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INFLAMMATION AND IMMUNOPHARMACOLOGY

Nitric Oxide and Cyclic GMP Increase the Expression of Matrix Metalloproteinase-9 in Vascular Smooth Muscle

Departments of Medicine (M.M.-P., A.D.B., I.M.) and Oncology (KG., CC.) University of Alberta, Edmonton, Alberta, Canada; Department of Integrative Biology and Pharmacology, University of Texas Health Sciences Center (M.W.R.), Houston, Texas; and The Cross Cancer Institute (K.G., C.C.); and Pulmonary Research Group (M.M.-P., A.D.B.), Edmonton, Alberta, Canada

Received February 10, 2003; accepted August 5, 2003.

	Abstract

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Interactions and possible cross talk between inducible nitricoxide synthase (iNOS), cyclooxygenase-2 (COX-2) and matrix metalloproteinase-9 (MMP-9), were studied in rat aortic vascular smooth muscle cells stimulated with bacterial lipopolysaccharide (LPS), interferon- (IFN-), and phorbol 12-myristate13-acetate (PMA). The expression and activity of iNOS, COX-2, and MMP-9 were characterized at the transcriptional, protein, and enzyme activity levels. The NOS inhibitor N-nitro-L-arginine methyl ester (L-NAME) was used to investigate the effects of NO on COX-2 and MMP-9 at the transcriptional level. The measurements of mRNAs for these enzymes using real-time polymerase chain reaction (PCR) showed that COX-2 mRNA was up-regulated 2.3-fold, whereas MMP-9 mRNA up-regulation was 11.7-fold in the presence of LPS, IFN-, and PMA. Real-time PCR results indicated that L-NAME exerted an inhibitory effect on COX-2 and MMP-9 mRNA synthesis. Both superoxide dismutase (SOD) and the SOD mimetic Mn(III)tetrakis(1-methyl-4-pyridyl)porphyrin pentachloride (MnTMPyP) did not modify significantly the up-regulation of these enzymes, indicating that neither superoxide nor peroxynitrite are involved in this mechanism. Furthermore, NO-mediated up-regulation of MMP-9 was cGMP-dependent since 1H-[1,2,4] oxadiazolo[4,3-a]quinoxalin-1-one (ODQ), an inhibitor of soluble guanylate cyclase, blocked, in a concentration-dependent manner, the increased expression of MMP-9, an effect reversed by 8-bromo-cGMP, a soluble analog of cGMP. Our findings suggest that NO and cGMP are necessary to up-regulate the expression of MMP-9.

The deleterious effects of LPS are often mediated by the enzymatic activity of LPS-inducible proteins, such as inducible nitric-oxide synthase (iNOS), cyclooxygenase-2 (COX-2), and matrix metalloproteinase-9 (MMP-9). Initially, these enzymes are induced in a highly organized fashion to compensate for the damaging effects of LPS, maintain homeostasis, and to contribute to the systemic inflammatory response. However, the overwhelming concentrations of these inducible enzymes can become harmful to the body, contributing to multiorgan dysfunction and death. The increased expression of iNOS, for example, results in massive vasodilatation and hypotension (Gomez-Jimenez et al., 1995; Titheradge, 1999). Additionally, fast reaction of NO with superoxide leads to the highly reactive species peroxynitrite (ONOO^–) (Beckman et al., 1990; Beckman and Koppenol, 1996). The latter is thought to be responsible for many deleterious effects of NO, acting as an oxidant itself, or leading to the formation of other reactive species (Szabo et al., 1996). Moreover, COX-2 up-regulation leads to overproduction of thromboxane and prostaglandin E₂ (PGE₂), which have been implicated in the pathogenesis of septic shock (Ermert et al., 2000; Fischer et al., 2000; Strong et al., 2000). In the presence of LPS, cells like neutrophils secrete considerable amounts of latent MMPs, including MMP-9 (Albert et al., 2003). Reactive oxygen species produced by the same cell type lead to the chemical activation of MMP-9, and as a consequence, degradation of the vascular wall takes place (Opdenakker, 2001). Other lines of evidence from mice and humans have also correlated MMP-9 levels with shock conditions (Pugin et al., 1999; Dubois et al., 2002; Albert et al., 2003).

Despite solid evidence that iNOS, COX-2, and MMP-9 are involved in the pathogenesis of cellular damage caused by LPS, the interactions (cross talk) between these enzymes are unclear. We found that the simultaneous induction of these enzymes could be triggered by IFN-, phorbol 12-myristate13-acetate (PMA), and LPS in rat vascular smooth muscle (VSM) cells, allowing us to study the hypothesis that these three enzymes cross talk to each other. The production of these three enzymes was studied at the transcriptional, protein, and enzyme activity levels. The results of our experiments provide evidence that iNOS up-regulates COX-2 and MMP-9 gene expression in VSM.

	Materials and Methods

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Materials. Polyclonal antibodies against iNOS and COX-2 were obtained from BD Transduction Laboratories (Basingtoke, Hampshire, UK) and Santa Cruz Biotechnology, Inc. (Santa Cruz, CA), respectively. Monoclonal antibodies against MMP-9 were obtained from OncogeneScience (Cambridge, MA). RNeasy Mini Kit was obtained from QIAGEN (Mississauga, ON, Canada).

Cell Culture. Rat aortic VSM (A7r5) were obtained from American Type Culture Collection (Manassas, VA). Cells were cultured in 75-cm² flasks in a humidified atmosphere with 5% CO₂ at 37°C. The medium was Dulbecco's modified eagle's medium supplemented to contain 4.5 g/liter glucose, 1.5 g/liter sodium bicarbonate, 10% fetal bovine serum, gentamycin sulfate (0.05 mg/ml), penicillin G (0.06 mg/ml), and streptomycin sulfate (0.01 mg/ml). Cells were grown until they were confluent and then treated for 12 h with a cocktail containing 10 ng/ml IFN-, 1 nM PMA, and 10 µg/ml LPS in 15 ml of 1% fetal bovine serum (activating cocktail). When treating with dexamethasone (Dex), cells were preincubated with this chemical for 1 h before induction with the activating cocktail. Dex concentrations of 1.0, 3.0, and 10 µM were tested. At a concentration of 1 µM, Dex did not confer inhibitory effects as detectable by reverse transcription-polymerase chain reaction (RT-PCR). Because 3 µM Dex was found to be a sufficient amount to inhibit transcription of iNOS, COX-2, and MMP-9, all consecutive experiments were performed using 3 µM Dex. When either N-nitro-L-arginine methyl ester (L-NAME; 300 µM), 1H-[1,2,4]oxadiazolo[4,3-a]quinoxalin-1-one (ODQ; 0.05, 0.1, 0.5, 1.0, and 5 µM), or superoxide dismutase (SOD) (0.5, 1, 5, 10, 100 U/ml) were used, these compounds were included in the activating cocktail. Cell harvesting was conducted by scraping cells off plates in the presence of 1 ml of homogenizing buffer (50 mM Tris-HCl, 320 mM sucrose, 1 mM dithiothreitol, 10 µg/ml leupeptin, 10 µg/ml soybean trypsin inhibitor, 2 µg/ml aprotinin, pH 7.4).

Gelatin Zymography. This technique was used to measure pro and active MMP-2 and MMP-9 gelatinolytic activity as previously described (Radomski et al., 1998). After cell extraction, samples were immediately subjected to electrophoresis on 7% SDS-polyacrylamide gel electrophoresis copolymerized with gelatin (2 mg/ml) as the substrate. Independent experiments were performed and run on the same gel. Following electrophoresis, gels were washed in 0.1% Triton X-100 (3x for 20 min). The gels were then incubated for 72 h in the zymography buffer containing 25 mM Tris-Cl, 5 mM CaCl₂, 142 mM NaCl, and 0.5 mM Na₃N to determine the activity of secreted enzymes. After incubation, the gels were stained with 0.05% Coomassie Brilliant Blue G-250 in a mixture of methanol/acetic acid/water (2.5:1:6.5) and destained in 4% methanol with 8% acetic acid. The gelatinolytic activities were detected as transparent bands against the background of Coomassie Blue-stained gelatin. Enzyme activity was assayed by densitometry using a ScanJet 3c scanner and SigmaGel measurement software. The pro and active forms of MMP-9 were identified as bands at 92 and 88 kDa, respectively.

Inducible and Constitutive Nitric-Oxide Synthase Activity Assay (Citrulline Assay). Nitric-oxide synthase activity in A7r5 cell homogenate was assessed by measuring the formation of L-[¹⁴C]citrulline from L-[U-¹⁴C]arginine as previously described (Radomski et al., 1993). Briefly, samples were homogenized by sonication (VibraCell, Danbury, CT) in 1 ml of ice-cold homogenizing buffer followed by centrifugation at 10,000g for 20 min at 4°C. Following centrifugation, 40 µl of supernatant was incubated at 37°C for 20 min in assay buffer (pH 7.4) containing 50 mM KH₂PO₄, 1 mM MgCl₂, 0.2 mM CaCl₂, 1 mM L-citrulline, 20 µM L-arginine, 1.5 mM dithiothreitol, 1.5 mM NADPH, 10 µM tetrahydrobiopterin, 10 µM FAD, 10 µM FMN, and 0.5 µCi/ml L-[U-¹⁴C]arginine. The specificity of L-arginine conversion by NOS to L-citrulline was further confirmed using 1.2 mM N-nitro-L-arginine methyl ester, a selective inhibitor of NOS. Additionally, 1.5 mM EGTA, a calcium chelating agent, was used to differentiate between Ca²⁺-dependent and -independent isoforms of NOS. All enzyme activities were expressed as picomoles of product generated per minute per milligram of protein. The limit of detection of this method was 0.05 pmol/min/mg of protein.

Griess Assay. Formation of was measured as previously described (Gilchrist et al., 2002). Briefly, 50 µl of supernatant was measured by the Griess reaction. Results were expressed as micromolar concentration per 10⁶ cells following incubation for 6, 12, 24, 48, and 72 h. Equal volumes of cell-free supernatant and Griess reagent (1% sulfanilamine, 0.1% N-(1-naphyl)-ethylene-diamine dihydrochloride, and 2.5% H₃PO₄) were mixed. NaNO₂ was used as a standard. Plates were read on a V_max kinetic microplate reader (Molecular Devices Corp., Sunnyvale, CA) at 540 nm.

Live-Cell Fluorescence Determination of Intracellular NO. NO production by A7r5 was assayed using DAF-FM, a cell-permeable NO-sensitive fluorescent dye, as previously described (Grisham et al., 1999). Cells were incubated for 1 h with 10 µM DAF-FM, prior to visualization.

Cyclooxygenase-2 Enzyme Immunoassay. The activity of COX-2 was measured using a prostaglandin E₂ enzyme immunoassay kit (Amersham Biosciences Inc., Piscataway, NJ) according to manufacturer's instructions. The amounts of PGE₂ were expressed in picrograms per well of PGE₂ with each well containing a cell concentration of 10⁵.

Immunoblot Detection. The cells were harvested and homogenized in the homogenization buffer. The homogenates were subjected to 7% SDS-polyacrylamide gel electrophoresis (Radomski et al., 1998), and proteins were identified using anti-MMP-9 antibodies (0.2 µg/ml; OncogeneScience), anti-COX-2 antibodies (2 µg/ml; Santa Cruz Biotechnology, Inc.), and anti-iNOS antibodies (0.125 µg/ml; BD Transduction Laboratories).

RT-PCR and Quantitative Real-Time PCR. Experiments were performed as previously described (Szkotak et al., 2001). Total RNA was isolated using the Qiagen RNeasy kit. The RNA was reverse-transcribed with the use of Superscript II reverse transcriptase (Invitrogen, Carlsbad, CA) using oligo(dT) as primers. Thereafter, PCR was performed in 20-µl reactions with the primer pairs (25 µM) described in Table 1. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH)-specific primers were run in all reactions as the internal positive control. The PCR products were amplified for 30 cycles. The selected cycle number was chosen to stop the PCR reaction during its log phase to ensure availability of all reagents. Additional RT-PCR-related information is summarized in Table 1.

Quantitative Real-Time PCR. Real-time PCR experiments were performed using the TaqMan quantitative RT-PCR reaction (Applied Biosystems, Foster City, CA) as previously described (Ritzel et al., 2001). Briefly, an oligonucleotide probe labeled with a fluorescent tag at the 5'-end and a quenching molecule at the 3'-end is hybridized between two PCR primers at the beginning of the reaction. The 5'-nucleotidase activity of Taq polymerase, cleaves the fluorescent dye from the probe during each PCR cycle. The fluorescent signal generated is monitored in real time and is proportional to the amount of starting template in the sample. Real-time PCR products were cloned and sequenced to confirm the identity of the mRNAs. The primer and probe sequences are summarized in Table 2.

Reagents. Other PCR related chemicals such as 100-bp DNA ladder, Superscript II, TaqDNA polymerase, oligo(dT) primer, dNTP, and RNaseOut ribonuclease were obtained from Invitrogen. PMA, ethidium bromide, interferon-, lipopolysaccharide, dexamethasone crystalline, L-NAME (Alexis Corporation, San Diego, CA), N^G-monomethyl-L-arginine, ODQ, and superoxide dismutase were obtained from Sigma Diagnostics, Canada (Mississauga, ON, Canada). Mn(III)tetrakis(1-methyl-4-pyridyl)porphyrin pentachloride (MnTMPyP) and 8-bromo-cGMP were obtained from Calbiochem (Mississauga, ON Canada). L-[U-¹⁴C]arginine was obtained from Amersham Biosciences (Oakville, ON, Canada) and AG50W-X8 resin was obtained from Bio-Rad (Hercules, CA).

Statistics. Results are means ± S.E. of at least three independent experiments. They were analyzed using one-way analysis of variance, and when significant differences were found, the multiple comparison Tukey-Kramer test was used (GraphPad InStat). Values where P < 0.05 were considered statistically significant.

	Results

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Since the objective of this research was to investigate the interactions between COX-2, iNOS, and MMP-9 in VSM we have first tested various combinations of cytokine/inducing agents to cause simultaneous induction of these enzymes. The following combinations have been tested. A concentration gradient of 0.01, 0.1, 1, and 10 µg/ml LPS did not produce a band detectable by zymography or RT-PCR for MMP-9, but constitutive MMP-2 activity was constant for every treatment (data not shown). PMA (100 nM) produced detectable bands for MMP-9 through RT-PCR and zymography but did not activate iNOS mRNA as detectable by RT-PCR (data not shown). IFN- at a concentration of 1 mg/ml did not induce any of the mRNA signals for iNOS, COX-2, or MMP-9 (data not shown). After these preliminary experiments, a combination containing 0.1 ng/ml IFN-, 10 µg/ml LPS, and 1 nM PMA was found to be most effective in enzyme induction and all subsequent experiments were performed with cells stimulated with IFN-, LPS, and PMA. Initial experiments were done using this cocktail and a time course for MMP-9 activity indicated optimal expression by 12 h after activation (data not shown).

A time course of formation using the Griess reaction showed that there was a significant increase in nitrite formation after 24 h (13.9 ± 6.03 µM) and after 48 h (37.6 ± 1.2 µM) of cell activation. Nitrite formation was not detected in control conditions or when activated cells were cotreated with either L-NAME (300 µM) or 1400W (10 nM) (n = 3). A more sensitive fluorimetric assay using the fluorochrome DAF showed NO production by 12 h after cell induction (Fig. 1).

View larger version (131K): [in this window] [in a new window]   Fig. 1. Fluorescence detection of intracellular NO production. Cells were either serum-starved (A and B) or treated with a cocktail containing PMA, LPS, and IFN- (C and D) for 12 h then stained with DAF for 1 h. DAF fluorescence (white in the figure) was analyzed with confocal analysis. Differential interference contrast combined with DAF (B and D) shows cellular morphology (bars = 50 µm).

To study enzyme induction at the transcriptional level, RT-PCR was used. Primers were developed and analyzed using the BLAST sequencing program at GenBank to ensure unique complementation. When A7r5 cells were exposed to the activating cocktail, the mRNAs for iNOS, COX-2, and MMP-9 were markedly up-regulated (Fig. 2, A and B). The PCR products for all templates were identified at their predicted molecular weights of 241, 347, 664, and 528 bp for MMP-9, iNOS, COX-2, and GAPDH, respectively (Fig. 2A). The induction of iNOS, COX-2, and MMP-9 mRNAs was significantly inhibited by 3 µM Dex (Fig. 2, B and C). The housekeeping gene GAPDH was amplified for each experiment and used as the internal positive control. GAPDH mRNA levels were not significantly changed by different treatments (Fig. 2B).

View larger version (60K): [in this window] [in a new window]   Fig. 2. Characterization of the septic shock model at the transcriptional level. A, A7r5 cells were treated with a cocktail of pro-inflammatory substances (under Materials and Methods) to induce the production of iNOS, COX-2, and MMP-9. The 100-bp DNA ladder marker provides an estimate of the size of each PCR product (n = 6). B, RT-PCR was used to investigate the mRNA levels corresponding to each gene. Cells were untreated (Sham), induced with the activating cocktail (Activated), or incubated with dexamethasone (Dex) 1 h before activation. GAPDH was used as the positive control for each experiment (n = 3). C, densitometric analysis for RT-PCR experiments shown in panel B (n = 3). Statistical significance: P < 0.001 (***), ().

To investigate whether protein levels for these enzymes were affected by the treatment, Western blot experiments were conducted using the same experimental conditions. We found that the levels of iNOS, COX-2, and MMP-9 proteins were increased by the treatment with the activating cocktail, and this enhancement was abolished in the presence of dexamethasone (Fig. 3, A–C).

View larger version (30K): [in this window] [in a new window]   Fig. 3. Characterization of the septic shock model at the protein level. To further investigate the expression of MMP-9, iNOS, and COX-2, we estimated protein expression levels using immunoblot. A7r5 cells were untreated (Sham), activated with a cocktail (Activated), or preincubated with dexamethasone 1 h before activation. A, the pro and active form of MMP-9 at 92 and 88 kDa were detected. Active MMP-9 was significantly up-regulated in the activated cells. Pretreatment with dexamethasone significantly reduced MMP-9 induction (n = 3). B, iNOS appeared as a band at 130 kDa. Significant differences were found between the sham and activated groups. Also significant differences were obtained when iNOS induction was inhibited with dexamethasone (n = 3). C, similar results to those in panels A and B were obtained for the COX-2 protein at 74 kDa. A second band at a higher molecular weight also appeared only in activated conditions and was suspected to be due to antibody cross-reactivity (n = 3). Statistical significance: P < 0.01 (**), (); P < 0.001 (***), ().

The enzymatic activities of iNOS, COX-2, and MMP-9 were also characterized. The activity of Ca²⁺-independent NOS in unstimulated (Sham) VSM was 12.4 ± 0.6 pmol/mg of protein/min (Fig. 4A). Cell induction resulted in a significant increase in the activity of this enzyme to 24.4 ± 2.6 pmol/mg of protein/min. Ca²⁺-dependent NOS activity was not detectable under these conditions, indicating that neither nNOS or eNOS were expressed in A7r5.

View larger version (34K): [in this window] [in a new window]   Fig. 4. MMP-9, iNOS, and COX-2 enzymatic activity in A7r5 cells. A7r5 cells were untreated (Sham), activated with a cocktail (Activated), or preincubated with dexamethasone 1 h before activation. A, inducible NOS activity was measured using the citrulline assay. Significant changes between sham and activated conditions were found (P = 0.027), but for iNOS there were no significant changes between sham and Dex treatment. The constitutive NOS activity, also measured with this assay, was not detected after each treatment (n = 3). B, PGE2 formation was monitored using enzyme-linked immunosorbent assay. Significant changes were observed between sham and activated cells, as well as between activated and dexamethasone treatment (n = 4). C, zymography was performed to measure the gelatinolytic activity of MMP-9. The pro and active forms of MMP-9 were detected at 92 and 88 kDa, respectively. At 72 and 62 kDa, the pro and active forms of MMP-2 were also detected. MMP-2 is a constitutive metalloproteinase that remained relatively unchanged despite different treatments. Densitometric analysis for MMP-9 and MMP-2 revealed a highly significant up-regulation in activity of MMP-9 in activated cells, whereas Dex inhibited this up-regulation to near control levels (n = 3). Statistical significance: P < 0.05 (*), (); P < 0.01 (**), (); P < 0.001 (***), ().

The activity of COX-2, as measured by PGE₂ levels, was significantly increased following cell induction (Fig. 4B). The gelatinolytic activity of MMP-9, but not MMP-2, was markedly up-regulated by the activating cocktail (Fig. 4C). Dexamethasone abolished increases in iNOS, COX-2, and MMP-9 activities caused by cell induction (Fig. 4, A-C).

Effects of L-NAME on COX-2 and MMP-9 Transcription. Inhibition of NOS with L-NAME (300 µM) exerted no significant effect on iNOS mRNA expression in activated cells (Fig. 5A). In contrast, COX-2 and MMP-9 transcription was down-regulated by this inhibitor (Fig. 5A). L-NAME did not change the expression levels of the positive control GAPDH. To quantify the changes detected using RT-PCR, real-time PCR analysis was conducted for COX-2 and MMP-9 mRNAs. It was found that COX-2 was up-regulated by 2.3-fold (Fig. 5C) and that this up-regulation could be inhibited to lower than control levels by either Dex or L-NAME. MMP-9 mRNA levels were up-regulated with the activating cocktail by approximately 11.7 times, and they were significantly reduced in the presence of Dex or L-NAME (Fig. 5B).

View larger version (28K): [in this window] [in a new window]   Fig. 5. Effect of L-NAME treatment on levels of mRNA for iNOS, COX-2, and MMP-9. A, RT-PCR was performed using both Dex and L-NAME as pharmacological tools. The positive control GAPDH was constant for each treatment. L-NAME had an inhibitory effect obvious for COX-2 and for MMP-9 (n = 3). B, real-time PCR was performed to validate and quantify the RT-PCR findings. MMP-9 mRNA up-regulation was found to be of 11.7-fold. This effect was inhibited by both dexamethasone and L-NAME to levels relatively close to sham (n = 3). C, COX-2 up-regulation during activation was found to be 2.3-fold, and this effect was inhibited to lower than sham levels when treating with either dexamethasone or L-NAME (n = 3). Statistical significance: P < 0.05 (*), (); P < 0.01 (**), ().

Effects of SOD and MnTMPyP on MMP-9 mRNA. To investigate whether superoxide and possibly peroxynitrite (Patel et al., 1999) were involved in MMP-9 regulation by iNOS the scavenger of superoxide, SOD, was tested on A7r5 cells induced with the activating cocktail. SOD at concentrations of 100, 10, 5, 1, and 0.5 U/ml exerted no significant effects on MMP-9 mRNA induction (Fig. 6A). These concentrations of SOD effectively blocked superoxide generation by xanthine oxidase (Ohara et al., 1993) (data not shown). The membrane-soluble SOD mimetic MnTMPyP did not modify significantly the up-regulation of these enzymes indicating that neither superoxide nor peroxynitrite are involved in this mechanism (Fig. 6B).

View larger version (24K): [in this window] [in a new window]   Fig. 6. MMP-9 gene induction is not mediated by ONOO–. A, an SOD concentration dependence curve was created. Cells were left untreated (Sham) or induced with the activating cocktail containing different concentrations of SOD. No significant variations were observed when comparing maximum activation (0.0 U/ml SOD) with activation in the presence of SOD (0.5–100 U/ml SOD) (n = 3). B, the SOD mimetic MnTMPyP (0–20 µM) did not exert a significant reduction of MMP-9 mRNA relative abundance (n = 3). Statistical significance: P < 0.05 (*).

Effects of ODQ and 8-bromo-cGMP on MMP-9 mRNA Regulation. To study the involvement of cGMP in MMP-9 mRNA regulation, ODQ, an inhibitor of soluble guanylate cyclase was used. ODQ effectively decreased, in a concentration-dependent fashion, the levels of MMP-9 mRNA (Fig. 7A). 8-bromo-cGMP, a soluble analog of cGMP, reversed the effect of ODQ (Fig. 7B).

View larger version (19K): [in this window] [in a new window]   Fig. 7. Nitric oxide up-regulates MMP-9 via a soluble guanylate cyclase-dependent pathway. Relative abundance of MMP-9 mRNA was measured using real-time PCR. A, there was a significant up-regulation of the MMP-9 gene when inducing A7r5 cells with the activating cocktail (Activated). When cells were activated in the presence of ODQ (0.05–5 µM), this resulted in a significant decrease in the MMP-9 mRNA levels. It was shown that at a concentration of 0.5 µM ODQ the activating cocktail did not induce any significant changes in the MMP-9 mRNA levels (n = 3). B, exogenously added 8-bromo-cGMP (0–0.5 mM) reversed the effect of ODQ (0.5 µM). Statistical significance: P < 0.05 (*), ().

	Discussion

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We have investigated the interactions between iNOS, COX-2, and MMP-9, three pro-inflammatory enzymes that are induced in the vasculature during acute inflammatory reactions such as septicemia and septic shock (Beckman et al., 1990; Gomez-Jimenez et al., 1995; Beckman and Koppenol, 1996; Szabo et al., 1996; Pugin et al., 1999; Titheradge, 1999; Ermert et al., 2000; Fischer et al., 2000; Strong et al., 2000; Opdenakker, 2001; Dubois et al., 2002; Albert et al., 2003). We have studied these interactions using rat smooth muscle vascular cells, which were induced by LPS, IFN-, and PMA. Under the experimental conditions used, the inducing agents resulted in up-regulation of enzyme expression at transcriptional (mRNA), translational (protein), and activity (product generation or substrate degradation) levels. We have further validated our model of simultaneous induction of iNOS, COX-2, and MMP-9 using dexamethasone, a potent glucocorticoid known to inhibit many pathways of inflammation. As expected, this drug treatment efficiently reduced the expression of iNOS, COX-2, and MMP-9 (Figs. 2B and 3, A–C).

We then used RT-PCR to study the effects of blockade of NOS with L-NAME on MMP-9 and COX-2 mRNA levels. Inhibition of NOS with L-NAME down-regulated the expression of COX-2. This is not surprising since NO is known to stimulate the activity of cyclooxygenase (Salvemini et al., 1993) and NO stimulates COX-2 mRNA expression in rat mesangial cells (Tetsuka et al., 1996). Interestingly, we have found that MMP-9 mRNA up-regulation was inhibited by L-NAME showing that MMP-9 induction is NO-dependent by a NOS-dependent mechanism. This observation was confirmed and quantified by real-time PCR experiments (Fig. 5A).

Initially it was thought that NO effects could potentially be mediated through either nitrosylation, oxidation, nitration, or a combination of these reactions, since these nonspecific reactions are common in inflammatory settings. Peroxynitrite is one of the most reactive biochemical fates of NO and in many studies has been described as a signaling molecule that acts through tyrosine nitration (Patel et al., 1999). For this reason, SOD and membrane-soluble SOD mimetic MnTMPyP were used to investigate ONOO^– effects on the MMP-9 induction pathway. The treatments did not modify the MMP-9 gene control mechanism suggesting that ONOO^– was not involved in this process (Fig. 6, A and B). A second pathway by which NO could be acting was through soluble guanylate cyclase. Our results indicated that in the presence of ODQ, an inhibitor of soluble guanylate cyclase, the MMP-9 mRNA returned to its basal levels. Furthermore, the effect produced by ODQ was reversed by exogenous 8-bromo-cGMP, a soluble analog of cGMP (Fig. 7B). These data imply that the soluble guanylate cyclase was the main pathway for NO-dependent MMP-9 gene up-regulation.

Recently, in contrast to our results on enhancement of MMP-9, Eberhardt et al. (2000) using rat mesangial cells found that endogenous or exogenously provided NO decreased MMP-9 gene expression. They also showed that administration of another inhibitor of NOS, N^G-monomethyl-L-arginine increased MMP-9 mRNA expression. NO can reduce t_1/2 life of MMP-9 mRNA (Eberhardt et al., 2002) as well as S-nitrosylate nuclear factor-B (Marshall and Stamler, 2001), two mechanisms by which NO may modulate MMP-9 expression. By contrast NO has been found to increase MMP-9 activity through S-nitrosylation (Gu et al., 2002).

Interestingly, pharmacological studies using L-NAME (inhibitor of NOS) and AMD6221 [ruthenium(III) polyaminocarboxylate complex; NO scavenger] yielded contradictory data on MMP-9 activity. The administration of L-NAME to neonatal hyperoxic rats increased activity of MMP-9 (Radomski et al., 1998), whereas NO scavenging decreased MMP-9 activity elevated by extracorporeal circulation (Mayers et al., 2003). These complex effects of inhibition by NO of expression and activity of MMP-9 likely reflect the dual nature of NO as inflammatory-inhibitor and inflammatory-mediator (Droge, 2002).

The conflicting results on enhancement or inhibition of MMP-9 illustrate the complexity of the interactions of NO with cell-signaling cascades. Factors such as cell type, stimulation time, and stimulatory cocktail likely contribute to some of the differences observed. In our study, we examined the expression of MMP-9 mRNA after 12 h of treatment using a cocktail containing LPS, IFN-, and PMA and observed that serum-starved cells even in the absence of the stimulatory cocktail (control conditions) began to synthesize MMP-9 mRNA by 24 h. These results were further validated by a time course of MMP-9 activity, using zymography, in which 12 h was clearly the optimal induction time as described above. It is possible that in settings where cells have been stimulated for 24 to 72 h, as done in several of the studies outlined above (Eberhardt et al., 2000, 2002), the levels of NO are different than those we have observed. A time course of formation using the Griess reaction showed that there was a significant increase in nitrite formation after 24 and 48 h of cell activation, whereas nitrite formation could not be detected in control conditions or when activated cells were cotreated with either L-NAME or 1400W.

Thus, NO may function through alternative pathways to reduce MMP-9 levels, e.g., through oxidation of MMP-9 mRNA. Our findings could be interpreted in terms of early and late NO-mediated events, in which NO concentration and exposure time are critical in dictating outcomes. Future experiments must attempt to clarify the underlying causes of the contradictory results in the literature.

The list of genes that are up-regulated via soluble guanylate cyclase-mediated pathways includes only COX-2, tumor necrosis factor, PAI-1, FLT-1, MKP-1 (Pfeilschifter et al., 2001), and now MMP-9. The exact mechanism by which cGMP exerts its transcription regulatory functions has not been fully elucidated. Activated soluble guanylate cyclase synthesizes cGMP, which in turn alters the activity of three main target proteins: 1) cGMP-regulated ion channels, 2) cGMP-regulated phosphodiesterases, and 3) cGMP-dependent protein kinases (PKG). Several lines of evidence appear to involve PKG as the mediator of soluble guanylate cyclase action on the MMP-9 gene (Gudi et al., 1996, 1997, 1999). More recent studies have shown that PKG-mediated gene regulation takes place via activation of members of the mitogen-activated protein kinase protein family including Raf1, extracellular signal-regulated kinase 1/2, and synthesis of c-Jun. Interestingly, protein kinase C (PKC) has been shown to regulate transcription of MMP-9 gene via stimulation of extracellular signal-regulated kinase 1/2 pathway (Genersch et al., 2000; Lee et al., 2003). This PKC-like behavior may partially explain how increased soluble guanylate cyclase activity up-regulates MMP-9 gene. However, under the conditions of our experiments, stimulation of PKG but not PKC appears to be necessary for MMP-9 gene induction since, despite the continuous presence of PMA a known stimulator of PKC (Ron and Kazanietz, 1999), the blockade of soluble guanylate cyclase pathway with ODQ abolished MMP-9 gene transcription.

The precise mechanisms involved in soluble guanylate cyclase-mediated MMP-9 gene induction remain to be studied. It is important to note that our current understanding of regulatory mechanisms of the MMP-9 promoter is very limited. Transcription Elements Search System (TESS) analysis of 1,300 bp of the rat MMP-9 promoter reveals a complex picture of multiple binding sites for more than 200 different transcription factors. We are currently investigating the relevance of a novel PKG/protein kinase A target, which we have identified as a possible MMP-9 repressor.

The results of our experiments show that the NO-cGMP pathway plays a crucial role in MMP-9 gene regulation. The pharmacological significance of our findings remains to be studied.

	Acknowledgements

 
We thank Ada Chung, Dr. Anna Radomska, Dr. Paul Jurasz, Haide Razavy and Mark Gilchrist for invaluable technical help and support.

	Footnotes

 
This work was supported by funds from the Canadian Institutes for Health Research (M.W.R. is a CIHR scientist). A.D.B. is the holder of the AstraZeneca Canada, Inc. Chair in Asthma Research.

DOI: 10.1124/jpet.103.050385.

ABBREVIATIONS: LPS, lipopolysaccharide; NO, nitric oxide; iNOS, inducible NO synthase; COX-2, cyclooxygenase-2; MMP, matrix metalloproteinase; PGE₂, prostaglandin E₂; IFN-, interferon ; PMA, phorbol 12-myristate 13-acetate; VSM, vascular smooth muscle; Dex, dexamethasone; RT-PCR, reverse transcription-polymerase chain reaction; L-NAME, N-nitro-L-arginine methyl ester; ODQ, 1H-[1,2,4]oxadiazolo[4,3-a]quinoxalin-1-one; SOD, superoxide dismutase; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; bp, base pair(s); MnTMPyP, Mn(III)tetrakis(1-methyl-4-pyridyl)porphyrin pentachloride; DAF-FM, diaminofluorescein-fluorometry; PKG, cGMP-dependent protein kinase.

Address correspondence to: Dr. A. Dean Befus, Chair in Asthma Research, AstraZeneca Canada Inc., Glaxo-Heritage Asthma Research Laboratories, Department of Medicine, University of Alberta, Edmonton, AB, T6G 2S2, Canada. E-mail: dean.befus{at}ualberta.ca

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