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Originally published In Press as doi:10.1074/jbc.M410396200 on December 22, 2004

J. Biol. Chem., Vol. 280, Issue 10, 9291-9296, March 11, 2005
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A Key Role for Mast Cell Chymase in the Activation of Pro-matrix Metalloprotease-9 and Pro-matrix Metalloprotease-2*

Elena Tchougounova{ddagger}, Anders Lundequist{ddagger}, Ignacio Fajardo{ddagger}§, Jan-Olof Winberg¶, Magnus Åbrink{ddagger}, and Gunnar Pejler{ddagger}||

From the {ddagger}Swedish University of Agricultural Sciences, Department of Molecular Biosciences, BMC, Box 575, 75123 Uppsala, Sweden and the Department of Biochemistry, Institute of Medical Biology, University of Tromsø, 9037 Tromsø, Norway

Received for publication, September 10, 2004 , and in revised form, November 12, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Chymases, serine proteases exclusively expressed by mast cells, have been implicated in various pathological conditions. However, the basis for these activities is not known, i.e. the in vivo substrate(s) for mast cell chymase has not been identified. In this study we show that mice lacking the chymase mouse mast cell protease 4 (mMCP-4) fail to process pro-matrix metalloprotease 9 (pro-MMP-9) to its active form in vivo, whereas both the pro and active form of MMP-9 was found in tissues of wild type mice. Moreover, the processing of pro-MMP-2 into active enzyme was markedly defective in mMCP-4 null animals. Histological analysis revealed an increase in collagen in the ear tissue of mMCP-4-deficient animals accompanied by increased ear thickness and a higher content of hydroxyproline. Furthermore, both lung and ear tissue from the knock-out animals showed a markedly increased staining for fibronectin. MMP-9 and MMP-2 are known to have a range of important activities, but the mechanisms for their activation in vivo have not been clarified previously. The present study thus indicates a key role for mast cell chymase in the regulation of pro-MMP-2 and -9 activities. Moreover, the results suggest an important role for mast cell chymase in regulating connective tissue homeostasis.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
When mast cells (MCs)1 are activated, they degranulate and thereby release a panel of powerful preformed inflammatory mediators, including histamine, cytokines such as tumor necrosis factor-{alpha}, proteoglycans, and various MC-specific proteases (1, 2). The MC proteases are divided into three main subclasses, tryptases, chymases, and carboxypeptidase A (3–5), all of which are stored in the MC granule in complex with heparin proteoglycan (6, 7). Chymases, serine proteases with chymotrypsin-like substrate specificities, have potent pro-inflammatory properties (8) and have been implicated in a variety of pathophysiological conditions, e.g. angiogenesis (9), heart failure (10), and fibrosis (11). However, it has not been possible to determine the mechanism by which chymases influences these processes, i.e. the physiological substrate(s) for chymase has not been identified.

Matrix metalloproteases (MMPs) are known to be involved in a variety of physiological and pathological processes and are currently attracting a large clinical interest as potential drug targets in therapeutic intervention with various diseases (12–16). The MMPs, similar to most proteolytic enzymes, are synthesized with an N-terminal propeptide that needs to be removed to achieve proteolytic activity. Thus, the physiological processes that lead to propeptide cleavage are imperative in terms of regulating the activity of most proteases (17). The MMP family currently comprises >20 members (18). The members all share common structural features but differ in regard to substrate specificities, although overlapping substrate specificities between certain members of the MMP family occur. Thus, MMP-2 and -9 share the ability to degrade denatured collagen (gelatin) and are therefore also denoted gelatinases A and B, respectively. Gene-targeting studies have implicated MMP-2 (14) and MMP-9 (19) in angiogenesis and tumor metastasis. However, it is not known how these proteases are activated in vivo. A number of in vitro studies have identified potential pro-MMP-9 processing proteases, including MMP-13, MMP-3, tissue kallikrein, MMP-2, plasmin, and MC chymase (20, 21), and it has been shown that a membrane-type MMP (MT1-MMP) can activate pro-MMP-2 (22). For most of these proteases, the in vivo relevance of their ability to activate pro-MMP-2 and -9 is not clear. However, in the case of pro-MMP-2 it appears that MT-MMPs are important activators in tissues and that tissue inhibitor of metalloproteinase-2 (TIMP-2) is required for efficient activation in vivo (23).

Because MCs are often found close to sites where MMP activity is operating, e.g. in wound healing areas and the vicinity of neoplastic tissue, MC chymase would be well suited for an in vivo role in pro-MMP-2 and -9 activation. Indeed, MCs have, in addition to their well established role in immediate hypersensitivity (24), been strongly implicated in various processes in which MMP activity is involved, e.g. angiogenesis (25), wound healing (26), arthritis (27), and tumor metastasis (28), but their mechanism of action under these circumstances has not been clarified. The aim of the present investigation was therefore to address the relative importance of MCs and the MC proteases, in particular chymase, in the activation of pro-MMP-2 and -9 in vivo.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Mice—Mice deficient in N-deacetylase/N-sulfotransferase 2 (NDST-2; congenic C57BL/6 background) and mouse MC protease 4 (mMCP-4; mixed genetic background C57BL/6-129SVJ; F1 from N2 and F1 from N6 x N7; mice at different stages of back-crossing toward C57BL/6 produced similar results) were generated by gene targeting (6, 29). In all experiments, age-matched litter mates (12–16 weeks old) were used.

Cell Culture and Purification of MCs—Peritoneal cells were collected by peritoneal washing with 10 ml of cold phosphate-buffered saline (pH 7.4). Cells were centrifuged (300 x g at 4 °C for 10 min) and cultured in serum-free medium, HybridoMed DIF (Biochrom AG, Berlin, Germany), supplemented with 50 µg/ml gentamycin (Invitrogen). The cells were distributed in 24-well plates (~1.0 x 106 cells in 0.5 ml/well; Nunc). Cells were incubated at 37 °C in a humidified atmosphere of 5% CO2. Conditioned media were collected at various time points. For the activation of cells, phorbol 12-myristate 13-acetate (final concentration 25 ng/ml) was added. After various times of incubation, conditioned media were collected.

MCs of ~95% purity, as judged by toluidine blue staining (the majority of the contaminating cells were red blood cells), were prepared by density gradient centrifugation on metrizamide. Mononuclear cells (MC-depleted) were recovered from the buffer/metrizamide interphase. Purified mononuclear cells (macrophages and lymphocytes in approximately equal proportions) and MCs were cultured either separately or in co-culture.

Zymography—Protein was extracted from homogenized ears, lungs, and hearts in two steps. First, homogenization (using a PT1200 Polytron device; Kinematica AG, Lucerne, Switzerland) of tissues was performed in 1.5 ml of low salt phosphate-buffered saline buffer (pH 7.4) containing 1 mM EDTA, 1% Triton X-100, 0.14 M NaCl and a protease inhibitor mixture (final concentrations were 2.3 mM 4-(2-aminoethyl)-benzenesulfonyl fluoride, 10 mM EDTA, 0.2 mM bestatin, 0.03 mM pepstatin A, and 0.03 mM E-64) (Sigma-Aldrich). Homogenates were centrifuged (13,400 rpm at 4 °C for 20 min), and supernatants were collected. Gelatinase activities in the low salt extracts were negligible, as assessed by gelatin zymography (not shown). Next, the pellets were extracted with 500 µl of a high salt phosphate-buffered saline buffer (pH 7.4) containing 1 mM EDTA, 1% Triton X-100, 2 M NaCl, and protease inhibitors as described above. Samples were rotated for 30 min at 4 °C and centrifuged (13,400 rpm at 4 °C for 20 min), and the resulting supernatants were collected for zymography analysis. Zymography was performed with samples normalized for protein concentration after measuring protein content using the Bio-Rad protein assay according to the instructions provided by the manufacturer. The samples were subjected to SDS-PAGE in gels containing 8% polyacrylamide and 1 mg/ml gelatin. After electrophoresis, gels were washed 2 x 10 min with 2% Triton X-100 and 50 mM Tris-HCl (pH 7.4) followed by 2 x 10 min of washing with 50 mM Tris-HCl (pH 7.4) and subsequent incubation at 37 °C for 20 h in 50 mM Tris-HCl (pH 7.4), 5 mM CaCl2, and 1% Triton X-100. Gels were stained with Coomassie Blue R-250 and destained. The positions of the different forms of MMP-2 and -9 in the zymograms were determined by comparison with the migration of pro and active MMP-2 and -9 present in conditioned medium from HT-1080 cells.

Histochemical Analysis—Ears, lungs, and hearts were frozen on dry ice and kept at –70 °C. Organs were mounted with Tissue-Tek (MICROM International, Walldorf, Germany), and cryosections of 7 µm were prepared. For toluidine blue staining, slides were fixed in ethanol for 3 min, stained with 0.1% toluidine blue in 50% ethanol (v/v) (pH 3) for 15 min, and then rinsed with water. In situ zymography was performed by incubating cryosections with 40 µg/ml fluorescein-conjugated gelatin (Molecular Probes, Eugene, Oregon) in 50 mM Tris-HCl (pH 7.4), 10 mM CaCl2, 150 mM NaCl, and 0.05% Brij-35 for 10 h at 37 °C. Sections were washed 3 times with H2O and mounted with Vectashield. Gelatinase activity was visualized using fluorescence microscopy. For visualization of collagen content, cryosections (5 µm) were prepared from paraffin-embedded organs and stained with a van Gieson stain (which stains collagen red). Immunohistochemical analysis of fibronectin was likewise performed on cryosections (5 µm) prepared from paraffin-embedded organs using an avidin-biotin complex-based technique (Vectastain Elite ABC Kit; Vector Laboratories, Burlingame, CA). Sections were incubated with a rabbit antiserum toward bovine fibronectin (cross-reacts with mouse fibronectin; gift from Staffan Johansson, Uppsala University) diluted 1:100 in Tris-buffered saline. As a secondary antibody, biotinylated anti-rabbit IgG was used (diluted 1:100 in Tris-buffered saline).

Amino Acid Analysis—For determination of amino acid content, wet samples (70–170 mg) were hydrolyzed for 24 h at 100 °C in 5 ml of 6 M HCl containing 5 µmol of norleucine as internal standard. Following hydrolysis, 1-ml aliquots were evaporated to dryness, and the pellets were dissolved in 5 ml of pH 2.2 sample application buffer. 50-µl aliquots were analyzed with a Biotronik LC-5001 amino acid analyzer using the extended physiological program with lithium citrate buffers and ninhydrin detection. Data collection was done with a Shimadzu CR2-AX integrator. The results were normalized on the basis of the weights of sample taken for analysis and the recovery of the internal standard, norleucine. Measurements of ear thickness were carried out with a Oditest device (Kroeplin, Germany) and conducted in a blinded fashion.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
First we addressed the importance of MCs in the processing of endogenous pro-MMP-9. Mouse peritoneal cell populations, typically composed of ~2–4% MCs with the remainder being macrophages and lymphocytes in approximately equal proportions, were recovered. MCs were separated from the macrophages/lymphocytes, and the two cell populations were cultured either separately or in co-culture. Culture supernatants were recovered and analyzed for the presence of pro and active MMP-9 by gelatin zymography. From Fig. 1A it is clear that culture supernatants recovered from the MC-depleted peritoneal cell population contained only the pro-form of MMP-9. However, when MCs were added, processing of pro-MMP-9 into its active form was seen, and the degree of pro-MMP-9 activation was dependent on the number of MCs added. These results thus indicate that the processing of pro-MMP-9 into active enzyme is MC-dependent.



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  		FIG. 1.
MC-dependent activation of pro-MMP-9. A, peritoneal cells depleted of MCs (1 x 106 cells in 0.5 ml/well) were cultured either alone or together with purified peritoneal MCs as indicated. MCs were also incubated alone. Conditioned media were recovered after 48 h of culture and analyzed by gelatin zymography. B, peritoneal cells were recovered from WT and NDST-2 null mice and cultured with or without phorbol 12-myristate 13-acetate (PMA). Conditioned media were collected after 96 h and analyzed by gelatin zymography. As a source of standard human pro-MMP-9, a sample of conditioned medium from the human cell line, HT-1080, was analyzed (far left lane in panel A). Note that the human pro-MMP-9 is somewhat smaller than the murine counterpart, which is consistent with previous studies (see, for example, Ref. 39).

 
To determine whether the pro-MMP-9 processing was catalyzed by any of the neutral proteases that are present in the MC secretory granule, we used peritoneal cells recovered from a mouse strain deficient in the gene for N-deacetylase/N-sulfotransferase 2. NDST-2 is an essential enzyme for the biosynthesis of MC heparin, and it was shown previously that the lack of heparin leads to a global reduction in the neutral proteases present in the peritoneal MC secretory granule A (6, 7). As shown in Fig. 1B, pro-MMP-9 processing was clearly seen in wild type (WT) peritoneal cells but was undetectable in peritoneal cell populations recovered from NDST-2–/– animals, indeed supporting a central role for the MC neutral proteases in the activation of pro-MMP-9. MMP-2, either in pro or active form, was not clearly visible in conditioned media from peritoneal cells recovered from WT or NDST-2 null animals.

To investigate whether MC chymase is of physiological relevance for MMP-2 and -9 activation, we made use of a mouse strain carrying a targeted inactivation of the gene for mMCP-4 (29). mMCP-4 constitutes the dominating chymotrypsin-like enzyme in connective tissue type MCs and, importantly, mMCP-4 may be the functional counterpart to the only human chymase identified (29). Indeed, only the pro-form of MMP-9 was detected in peritoneal cell culture supernatants from mMCP-4–/– mice, whereas both pro and active MMP-9 was present in supernatants from WT cells (Fig. 2A). To examine the in vivo relevance of these findings, we investigated whether the lack of mMCP-4 caused alterations in the ratio of pro versus active MMP-9 in various tissues. Extracts were prepared from ears, lungs, and hearts and analyzed by gelatin zymography. As is evident from Fig. 2, B–D, only the pro-form of MMP-9 was detected in mMCP-4–/– tissues, whereas both the pro and active forms of MMP-9 were present in WT tissues. Interestingly, the degree of pro-MMP-9 processing in WT tissues appeared to be correlated with the number of MCs present in the respective tissues. Thus, ear tissue, which is rich in connective tissue type MCs, predominantly contained the active form of MMP-9, whereas lung tissue, a tissue with low numbers of MCs, had lower levels of active MMP-9.



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  		FIG. 2.
pro-MMP-9 processing in wild type and mMCP-4 null mice. A, gelatin zymography analysis of conditioned media from peritoneal cell cultures of WT and mMCP-4–/– mice collected after the time points indicated. B–D, tissue homogenates were prepared from ears (B), lungs (C), and hearts (D) from WT and mMCP-4 null animals and analyzed by gelatin zymography. Arrows indicate the position of the pro and active forms of MMP-9. Samples from at least four different animals of each genotype were analyzed. Shown are the results obtained from representative samples taken from different animals. As a source of standard human pro-MMP-9, a sample of conditioned medium from the human cell line, HT-1080, was analyzed (far left lane in panel A).

 
We also assessed whether mMCP-4 was of importance in the processing of MMP-2. As shown in Fig. 3, both the pro and active forms of MMP-2 were found in the various WT tissues. Both forms of MMP-2 were also detected in mMCP-4–/– tissues. However, the level of active MMP-2 was markedly lower than in WT tissues, indicating that mMCP-4 is important but not essential for the activation of pro-MMP-2 in vivo.



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  		FIG. 3.
pro-MMP-2 processing in wild type and mMCP-4 null mice. A, gelatin zymography analysis of conditioned media from peritoneal cell cultures of WT and mMCP-4–/– mice collected after the time points indicated. B–D, tissue homogenates were prepared from ears (B), lungs (C), and hearts (D) from WT and mMCP-4 null animals and analyzed by gelatin zymography. Arrows indicate the position of the pro and active forms of MMP-2. Samples from at least four different animals of each genotype were analyzed. Shown are the results obtained from representative samples taken from different animals. As a source of standard human pro-MMP-2, a sample of conditioned medium from the human cell line, HT-1080, was analyzed (far left lane in panel A).

 
The effect of mMCP-4 on gelatinolytic activity was also studied by in situ gelatin zymography. In ear tissue, gelatinolytic activity was observed in the epidermis of both WT and mMCP-4–/– mice (Fig. 4, C and D), possibly due to MMP-9 expression and non-chymase-dependent activation in keratinocytes. In the dermis region, by contrast, strong gelatinolytic activity was found in the WT but not in mMCP-4–/– tissue. In lung tissue of WT mice, strong gelatinolytic activity was found in the vicinity of blood vessels and bronchioli but was essentially absent at the corresponding sites in mMCP-4–/– lung tissue (Fig. 4, A and B). The in situ gelatinolytic activity was completely blocked by EDTA, an inhibitor of metalloprotease activities (not shown), but it was not blocked by Pefabloc SC, a general serine protease inhibitor (not shown). In Fig. 4 it is also shown that MCs, as judged by staining with toluidine blue, are localized to regions in which gelatinolytic activity is present (Fig. 4, E–H).



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  		FIG. 4.
A–D, in situ gelatin zymography of lung (A and B) and ear sections (C and D) prepared from of mMCP-4+/+ and mMCP-4–/– mice. Sections were incubated with fluorescein-conjugated gelatin as a substrate, and the fluorescence seen corresponds to gelatinase activity. The red arrow in lung tissue indicates the presence of gelatinase activity in bronchioli, whereas the white arrow indicates gelatinase activity around veins. Note that gelatinase activity at these sites is seen in tissue obtained from WT mice (A) but is absent or diffuse in sections from mMCP-4–/– animals (B). Strong gelatinase activity was observed in the dermis region in WT mice (C) but was essentially absent in dermis of mMCP-4 null mice (D). E–H, lung (E and F) and ear (G and H) tissue sections from WT and mMCP-4 null animals were stained with toluidine blue in order to detect MCs. Note the large number of MCs in ear tissue from both WT and mMCP-4–/– mice and the low number of MCs in lung tissue. Note also the similar numbers and staining properties of MCs from WT and mMCP-4 null animals. Scale bars, 100 µm. KO, knock-out.

 
Because MMPs have been implicated in various connective tissue remodeling processes, we next studied whether the lack of mMCP-4 and the consequent reduction in active MMPs caused alterations in connective tissue composition. Analysis using the van Gieson stain showed an enhanced staining in ear tissue from mMCP-4 null animals as compared with controls, indicating an increase in collagen deposition (Fig. 5, A and B), and the increase in staining was accompanied by a significant increase in ear thickness (Fig. 5E). Moreover, amino acid analysis of ear tissue revealed a significant increase in the content of hydroxyproline, an amino acid modification that is specific for collagen, in ear tissue from mMCP-4 null animals (Fig. 5F). In contrast, no significant alterations in the levels of other amino acid residues apart from that of hydroxyproline were seen (not shown). There was also a tendency of increased collagen deposition in the vicinity of bronchioli (Fig. 5, C and D), although there was no significant increase in hydroxyproline content of total lung tissue (not shown). We also stained for fibronectin, another prominent connective tissue component. As can be seen in Fig. 5, G–J, there was a markedly stronger staining for fibronectin in mMCP-4 null lung and ear tissue as compared with control, indicating a higher degree of fibronectin deposition.



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  		FIG. 5.
Effect of the mMCP-4 knockout (KO) on connective tissue. A–D, cryosections from paraffin-embedded ears (A and B) and lungs (C and D) from mMCP-4+/+ (A and C) and mMCP-4–/– (B and D) animals were stained with van Gieson stain for visualization of collagen. E, measurement of ear thickness in mMCP-4++ and mMCP-4–/– animals. Data are shown as mean ± S.D., n = 6. *, p < 0.05. F, quantification of hydroxyproline in ears of mMCP-4++ and mMCP-4–/– animals. Data are shown as mean ± S.D., n = 2 (WT) or 6 (–/–). *, p < 0.05. G–J, fibronectin content in WT and mMCP-4–/– tissues. Cryosections from paraffin-embedded lungs (G and H) and ears (I and J) from mMCP-4++ (G and I) and mMCP-4–/– (H and J) animals were stained with an antiserum toward fibronectin. Scale bars, 250 µm.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The data presented here indicate a central role for mMCP-4 in the regulation of MMP-2 and -9 activities. Because mMCP-4 most likely represents the functional homologue to the only human chymase identified (29), we may propose a similar role for the human chymase in the regulation of the corresponding human MMPs. Although the present data establishes that MMP-9 and -2 activation is strongly chymase-dependent, we cannot with certainty define the mechanism by which this occurs. The most straightforward explanation would be that pro-MMP-9 and pro-MMP-2 are substrates for mMCP-4. Indeed, because several studies have shown that pro-MMP-9 is a substrate for chymase in vitro (21, 28), we favor the notion that pro-MMP-9 is also a substrate for mMCP-4 in vivo.

It was shown previously that dog {alpha}-chymase activates the canine pro-MMP-9 by cleaving the Phe88–Gln89 and Phe91–Glu92 bonds (21). Cleavage after aromatic side chains is a characteristic feature of chymotrypsin-like enzymes such as the MC chymases. Importantly, the Phe88–Gln89 and Phe91–Glu92 sequences, as well as the amino acid sequences surrounding these sites, are conserved between dog, human, and mouse pro-MMP-9. Hence, we may propose that mMCP-4 is likely to catalyze the corresponding cleavages in murine pro-MMP-9. Interestingly, it is known that pro-MMP-9 activation, e.g. catalyzed by stromeolysin, matrilysin collagenase, and trypsin, can also occur through cleavage of the Arg87–Phe88 bond (30, 31). Apparently, enzymatic activation can thus be accomplished by cleavage at either of these neighboring sites. In case of pro-MMP-2, a number of studies have shown that activation of the protease is accomplished by cleavage of the Asn80–Tyr81 or Asn82–Phe83 bonds (32–34). Because cleavage after an Asn residue is not consistent with the chymotrypsin-like substrate specificity carried by the MC chymases, it is not likely that mMCP-4 causes activation of pro-MMP-2 by cleavage of this bond. It is more likely that mMCP-4 cleaves at the Tyr81–Asn82 bond downstream of the Asn80–Tyr81 site, implying that cleavage of this bond also generates active protease. However, we cannot exclude the possibility that mMCP-4-dependent activation of pro-MMP-2 is indirect, i.e. that another protease than mMCP-4 performs the activating cleavage but that this protease, in turn, is dependent on mMCP-4. Clearly, further work will be required to determine the mechanism behind the partial dependence of pro-MMP-2 on mMCP-4 for activation.

A plausible explanation for the enhanced levels of connective tissue proteins in the mMCP-4–/– tissues is that MMPs normally control the levels of these components, either directly by proteolysis or indirectly by activating other enzymes that execute proteolytic processes, and that the reduction in the levels of active MMPs thus causes an accumulation of connective tissue proteins. In line with an indirect scenario, it has been shown that MMP-2 can participate in the activation of procollagenase-3 (35). However, we cannot exclude MMP-independent effects of mMCP-4 on the connective tissue components. For example, we have shown previously that fibronectin is a substrate for mMCP-4 (36), and it has been shown that MC chymase can degrade type I procollagen (37). Furthermore, chymase has been reported to activate human interstitial procollagenase by cleavage of the Leu83–Thr84 bond (38). The present data thus indicate a role for MC chymase in maintaining connective tissue homeostasis by regulating, either indirectly or directly, the levels of various connective tissue components.

MC chymases have, as noted above, been implicated in a variety of disorders. However, the mechanism of action in vivo has not been determined for any of the MC proteases, i.e. the in vivo substrates for the MC proteases have not been identified. The present report provides, to our knowledge for the first time, information on the molecular level on how any of the MC proteases acts in vivo. Because MMP-2 and -9 have been strongly implicated in similar disorders as those that have been linked to chymase, mMCP-4-mediated activation of pro-MMP-2 and -9 may provide a link between MCs and processes that are regulated by MMPs. Furthermore, we may propose that in therapeutic regimens aiming at the inhibition of MMP activities, an even more effective treatment may include targeting of the MMP activators, e.g. chymase.


   FOOTNOTES
 
* This work was supported by grants from the AgriFunGen program at the Swedish University of Agricultural Sciences, the Swedish Research Council, Formas, and the King Gustaf V 80th Anniversary Fund. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§ Present address: Dept. of Molecular Biology and Biochemistry, Faculty of Sciences, University of Malaga, Campus de Teatinos, s/n 29071 Malaga, Spain. Back

|| To whom correspondence should be addressed. Tel.: 46-18-4714090; Fax: 46-18-550762; E-mail: Gunnar.Pejler{at}bmc.uu.se.

1 The abbreviations used are: MC, mast cell; MMP, matrix metalloprotease; mMCP, mouse mast cell protease; NDST, N-deacetylase/N-sulfotransferase; WT, wild type. Back


   ACKNOWLEDGMENTS
 
We are grateful to Eva Hellmén for helpful discussions.



   REFERENCES
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 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

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