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CELLULAR METABOLISM
1Graduate Institute of Biochemistry and Molecular Biology, College of Medicine, National Taiwan University, Taipei, Taiwan; 2Osaka Prefecture University, Graduate School of Veterinary Medicine, Department of Toxicology, Sakai, Osaka, Japan; and 3Department of Oncology, Lombardi Cancer Center, Georgetown University Medical Center, Washington, District of Columbia
Submitted 7 December 2006 ; accepted in final form 28 February 2007
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ABSTRACT |
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 TOP ABSTRACT EXPERIMENTAL PROCEDURESRESULTSDISCUSSIONGRANTSREFERENCES |
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hepatocyte growth factor activator inhibitor 1; protease activation; low-density lipoprotein
Deregulation of the control of matriptase activation can result from overexpression of the enzyme and alteration of the ratio of the protein relative to HAI-1 and may also be due to altered glycosylation mediated by 
1-6-N-acetylglucosaminyltransferase V (10, 11), which causes 1-6 N-acetylglucosamine branching. This oligosaccharide modification results in enhanced matriptase stability. Altered subcellular localization of matriptase from cell-cell junctions in mammary epithelial cells to the membrane ruffles in breast cancer cells also results in uncontrolled activation (2, 4, 9). Matriptase is activated by a cleavage at its canonical activation motif, converting a single-chain zymogen into a two-chain active protease (2). Because the functional active site triad of matriptase is required for the activation of matriptase (mutation of these amino acids blocks activation), the activational cleavage of matriptase is not carried out by other active proteases, as is the case for most other serine proteases. In addition, the normal posttranslational modifications of the protein, such as NH2-terminal processing via cleavage at Gly-149 within the SEA domain and N-linked glycosylations and the intact LDL receptor class A domains of matriptase, are also required for its activation. Paradoxically, HAI-1 via its LDL receptor class A domain is also involved in matriptase activation (40). Therefore, we have proposed that autoactivation, via the interactions of matriptase zymogens, HAI-1, and other components not yet identified, is the mechanism for activation of this membrane-bound serine protease (40). In mammary epithelial cells, matriptase activation can be induced by sphingosine 1-phosphate (S1P), a blood-borne lysophospholipid, and occurs at cell-cell junctions (2, 3, 9). In contrast, breast cancer cells constitutively activate matriptase on the cell surfaces in the absence of contact with other cells and concentrate activated matriptase at membrane ruffles in response to EGF treatment (4). Furthermore, translocation and accumulation of both matriptase and HAI-1 at "activation foci" has been observed in 184 A1N4 immortal mammary epithelial cells during induced activation of matriptase (19). In these cells, matriptase is initially located primarily in perinuclear regions, where its activation does not occur. Upon the stimulation of cells with serum or S1P, matriptase translocates and accumulates at the cell-cell junctions (2, 3). The onset of activation of matriptase occurs at the activation foci at cell-cell junctions as tiny spots, after which the activation expands and elongates along the cell-cell junctions (9). The activation foci have also been observed inside the cells, different from cell-cell junctions, when the cells are exposed to suramin, a chemical inducer for matriptase activation (9, 19). The accumulation of matriptase at activation foci is consistent with the hypothesis that autoactivation occurs where dimerizations or oligomerizations of matriptase and other proteins are concentrated at particular cellular locations for interactions with each other and the occurrence of an activational cleavage. Despite the understanding of the processes described above, the detailed mechanisms of how matriptase activation occurs, how it is regulated, and what other proteins, besides HAI-1, are involved in matriptase activation, remain unknown. One of the major obstacles is the lack of a reliable and controllable cell-free system for biochemical characterization of matriptase activation. In the current study, we set out to establish and characterize such an in vitro system for further biochemical study of matriptase activation.
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EXPERIMENTAL PROCEDURES |
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Cell lines and culture conditions. Immortalized 184 A1N4 human mammary epithelial cells were a gift from Dr. Martha Stampfer (Lawrence Berkeley National Laboratory, Berkeley, CA) and routinely maintained, as previously described (2).
Monoclonal antibodies. Human matriptase protein was detected using the M32 monoclonal antibody (MAb), which recognizes both the latent (one chain) and activated (two chain) forms of the protease; the activated matriptase was detected using the M69 MAb, which recognizes an epitope present only in the activated (two chain) form of the enzyme (2, 3). Human HAI-1 was detected using the HAI-1-specific MAb M19 (22). We have summarized the forms of matriptase and HAI-1 and the interactions of these MAbs with matriptase and HAI-1 in Fig. 1.
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Figs. 5–9 were obtained by using two almost identical nitrocellulose membranes prepared by diffusion transfer methodology but not by reprobing the same nitrocellulose membrane. The HRP-conjugated M32 or M69 MAbs were used for immunoblot analysis (see Fig. 10).
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RESULTS |
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TOPABSTRACTEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONGRANTSREFERENCES |
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By systematic evaluation of various assay conditions, such as pH and the salt concentrations in buffer systems, we discovered that matriptase activation can occur spontaneously in cell homogenates (Fig. 2). We used 184 A1N4 immortal human mammary epithelial cells for our model system; however, we have demonstrated that in vitro matriptase activation assays can be conducted in various other cell lines that endogenously express matriptase and HAI-1 (data not shown). 184 A1N4 cells express high levels of matriptase, but when grown in culture media supplemented with 0.5% FBS for 2 days, they become devoid of matriptase activation (2). Following cellular homogenization by Dounce homogenizer in phosphate-citric acid buffer (pH 6.0), the resultant homogenates were incubated at room temperature for 10 min. Under these conditions, matriptase activation occurred spontaneously (Fig. 2, lanes 1). However, matriptase activation did not occur when the homogenates were additionally lysed by the addition of a nonionic detergent such as Triton X-100 (Fig. 2, lanes 2), subjected to sonication for few seconds on ice using a Tekmar Sonic Disruptor at setting 80 for microtip (Fig. 2, lanes 3), or incubated on ice at 4°C (Fig. 2, lanes 4). Activation of matriptase was evaluated by determining the levels of activated matriptase detected using the MAb M69, which is specifically directed against the two-chain activated matriptase but not the single-chain zymogen (2). We also determined the levels of total matriptase by using the MAb M32, which is directed against the third LDL receptor class A domain of the protease (9) [and is thus able to interact with both two-chain and single-chain matriptases containing this noncatalytic domain (19)].
In live cells, activation of matriptase induced by S1P or suramin is immediately followed by the inhibition of the protease by its cognate inhibitor HAI-1, resulting in a 120-kDa matriptase-HAI-1 complex, due to the requirement for HAI-1 in matriptase activation (40) and its presence with matriptase in activation foci (19, 37). Interestingly, the simultaneous HAI-1-mediated inhibition of active matriptase also occurred in this in vitro setting, since the majority of activated matriptase was detected in a 120-kDa complex (Fig. 2, lanes 1). As expected, the anti-total matriptase M32 MAb also recognized this 120-kDa matriptase-HAI-1 complex, in addition to the 70-kDa latent form of matriptase (Fig. 2). Concomitant with the appearance of the 120-kDa matriptase-HAI-1 complex, the levels of 70-kDa latent matriptase decreased. The 120-kDa matriptase-HAI-1 complex was also detected by anti-HAI-1 M19 monoclonal antibody (see Fig. 5). These results suggest that the in vitro matriptase activation model shares some characteristics with the induced matriptase activation in live cells, such as simultaneous HAI-1-mediated inhibition, formation of HAI-1 complex, and the involvement of membrane structures. Because active matriptase is inhibited by HAI-1 following its activation, it is not feasible to express the level of matriptase activation in terms of matriptase proteolytic activity by using synthetic substrates or gelatin zymography (2).
We further fractionated the cell homogenates by sequential centrifugations at 1,000 g for 10 min, which yielded insoluble pellets that contained mainly nuclei encompassed with cytoplasmic filaments and endoplasmic reticulum, and at 15,000 g for 10 min, yielding insoluble membrane pellets, which contained mainly membrane vesicles, mitochondria, and lysosomes (post-nuclear fraction), as well as the supernatant, which contained soluble cytosolic factors and proteins. The bulk of matriptase was detected in the insoluble pellets with a very small portion of the protease in the post-nuclear fraction, based on the yield of matriptase in both fractions (data not shown). When the insoluble pellets and the post-nuclear fraction were resuspended in phosphate-citric acid buffer and incubated at room temperature for in vitro matriptase activation, the levels of matriptase activation were examined by normalization of roughly equal amounts of total matriptase from both fractions (Fig. 3A). Activation of matriptase occurred in the insoluble pellets but not in the post-nuclear fraction (Fig. 3A). These data suggest the soluble cytosolic molecules in cell homogenates are not necessary for matriptase activation and that the insoluble pellets contain all the required elements for matriptase activation in vitro. Furthermore, although cell membranes are required for matriptase to undergo activation (Fig. 2), the simple presence of matriptase in membrane vesicles is not sufficient for its in vitro activation.
Although the soluble cytosolic molecules were not required for matriptase activation, it was interesting to note that their removal by separation of the soluble cytosolic fractions from the insoluble pellets significantly increased the rate of matriptase activation in the insoluble pellets (Fig. 3B, compare lanes 2 and 4). These data suggest that matriptase activation may be regulated by two independent mechanisms: a soluble cytosolic suppressor and insoluble autonomous activation machinery. Homogenization of cells in the absence of detergent apparently preserves the function of the activation machinery and may also sufficiently dilute the cytosolic suppressor to allow matriptase activation to proceed spontaneously. Removal of the cytosolic suppressor from the insoluble activation machinery by centrifugation allowed matriptase activation to proceed to a much greater extent, as illustrated by comparing the ratios between activated matriptase (in the 120-kDa HAI-1 complex) relative to the latent matriptase (the 70-kDa band). In the absence of the cytosolic suppressor(s), about one-third to one-half of the matriptase underwent activation within 10 min at room temperature (see Figs. 3 and 5–10). In contrast, in the presence of the cytosolic suppressor(s), <10% of the matriptase underwent activation under the same conditions (Figs. 2 and 3B). In addition to 184 A1N4 mammary epithelial cells, in vitro activation of matriptase and a soluble suppressor of matriptase activation were observed in a variety of breast cancer cells, including T-47D and MCF-7 cells, and in prostate cancer cells, including LNCaP cells (data not shown), suggesting the ubiquity of these mechanisms governing matriptase activation.
Biochemical characterization of matriptase activation in vitro. In this study, we have focused on the biochemical characterization of the activation machinery. Characterization of the nature and identity of the cytosolic suppressor of matriptase activation must be the subject of a future study.
We began our characterization by examining the location of the protease and where its activation occurred in the insoluble pellets by immunofluorescent staining (Fig. 4). By phase-contrast microscopy, the insoluble pellets were observed mainly as the broken cells that contained nuclei surrounded by cytoplasmic filaments (perinuclei) (Fig. 4, blue color for nuclei in merged image). Total matriptase (Fig. 4, M32, green) was detected at the cytoplasmic filament area surrounding the nuclei. Activated matriptase, detected by MAb M69 (Fig. 4, M69, red) also was observed in the perinuclear area only for those insoluble fractions incubated at room temperature (Fig. 4B, M69, red), not for the controls, which were kept on ice (Fig. 4A, M69, red). In live cells, the majority of the matriptase was detected in the perinuclear, presumably Golgi-endoplasmic reticulum (ER), area (Fig. 4B, inset bottom right, green) (9, 19, 24, 37). This specific perinuclear distribution of matriptase in cellular homogenates simply reflected its perinuclei localization in live cells. The spontaneous activation of matriptase in the insoluble pellets suggests that matriptase and its activation machinery were still well preserved after cell fractionations (cellular homogenization).
Time course of matriptase activation in vitro. Activation of matriptase began to occur as soon as 5 min after incubation at room temperature (Fig. 5). This in vitro activation of matriptase apparently occurred in a robust fashion, with more than one-third of latent matriptase becoming activated after 10 min of incubation at room temperature and more than one-half after 30 min, comparing the ratio of the level of the 120-kDa matriptase complex to the 70-kDa latent matriptase form (Fig. 5, total matriptase). The appearance of activated matriptase, mainly in the 120-kDa matriptase-HAI-1 complex, was at the cost of the 70-kDa latent matriptase form; increased levels of the 120-kDa matriptase-HAI-1 complex were accompanied with a decrease in the 70-kDa latent form. As described above, the activated matriptase was detected mainly in its 120-kDa complex with its cognate inhibitor HAI-1 by using MAb M69, which can specifically recognize the two-chain form of matriptase. We further confirmed that this 120-kDa band contains HAI-1 by using anti-HAI-1 MAb M19, which recognizes both the 120-kDa complex and the 55-kDa full-length HAI-1 (Fig. 5, HAI-1). Although more than one-half of the total matriptase was activated after 30 min of incubation, only a small portion of HAI-1 was detected in the matriptase-HAI-1 complex. Since the stoichiometry of the interaction between active matriptase and HAI-1 is likely to be 1:1, based on the size of their complex (120 kDa) and the fact that only the Kunitz domain 1 of HAI-1 can inhibit matriptase (16), the relatively small proportion of the available HAI-1 in the 120-kDa complex suggests that HAI-1 is present in considerable excess relative to matriptase levels in the intracellular pools. Together, these results suggest that matriptase activation in vitro shares many features with its counterpart in live cells, such as rapidity of activation (occurrence within 10 min) (3, 19) and the maintenance of a significant molar excess of HAI-1 relative to matriptase (19).
Matriptase activation in vitro occurs in narrow pH range. To determine the optimal pH range for matriptase activation, we first tested a broad range of pH. Matriptase activation only occurred over a narrow range from pH 5 to 8. Thus a buffer system spanning pH 4.8 to 7.4, with intervals of 0.2 pH unit, was established by mixing 0.1 M citric acid with 0.2 M disodium phosphate (32). Although matriptase activation did not occur at pH 5 and occurred only slightly at pH 5.2, a sharp increase in activation was observed at pH 5.4 (Fig. 6). The optimal pH for matriptase activation was found to be between 5.8 and 6.0. A gradual decrease in activation was observed with increased pH between 6.4 and 7.2.
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Anti-matriptase MAbs M32 and 21-9 inhibit matriptase activation in vitro. In previous studies (9, 37), our group demonstrated that the mouse-derived, anti-matriptase MAb M32, which recognizes an epitope at the third LDL receptor class A domain of matriptase, was able to inhibit S1P-induced in vivo matriptase activation in mammary epithelial cells. A similar effect on in vivo S1P-induced matriptase activation was also demonstrated with another rat-derived, anti-matriptase MAb, 21-9 (data not shown). Although the epitope recognized by this rat-derived anti-matriptase MAb has not been determined, MAb 21-9 likely recognizes the LDL receptor class A domains of the protease, based on its interaction with a 40-kDa matriptase fragment, which presumably contains the serine protease domain and LDL receptor class A domains (24), but lack of interaction with the serine protease domain of matriptase (data not shown). Pretreatment of insoluble pellets with either of these anti-matriptase MAbs for 2 h at 4°C inhibited subsequent in vitro matriptase activation in a dose-response manner (Fig. 10). Monoclonal antibody 21-9 at 5 µg/ml (30 nM) completely inhibited matriptase activation in vitro, with M32 being slightly less potent. This minor difference in the inhibitory potency between both MAbs could result from their binding affinities or localizations of the epitopes recognized by these two MAbs in LDL receptor class A domains. Control mouse IgG at 5 µg/ml produced no inhibition. Directly HRP-labeled M69 MAb was used to detect activated matriptase in this study to avoid the use of an anti-mouse IgG secondary antibody, which would have interacted with the antibodies added to the insoluble pellets, generating problems with background bands. The blockage of matriptase activation by anti-matriptase MAbs further supports the idea that protein-protein interactions between matriptase zymogens are involved in matriptase activation.
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DISCUSSION |
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TOPABSTRACTEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONGRANTSREFERENCES |
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pH 5) (14, 29, 33, 45), activation of matriptase may occur during its intracellular trafficking to cell surfaces, particularly during its transit through the Golgi, since the optimal pH for matriptase activation is around 6 (Fig. 6). Therefore, interference with pH may represent one of the mechanisms for suppressing matriptase activation in ER during intracellular trafficking. However, the presence of other activation suppressor seems likely to be required during its trafficking. In contrast to preventing premature matriptase activation during trafficking, the neutral pH at cell-cell contacts or the cell surfaces may present a barrier to matriptase activation that must be overcome by S1P-induced activation of matriptase in the context of whole cells. Matriptase activation in vitro occurs only in the insoluble fractions. Solubilization of matriptase from the insoluble fractions by the nonionic detergent Triton X-100 completely inhibited matriptase activation. This result suggests that matriptase and its activation machinery must be anchored on lipid bilayers in order for activation to proceed. However, the anchoring of matriptase in lipid bilayers is not sufficient for its activation, since matriptase, when anchored in membrane vesicles in the post-nuclear fractions, failed to undergo in vitro activation (Fig. 3). Furthermore, physical disruption of the insoluble fractions by sonication also completely abolished matriptase activation (Fig. 2). Therefore, simple lipid bilayers may provide necessary platforms for the anchoring of matriptase and its activation machinery, whereas more complicated, higher order structures of lipid bilayers may be required for efficient matriptase activation. Because of the likely involvement of lipid bilayers in matriptase activation, the fluidity of the lipid bilayer may also affect activation. Since temperature is a major factor affecting the fluidity of lipid bilayers, the effect of temperature on matriptase activation in vitro, which occurred at very low rate at 12°C but at a much higher rate at 13°C and above (Fig. 7), may be the result of a phase transition of membranes from a gel (frozen) state to a liquid state. In the gel state, it would be predicted that the movement of the matriptase enzyme and other proteins would be much slower, interfering with the protein-protein interactions required for activation.
The involvement of lipid bilayers and higher order membrane structures in matriptase activation may explain the atypical biochemical characteristics of in vitro matriptase activation. If matriptase activation depended on the activity of other proteases, the protease inhibitors (including the protease inhibitor cocktail), would have been expected to inhibit matriptase activation more effectively (Fig. 9), and matriptase activation would likely have occurred in the presence of Triton X-100 or following the sonication of cell homogenates (Fig. 2). Moreover, for a classic protease-catalyzed reaction, a symmetrical, bell-shaped profile would be expected for the plots of activity vs. pH, because the enzymatic activity would drop on either side of the optimal pH due to ionization or protonation of the amino acids participating in catalysis with the increase or decrease of pH, respectively. In vitro activation of matriptase occurs within a relatively narrow pH range with an asymmetrical profile of activation rate vs. pH. This unique pH effect may result from interference with the protein-protein interactions or the higher order structure of the lipid bilayers, rather than from interference with the amino acid residues actually participating in the subsequent proteolytic cleavage. This hypothesis is also supported by the effects of ionic strength on matriptase activation, where a sharp decrease in matriptase activation was observed with an increase in NaCl concentration from 80 to 100 mM. This modest increase in ionic strength again may disrupt protein-protein interactions or the higher order structure of the lipid bilayers. The marginal inhibition of matriptase activation by protease inhibitor cocktail further supports the suggestion that activation of matriptase is not via the activity of other proteases (Fig. 9). Taken together, these unique biochemical features of in vitro matriptase activation are consistent with autoactivation of matriptase in which the cleavage of the activation motif of matriptase is proposed to be carried out by the intrinsic, weak proteolytic activity of matriptase zymogens. It is conceivable that dimerization or oligomerization of matriptase zymogens is required for the activational cleavage. The anchor of matriptase on lipid bilayers could facilitate the interactions between matriptase zymogen molecules. The unidentified proteins could play roles in facilitating the contacts between two matriptase zymogen molecules. The noncatalytic domains and posttranslational modification of matriptase could provide structural basis for the protein-protein interactions required for matriptase activation. For example, the CUB domain possesses self-assembly potential, and the LDL receptor class A domain could interact with other proteins.
The essential roles of protein-protein interactions and the maintenance of proper lipid bilayer structures in matriptase activation provide attractive targets for novel strategies for the development of anti-matriptase inhibitors. Classic protease inhibitors, which inhibit the activity of the catalytic domain, often present serious problems in terms of selectivity for the protease that they are designed to inhibit due to the overlapping specificity of serine protease catalytic domains. The unconventional activation mechanism for matriptase may provide a unique opportunity to inhibit the protease function before its activation. Indeed, we have shown that interference with these protein-protein interactions by using the anti-matriptase MAbs was indeed able to inhibit matriptase activation. In future studies, high-throughput screening of small molecules could yield inhibitors of matriptase activation.
In conclusion, based on these biochemical characterizations, matriptase activation can be considered to be a complex process, probably involving multiple steps of protein-protein interactions. Anchoring matriptase, HAI-1, and other components not yet identified in cell membranes, as well as the organization of theses membrane-anchored proteins into subcellular, higher order lipid-bilayer membrane structures, may be critical to ensure appropriate protein-protein interactions. The in vitro matriptase activation system seems to faithfully replicate many features of the inducible matriptase activation found in whole cells, including a rapid reaction profile, involvement of cell membranes, HAI-1-mediated inhibition, and the formation of 120-kDa matriptase-HAI-1 complex as an end product. This new activation system may not only shed light on how cells regulate matriptase activity but may also provide a new strategy for development of matriptase activation inhibitors.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
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