critical for cell invasion through Matrigel (IV) MT1-MMP has transmembrane and cytoplasmic domains, which may target it to invasive fronts (Nakahara et al., 1997). To study the effects of the cytoplasmic domain of MT1-MMP on its own protein expression, proMMP-2 activation and

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cell invasion, we expressed MT1-MMP mutants with progressively truncated C-termini in human Bowes melanoma cells. By immunofluorescence analysis we found that wt MT1-MMP was clustered at the leading edge of cells migrating in chemotactic FGF-2 gradient. ProMMP-2 was activated in all transfected cell clones regardless of cytoplasmic deletions. However, the localization of MT1-MMP to the leading edge of migrating cells and cell invasion through matrigel were strongly enhanced only in cells expressing either wt or truncated MT1-MMP lacking 6 C-terminal amino acid residues (∆577). Truncations of 10 or 16 amino acid residues in the cytoplasmic domain (∆567 and ∆573, respectively) disturbed the clustering of MT1-MMP and gradually decreased MT1-MMP induced invasion. To find out if the increased invasion of MT1-MMP transfected cells was due to enhanced MMP-2 activity, we compared the effects of a broad spectrum MMP inhibitor BB-3103 to those of the cyclic peptide CTTHWGFTLC that inhibits MMP-2 and MMP-9, but not MT1-MMP (Koivunen et al., 1999). A serine proteinase inhibitor aprotinin was used as a control. The invasion of cells expressing wt MT1-MMP was decreased to the level of control cells by BB-3103. The CTTHWGFTLC peptide partially inhibited the invasion of MT1-MMP overexpressing cells (56 % inhibition) while aprotinin had only a moderate effect.

Endocytosis in clathrin-coated pits regulates cell surface expression of MT1-MMP (V) We found that full length MT1-MMP was localized mainly to intracellular compartments in various types of cells. ConA, which induces proMMP-2 activation in HT1080 cells, shifted the majority of endogenous MT1-MMP from intracellular compartments to the cell surface. The cytoplasmic truncation mutant MT1∆C mimicked this phenotype. In transfected cells, it induced robust proMMP-2 activation and was expressed at high levels on the cell surface, while wt MT1-MMP localized predominantly to the intracellular compartments and induced modest proMMP-2 processing to the intermediate and active forms. In addition, ConA increased proMMP-2 activation in wt MT1-MMP transfectants, whereas MT1∆C was resistant to this increase, suggesting that ConA may regulate MT1-MMP activity by affecting its cytoplasmic domain mediated trafficking. Indeed, MT1-MMP was co-localized with clathrin on the plasma membrane and Early Endosomal Antigen 1 (EEA1) in endosomes. Internalization experiments revealed that MT1-MMP was internalized rapidly in clathrin coated vesicles while MT1∆C remained on cell surface. Furthermore, co-expression of a dominant negative mutant of dynamin, K44A, resulted in elevation of MT1-MMP activity by interfering with the endocytotic process. Therefore, inhibition of internalization by three different means, i.e., ConA treatment (Luttrell et al., 1997), cytoplasmic domain deletion, and co-transfection with dominant negative mutant of dynamin K44A, all enhanced the relative cell surface levels of MT1-MMP accompanied by elevated proMMP-2 activation.

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Discussion

Cell surface association of secreted proteinases has been recognized as a key mechanism to focus degradative processes (Chen, 1992; Mignatti and Rifkin, 1993). Indeed, in tissues ECM remodeling only occurs in immediate pericellular environment. When this work was initiated the first membrane-bound MMP, MT1-MMP, had recently been discovered and found to induce cell surface proMMP-2 activation and invasive cell phenotype. It had been detected in normal tissues and in some forms of cancer. Subsequent studies soon indicated that MT1-MMP was the first member discovered belonging to the MT-MMP subfamily, but only little was known about the regulation of these unique enzymes. In this work we first addressed the regulation of MT1-MMP mRNA expression and then I concentrated on characterizing posttranslational mechanisms that regulate the function of MT1-MMP at the cell surface. The results indicate that rapid zymogen activation by removal of the prodomain (II), cell surface events like oligomerization (III), clustering (IV), and inactivation by intermolecular autocatalytic cleavage of the catalytic domain (I, II, IV), as well as endocytosis of MT1-MMP from the cell surface in clathrin coated pits (V) provide cells with efficient mechanisms to regulate pericellular proteolysis by MT1-MMP.

Correlation between MT1-MMP expression and proMMP-2 activation (I) MMP-2 was originally characterized in a malignant mouse tumor as a type IV collagen-degrading activity, whose expression in the stromal side of the invasive front correlated with metastatic potential and cell transformation (Liotta et al., 1979; Liotta et al., 1980; Liotta et al., 1981; Salo et al., 1982; Salo et al., 1983). During development and tissue remodeling it is expressed by mesenchymal cells (Reponen et al., 1992). In addition to the elevated expression levels, the activation of proMMP-2 correlates with the invasive cell phenotype (Yamamoto et al., 1996) and is likely to be a critical step, which determines the total MMP-2 activity. Impaired proMMP-2 activation in MT1-MMP deficient mice indicates the physiological role of MT1-MMP in the activation (Zhou et al., 2000).

On the cell surface, MT1-MMP can co-operate with MMP-2, which is secreted by the same cell or by adjacent cells. Mesenchymal cells express both these proteinases during development, whereas MT1-MMP may be specifically upregulated in endothelial cells and in neoplastic epithelium. Based on biochemical and cell culture studies, the proMMP2 activation by the two-step proteolytic removal of the prodomain involves the formation of cell surface complexes that contain MT1-MMP, TIMP-2, and proMMP-2 (Strongin et al., 1995; Emmert-Buck et al., 1995; Imai et al., 1996; Overall et al., 1999; Overall et al., 2000). We compared MT1-MMP and TIMP-2 expression levels to proMMP-2

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expression and activation using human CCL-137 embryonic lung fibroblasts and HT-1080 fibrosarcoma cells as cell culture models. Both cell types had similar baseline expression levels of MT1-MMP and TIMP-2 mRNAs, while the relative expression level of MMP-2 was higher in fibroblasts than in HT-1080 cells. MT1-MMP mRNA expression could be regulated by treatments with growth factors or chemicals: PMA increased this expression in both cell types, whereas TNF-α was effective only in HT-1080 cells and ConA only in fibroblasts. TIMP-2 and MMP-2 mRNA levels remained virtually constant during these treatments. However, proMMP-2 activation occurred only in PMA or ConA treated HT-1080 cells. Thus, the activation did not directly correlate with increased MT1-MMP mRNA expression even in fibrosarcoma cells suggesting that regulation by other mechanisms is also important.

Role of MT1-MMP processing/activation in proMMP-2 activation (II) In the trimolecular cell surface complexes the N-terminal inhibitory domain of TIMP-2 binds to the catalytic site of MT1-MMP (Strongin et al., 1995; Zucker et al., 1998; Fernandez-Catalan et al., 1998; Hernandez-Barrantes et al., 2000), while the C-terminal region of TIMP-2 interacts with the C-terminal domain of proMMP-2 (Overall et al., 1999). Therefore, proMT1-MMP activation by either proteolytic removal of the prodomain or conformational change to expose the catalytic site of MT1-MMP is a prerequisite for the complex formation. In the first characterization of the complexes the purified endogenous MT1-MMP had N-terminal Tyr112 (Strongin et al., 1995). Thus, the prodomain had been removed. Later, an intracellular activation mechanism by Golgi-associated proprotein convertase furin was characterized for the soluble truncated MT1-MMP (Pei and Weiss, 1996; Sato et al., 1996). We found that in COS-7 cells, which express endogenous furin, the transiently transfected full length MT1-MMP was detectable only as the 63-kDa zymogen (I). In addition, the requirement of the proteolytic removal of the N-terminal prodomain of MT1-MMP for cell surface MMP-2 activation has been questioned (Cao et al., 1996; Cao et al., 1998; Cao et al., 2000). However, fibrosarcoma cells and fibroblasts differed from COS-7 cells in that both endogenous and transfected MT1-MMP was rapidly processed to the activated 60-kDa MT1-MMP with N-terminal Tyr112 beginning right after the furin recognition sequence. This processing/activation of MT1-MMP was prevented by ionomycin. Concomitantly, the binding of TIMP-2 and MMP-2, and proMMP-2 activation were inhibited, suggesting that the MT1-MMP complex formation and subsequent proMMP-2 activation depended on MT1-MMP processing. Since these observations, substantial evidence supporting the activation of also full-length MT1-MMP during or upon transport to cell surface by furin-like proteinases has emerged (Maquoi et al., 1998; Kurschat et al., 1999; Yana and Weiss, 2000; Aznavoorian et al., 2001; Rozanov et al., 2001a). In addition, the endogenous MT1-MMP is rapidly activated and detectable

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predominantly in the activated 60-kDa form in all cultured cells that we have tested. Therefore, it is likely that most cells, which express MT1-MMP, have the capacity to constantly process it to the activated form, which can then act with TIMP-2 as a receptor for proMMP-2.

Several reports have indicated that at low concentrations TIMP-2 induces MT1-MMP mediated MMP-2 activation, whereas at high concentrations it prevents the activation (Strongin et al. 1995; Butler et al., 1998; Kinoshita et al., 1998; Hernandez-Barrantes et al., 2000). MT1-MMP is capable for the initial cleavage in MMP-2 prodomain that results in 68-kDa intermediate form of MMP-2, whereas the final activation occurs through autocatalytic MMP-2 cleavage. Because TIMP-2 inhibits the catalytic site of the “receptor” MT1-MMP, neighboring TIMP-2 free MT1-MMP is likely to do the initial cleavage in the MMP-2 prodomain. In this model excess of TIMP-2 inhibits the proteolytic activity of all MT1-MMP molecules and, hence, proMMP-2 activation. The TIMP-2 mRNA was expressed at similar levels in HT-1080 cells and fibroblasts, and was not affected by PMA or ConA treatment. However, the levels of both cell-bound TIMP-2 and proMMP-2 protein were higher in fibroblasts than in HT-1080 cells. This was reflected by accumulation of activated 60-kDa MT1-MMP more potently in PMA and ConA-treated fibroblasts than in respective HT-1080 cells. Thus, it is likely that binding of TIMP-2 and proMMP-2 to the activated MT1-MMP on the surface of fibroblasts stabilizes activated MT1-MMP and exhausts free active MT1-MMP available for proMMP-2 activation. Accordingly, other studies have confirmed that the formation of the trimolecular complexes is not sufficient for activation, rather the activation of bound proMMP-2 requires overexpression of MT1-MMP, treatment of cells with ConA, phorbol esters, or cultivation of cells in the presence of special ECM components (Overall et al., 2000). In certain conditions an alternative mechanism to activate MT1-MMP bound proMMP-2 may be through serine proteinases like thrombin and cell surface uPAR/uPA/plasmin pathway (Schwartz et al., 1998; Lafleur et al., 2001a; Lafleur et al., 2001b; Monea et al. 2002).

Autocatalytic MT1-MMP inactivation (I, II, IV) We found one direct correlation between MT1-MMP protein expression and

proMMP-2 activation in HT-1080 cells. In these cells enhanced MT1-MMP mRNA expression did not correlate with accumulation of the active enzyme, but it resulted in further N-terminal processing of the activated 60-kDa MT1-MMP to the 43-kDa form within 1-4 h after translation (I, II). During this cleavage soluble, ~20 kDa N-terminal fragment containing the catalytic site was released to the conditioned medium (IV). Neither of these fragments retained gelatinolytic activity detectable by zymography. Since our primary reports, others have also reported the existence of 40-45 kDa cellular forms of MT1-MMP, usually in association with proMMP-2 activation (Stanton et al., 1998; Ellerbroek et al., 1999; Theret et al., 1999; Annabi et al., 2001a; Annabi et al., 2001b; Gingras et

53

al., 2000; Maquoi et al., 2000; Ellerbroek et al., 2001; Rozanov et al., 2001a; Zhuge and Xu, 2001). Hernandez-Barrantez et al. (Hernandez-Barrantes et al., 2000) found that in their vaccinia virus expression system either co-expression or addition of exogenous TIMP-2 is actually required for the detection of the active MT1-MMP as well as for proMMP-2 activation. Otherwise, MT1-MMP is processed to 40-44 kDa membrane bound forms. A portion of soluble mutant form of MT1-MMP was proteolytically processed in a manner similar to that of the wild-type enzyme in HT-1080 cells (II). We determined the N-terminal sequence starting at Ile256 for this inactive 43-kDa form of soluble MT1-MMP. On the other hand, the 44-40 kDa MT1-MMP bands of wt MT1-MMP from vaccinia virus infected BS-C-1 cells starts at Gly285, and is therefore lacking the whole catalytic domain. In both cases the ~43 kDa forms represent catalytically inactive fragments. Lack of catalytic activity for the soluble 20-kDa remnant suggests that the 43-kDa form with Ile256 is the predominant form.

Stanton et al. (Stanton et al., 1998) demonstrated that at least soluble MT1-MMP, MMP-2, MMP-3 and MMP-13 can process membrane bound MT1-MMP to the inactive form in vitro. In addition, synthetic metalloproteinase inhibitors and TIMP-2 inhibit the processing (Stanton et al., 1998; Ellerbroek et al., 1999), while TIMP-1 that is a poor inhibitor of MT1-MMP (Will et al., 1996) has no effect on processing. We found that E240A point mutation in the catalytic domain of MT1-MMP prevented the inactivating cleavage in human Bowes melanoma cells (IV). These results suggest that MT1-MMP is involved in the processing, either directly or through activation of another proteinase. Bowes cells do not express detectable levels of MMP-13 (Airola et al. 1999), which has also been characterized as a potential substrate for MT1-MMP (Knäuper et al., 1996b). Activated MMP-2 and MT1-MMP itself were therefore the best candidate proteases responsible for the cleavage. However, the gelatinase specific inhibitory peptide (Koivunen et al., 1999) did not prevent MT1-MMP processing, and the processing of C-terminally truncated MT1-MMP mutants did not correlate with proMMP-2 activation, suggesting that MMP-2 was not involved in MT1-MMP inactivation (IV). In vitro experiments with recombinant soluble forms of wt and E240A MT1-MMP confirmed that the catalytically active MT1-MMP was able to cause the cleavage (IV). This cleavage was concentration dependent suggesting that it is an intermolecular event (data not shown). Based on these results we proposed that at the sites of high focal activities a putative mechanism for negative regulation of MT1-MMP activity is an autocatalytic inactivating cleavage. At the same time Hernandez-Barrantes (Hernandez-Barrantes et al., 2000) also reported that the processing occurs independently of MMP-2.

Identification of oligomeric MT1-MMP complexes (III) The primary evidence for the presence of homo-oligomeric MT1-MMP

complexes in HT-1080 cells came from crosslinking experiments of endogenous and transfected wt MT1-MMP with the identification of the cell surface ~200-

54

240-kDa complexes composed of mainly the 60-kDa and 43-kDa species. The complex sizes suggest assembly of MT1-MMP to tetramers. However, the apparent molecular weight of oligomers in gels did not necessarily increase exactly according to the molecular weights of the monomers. In addition, the complex sizes based on the migration in SDS-PAGE were determined under nonreducing conditions, where MT1-MMP monomers migrate slightly faster than under reducing conditions. The 200-240 complexes could therefore also correspond, for example, to pentamers or hexamers of MT1-MMP. Alternatively, they may contain other protein(s) that have long half-lives and are not efficiently biotinylated and therefore not detected by our metabolic labeling and cell surface biotinylation experiments. By co-purification analysis of wt and C-terminally truncated MT1-MMP forms with N-terminally truncated MT1-MMP, we demonstrated that both hemopexin and cytoplasmic domains, when attached to the cell membrane by the MT1-MMP transmembrane domain, formed stable homophilic interactions. Based on these results we proposed a model where MT1-MMP forms oligomers, apparently dimers and tetramers, by interactions through the hemopexin and cytoplasmic domains. We designated these as type I and II interactions, respectively (Fig. 3).

The formation of disulfide bonds between Cys574 residues on the cytoplasmic tails of neighboring MT1-MMP monomers may stabilize type II interactions as reported recently (Rozanov et al., 2001a). However, in our experimental conditions, without crosslinking, the cell surface biotinylated MT1-MMP proteins migrated on SDS-PAGE mainly as monomers even under nonreducing conditions, and only a minor proportion was detectable as dimers or other complexes (data not shown). The purified ∆Cat/Pex- and ∆Cat/MT1-MMP complexes migrated under nonreducing conditions mainly as dimers indicating stabilization by disulfide bonds. However, it is possible that these bonds were formed by spontaneous oxidation during the cell extraction and purification processes. Nevertheless, our results do not rule out the possibility that MT1-MMP would be regulated on the cell surface by transient intermolecular disulfide bond formation between the cytoplasmic domains.

It remains unclear whether the wt MT1-MMP molecules form oligomeric complexes on the control HT-1080 cells via both types of interactions. In contrast to ∆Cat, the ∆Cat/Pex that lacks most of the hemopexin domain did not co-localize or co-purify with endogenous MT1-MMP in these cells without PMA or ConA stimulation. Type I interactions through hemopexin domain could also be multivalent supporting complexing of more than two molecules. Therefore, it is possible that type II interactions through the cytoplasmic domain only transiently regulate the function of these complexes in stimulated cells. An associated event, such as binding or release of another protein, phosphorylation or other modification of the cytoplasmic tail may modulate type II interactions. Cytoplasmic domain interaction may then induce conformational changes in MT1-MMP, which may in turn affect its proteolytic activity. Alternatively, extracellular events such as association or dissociation of the ternary complex,

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interactions with ECM or cell surface proteins, and autocatalytic cleavage of MT1-MMP to the 43-kDa form may induce changes, which in turn affect type II interactions. In any case, these observations suggest an intriguing role for the 43-kDa form, which lacks the catalytic site. Instead of being only a side product as previously suggested (II, Overall et al., 2000), it may also regulate adjacent active MT1-MMP.

ProMMP-2 activation and autocatalytic MT1-MMP inactivation in the oligomeric complexes (II, III, IV) Two proteolytic events associated with MT1-MMP, namely the initial cleavage of proMMP-2 in MT1-MMP/TIMP-2 receptor to intermediate form, and the intermolecular autocatalytic MT1-MMP cleavage to the 43-kDa form, are presumably dependent on a close proximity of at least two MT1-MMP molecules. We found that the interactions between the hemopexin domains (type I interactions) were critical for the MT1-MMP mediated proMMP-2 activation, and those between its transmembrane/cytoplasmic domains (type II interactions) regulated the turnover of the enzyme to the 43-kDa form (III). Itoh et al. (2001) also provided evidence for homophilic MT1-MMP hemopexin domain interactions and their importance for proMMP-2 activation and cell invasion.

In the MT1-MMP oligomers, a portion of MT1-MMP molecules presumably binds TIMP-2 and proMMP-2 forming the trimolecular receptor complex (Strongin et al., 1995). Three proteolytic events may then occur in these complexes (see Fig. 3): [1] MT1-MMPs cleave the proMMP-2 molecules bound to the neighboring MT1-MMP to the 64-kDa intermediate form (Butler et al., 1998; Kinoshita et al., 1998); [2] autocatalytic processing of two 64-kDa MMP-2 molecules generate the 62-kDa form (Atkinson et al., 1995; Will et al., 1996), and; [3] MT1-MMP cleaves the other TIMP-free MT1-MMP to the 43-kDa form (I, II, IV). The effects of ∆Cat/Pex and ∆Cat on these proteolytic steps support this model (Fig. 3; III). ∆Cat/Pex, which is capable of type II interactions but not of type I interactions competitively inhibits the formation of normal oligomeric complexes and prevents the processing of MT1-MMP to the 43-kDa form. However, ∆Cat/Pex cannot prevent MT1-MMP dimerization through hemopexin domains (type I interactions) or proMMP-2 activation, although proMMP-2 activation is reduced to some extent. ∆Cat, on the other hand, prevents the formation of the normal oligomeric complexes by competing with both type I and type II interactions in a dominant negative fashion. Consequently, both MT1-MMP processing to the 43-kDa form and MMP-2 activation are completely blocked.

We were unable to identify TIMP-2 and MMP-2 from the crosslinked cell-surface complexes (III). While faint biotinylated protein bands possibly corresponding to the co-precipitated TIMP-2 and MMP-2 were detectable with long exposures of the reduced samples in the crosslinking analyses (data not shown), our immunoblotting analyses of the same immunoprecipitates were not

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sensitive enough to identify them directly. Previously, we have observed co-precipitation of MMP-2 with MT1-MMP from HT-1080 and CCL-137 cells without crosslinking as assessed by gelatin zymography. Together, these experiments suggest that only a minor proportion of MT1-MMP molecules form ternary complexes with TIMP-2 and MMP-2 in these cells or that the half-life of such complexes is relatively short.

The concentration dependence of the MT1-MMP cleavage to the 43-kDa form and its prevention by dominant negative mutants suggest that it is an intermolecular event (III, IV). This is also supported by the observation that N-terminally truncated ∆Cat mutant lacking the catalytic domain was also cleaved by wt MT1-MMP (data not shown). In the cell surface oligomers, homophilic hemopexin domain interactions appear to facilitate this autocatalytic cleavage (Fig. 3). The release of active MMP-2 to the culture medium and the accelerated turnover of MT1-MMP via the 43-kDa form in HT-1080 cells have been suggested to result from in trans cleavage of the “receptor” MT1-MMP by the neighboring free MT1-MMP (Overall et al., 2000). TIMP-2 inhibits autocatalytic MT1-MMP cleavage and the detection of 43-kDa form correlates with the down-regulation of TIMP-2 by degradation through MT1-MMP-dependent mechanism (Maquoi et al., 2000). However, whether TIMP-2-bound MT1-MMP can be processed at low TIMP-2 concentrations to the 43-kDa form by adjacent MT1-MMP, is unclear.

In the 200-240-kDa oligomers, only a minor proportion of MT1-MMP appears to be occupied by TIMP-2 (III), but the exact relationship between TIMP-2 binding and oligomerization is not known. Within the oligomer, the catalytic sites of some MT1-MMP molecules may be buried from TIMP binding, thus positioning more than one TIMP-2 free MT1-MMP molecules close to each other. TIMP-2 binding may also prevent or affect the level of MT1-MMP oligomerization. In addition, interactions with αVβ3 integrin may modulate the assembly and function of MT1-MMP complexes. Indeed, the 43-kDa form of MT1-MMP has been co-purified with αVβ3 integrin from surfaces of crosslinked MCF-7 cells, which had been co-transfected to express MT1-MMP and β3 integrin (Deryugina et al. 2001a). The intermediate form of MMP-2 can bind to αVβ3 integrin and MT1-MMP can also process αV integrin (Deryugina et al. 2001b).

Of the characterized autocatalytic cleavage sites the Gly284-Gly285 bond (Hernandez-Barrantes et al., 2000) between the catalytic domain αC helix and the flexible hinge region may be exposed for cleavage of TIMP-2 bound MT1-MMP, while the Ala255-Ile256 bond near the Met-turn in the catalytic domain is shielded by interacting TIMP-2 (Fernandez-Catalan et al., 1998; Overall et al.1999). In the vaccinia virus gene transfer system MT1-MMP rapidly undergoes processing to 40-44 kDa forms without co-expressed TIMP-2 (Hernandez-Barrantes et al., 2000). This processing is only partially inhibited by exogenously added TIMP-2. When the expression levels of transfected MT1-MMP are very high in individual cells, the normal machinery for post-translational modifications and transport of the protein may be overloaded. Consequently, most of the protein detected may

57

TIMP-2

MMP-2

MT1

II II

I

[1]

[3]

wt ∆

Cat

II II

I

∆

Cat / Pex

II II

I I

II II

I I

A B

[1]

Figure 3. Model for MT1-MMP oligomerization. A) Wild-type MT1-MMP (wt) exists mainly as oligomers, potentially tetramers bound together by two types of interactions on the HT-1080 cell surface. Homophilic type I (I on figure) interactions through the hemopexin domains are co-operative with type II (II) interactions, which occur through the cytoplasmic domains. Type II interactions may involve regulation by disulfide bond formation between Cys574 residues in the cytoplasmic domain, as suggested recently. Type II interactions may also bring together two dimers to a tetrameric conformation or stabilize dimers bound together by type I interactions. Type I interactions may also be multivalent. Some of the cell surface MT1-MMP molecules bind TIMP-2 through the catalytic center, and this TIMP-2 binds proMMP-2 forming a ternary complex. The three proteolytic events occurring in the complexes are the following: [1] MT1-MMPs cleave the proMMP-2 molecules bound to the neighboring MT1-MMP to their 64-kDa intermediate forms; two 64-kDa MMP-2 molecules cleave each other in an intermolecular autocatalytic event to the 62-kDa form, and; the TIMP-free MT1-MMP cleaves another free MT1-MMP to the 43-kDa inactive form. Alternatively, autocatalytic MMP-2 maturation may in some cellular systems also involve interactions of intermediate MMP-2 forms with avb3 integrin (Brooks et al. 1996). B) These proteolytic events are affected by the non-catalytic MT1-MMP mutants as follows: ∆Cat/Pex, which is capable of type II interactions but not of type I competes with the formation of normal oligomeric complexes and inhibits MT1-MMP processing to 43-kDa form. However, ∆Cat/Pex cannot prevent MT1-MMP dimerization through the hemopexin domains (I) or proMMP-2 activation. ∆Cat, which can form complexes through both type I and II interactions prevents the formation of dimeric and higher oligomeric complexes of wt MT1-MMP by competition in a dominant negative fashion. Therefore, MMP-2 activation is completely prevented (modified from III).

represent functionally inactive protein that accumulates and may be degraded along the secretory pathway. Furthermore, the ~20-kDa fragment exhibits no gelatinolytic activity in zymography, in contrast to the shed soluble 56 kDa MT1-MMP, suggesting that the cleavage to this form occurs within the catalytic domain. Initial cleavage in other exposed parts of the catalytic domain could also induce conformational change to expose the Ala255-Ile256 bond for cleavage as discussed (English et al., 2001a). However, at least mutations in the MT-loop region have no effect in the formation of ~43-kDa form (English et al., 2001a). In addition, the release of the single ~20-kDa soluble fragment recognized by the Ab-3 (raised against amino acids corresponding to the MT-loop region) does not

58

support this option. Simultaneous detection of two distinct autocatalytically truncated cell-surface MT1-MMP forms has also been reported suggesting that the cleavage may occur at distinct sites (Rozanov et al., 2001a).

MT-MMP targeting and trafficking during cell migration (IV, V) Interactions of MT1-MMP with cytoskeletal protein complexes or aggregation with integrins can potentially induce MT1-MMP clustering to specific sites at the cell surface. In contrast to wt MT1-MMP chimeric MT1-MMP containing the transmembrane and cytoplasmic domains of IL2 receptor α-chain does not localize to specialized cell surface protrusions called invadopodia or induce invasion of RPMI7951 melanoma cells (Nakahara et al., 1997). We found that MT1-MMP was clustered at the leading edge of human Bowes melanoma cells migrating in chemotactic FGF-2 gradient, and that the truncation of the cytoplasmic domain abolished both the clustering of the enzyme and MT1-MMP-induced cell invasion through reconstituted basement membrane (Matrigel) matrixes (IV). The results suggest that the cytoplasmic domain, and especially the amino acid residues between Thr567 and Ser577 in the middle part of MT1-MMP, were essential for the regulation of the MT1-MMP clustering and invasion promoting activity. Later studies have confirmed the cytoplasmic domain dependent mechanism to promote cell invasion through reconstituted basement membrane matrixes (Gingras et al., 2001; Rozanov et al., 2001a; Uekita et al., 2001). By contrast, direct MMP-2-independent MT1-MMP activity against fibrin and type I collagen in transfected MDCK cells is dependent on transmembrane localization, but not on cytoplasmic domain (Hiraoka et al., 1998; Hotary et al., 2000).

Membrane trafficking is likely to be one key mechanism to regulate MT1-MMP activity. The cytoplasmic domain of MT1-MMP has a C-terminal valine that may affect the activation of MT1-MMP and its transport from endoplasmic reticulum to the plasma membrane (Urena et al., 1999). However, C-terminally truncated MT1-MMP mutants (∆567, ∆573 and ∆577), all lacking the C-terminal valines, were transported to the cell surface, and we observed no intracellular accumulation of 63-kDa proenzyme or glycosylated 80-84 kDa forms, as was reported by Urena et al. (Urena et al., 1999). In fact, the cell surface levels of the C-terminally truncated mutants were comparable or higher as compared to the wt MT1-MMP. On the other hand, a variety of PDZ-domain containing proteins, which interact with receptors having C-terminal valines are involved in cytoplasmic protein complexes. These proteins may modulate various trafficking, clustering, and signaling events. Therefore, identification of specific interactions and their potential effects on MT1-MMP function in different cell types and in response to different stimuli are required to characterize the role of the C-terminal valine in MT1-MMP function.

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Interestingly, we found that endocytosis in clathrin coated pits provides cells with a mechanism to down-regulate cell surface levels of MT1-MMP. Inhibition of internalization by ConA treatment (Luttrell et al., 1997), cytoplasmic domain deletion, or co-transfection with dominant negative mutant of dynamin K44A enhanced the cell surface levels of MT1-MMP accompanied by elevated proMMP-2 activation. Therefore, it remains possible that transient increase in proteolytic activity can be achieved by temporary inhibition of MT1-MMP endocytosis, perhaps, mediated by the cellular machinery through its cytoplasmic domain. On the other hand, excessive MT1-MMP activity may be disposed of rapidly by enhancing its internalization through the same cytoplasmic domain. In essence, an “on/off” or “up/down” switch might have been built into the cytoplasmic domain to regulate MT1-MMP activity. Thus, the cytoplasmic domain appears to play a critical role in MT1-MMP mediated proteolysis by controlling not only its localization but also its magnitude. Uekita et al. (2001) subsequently reported that the dileucine and tyrosine residues in the MT1-MMP cytoplasmic domain were important for interaction with µ2 subunit of adaptor protein 2, and internalization in clathrin coated pits. Consistent with our results about the effects of cytoplasmic truncations on cell invasion (IV) they found that cytoplasmic mutations, which prevent internalization, also abolish the invasion promoting function of MT1-MMP suggesting that dynamic turnover of MT1-MMP at the edge of the cell is important for migration through matrigel. On the other hand, they also confirmed that transmembrane anchorage was sufficient for the invasion in fibrillar collagen matrix.

To bring all this to a context of a fibroblastic cell (see Fig. 4), under baseline situation MT1-MMP is slowly synthesized (I, II), it is constitutively activated during transport to cell surface (I, II), where it may interact with adjacent MT1-MMP molecules to form oligomers (III, Itoh et al., 2001), most likely tetramers, and with TIMP-2 and proMMP-2. Cytoplasmic interactions with clathrin-coated pit components direct MT1-MMP for internalization (V, Uekita et al., 2001). During overexpression in stimulated or transfected cells, the MT1-MMP synthesis, activation, and transport to cell surface are accelerated, and the extracellular TIMP-2 levels are decreased (Maquoi et al. 2000). Consequently, MT1-MMP activates MMP-2 and inactivates adjacent MT1-MMP molecules at the cell surface complexes (III). When the cells receive signals for directional migration like in chemotactic FGF-2 gradient, MT1-MMP is clustered at the cell leading edge through a mechanism that depends on cytoplasmic, and potentially extracellular interactions (IV). In addition to the potential interactions with multi-protein complexes of cytoplasmic and transmembrane proteins, interactions of the hemopexin domain with ECM molecules such as collagen may modulate MT1-MMP assembly. Recent reports have also characterized novel extracellular proteins that may positively or negatively regulate MT1-MMP mediated proMMP-2 activation (Miyamori et al., 2001; Nakada et al., 2001). At the cell surface complexes, the autocatalytically cleaved 43-kDa forms may negatively regulate adjacent active MT1-MMP molecules (III, Itoh et al., 2001), and overexpression of corresponding dominant negative mutant also inhibit cell

60

migration (Itoh et al., 2001). However, in the case of endogenous MT1-MMP or transfected wt MT1-MMP the accelerated turnover to 43-kDa form and elevated internalization correlates well with enhanced MT1-MMP activity and cell migration. Therefore, the accelerated MT1-MMP turnover and internalization during cell migration may be a mechanism for cells to provide continuous proteolysis to the sites of invasion.

Figure 4. Function of MT1-MMP in fibroblasts. MT1-MMP zymogens are constitutetively synthesized in endoplasmic reticulum. They are constantly activated during transport through Golgi networks to the cell surface (I, II), where the activated enzyme mainly exists as oligomers (III, Itoh et al., 2001), and interact with TIMP-2 and MMP-2. In the oligomers MT1-MMP activates proMMP-2 and inactivates adjacent MT1-MMP molecules (III). In migrating cells cytoplasmic and potential extracellular interactions direct MT1-MMP oligomers to the leading edge. MT1-MMP is then rapidly internalized in clathrin-coated vesicles (V, Uekita et al., 2001). Potential MT1/MMP dimerization through cytoplasmic disulfide bonds, and recycling and caveolae mediated compartmentalization events are indicated with question marks.

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Perspective

MT1-MMP and potentially also other MT-MMPs are important for normal development and cancer. They are, therefore, being studied extensively. Several recent findings have begun to reveal the complexity of the temporally and spatially coordinated regulation of proteolytic, adhesive, and signaling events occurring at both sides of the plasma membrane during morphogenic and invasive processes. In the current work we found that MT1-MMP activity may be regulated by various posttranslational mechanisms including autocatalytic inactivation, assembly to oligomers and clusters, and membrane trafficking. However, the details of these and other regulatory processes and the regulation in vivo are poorly understood. Future work to compare these processes involving type I transmembrane or GPI-anchored MT-MMPs as well as related families of membrane-anchored proteinases should shed light on the mechanistic and functional aspects of pericellular proteolysis. In addition, new data about the structural and biochemical properties of distinct MT-MMPs, and their complexes with inhibitors, substrates, and other specific binding proteins should help to understand their structure-function relationships.

Currently it is unclear, which factors determine whether MT1-MMP acts directly towards ECM components or activates MMP-2 to act during different physiological processes. The initial characterization of the MT1-MMP deficient mice suggested a significant role for the enzyme in connective tissue collagenolysis, which is further supported by substantial in vitro evidence. However, given the long and continuously growing list of potential MT1-MMP and MMP-2 substrates it is possible only to speculate of the specific proteolytic events, which modulate, for example, vascular invasion and chondrocyte proliferation. Therefore, one of the most challenging future goals in MMP research is the definition of significant in vivo substrates for these enzymes. Towards this end, the generation of transgenic animals with either tissue-specific overexpression or gene inactivation of distinct MT-MMPs and their combinations is important. Detailed characterization of these mice and comparison to other transgenic mice with related phenotypes should bring valuable new information. In addition, although I have here emphasized the regulation by posttranslational events, MT1-MMP expression is clearly upregulated in various forms of cancer and during developmental processes. In the near future, DNA and protein microarrays will provide large amounts of expression data, which should help to resolve the interplay between the regulatory cascades that determine cell phenotype.

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Acknowledgements This work was carried out at the Departments of Virology and Pathology, Haartman Institute, University of Helsinki. I wish to express my gratitude to professor Antti Vaheri, the head of the Department of Virology and to professors of the Department of Pathology, Eero Saksela and Leif C. Andersson for the excellent working facilities.

I warmly thank my supervisor professor Jorma Keski-Oja for guidance, encouragement and support during the years in his laboratory. His unfailing optimism and patience have been invaluable for completing this work.

I thank Jouko Lohi for friendship, help, and cooperation throughout the course of my thesis work. I am grateful for his guidance in the beginning of this work and for his time and support ever since. Discussions with Jouko have been a great source of inspiration and information, and his enthusiastic attitude has made the work in laboratory especially enjoyable. Jussi Taipale is thanked for the inspiring comments and ideas, which helped me to get going with the first projects. I thank Sami Starast for excellent technical assistance and Terhi Kulmala for secretarial help.

I thank Marko Hyytiäinen Katri Koli, Sara Wikström, Sara Illman, Piia Vehviläinen, Sami Starast, Heli Valtanen, Juha Saharinen, Christine Unsöld, Carita Koski, Minna Juntunen, Mariliina Liflander, Samu Heinaro, Pia Luiskari, Hannu Ojanperä, and Krista Weikkolainen for the time we have spent together, for creating a pleasant working atmosphere in the laboratory and for help and discussions.

I wish to thank all my coauthors. I owe my special thanks for Heli Valtanen and Sara Wickström for their help and contribution while I was not in the laboratory. I am grateful for Dr. Duanqing Pei for fruitful and educative collaboration and for encouragement.

I thank professor Kari Alitalo, the head of the molecular and cancer biology program, professor Tomi Mäkelä, and docent Marikki Laiho for cooperation and for creating excellent working facilities for this program in Biomedicum.

Docent Erkki Koivunen and professor Leif C. Andersson are thanked for the review of this thesis.

Very special thanks go to my parents and parents in-law; my brothers, sister in-law, brother in-law and all their families; my grandmother; and all my friends for their encouragement and help and for joyful moments outside the laboratory.

My dearest thanks belong to my husband Juha, who has stood by me every day during this work. Finishing it would have been much harder without his love, support, and understanding. The births of our daughter Janina and son Markus gave a new meaning for everything. They are my greatest source of happiness and strength.

I have been financially supported by the Academy of Finland, Helsinki University Hospital, University of Helsinki, the Finnish Cancer Organizations, Farmos Research and Science Foundation, Ella and Georg Ehrnrooth Foundation, Ida Montin Foundation, Emil Aaltonen Foundation, and Helsinki Graduate School in Biotechnology and Molecular Biology.

Helsinki, April 2002 Kaisa Lehti

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