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Vol. 17, Issue 12, 5390-5399, December 2006
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*Department of Matrix Biology, Kennedy Institute of Rheumatology Division, Faculty of Medicine, Imperial College London, London W6 8LH, United Kingdom; and Division of Cancer Cell Research, Institute of Medical Science, The University of Tokyo, 4-6-1 Shirokanedai, Minato-ku, Tokyo, 108-8639, Japan
Submitted August 23, 2006;
Revised September 22, 2006;
Accepted October 5, 2006
Monitoring Editor: Jean Schwarzbauer
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). The interstitial collagens types I, II, and III further polymerize to form fibrils that serves as stabilizing scaffolds in ECM. During tissue remodelling, collagen degradation is an essential process in that the collagen is a structural frame work of tissues and a physical barrier for migrating cells (Cawston, 1996; Sternlicht and Werb, 2001; Visse and Nagase, 2003). Because of its triple helical structure, collagens are resistant to most proteinases at neutral pH. However, collagenases belonging to the matrix metalloproteinase (MMP) family can initiate degradation of triple helical collagen cleaving a single site about 
away from the N-termini.
Among the 23 members of human MMPs, there are at least five collagenases including MMP-1 (collagenase I), MMP-8 (collagenase 2), MMP-13 (collagenase 3), MMP-2 (gelatinase A), and MMP-14 (membrane type-1 MMP, MT1-MMP; Visse and Nagase, 2003). These MMPs consist of a propeptide, a catalytic domain, a hinge (or linker) region, and a hemopexin (Hpx) domain. In addition, MMP-2 has three repeats of fibronectin type II modules inserted in the catalytic domain, and MT1-MMP has a transmembrane and cytoplasmic domains at the C-terminus. The catalytic domain of these MMPs alone can cleave peptides or noncollagenous proteins, but collagenolytic activity requires the Hpx domain (Clark and Cawston, 1989; Murphy et al., 1992; Knäuper et al., 1993; Ohuchi et al., 1997; Patterson et al., 2001). The mechanism of action as to how collagenase cleave triple helical collagens together with the Hpx is not clear because the crystal structures of collagenases (Bode et al., 1994; Borkakoti et al., 1994) have indicated that the binding site of the catalytic domain is too narrow to accommodate the triple helical structure. Recent studies of Chung et al. (2004) demonstrated that collagenases interact with collagen and locally unwind the triple helical structure before they hydrolyze the peptide bonds of the three polypeptides chains. This action occurs cooperatively with the catalytic domain and the Hpx domain together. MMP-1, -2, -8, and -13 are secreted from the cells as soluble inactive zymogens (proMMPs) which will be activated in the tissue. Thus most collagenase studies were conducted with both collagen and proteinases in solution. However, MT1-MMP is a unique collagenase in that is activated intracellularly and anchored on the cell surface.
MT1-MMP is the only membrane-anchored collagenase. It is involved in many physiological and pathological events such as wound healing (Okada et al., 1997), angiogenesis (Hiraoka et al., 1998; Zhou et al., 2000; Chun et al., 2004), bone development (Holmbeck et al., 1999), and cancer cell growth (Hotary et al., 2003), invasion and metastasis (Sato et al., 1994; Seiki, 1999). Besides cleaving collagens types I, II, and III, MT1-MMP degrades a range of extracellular macromolecules including fibronectin, laminin 1 and 5, vitronectin, fibrin, and aggrecan (d'Ortho et al., 1997; Ohuchi et al., 1997; Koshikawa et al., 2000). Cell surface proteins such as transglutaminase (Belkin et al., 2001), integrin 
V (Ratnikov et al., 2002), CD44 (Kajita et al., 2001), and syndecan 1 (Endo et al., 2003). It also activates proMMP-2 (progelatinase A; Sato et al., 1994) and proMMP-13 (procollagenase 3) on the cell surface and enhances pericellular matrix proteolysis. (Knäuper et al., 1996). Although MT1-MMP has a wide variety of functions, its collagenolytic activity appears to be one of the most important functions in vivo. Deletion of the MT1-MMP gene in mice revealed a severe phenotype in bone development that was concluded to be due to inadequate collagen turnover (Holmbeck et al., 1999). Its collagenase activity is also essential for branching morphogenesis of Madin-Darby canine kidney (MDCK) cells in a collagen gel (Kadono et al., 1998), for cancer cells to grow within a three-dimensional (3D) collagen matrix (Hotary et al., 2003), and for neovessel formation in a collagen-based matrix (Chun et al., 2004).
We and others have previously shown that MT1-MMP forms a homodimer through its Hpx domain (Itoh et al., 2001; Lehti et al., 2002) and that this is essential for the activation of proMMP-2 on the cell surface. One of the two MT1-MMP molecules binds to its endogenous inhibitor, TIMP-2 to form an MT1-MMP-TIMP-2-proMMP-2 complex and the other MT1-MMP activates the proMMP-2 (Itoh and Seiki, 2006). MT1-MMP dimers form higher order complexes, but the functional significance of the oligomer is not known (Itoh et al., 2001; Lehti et al., 2002).
In the present work, we first addressed whether the homodimer formation of MT1-MMP on the cell surface is required for expression of its collagenolytic activity. This requirement for collagen degradation on the cell surface was also tested for MMP-13, a soluble collagenase, by expressing it as a membrane-bound form. Our data shows that dimer formation of membrane-anchored collagenases is essential to cleave insoluble collagen. Such molecular assembly may help to locate collagenolytic activity at the correct site of the cell to interact with collagen fibrils during cell migration in the tissue.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Antibodies
Mouse anti-FLAG M2 antibody and alkaline phosphatase–conjugated anti-(mouse IgG) antibodies were purchased from Sigma-Aldrich (Dorset, United Kingdom). Mouse anti-(phospho-tyrosine) antibody (PY-20) was purchased from ICN Biochemicals (Costa Mesa, CA), Alexa-488– and Alexa-568–conjugated anti-mouse and anti-(rabbit IgG) antibodies were from Molecular Probes (Cambridge, United Kingdom). Anti-human MT1-MMP catalytic domain antibody was generated in rabbit as described previously (Itoh et al., 2001).
Construction of MT1-MMP Mutants
FLAG (DYKDDDDK)-tagged MT1-MMP (MT1F) was constructed as previously described (Itoh et al., 1999) and subcloned into pSG5 (Stratagene, La Jolla, CA). A FLAG tag was inserted at the end of the propeptide (between Arg111 and Tyr112), and properly activated enzyme will have the FLAG tag at its N-terminus and thus can be recognized by anti-FLAG M1 antibody (Itoh et al., 1999). MT1F-
Cat is a FLAG-tagged mutant MT1-MMP in which the region of Tyr112 to Pro312 was deleted. MT1F-CatTM is a FLAG-tagged mutant MT1-MMP in which the region of Tyr112 to Gly288 and also Ala536 to Val582 were deleted. MT1-Hpx is a Hpx domain deletion mutant of MT1-MMP without a FLAG tag where Phe336-Gly535 was deleted. MT1-Cat is the same as MT1F-Cat but does not have a FLAG tag. MT13F is a chimera mutant of human MMP-13 and MT1-F in which Tyr112 to Cys508 in MT1F was replaced with Tyr104 to Cys471 of MMP-13. MT13F-HPXMT1 is a chimera mutant in which Tyr112 to Gly284 in MT1-F was replaced with Tyr104 to Cys471 of MMP-13. These mutants were generated by the PCR extension method as described by Ho et al. (1989). Chimera mutants of the ectodomain of MT1-F, MT13-F, and MT13F-HPXMT1 and transmembrane/cytoplasmic domain of NGFR (MT1-F/NGFR, MT13F/NGFR, and MT13F-HPXMT1/NGFR, respectively) were also generated by PCR and subcloned into pSG5. The mutant is derived from sequences corresponding to Met1 to Asp515 of MT1-MMP and Glu384 to Gly790 of NGFR. The other chimera mutants were also generated at the corresponding sites. All the PCR-generated fragments were confirmed by DNA sequencing and subcloned into the pSG5 vector. MT1F-Cat and MT1F-CatTM inserts were also subcloned into pCEP4 vector (Invitrogen, Paisley, United Kingdom) to establish stable MDCK cell lines.
Western Blotting and Zymography
Western blotting was carried out as described previously (Itoh et al., 1998). Total cell lysates were prepared by addition of 1x SDS-PAGE loading buffer containing 2-mercaptethanol to cells in the culture plate and subsequent boiling for 20 min. Zymography was carried out as described previously (Itoh et al., 1999).
In Situ Collagen Degradation Assay
Six-well culture plates were coated with a thin layer of chilled neutralized Vitrogen collagen (Nutacon, Leimuiden, Netherland) at 2.7 mg/ml in 1x RPMI medium (typically 100 µl/well) and incubated for 60 min at 37°C for fiber formation, and COS7 cells (4 x 105/well) were then seeded on the film. Eighteen hours later, cells were transfected with the expression plasmids in the growth medium (10% FBS/DMEM) using FUGENE6 according to the manufacturer's instruction. The following day, culture medium was changed to serum-free DMEM, and cells were cultured for a further 3 d at 37°C. The remaining collagen film was exposed by removing cells using repeated treatment with PBS containing 0.5 mg/ml trypsin and 1 mM EDTA. The collagen film was then fixed with 3% paraformaldehyde for 20 min at room temperature (RT). Collagen was visualized by staining with Coomassie brilliant blue R250, and the images were captured by a CCD camera-equipped microscope (Nikon TE-2000; Melville, NY) with a 20x objective lens. Degraded areas were visualized as a white, unstained, noncollagen-containing zone. In this assay, stained collagen was trypsin-resistant, suggesting that they were intact fibrillar collagen.
In Situ Gelatin Degradation Assay
Four-well chamber slides (Nunc, Napierville, IL) were coated with Alexa-488–conjugated gelatin (F-gelatin) prepared with an Alexa-488–labeling kit (Molecular Probes) as described previously (Itoh et al., 2001). Transfected COS7 cells were cultured in the chamber slides for 16 h. Cells were then fixed with 3% paraformaldehyde in PBS and analyzed using Ultraview confocal microscopy (PerkinElmer Life and Analytical Sciences, Monza, Italy). The degraded area was visualized as a dark, nonfluorescent zone.
Expression and Purification of Recombinant MT1-MMP Hemopexin Domain
The cDNA fragment encoding Asn317-Gly511 was generated by PCR. The sequence encoding Met was included in the 5'end and 6xHis at 3'end, followed by a termination codon. The sequence of the PCR fragment was confirmed by DNA sequencing and subcloned into the pET3a Escherichia coli expression vector (Stratagene). BL21(DE3)pLysS cells (Stratagene) were transformed with the constructs, and protein expression was induced by 0.4 mM IPTG. Proteins were purified from inclusion bodies and folded as described previously (Itoh et al., 2001) according to the method of Huang et al. (1996). Formation of disulfide bonds was confirmed by subjecting the purified protein to SDS-PAGE under reducing and nonreducing conditions.
Indirect Immunofluorescence Staining
To localize cell surface MT1-F, MT13-F, or MT13F-HPXMT1, transfected COS7 cells cultured on four-well Permanox slide chambers (Nalge Nunc) coated with collagen film or four-well glass slide chambers (Nalge Nunc) coated with F-gelatin were fixed with 3% paraformaldehyde in PBS. After blocking with 5% goat serum and 3% bovine serum albumin in TBS for 1 h at RT, cells were incubated with an anti-FLAG M1 antibody (5 µg/ml) at RT for 2 h without permeabilizing cells. CaCl2, 1 mM, was included throughout the procedure of washing and incubation for the staining with the anti-FLAG M1 antibody. Alexa-488–conjugated goat anti-mouse IgG was used to visualize the antigen signal. Note that anti-FLAG M1 antibody can only recognize FLAG tag at the N-terminus of molecule (Itoh et al., 1999); thus only active forms of the enzyme can be stained with this procedure. The signals were analyzed by Ultraview confocal microscopy (PerkinElmer).
Immunoprecipitation
Transfected COS7 cells were lysed in RIPA buffer (1% Triton X-100, 0.1% SDS, 1% deoxycolate, 50 mM Tris-HCl, pH 8.0, 150 mM NaCl, 0.02% NaN3) containing protease inhibitor cocktail (Sigma) and 10 µM GM6001. Three 35-mm dishes per constructs were used. FLAG-tagged enzymes were immunoprecipitated from the lysate using anti-FLAG M2-conjugated agarose beads (Sigma, 100 µl, 50% gel suspension) by reacting at 4°C for 2 h with gentle rotation. Enzymes bound to the beads were eluted by FLAG peptide (Sigma) in TBS (200 µg/ml, 100 µl) without any inhibitors.
3D Culture of MDCK Cells for Branching Tube Formation
MDCK cells were transfected with pCEP4 constructs, and transfectants were selected by treatment of 800 µg/ml hygromycin B (PAA Laboratories, Karlsruhe, Germany). A population of hygromycin-resistant cells were trypsinized and suspended in the neutralized type-I collagen (2 mg/ml, Cellmatrix type 1-A, Nitta Gelatin, Osaka, Japan) at 2 x 104/ml. Cells were cultured in the presence or absence of 50 ng/ml hepatocyte growth factor (HGF, PeproTech EC, London, United Kingdom). Bright-field images of the cells were taken by CCD-equipped microscope using 10x objective lens at the 7th day.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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When cultured on a collagen film, cells expressing membrane-anchored MT1-MMP (Figure 1A) degraded solid-phase collagen, producing visible holes in the film (Figure 1B, top panel). The COS7 cells used in this study do not produce proMMP-2 or proMMP-13 that degrade type I collagen when activated by MT1-MMP (data not shown). Thus, this collagenolytic activity is most likely due to MT1-MMP. This was further confirmed by transfection dependency, and inhibition by TIMP-2 (0.5 µM), but not by TIMP-1 (0.5 µM). These cells also degraded fluorescence-labeled gelatin (F-gelatin, Figure 1B, bottom panel) that was also inhibited by TIMP-2 but not by TIMP-1. It was therefore concluded that collagenolytic activity and gelatinolytic detected in this system is a direct action of MT1-MMP.
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), which is required for proMMP-2 activation. We therefore examined whether the Hpx-dependent dimerization of MT1-MMP is required for the collagenolytic activity. This possibility was tested by coexpressing MT1-MMP and the catalytic domain-deletion mutant of MT1-MMP (MT1F-Cat) in COS 7 cells (Figure 2A), which express the Hpx domain on the cell surface. As shown previously (Itoh et al., 2001) overexpression of MT1F-Cat on the cell surface effectively inhibited homodimerization of MT1-MMP, as evidenced from inhibition of activation of exogenously added proMMP-2 (Figure 2B; also see Figure 3C below). Under these conditions, the collagenolytic activity of MT1-MMP was also inhibited in MT1F-Cat expression–dependent manner (Figure 2C), whereas MT1F-Cat did not inhibit general catalytic activity of MT1-MMP or cell surface expression of the full-length MT1-MMP because these cells degraded fluorescence-labeled gelatin (F-gelatin) as effectively as cells expressing the full-length enzyme alone. These were unexpected results because collagenases are thought to act on collagen as a monomer. To further examine if the inhibitory activity of MT1F-Cat was truly the Hpx domain–dependent phenomenon, the effect of a soluble Hpx domain construct MT1-MMP (MT1F-CatTM) was tested (Figure 3A). As shown in Figure 3B, the expression of MT1F-CatTM also inhibited cell surface collagenolysis of MT1-MMP in a dose-dependent manner, although with a lesser efficacy (see Figure 2C for a comparison). As we described previously (Itoh et al., 2001), a chimera consisting of the MT1-MMP ectodomain and the transmembrane/cytoplasmic domain of the nerve growth factor receptor (MT1F/NGFR) allowed us to monitor ectodomain-driven homodimer formation of MT1-MMP by analyzing autophosphorylation at tyrosine residues in the NGFR cytoplasmic domain. As shown in Figure 3C, MT1F/NGFR expressed in COS7 cells was strongly phosphorylated, indicating that the ectodomains of MT1-MMP do form a dimer (lane 1). When MT1F-CatTM or MT1F-Cat was coexpressed, the level of phosphorylation was decreased in an expression level–dependent manner (lanes 2–7). Calculating relative intensity of the phosphotyrosine (PY) bands by calibrating with relative band intensity of the anti-FLAG bands, MT1F-CatTM reduced PY signal by 46% and MT1F-Cat by 93% at the highest expression (lanes 4 and 7). These results indicate that both constructs prevent dimerization of MT1-MMP.
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have reported that MT1-MMP binds to type I collagen through the Hpx domain for collagenolysis and that the soluble Hpx domain inhibits collagenolytic activity of MT1-MMP by competing for its binding to collagen. We therefore considered whether inhibition of collagenolysis by the MT1F-CatTM can be due to its direct binding to collagen. To test this, COS7 cells expressing MT1F-CatTM mutant were cultured either on plastic or on a collagen film, and the level of the Hpx domain released into the culture medium from these cells was compared. If soluble Hpx domain has significant affinity to collagen to be able to inhibit MT1-MMP-collagen interaction, MT1F-CatTM should be deposited to collagen matrix elsewhere on the bottom of culture, which would result in decreased levels in the culture medium when cells are on collagen matrix. As shown in Figure 4A, similar levels of MT1F-CatTM were found in the culture medium from cells cultured on plastic and collagen I (lanes 2 and 6, FLAG-Sup), suggesting that the soluble Hpx domain does not bind to the collagen film under these conditions. Coexpression of wild-type MT1-MMP (without a FLAG-tag) decreased the level of MT1F-CatTM secreted into the culture medium and increased the level in the cell lysates (Figure 4A, lanes 3 and 7, FLAG-Sup and Cell). To further test whether there is any binding of the Hpx domain to the collagen matrix, MT1F-CatTM was immunolocalized on cells cultured on a type I collagen gel without permeabilization. As shown in Figure 4B, a low level of the signal was found on the cell surface transfected with MT1F-CatTM, but no signal was detected on the collagen matrix. When wild-type MT1-MMP was coexpressed with MT1F-CatTM, the level of cell surface-associated soluble Hpx domain (MT1F-CatTM) was significantly increased, but no signal was detected on collagen film (Figure 4B). A similar effect was also observed when the catalytic domain deletion mutant without a FLAG-tag (MT1-Cat) was coexpressed, but not with the Hpx domain deletion mutant (MT1-Hpx, Figure 4B), suggesting that soluble Hpx domain was bound to cell surface when the membrane-bound MT1-MMP or Hpx domain (MT1-Cat) is present, by interacting with its Hpx domain. These results indicate that soluble Hpx domain inhibits collagenolysis by preventing dimerization of MT1-MMPs through their Hpx domains on the cell surface.
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). Under these conditions, rHpx failed to inhibit collagenolytic activity of MT1-MMP on the cell surface (Figure 5C). Therefore, to inhibit dimerization and collagenolysis the Hpx domain needs to be coexpressed with the full-length enzyme in the cell where hetero complex of MT1-MMP and the Hpx domain can form during secretion before they form homo-dimers.
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Cat, the signal of phosphotyrosine was reduced in a manner dependent on the level of MT1F-Cat expression (Figure 7C). Furthermore, collagen degradation by both MT1-F and MT13F-HpxMT1 were effectively inhibited by the coexpression of MT1F-Cat and MT1F-CatTM (Figure 7D), suggesting that dimerization through the Hpx domain of MT1-MMP is essential for MT13-HpxMT1 to degrade collagen on the cell surface.
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Cat and MT1F-CatTM in Tube Formation of MDCK Cells in 3D Collagen Gels; Hotary et al., 2000). MDCK cells stably expressing MT1F-Cat or MT1F-CatTM (see Figure 8A) were tested for the ability to form branching tube structures. As shown in Figure 8B, mock cells form tube structures upon stimulation with hepatocyte growth factor (HGF). Formation of such structures was inhibited by addition of TIMP-2 (0.5 µM) or GM6001 (10 µM), but not with TIMP-1 (0.5 µM) as reported previously (data not shown), confirming that the process is MT1-MMP-dependent. When MT1F-Cat expressing cells were cultured in the presence of HGF, none of the cells were able to extend structures into collagen (Figure 8B, MT1F-Cat, +HGF). The effect of MT1F-CatTM expression was not as strong as MT1F-Cat: some populations of the cells were completely blocked for tube formation, but some were still able to scatter around, although they did not form tube structures (Figure 8B, MT1F-CatTM, +HGF). This may be due to the different expression level of each population of the cells and corresponds to the level of inhibition of collagenolytic activity on the cell surface (Figure 3). These results indicate that both MT1F-Cat and MT1F-CatTM inhibit collagenolytic activity of endogenous MT1-MMP in MDCK cells and that dimer formation of MT1-MMP is crucial for the degradation of collagen gels during tube morphogenesis.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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has demonstrated that unwinding of triple-helical collagen occurs locally around the cleavage site. Thus the interaction of the collagen molecule and collagenase for unwinding is expected to occur near or at the cleavage site. In contrast to the collagenolysis in solution, degradation of solid phase collagen on the cell surface by membrane-anchored MT1-MMP is spatially and temporally restricted because the enzyme cannot diffuse into the substrate by itself, and the half-life of the enzyme is <30 min on the cell surface because it is constantly internalized through clathrin- and caveolae-mediated pathways (Jiang et al., 2001; Uekita et al., 2001; Remacle et al., 2003; Anilkumar et al., 2005). Therefore, cell surface collagenolysis can be inefficient compared with that in solution. Here we have shown that homodimerization of MT1-MMP is one of the mechanisms that facilitates collagen degradation on the cell surface.
When dimerization of MT1-MMP was inhibited by coexpression of MT1F-Cat, cell surface–associated collagen degradation was effectively inhibited. Expression of MT1F-Cat does not inhibit its catalytic activity or cell surface expression of the full-length enzyme as observed with F-gelatin degradation (Figure 2). To inhibit dimer formation and collagenolytic activity of MT1-MMP on the cell surface, the Hpx domain does not have to be expressed as a membrane-anchored form (MT1F-Cat), because the soluble Hpx domain (MT1F-CatTM) could also inhibit it, albeit less efficiently. In the latter case the MT1F-CatTM needs to be coexpressed with full-length MT1-MMP. Addition of purified recombinant Hpx domain to the cells (Figure 5) or coculturing cells expressing MT1F-CatTM with those expressing MT1F does not inhibit collagen degradation (data not shown). This is most likely because the Hpx domains form tight homodimers during synthesis or in the secretory pathway, and it is difficult to exchange the dimer partner with exogenously added Hpx domain, which is also in dimer form (Itoh et al., 2001). It has recently been proposed that the Hpx domain of MT1-MMP binds to collagen I and that soluble Hpx domain can inhibit collagen degradation by the soluble form of recombinant MT1-MMP by competing for the binding to collagen (Tam et al., 2002). However, in cell-based experiments, we were unable to confirm binding of Hpx domain to collagen I fibrils. The soluble Hpx domain (MT1F-CatTM) was rather associated with the cell surface under the condition where MT1F-CatTM inhibits collagenolytic activity of the full-length enzyme. This discrepancy may be due to the expression system of the recombinant Hpx domain and their folding, but we concluded that the Hpx domain of MT1-MMP does not have significant affinity to collagen I fibrils at least under the conditions where MT1F-CatTM inhibits collagenolytic activity of full-length enzyme on the cell surface.
The exact reason why MT1-MMP needs to form a dimer to degrade collagen on the cell surface is not clear at present, but one possible explanation may be that the freedom of the ectodomains of the enzyme (catalytic domain, linker-1, and Hpx domain) on the cell surface is too restricted to carry out "unwinding" and "cutting" of triple helical collagen and requires collaboration of at least two molecules to cleave a single collagen molecule. Another possibility may be that dimeric, but not monomeric, form of MT1-MMP interacts with other cell surface molecules, which may help to present MT1-MMP to fibrillar collagen on the cell surface. Soluble enzyme can access the collagen cleavage site by diffusion, thus the orientation of the enzyme relative to the collagen molecule can be readily arranged. For immobilized fibrillar collagen to be cleaved by a membrane-anchored collagenase, correct positioning of the enzyme in relation to the cleavage site within long filamentous molecules is limited. Dimerization of the enzyme may therefore enhance the chances for the collagenase to act on collagen fibrils. Indeed when MMP-13 is expressed as a transmembrane enzyme, it failed to cleave solid-phase collagen on the cell surface. The cell surface MMP-13 fused with the Hpx domain of MT1-MMP regains the ability to cleave it. MT1-MMP forms not only dimers, but also oligomeric complexes (Itoh et al., 2001; Lehti et al., 2002). Oligomerization occurs in a concentration-dependent manner at least in vitro (Itoh et al., 2001). At the present time, it is not known if a dimeric form is sufficient to degrade collagen or whether a higher order of complexes is required. It is quite possible that the collagenolytic efficiency would be increased by clusterization of dimeric MT1-MMP on the cell surface.
Collagen-degrading activity is considered to be one of the most important biological activities of MT1-MMP during development (Holmbeck et al., 1999, 2004). MT1-MMP null mice showed dwarfism, osteopenia, and development of spontaneous arthritis and progressive fibrosis (Holmbeck et al., 1999). These phenotypes are considered to be a result of inadequate collagen turnover. The collagenolytic activity of MT1-MMP is also important in tumor growth (Hotary et al., 2003), neovessel formation (Chun et al., 2004), and cancer invasion into a collagen-based matrix (Sabeh et al., 2004). When cells are migrating through collagen, the barrier collagen must be removed by positioning the dimeric MT1-MMP at the leading edge. Because Rac1 small GTPases can regulate the dimerization (Itoh et al., 2001), it is most likely that localization and dimerization of MT1-MMP are regulated according to reorganization of the cytoskeleton. As MT1-MMP is reportedly colocalized with 
1 integrin (Ellerbroek et al., 2001; Wolf et al., 2003), there may be a regulatory link between integrin and MT1-MMP in order to sense a barrier matrix, to position the enzyme at the leading edge, and to facilitate the formation of a functional dimer.
MT1-MMP is essential during branching morphogenesis of MDCK cells in a 3D-collagen gel (Kadono et al., 1998). Expression of either MT1F-Cat or MT1F-CatTM inhibited the tube formation by these cells, but MT1F-CatTM exhibited a weaker inhibitory phenotype than that of MT1F-Cat. The difference is most likely attributed to differences in their membrane anchoring because the membrane-anchored MT1F-Cat is likely to have a tighter interaction with wild-type enzyme. Nevertheless, our data suggest that dimerization is also required for endogenously expressed MT1-MMP. Recently we have also reported that expression of MT1F-Cat effectively inhibited gastric cancer progression (Nonaka et al., 2005). It is not known whether this inhibitory activity was attributed to inhibition of collagenolysis, but it suggests that inhibition of dimerization of MT1-MMP and/or protein interaction through the Hpx domain is an effective means of inhibiting the biological function of MT1-MMP. Human MT1F-Cat can inhibit zebra fish MT1-MMP (zMT1-MMP and zMT1-MMP) activities in vitro and in vivo (Bai et al., 2005). These studies suggest that MT1-MMP derived from human and zebra fish can form a complex through their Hpx domains and also that dimerization may be an evolutionally conserved regulatory mechanism for MT1-MMP at least from zebra fish to human.
Soluble collagenases and membrane-tethered collagenase (MT1-MMP) are utilized dependent on the situations in vivo. For collagenolysis in a broader area of tissues, soluble collagenases may function more readily. For cells to migrate in collagen-enriched tissues, it is more advantageous to utilize membrane-anchored MT1-MMP. Our study emphasizes that these collagenolytic processes are regulated differently. Soluble collagenases can be regulated by expression, activation of zymogens, and inhibition by endogenous inhibitors. In the case of membrane-anchored MT1-MMP, its expression and activation produce biochemically active MT1-MMP, but they are not sufficient to make the enzyme functionally active to degrade collagen and to participate cellular migration in the tissue. It further requires dimerization, localization to specific membrane structures, and association and coordination with cell adhesion molecules and other cellular signaling molecules. Because cell migration is a dynamic process that requires orchestration of many different molecules, MT1-MMP, as one of the cell migration machineries, is perhaps required to be regulated in a such multidimensional manner.
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ACKNOWLEDGMENTS |
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Footnotes |
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Address correspondence to: Yoshifumi Itoh (y.itoh{at}imperial.ac.uk)
Abbreviations used: Hpx, hemopexin domain; MDCK, Madin-Darby canine kidney epithelial cells; MMP, matrix metalloproteinase; MT1-MMP, membrane-type 1 matrix metalloproteinase; NGFR, nerve growth factor receptor; TIMP, tissue inhibitor of metalloproteinase; PY, phosphotyrosine.
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REFERENCES |
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