Article |
Address correspondence to Stephen L. Nishimura, Bldg. 3, Rm. 207, San Francisco General Hospital, 1001 Potrero Ave., San Francisco, CA 94110. Tel.: (415) 206-5906. Fax: (415) 206-3765. E-mail: cdog{at}itsa.ucsf.edu
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Abstract |
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Key Words: integrins; transforming growth factor ß; metalloprotease; cell cycle; homeostasis
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Introduction |
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TGF-ß1 is normally maintained in a latent or inactive state by the noncovalent association of the bioactive peptide of TGF-ß1 with its NH2-terminal propeptide, latency-associated peptide (LAP)*-ß1 (Munger et al., 1997). Therefore, normal TGF-ß function is thought to be largely controlled by its activation from the latent state, a process that is not understood completely (Munger et al., 1999). However, recent evidence suggests that cell surface molecules or secreted extracellular molecules can activate TGF-ß. Specifically, the integrin vß6 and the secreted extracellular matrix molecule thrombospondin (TSP)-1 have been implicated in activation of TGF-ß1 through nonproteolytic mechanisms (Crawford et al., 1998; Munger et al., 1999). In addition, plasmin or the cell surface localization of matrix metalloprotease MMP-9 by CD44 has been proposed to lead to activation of TGF-ß through proteolytic degradation of LAP-ß1 and LAP-ß2, respectively (Lyons et al., 1990; Yu and Stamenkovic, 2000). Although these mechanisms may be important to activation of TGF-ß, particularly in response to injury (Jirtle et al., 1991; Munger et al., 1999; Murphy-Ullrich and Poczatek, 2000) or during neoplastic progression (Yu and Stamenkovic, 2000), they individually do not explain the activation of TGF-ß1 in normal tissues. Indeed, mice deficient in TSP-1 (Crawford et al., 1998), plasminogen (Bugge et al., 1995), CD44 (Protin et al., 1999), or
vß6 (Munger et al., 1999) are all born viable and are able to reproduce, in marked contrast to the uniform lethality of TGF-ß1null mice (Shull et al., 1992).
The propeptide of TGF-ß1, LAP-ß1, contains an RGD motif that is recognized by a subset of integrins sharing in common the v integrin subunit (Munger et al., 1998). Thus, three of the five
v integrins,
vß1,
vß5, and
vß6, have been shown to bind to LAP-ß1, and of these only
vß6 can mediate TGF-ß activation (Munger et al., 1998, 1999). Recently, evidence suggests that
vß6-mediated activation of TGF-ß1 plays an important role in response to injury (Munger et al., 1999). Of the two remaining
v integrins,
vß3 does not bind to LAP-ß1 or mediate activation of TGF-ß1 (Munger et al., 1998), and
vß8 has not been investigated since, until recently, a system has not been available to study its function (Cambier et al., 2000). The
vß8 integrin is of particular interest, since it has been identified recently as an epithelial growth inhibitory molecule (Cambier et al., 2000).
vß8 is expressed in the normal human airway epithelium but is lost in its malignant counterpart, suggesting a role in epithelial homeostasis (Cambier et al., 2000). Furthermore, heterologous expression of
vß8 inhibits lung carcinoma cell growth both in vivo and in vitro (Cambier et al., 2000). Since
vß8 and TGF-ß1 are coexpressed in normal tissues, such as the human airway (Crawford et al., 1998; Cambier et al., 2000), we considered the possibility that
vß8 may participate in the maintenance of airway homeostasis through activation of TGF-ß.
In this article, we demonstrate a novel mechanism of cell growth regulation mediated by activation of TGF-ß1 via the integrin vß8. We show that
vß8 can bind LAP-ß1 and that the consequence of this interaction is activation of TGF-ß1. This mechanism of activation of TGF-ß1 differs from other reported mechanisms because it is regulated through the coordinated interactions of integrins, TGF-ß, and MMPs on the cell surface. Furthermore, we show that when lung cancer cells are reconstituted with
vß8, which is normally present on the epithelial cells from which they are derived, growth is now inhibited by TGF-ß1. These data provide novel insights into the mechanisms underlying cellular homeostasis.
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Results |
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The integrin vß8 mediates activation of SLC
To determine the functional consequence of LAPvß8 interactions, we assessed the ability of
vß8 to activate the endogenous SLC present in coculture systems. These systems consisted of ß8-expressing or mock-transduced cells cocultured with reporter cell lines (TMLC [Abe et al., 1994] or HepG2-[SBE]4-Lux [Jonk et al., 1998]) responsive to active TGF-ß. We found that the TMLC reporter cell system was a more specific bioassay system for TGF-ß activity than the HepG2- (SBE)4-Lux system and was therefore used for most of these studies. The TMLC system consists of mink lung epithelial cells stably transfected with a TGF-ß responsive fragment of the plasminogen activator inhibitor-1 promoter driving the luciferase gene (Abe et al., 1994). TMLC cells are highly responsive to TGF-ß and produce a very low background of TGF-ß activation. TMLC cells can thus be used in coculture with other cell lines or cell-free fractions to test for the presence of active TGF-ß using luminescence as a readout.
In the HT1080, SW480, and H647 cell lines, heterologous expression of ß8 had either no effect or a slight effect on the cell surface expression of the other integrin ß subunits known to interact with the RGD motif (Table I). The only significant differences were a reduction of surface expression of the ß5 subunit in ß8-transduced compared with mock-transduced HT1080 and SW480 cells. It is possible that these slight reductions in surface expression of vß5- on ß8-expressing HT1080 and SW480 cells could potentially reduce the magnitude of the ß8 effect on adhesion to LAP-ß1 or influence the activation of TGF-ß. However, this is unlikely, since the
vß5 integrin binds very weakly and does not mediate adhesion to SLC (Fig. 1 f, mock).
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Evidence that the ß8-cytoplasmic domain is not required for vß8-mediated activation of TGF-ß
The mechanism of integrin vß6-mediated activation of TGF-ß is likely to depend on the transduction of mechanical forces to induce conformational changes of SLC (Munger et al., 1999). Thus,
vß6-mediated activation of TGF-ß is critically dependent on specific sequences within the ß6 cytoplasmic domain (Munger et al., 1999). However, the ß8 cytoplasmic domain has no similarity with the cytoplasmic domain of ß6 or any other integrin ß subunit (Moyle et al., 1991). We have shown previously that the ß8 cytoplasmic domain is incapable of linking to the cytoskeleton to stabilize cell adhesion (Nishimura et al., 1994; Cambier et al., 2000). Therefore, we sought to determine whether the ß8 cytoplasmic domain would influence interactions with LAP-ß1. We expressed and tested a series of ß8 cytoplasmic deletion mutants (Fig. 3, ac) for their ability to mediate adhesion to LAP-ß1 (Fig. 3 d) and to activate TGF-ß (Fig. 3, e and f). The complete (TM) cytoplasmic deletion mutant was expressed at sixfold lower surface levels than the partial (759) cytoplasmic deletion mutant or the wild-type (FL) subunit (Fig. 3 c). Low levels of surface expression of the TM mutant could be due to a decreased ability to associate with the
v subunit, alterations in intracellular transport, or increased degradation. SW480 cells expressing sixfold lower surface levels of the TM mutant compared with SW480 cells expressing the 759 mutant or the FL subunit adhered only slightly less well to LAP-ß1 (TM adhesion saturation reached at 5 compared with 2.5 µg/ml coating concentration for 759 and FL) (Fig. 3 d). Unlike ß6-transduced SW480 cells (Munger et al., 1999), SW480 cells transduced with full-length or mutant forms of ß8 failed to spread appreciably on LAP-ß1 (unpublished data).
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Together these findings suggest that the cytoplasmic domain of ß8 is not required for either adhesion to LAP-ß1 or activation of TGF-ß. Thus, it is likely that the mechanism of vß8-mediated activation of TGF-ß1 is distinct from the mechanosignal transduction mechanism described for
vß6 (Munger et al., 1999).
vß8-mediated activation of TGF-ß requires localization to the cell surface and metalloprotease activity
The fact that the cytoplasmic domain of ß8 is not required for activation of TGF-ß1 suggests that vß8-mediated activation of TGF-ß might be regulated extracellularly either in the extracellular space or on the cell surface. To test these possibilities, we first tested the ability of soluble secreted
vß8 to activate TGF-ß. We found no evidence, using a variety of receptor preparations (supernatant containing secreted
vß8 or lectin- or antibody-purified receptor), that soluble secreted
vß8 could activate TGF-ß (with supernatant containing secreted
vß8 or media control; relative luciferase units:
vß8, 7.8 ± 0.1; media control, 11.8 ± 1.2, p > 0.05). This suggests that cell surface localization of
vß8 is required for TGF-ß activation.
Because proteolytic cleavage is a common mechanism of regulating cytokine activity, we tested the ability of protease inhibitors to block vß8-mediated activation of TGF-ß. GM6001, a member of the hydroxamate class of protease inhibitors specific to metalloproteases, but not a control peptide lacking the metal-binding hydroxamate modification (C1006), significantly blocked
vß8-mediated TGF-ß activation in SW480 (Fig. 4 a) and HT1080 cells (59.0 ± 10.0% inhibition using 5 µM GM6001; 0.0 ± 0.9% using 5 µM C1006, p < 0.01). This inhibition was specific to
vß8 because
vß6-mediated activation of TGF-ß was not inhibited by GM6001 (5 µM) in SW480 cells (Fig. 4 a) or HT1080 cells (1.0 ± 0.1% inhibition, p > 0.05). Finally, other peptide and chemical inhibitors of aspartyl (pepstatin A), serine (PMSF, CK-23, aprotinin, and leupeptin), or cysteine (leupeptin and E64) proteases had no effect on
vß8-mediated activation of TGF-ß1 when used at the maximal nontoxic doses (Fig. 4 b). Together these data suggest a novel mechanism of
vß8-mediated activation of TGF-ß1 requiring both the cell surface and metalloprotease activity.
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Overexpression of MT1-MMP is sufficient to cleave and inactivate LAP-ß1
To determine if proteolysis of LAP-ß1 could be a mechanism of vß8-mediated activation of TGF-ß, we incubated recombinant LAP-ß1 with ß8-overexpressing H1264 cells transduced with either MT1-MMP,
MT1-MMP, or vector alone. After incubation of LAP-ß1 with mock or
MT1-MMPexpressing H1264 cells, LAP-ß1 remained intact (32 kD). In contrast, we found that almost all of the LAP-ß1 incubated with MT1-MMPexpressing H1264 cells was smaller (2628 kD) than intact LAP-ß1, suggesting proteolytic cleavage (Fig. 9 a, lane 4). LAP-ß1 cleavage was dependent on the metalloprotease activity of MT1-MMP, since the metalloprotease inhibitor GM6001 but not a control peptide C1006 completely blocked cleavage (Fig. 9 a, lanes 5 and 6). To determine if LAP-ß1 cleavage was also dependent on
vß8, we developed a peptide based on the human LAP-ß1 sequence, which was a relatively specific inhibitor of
vß8LAP-ß1 interactions. In the H1264 system, the GRRGDLATIH peptide completely blocked
vß8LAP-ß1 interactions while having a minimal effect on the binding of other RGD-dependent integrins to VN or fibronectin (Fig. 9 b). Using these peptide inhibitors, we determined that LAP-ß1 cleavage was also dependent on
vß8, since GRRGDLATIH but not the RGE mutant peptide inhibited LAP-ß1 cleavage (Fig. 9 c). Thus, in the H1264 cell system,
vß8 and MT1-MMP together are required for LAP-ß1 cleavage. Finally,
vß8-, MT1-MMPdependent cleavage of LAP-ß1 is functionally relevant, since LAP-ß1, after cleavage, loses the ability to inhibit the function of the recombinant mature TGF-ß1 peptide (Fig. 9 d).
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Discussion |
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Two molecular mechanisms have been proposed that may lead to the activation of TGF-ß1: conformational change leading to activation of the SLC complex (Crawford et al., 1998; Munger et al., 1999) or proteolysis of LAP-ß1 leading to the release of active TGF-ß1 (Munger et al., 1997; Yu and Stamenkovic, 2000). Our data demonstrate that a mechanism of conformational change leading to activation of TGF-ß, as proposed for the vß6 integrin (Munger et al., 1999) or TSP-1 (Crawford et al., 1998), is not responsible for
vß8-mediated activation of TGF-ß1. Specifically,
vß8-mediated activation of SLC does not require the ß8 cytoplasmic domain in contrast to the mechanism of
vß6-mediated activation of TGF-ß, which requires the ß6-cytoplasmic domain (Munger et al., 1999). Furthermore,
vß8 is unlikely to bind directly or indirectly to LAP-ß1 through a TSP-1dependent mechanism because
vß8 lacks the defined TSP-1 binding site for LAP-ß1 (Crawford et al., 1998) and
vß8 does not bind to TSP-1 (unpublished data). Moreover, unlike secreted TSP-1 (Crawford et al., 1998) secreted
vß8 cannot activate TGF-ß1. Thus, the mechanism by which
vß8 activates TGF-ß1 is not dependent on conformational changes, resulting from "inside-out" signal transduction as mediated by the ß6 cytoplasmic domain (Munger et al., 1999) or direct physical interaction as mediated by TSP-1(Crawford et al., 1998).
Our findings support a biologically relevant mechanism whereby SLC binds with high affinity to vß8 on the cell surface, which results in the metalloprotease-dependent release of active TGF-ß. Evidence to support this mechanism follows: (a) secreted
vß8 binds to LAP-ß1 with a high affinity with a dissociation constant similar to other TGF-ß receptors (Tucker et al., 1984); (b) both synthetic and endogenous MMP inhibitors block
vß8-mediated activation of TGF-ß1; (c) reconstitution of MT1-MMP into the H1264 MT1-MMPdeficient cell line rescues
vß8-mediated TGF-ß activation; (d)
vß8 and MT1-MMP specifically colocalize in LAP-ß1 substrate contacts; (e) consistent with a proteolytic event, active TGF-ß is liberated by an
vß8-dependent mechanism into the supernatants of tumor cell lines and into the aqueous phase of lung cancer xenografts; (f) the proteolytic substrate of
vß8-, MT1-MMPdependent activation of TGF-ß1 is likely to be LAP-ß1, since ß8-overexpressing, MT1-MMPexpressing H1264 cells cleave and inactivate LAP-ß1, whereas ß8-overexpressing, MT1-MMPdeficient H1264 cells do not; (g) cleavage of LAP-ß1 requires the concomitant activity of both ß8 and MT1-MMP, since ß8-specific RGD inhibitors and metalloprotease inhibitors both block cleavage. Precedent for such a proteolytic mechanism is that plasmin (Lyons et al., 1990) and MMP-9 (Yu and Stamenkovic, 2000) have each been shown to activate TGF-ß1 and TGF-ß2, respectively, by cleavage of LAP.
It is also possible that MT1-MMP acts indirectly by proteolytically modifying the activity of vß8 as suggested recently for the MT1-MMPdependent modification of the integrin
vß3 (Deryugina et al., 2000). However, this is unlikely because of the following: (a) cell lines expressing
vß8 attach to LAP-ß1 equally well whether or not they express MT1-MMP (unpublished data), suggesting that coexpression of MT1-MMP does not modify the activity of
vß8; (b) flow cytometry of H1264 cells overexpressing both ß8 and MT1-MMP using two different anti-ß8 monoclonal antibodies shows no alteration in surface expression of
vß8, indicating that antibody epitopes are preserved along with adhesive capability; (c) immunoprecipitations or Western blots of cells coexpressing
vß8 and MT1-MMP, using polyclonal antibodies against the cytoplasmic domain of ß8, show no electrophoretic shift or proteolytic degradation products. Therefore, we have no evidence of modification of
vß8 by MT1-MMP.
How does MT1-MMP interact with the vß8TGF-ß1 complex? Our data suggest that upon ligation of
vß8 with SLC,
vß8 and MT1-MMP become closely associated to form a complex on the cell surface. The cell surface appears to be required for productive interactions, since the secreted forms of
vß8 and MT1-MMP do not mediate activation of TGF-ß. Evidence for a physical association on the cell surface is that
vß8 and MT1-MMP colocalize in substrate contacts specifically on LAP-ß1. The nature of the MT1-MMPß8 interaction awaits elucidation by coimmunoprecipitation and domain interaction studies. Because the localization of MT1-MMP in LAP-ß1 substrate contacts is dependent on the presence of ß8, it is likely that
vß8SLC interactions are required to initiate the recruitment of MT1-MMP. The dynamic recruitment of MT1-MMP to
vß8TGF-ß complexes could provide a basis for the homeostatic regulation of TGF-ß activity in cellular microenvironments.
Although reconstitution of wild-type MT1-MMP is sufficient to support vß8-mediated activation, other metalloproteases could potentially be involved. For instance, MT1-MMP binds to and is potently inhibited by TIMP-2 (Brew et al., 2000), but MT1-MMPTIMP-2 complexes also serve as a cell surface receptor for MMP-2, and the function of this complex is activation of MMP-2 (Strongin et al., 1995). As such, it is not inconceivable that MMP-2 could also be involved in
vß8-mediated activation of TGF-ß. However, in H1264s cells MMP-2 is unlikely to be involved, since TIMP-1, a potent inhibitor of MMP-2 and weak inhibitor of MT1-MMP (Brew et al., 2000), has no effect on
vß8-mediated activation of TGF-ß. In contrast, ß8-mediated TGF-ß activation is inhibited by TIMP-2, suggesting that MT1-MMP may alone be sufficient to support ß8-mediated activation of TGF-ß. Although formally we cannot exclude additional roles for other MMPs or related metalloproteases such as ADAMs or ADAMTS, family members in
vß8 mediated activation of TGF-ß in other systems or cell types.
The ß8 subunit appears to be the only integrin subunit capable of coordinating metalloprotease activity with SLC bound to the cell surface because the other LAP-ß1 binding integrins are either incapable of activating TGF-ß (Munger et al., 1998) or, in the case of vß6, activating TGF-ß via a metalloprotease-independent pathway (Munger et al., 1999). Furthermore,
vß8-mediated TGF-ß activation is solely dependent on metalloproteases and not other proteases because inhibitors of aspartyl, serine, and cysteine proteases do not inhibit activation. Thus,
vß8-mediated activation of TGF-ß1 is not dependent on other proteases that have been implicated in SLC activation, including plasmin (Lyons et al., 1990), calpain (Abe et al., 1998), and cathepsin (Lyons et al., 1988).
Integrins (Brooks et al., 1996) and other cell surface molecules (Yu and Stamenkovic, 1999) have also been shown to localize MMP activity to the cell surface. For instance, the integrin vß3 has been shown to form an SDS stable cell surface complex with MMP-2 (Brooks et al., 1996) and to colocalize with MT1-MMP (Deryugina et al., 2001), whereas CD44 has been shown to mediate localization of MMP-9 (Yu and Stamenkovic, 2000) to the cell surface. However,
vß3 and CD44 are unlikely to be required for
vß8-mediated activation of TGF-ß because
vß3 is not expressed in multiple cell lines that support
vß8-mediated activation of TGF-ß (Table I) and because anti-CD44 antibodies do not inhibit
vß8-mediated activation of TGF-ß (unpublished data).
The selective MMP dependence of vß8- but not
vß6-mediated activation of TGF-ß1 clearly demonstrates that the mechanisms of
vß8- and
vß6-mediated activation of TGF-ß1 are different. A structural basis for these different mechanisms may be the striking difference in the predicted secondary structure of the extracellular domains of the ß8 and ß6 subunits (Moyle et al., 1991). Different integrin-mediated mechanisms of TGF-ß activation may have evolved to support distinct biologic functions. For instance, in the airway epithelium, a site where ß8 is normally expressed (Cambier et al., 2000), a mechanism to support a low and persistent level of activation of TGF-ß1 is necessary for homeostasis (Crawford et al., 1998). We speculate that
vß8 could sequester SLC to the cell surface where, in response to an environmental cue, changes in the local balance of MMP/TIMP activity could lead to
vß8-dependent liberation of active TGF-ß1. Thus,
vß8-mediated activation of TGF-ß1 might liberate the low levels of active TGF-ß1 sufficient to promote local paracrine effects but insufficient for undesirable local and systemic fibrogenic effects of TGF-ß1 (Border and Noble, 1994). Conversely, if
vß6 were to liberate TGF-ß by an MMP-dependent mechanism undesirable pathologic levels of TGF-ß might be released locally and into the systemic circulation because after injury expression of
vß6 (Breuss et al., 1993; Pilewski et al., 1997) and MMPs (Holgate et al., 1999) are both strongly and rapidly induced.
In summary, abundant evidence implicates the cytokine TGF-ß1, integrins, and MMPs as important mediators of homeostatic cell behaviors. This article provides the first evidence of the coordination of activity of members of these three major multigene families in the maintenance of homeostasis.
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Materials and methods |
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The following previously characterized antibodies were used: anti-ß8, SN1 (Nishimura et al., 1994), anti-ß6, 10D5 (Munger et al., 1999), E7P6 (Weinacker et al., 1994), polyclonal affinity purified rabbit anti-ß8 (Nishimura et al., 1994), antiLAP-ß1 (VB3A9) (Munger et al., 1998), and antiCD-44 (Picker et al., 1989) (Developmental Studies Hybridoma Bank). The following commercial antibodies and conjugates used were: anti-5 (P5D10; Chemicon), anti-ß1 (P5D2; Chemicon), pan-antiTGF-ß (1D11; R&D Systems), mouse or rabbit antiMT1-MMP (Calbiochem and Chemicon), mouse anti-BrdU (Dako), phycoerythrin goat antimouse, rhodamine donkey antirabbit (Chemicon), phycoerythrin goat antimouse, rhodamine goat antimouse (Jackson ImmunoResearch Laboratories), HRP-conjugated sheep antimouse (Amersham Pharmacia Biotech), HRP antirat (Cappel), HRPprotein A (Amersham Pharmacia Biotech). VN was prepared from outdated fresh frozen human plasma (Yatohgo et al., 1988). Collagen type 1 was prepared from rat tails (Montesano et al., 1983). Peptides (GRGDSNK and GRGESNK) were purchased (BioMol) or were commercially synthesized (GRRGDLATIH and GRRGELATIH) (BioSyn). The following antibiotics were used: puromycin, chloroquine (Sigma-Aldrich), geneticin (G418; GIBCO BRL), hygromycin (Calbiochem), and Fungizone, penicillin, and streptomycin (University of California at San Francisco cell culture facility).
Retroviral vectors, constructs, and RT-PCR
Retroviral vectors used were pLXSN (CLONTECH Laboratories, Inc.), pBabe Puro (a gift from Dr. Hartmund Land, Imperial Cancer Research Fund, London, UK) (Morgenstern and Land, 1990), pBabeß8Puro, pLXSNß8Neo (Cambier et al., 2000), and pBabeß6Puro. The ß6 cDNA was subcloned from ß6-Peak10 (a gift from R. Pytela, University of California at San Francisco) into pBabe Puro. Plasmids were purified using the QIAGEN plasmid purification system. ß8 truncation mutants were constructed using a PCR strategy introducing COOH-terminal truncations (amino acids 712 and 758 of the ß8 ORF [Moyle et al., 1991]), which replace the cytoplasmic domain of full-length ß8 cDNA in pcDNAIneo (Invitrogen). The mutant constructs were subcloned into pLXSN (CLONTECH Laboratories, Inc.). Truncated ß8 with a COOH-terminal AP tag (AP-vß8) was produced by in-frame blunt end ligation of the BspE1-Xho1 placental fragment from AP tag (Flanagan and Leder, 1990) into a chimeric pcDNAIß8/3neo construct (Nishimura et al., 1994). Full length MT1-MMP or transmembrane and cytoplasmic domain deleted MT1-MMP (
MT1-MMP) (a gift from Stephen Weiss, University of Michigan, Ann Arbor, MI) were subcloned into pBabe Puro or pLXSN. A construct creating a MT1-MMPGFP fusion protein (MT1-MMPpLEGFP) was created by destroying the stop codon of MT1-MMP in PCR3.1 by PCR mutagenesis to create a unique HpaI site. The HindIII, HpaI MT1-MMP fragment was then subcloned in-frame into pLEGFP (CLONTECH Laboratories, Inc.) between a HindIII and Klenow-treated BamHI site. Sequencing in both orientations was performed to verify the fidelity of each construct. Transfection of packaging cells and retroviral transduction was performed as described (Kinsella and Nolan, 1996). Pools of ß8-expressing cells were either used within 72 h for short term experiments or were sorted for uniform expression of ß8 and propagated on type I collagen-coated plates for long term experiments (Cambier et al., 2000). The (SBE)4-Lux reporter, which contains 4 CAGACA repeats of the SMAD binding element of the JunB promoter (Jonk et al., 1998), was a gift of Peter ten Dijke (Ludwig Institute for Cancer Research, Uppsala, Sweden). Total cellular RNA was harvested using a commercial kit (QIAGEN), and RT-PCR was performed as described (Nishimura et al., 1994). Primers, based on published sequences (Giambernardi et al., 1998; McCulloch et al., 2000) to MMP-2, -7, and -9, MT1-MMP, ADAMS-9, -10 and -17, and ß-actin were purchased (Operon Technologies). Annealing temperatures were based on published reports (Giambernardi et al., 1998; McCulloch et al., 2000).
Affinity chromatography, ligand binding, and adhesion assays
125I surface labeling and affinity chromatography was performed as described (Nishimura et al., 1994). Plasmids containing inserts for AP-tagged truncated secreted ß8 and truncated secreted v in pCDM8 (Nishimura et al., 1994) were stably expressed in 293 cells and characterized by immunoprecipitation (Nishimura et al., 1994). Serum-free supernatant containing AP-tagged
vß8 (
vß8-AP) was applied to 96-well plates precoated with LAP-ß1 (wild-type and mutant), VN, or BSA for 1 h at 37°C in the presence and absence of monoclonal antibodies. Bound receptor was detected colorimetrically with pNPP (Sigma-Aldrich) at A405. For affinity measurements, serum-free supernatant containing secreted AP-
vß8 was concentrated 40-fold (Vivaspin 100; Vivascience) and was applied to 10 µl of LAP-ß1Sepharose, VN-Sepharose, or IgG-Sepharose (Amersham Pharmacia Biotech) in the presence or absence of 1 mg/ml RGD peptide and incubated overnight at 4°C. The receptor concentration was determined against a standard curve generated using placental AP (Applied Biosystems). The samples were washed three times in wash buffer (Tris, 50 mM, pH 7.4, NaCl, 150 mM, CaCl2, 1 mM). Luminescence was determined using disodium 3-(4-methoxyspiro(1,2-dioxetane-3,2'-[(5'-chloro)tricyclo(3,3.1.13.7)decan]-4-yl) phenyl phosphate (CSPD) as a substrate (Tropix; Applied Biosystems) according to the manufacturer's instructions. Specific binding was defined as binding that remained after incubation with a 200-fold excess of RGD peptide. Binding curves were generated using nonlinear regression (Prism; GraphPad Software) from three independent experiments. Adhesion assays were performed as described (Nishimura et al., 1994).
Production of monoclonal and polyclonal ß8 antibodies
Balb/C mice were immunized by standard protocols with truncated secreted vß8 according to the University of California at San Francisco Committee on Animal Research guidelines. Splenocytes were fused with SP 2/0 myeloma cells using commercial protocols (Boehringer). Clones were screened by immunoprecipitation and flow cytometry to detect
vß8 (Nishimura et al., 1994). Antibodies were purified by ionic exchange chromatography using FPLC or used as supernatants for flow cytometry. Polyclonal anti-ß8 antiserum was generated and characterized as described (Nishimura et al., 1998) by immunization of rabbits with a cytoplasmic peptide (TRAVTYRREKPEEIKMDISK) corresponding to amino acids 740759 of the ß8 ORF (BioSyn).
Fluorescence activated cell analysis, sorting, and immunocytochemistry
For FACS®, ß8-expressing and mock-transduced cells were detached using 7 mM EDTA in DME, incubated with primary antibodies for 30 min at 4°C, and detected with phycoerythrin-conjugated secondary antibodies (Chemicon). Stained cells were analyzed using a FACsort® flow cytometer and CellQuest software (Becton Dickinson). Immunofluorescence microscopy was performed essentially as described with the following modifications (Cambier et al., 2000). HT1080 cells transduced with either pBabe Puro-ß8 or pBabe Puro underwent a second transduction with either MT1-MMPpLEGFP or pLEGFP and were selected with G418. LAP-ß1 was used to coat glass chamber slides at a concentration of 10 µg/ml, and cells were allowed to attach to coated slides for 4 h before fixation. Antibodies used were a polyclonal anti-ß8 antiserum and a monoclonal anti-GFP antibody (CLONTECH Laboratories, Inc.). Secondary reagents were AlexaFluor 595 goat antirabbit (Molecular Probes) and biotinylated sheep antimouse (Amersham Pharmacia Biotech) followed by Oregon green Streptavidin (Molecular Probes). Confocal images were obtained using a Bio-Rad Laboratories MRC-1024 laser scanning confocal system and LaserSharp2000 software (Bio-Rad Laboratories). Pseudocolored images and composites were generated in Adobe Photoshop® (v. 6.0).
Immunoprecipitation, Western blotting, and zymography
ß8-expressing or mock-transduced cells in confluent 10-cm dishes were either surface labeled with 125I, biotin, or were directly lysed in PBS with 1% Triton X-100 with 1 mM PMSF as described (Milner and Ffrench-Constant, 1994; Nishimura et al., 1994). Immunoprecipitations and Western blots were performed as described (Milner and Ffrench-Constant, 1994; Nishimura et al., 1994). For LAP-ß1 degradation experiments, LAP-ß1 (10 µg/ml) was added to individual wells of a 96-well plate containing ß8-overexpressing, MT1-MMP, MT1-MMP, or mock-transduced H1264 cells (4 x 104) in complete medium. For GM6001 and C1006 inhibition experiments, 104 cells were added, and for RGD peptide blocking experiments 2 x 104 cells were added. After a 1216-h incubation, the medium was collected and either added to TMLC reporter cells in the presence or absence of recombinant TGF-ß1 (10 pM) or was subjected to 12.5% SDS-PAGE and Western blotting as above. For zymography, cells were incubated in serum-free media overnight in the presence or absence of proMMP-2 (10 ng/well). Supernatants were harvested and loaded without heating or reduction and resolved by 10% SDS-PAGE (1 mg/ml gelatin). After three washes in 2.5% Triton X-100, the gels were incubated in substrate buffer (50 mM Tris-HCL, 5 mM CaCl2, 0.01% NaN3, pH 8.0) and incubated overnight at 37°C. Lucent bands of gelatinolytic activity were revealed by Coomassie staining. Digital images were acquired using Eastman Kodak Co. 1D 3.5.3 Imaging system. Composites were assembled in Adobe Photoshop® (v. 6.0).
TGF-ß bioassay
To determine TGF-ß activation in a coculture assay, TMLC cells were cultured with ß8-expressing or mock-transduced cells in the presence or absence of antiTGF-ßblocking antibody (10 µg/ml, 1D11; R&D Systems), anti-ß8 (100 µg/ml, 37E1) or anti-ß6 (150 µg/ml, 10D5) as described (Abe et al., 1994; Munger et al., 1999). To measure active TGF-ß in tumor tissue, equal weights of tumors were minced and incubated in sterile DME for 30 min at 4°C. The supernatants containing active TGF-ß were harvested after centrifugation (20 g) at 4°C. The pellets were then incubated in serum-free DME for 20 min at 80°C to activate SLC after which the supernatants were harvested. The supernatants containing active or heat-activated (latent) TGF-ß were then added to preplated TMLC cells with or without 1D11. For protease inhibitor assays, inhibitors were added at the initiation of the coculture. The maximal dose of each inhibitor was defined as the highest concentration that did not inhibit the ability of the TMLC cells to respond to recombinant active TGF-ß. To measure soluble TGF-ß activity from cultured cells, 106 HT1080 or SW480 cells, either ß8-expressing, ß6-expressing, or mock-transduced, were incubated in 100 µl of complete medium with or without 37E1 or 10D5 for 1 h at 37°C with gentle rotation. Cell-free supernatants were harvested by centrifugation (20 g) for 5 min at 4°C and then added to preplated TMLC cells in the presence or absence of 1D11. For soluble receptor assays, conditioned medium obtained from overnight cultures of 293 cells expressing truncated vß8 (Nishimura et al., 1994, 1998) was used. Relative luciferase units were defined as activity minus the background activity of the TMLC reporter cells. In some experiments, the TMLC reporter cells themselves activated a small amount of TGF-ß as determined by inhibition with antiTGF-ßblocking antibodies. In these experiments when the test cells did not activate TGF-ß the relative luciferase units (sample minus the TMLC background) were less than zero.
Cell proliferation assays and lung tumor xenographs
Cell cycle analysis was performed as described previously (Cambier et al., 2000) with the exception that some cells were treated overnight with 10 µg/ml LAP-ß1. BrdU incorporation assays were performed as described (Cambier et al., 2000). H647 tumor xenografts were established in nude mice as described (Cambier et al., 2000), and experiments were performed in full compliance with institutional guidelines and the University of California at San Francisco Committee on Animal Research.
Statistical analysis
Student's t test was used for comparison of two datasets; analysis of variance (ANOVA; for parametric data) or the Kruskal-Wallis test (for nonparametric data) were used for more than two datasets. Tukey's or Dunn's test was used for parametric or nonparametric data, respectively, to determine where the differences lay. Significance was defined as p < 0.05. Data are shown as means ± 1 SEM unless otherwise noted. Statistical software used was InStat v2.03 (GraphPad Software, Inc.).
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Footnotes |
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Acknowledgments |
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Submitted: 26 September 2001
Revised: 18 March 2002
Accepted: 18 March 2002
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References |
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