Pulmonary and Critical Care Division, Department of Medicine, Atlanta Veterans Affairs Medical Center and Emory University School of Medicine, Atlanta, Georgia 30033
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ABSTRACT |
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Mycobacterium tuberculosis
(Mtb) infection induces the expression of matrix metalloproteinase-9
(MMP-9) in mouse lungs. In cultured human monocytic cells, Mtb bacilli
and the cell wall glycolipid lipoarabinomannan (LAM) stimulate high
levels of MMP-9 activity. Here, we explore the cellular mechanisms
involved in the induction of MMP-9 by Mtb. We show that infection of
THP-1 cells with Mtb caused a fivefold increase in MMP-9 mRNA that was associated with increased MMP-9 activity. MMP-9 induction was dependent
on microtubule polymerization and protein kinase activation and was
associated with increased DNA binding by the transcription factor
activator protein-1 (AP-1), which appeared to be important for MMP-9
expression. We then explored the surface molecules potentially involved
in Mtb induction of MMP-9, focusing on ligands of the mannose and
-glucan receptors. MMP-9 activity was induced by the mannose
receptor ligands mannan, zymosan, and LAM, whereas the
-glucan
receptor ligand laminarin was not effective. The most active inducers
of MMP-9 activity were the particulate ligand zymosan and LAM.
Pretreatment of cells with an anti-mannose receptor monoclonal
antibody, but not anti-complement receptor 3, decreased the induction
of MMP-9 activity by Mtb bacilli. Together, these results suggest that
MMP-9 induction by Mtb occurs by receptor-mediated signaling mechanisms
involving the binding of mannosylated ligands to mannose receptors, the
modulation by cytoskeletal elements such as microtubules, the
activation of protein kinases, and transcriptional activation by AP-1.
matrix metalloproteinases; mannose receptor; protein kinases; cytoskeleton; activator protein-1; Mycobacterium
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INTRODUCTION |
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ONE OF THE MOST DEVASTATING consequences of infection with Mycobacterium tuberculosis (Mtb) is the formation of caseating granulomas followed by tissue destruction with liquefaction causing cavity formation (19, 26). Despite the predominant role these events play in the progression of disease and dissemination of mycobacteria, very little is known about how Mtb infection elicits this destructive process. Early studies have indicated that cellular hypersensitivity responses to Mtb are responsible for the massive caseous tissue necrosis and liquefaction observed during Mtb infection (13, 69). Proteolytic damage by macrophage-secreted proteases has been implicated because macrophages are the predominant cell in Mtb granulomas and contain a large number of proteases capable of degrading the extracellular matrix (21, 22, 47, 59, 61, 62). Together, these studies suggest that, on entering the lung, Mtb elicits a host response characterized by the expression and secretion of proteolytic enzymes capable of tissue remodeling.
Recently, we demonstrated that infection of C57BL6 mice with Mtb Erdman induced the differential expression in lung of the matrix metalloproteinases (MMPs) MMP-9 (gelatinase B) and MMP-2 (gelatinase A) (46). These gelatinases are members of the MMP family that constitute a group of zinc- and calcium-dependent endopeptidases capable of degrading connective tissue matrixes (6, 37, 38). Many cell types, including monocytes and macrophages, secrete MMPs as inactive zymogens that require activation. Extracellularly, the activity of MMPs can be inhibited by a related group of molecules termed the tissue inhibitors of matrix metalloproteinases (TIMPs). In addition to working in connective tissue degradation, these proteases have been found to be involved in other related processes, including regulation of cell migration and angiogenesis (6, 38). Alterations in the relative expression and/or function of MMPs and TIMPs have been described in many lung disorders characterized by inflammation, tissue fibrosis, and destruction, including emphysema (14), cystic fibrosis (15), sarcoidosis (17, 39), bronchiectasis (52), hypersensitivity pneumonitis (34), and idiopathic pulmonary fibrosis (34).
The expression and function of MMPs in pulmonary tuberculosis have not been studied extensively. We reported that the production and activation of the gelatinases MMP-2 and MMP-9 increased in the lungs of mice infected with Mtb (46). Furthermore, we demonstrated that inducers of MMP-9 expression in Mtb are found in long-term culture filtrates and detergent extracts of bacilli and that they could be separated by column chromatography. One cell wall component that is capable of inducing MMP-9 activity is the cell wall glycolipid lipoarabinomannan (LAM) (10, 46). These studies suggest that the induction of host MMPs by Mtb could play significant roles in the pathogenesis of mycobacterial infection.
The present study explores the cellular mechanisms by which Mtb bacilli induce MMP-9 expression in human cells of monocyte and macrophage lineage (i.e., THP-1 cells). We demonstrate that the induction of MMP-9 by Mtb requires the arrangement of microtubular elements, the activation of protein kinase-dependent pathways, and the binding of the transcription factor activator protein-1 (AP-1) to promoter elements in the MMP-9 gene. These events are triggered by the binding of mannosylated glycans of the cell envelope of Mtb to surface receptors on THP-1 cells, such as mannose receptors.
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MATERIALS AND METHODS |
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Reagents.
The mitogen-enhanced kinase 1 (MEK1) inhibitor PD-98059 was purchased
from New England Biolabs (Beverly, MA). Colchicine, cytochalasin D,
calphostin C, zymosan, yeast mannan, and laminarin were purchased from
Sigma Chemical (St. Louis, MO). Monoclonal antibody mouse anti-human
mannose receptor was purchased from RDI (Flanders, NJ), and monoclonal
antibody mouse anti-human complement receptor 3 (CR3; CD11b -chain)
was purchased from Santa Cruz Biotechnology (Santa Cruz, CA). LAM
glycolipids were obtained from Dr. John Belisle (Colorado State Univ.,
Fort Collins, CO) through National Institutes of Health contract TB
Research Materials and Vaccine Testing. MMP-9 and MMP-2 were isolated
and purified by gelatin chromatography as described before
(60). Polystyrene beads (5.3 µm) were purchased from
Duke Scientific (Palo Alto, CA). All other reagents were purchased from
Sigma Chemical or Fisher Scientific (Pittsburgh, PA) unless otherwise specified.
Bacterial cultures and growth conditions.
The Mtb laboratory strains H37Ra (ATCC 25177) and Erdman (ATCC 35801)
were obtained from the American Type Culture Collection (Rockville,
MD). Mycobacteria were grown in Middlebrook 7H9 broth supplemented with
DADC enrichment media (BD Biosciences) and 2% glucose and containing
1% glycerol. Growth occurred at 37°C with slow shaking to
mid-log growth phase (7-10 days) or to stationary phase (21 days). Bacteria were passaged in broth cultures once and
quantified with McFarland equivalence turbidity standards (Remel,
Lenexa, KS). Cultures of 106-108
bacilli/ml were used immediately or stored at 70°C in 7H9 broth containing 20% glycerol.
Cell culture and Mtb infection.
The THP-1 human monocytic cell line (ATCC) was grown at 37°C and 5%
CO2-95% O2 in RPMI 1640 supplemented with 10%
heat-inactivated fetal bovine serum and 1% antibiotic-antimycotic (100 U/ml penicillin G sodium, 100 U/ml streptomycin sulfate, and 0.25 g/ml
amphotericin B). The cells (2 × 106/ml) were washed
in serum-free media and seeded in 15-ml T flasks containing serum-free
media (Cellgro). Cells were infected with attenuated Mtb H37Ra or
virulent Mtb Erdman strains at a multiplicity of infection (MOI) of 10. Cells and Mtb bacilli were incubated for 24 h at 37°C and 5%
CO2-95% O2. Afterward, a 20-µl aliquot of
culture was collected to determine cell viability by trypan blue
exclusion. The infected cell cultures were centrifuged at 500 g, and the culture supernatant was collected and filtered through a 0.22-µm filter. Cell pellets were resuspended in 0.5 ml of
3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate buffer at
4°C, sonicated, and centrifuged at 12,000 g for 5 min, and
the supernatants were collected and filtered through a 0.22-µm filter. Total protein in the culture supernatants and detergent cell
extracts was determined by the Bradford method (8), and aliquots were stored frozen at 70°C until analyzed for
gelatinolytic activity by gelatin zymography.
Treatment of THP-1 cells. In some experiments, THP-1 cells were pretreated with 6.25 µM cytochalasin D (an inhibitor of actin microfilament polymerization), 50 µM colchicine (a tubulin inhibitor), 0.1 nM of calphostin C (protein kinase C inhibitor) (29), or 50 µM of PD-98059 MEK1 inhibitor (2) for 1 h before infection with Mtb. The doses for the above agents were chosen based on optimal doses recommended in the literature. In other experiments, THP-1 cells (2 × 106 cells in 1.0 ml) were seeded onto 24-well Costar plates in serum-free media and treated with mannan or laminarin (100 µg/ml to 1.0 mg/ml), zymosan (1.0-100 µg/ml), LAM (0.25-5.0 µg/ml), or uncoated polystyrene beads (0.1-1 mg/ml) for 24 h. Similar experiments were done in which THP-1 cells were incubated with the various agents for 1 h and infected with Mtb bacilli for 24 h. For antibody-inhibition studies, cells were treated with monoclonal antibodies (anti-mannose receptor, anti-CD11b) or normal mouse IgG for 1 h and infected with Mtb bacilli (at MOI of 10) for 24 h. After incubation, cells were collected and centrifuged at 500 g, and the culture supernatant was processed for analysis by gelatin zymography.
Gelatin zymography. Gelatin zymography was performed by using a 9% SDS-PAGE gel saturated with 1 mg/ml gelatin (Sigma Chemical, 300 bloom) as previously described (60). Samples with equal protein concentration (10 µg) were loaded onto the gel and electrophoresed at a constant 150 V for 1.5 h. The gels were incubated for 1 h at room temperature in 2.5% Triton X-100, followed by an overnight incubation at 37°C in gelatinase substrate buffer (50 mM Tris, 10 mM CaCl2, and 0.02% NaN2, pH 8.0). The gels were stained with 0.5% Coomassie blue followed by subsequent destaining with 50% methanol. The gels were dried onto cellophane and scanned under a densitometer for determination of gelatinolytic activity.
RT-PCR. The determination of mRNA levels was done by a semiquantitative bioluminescence-based RT-PCR assay as previously described (43, 57, 67). RNA was extracted from lung tissue and cultured cells by using the reagent RNAzol (Tel-test, Friendswood, TX) followed by purification with serial extractions in chloroform, isopropanol, and ethanol. Biotinylated PCR primers and digoxigenin-labeled probes for human MMP-9 and TIMP-1 were synthesized by Genosys Biotechnologies (The Woodlands, TX). Primers and probes for MMP-9 and TIMP-1 were synthesized based on GenBank published sequences. They were as follows: MMP-9 sense primer, CAGGGAGATGCCCATTTC; MMP-9 antisense primer, CTTTAGTGGTGCAGGCAG; MMP-9 probe, AGGTGAAGGGAAAGTGAC; TIMP-1 sense primer, ATCATCAGGAAGCTGGTG; TIMP-1 antisense primer, CAGCTACAAGAGACCCTG; TIMP-1 probe, GAATGGTGTGGTGATGCA.
Reverse transcription of mRNA was done by using the Promega Access RT-PCR kit (Promega, Madison, WI). The reaction mixture consisted of RNA sample (1 µg), 2.5 µl deoxynucleotide triphosphate mix (2.5 mM), 5.0 µl RT buffer (250 mM Tris · HCl, pH 8.3, 375 mM KCl, and 15 mM MgCl2), 2.0 µl dithiothreitol (0.1 M), 0.5 µl RNase inhibitor (20 U), 2.0 µl random hexamer oligonucleotides, and diethyl procarbonate-treated sterile water, in a total volume of 24 µl. The reaction mixture was heated at 70°C for 5 min and cooled in ice, and 1 µl (50-100 U) of Superscript II RT was added. The first-strand cDNA synthesis reaction was carried out at 42°C for 1 h. After cDNA synthesis, the reaction mixture was diluted by adding 175 µl of sterile water and a 5-µl aliquot withdrawn and added to PCR reaction tubes containing the following: 2 µl of 10× PCR buffer (100 mM Tris · HCl, pH 8.3, 500 mM KCl, 15 mM MgCl2, and 0.01% wt/vol gelatin), 2 µl of each biotinylated primer set (2 µM), 4 µl deoxynucleotide triphosphate mix (2.5 mM), and 0.5 µl (0.5 U) of Taq DNA polymerase (AmpliTaq, Perkin Elmer) in a total volume of 25 µl. The thermal cycling parameters consisted of 95°C (3 min); 35 cycles of 94°C (1 min), 54°C (0.5 min), and 72°C (1 min); and a final extension at 72°C (7 min). After amplification, the biotinylated PCR products were denatured by addition of denaturing solution (6.5 µl of 1 M NaOH, and 0.2 M EDTA), neutralized by addition of neutralizing buffer (31.5 µl of 0.15 M sodium phosphate, pH 6.0), and added to streptavidin-coated plates (Boehringer Mannheim). To each well, we added 100 µl of hybridization buffer (62.5 mM Na2HPO4, 94 mM citric acid, 15 mM sodium azide, 0.0625% BSA, 94 mM NaCl, 0.0125% Tween 20, and 10 mM MgCl2) containing 2 ng of the digoxigenin-labeled probes. Hybridization was done for 2 h at 42°C temperature. After hybridization, the plates were washed four times in wash buffer (PBS; 0.15% Tween 20 and 2 mM EDTA, pH 7.4) and 100 µl of assay buffer (25 M Tris base, 10 mM EDTA, 0.15 M KCl, 15 mM sodium azide, and 2 mg/ml BSA) containing 1 µl of anti-digoxigenin aequorin conjugate (AquaLite, Chemicon, Temecula, CA) was added to each well. The plates were incubated for 30 min at room temperature and washed in wash buffer four times, and 100 µl of trigger buffer were added to each well. Luminescence was measured on a Dynatech ML 3000 Microtiter Plate Luminometer after triggering with 0.1 M calcium. Results were plotted as relative fluorescent units. Standardization reactions were done for theScreening for lipopolysaccharide. Experimental reagents were reconstituted in lipopolysaccharide-free water (Sigma Chemical). All treatment materials and culture media were screened with a limulus-based endotoxin assay with a sensitivity of 0.06 ng/ml (Endotect-Schwarz/Mann Biotech, Cleveland, OH) (23). Reagents used were found to remain endotoxin free throughout all experiments. In contaminated samples, lipopolysaccharide was removed by described methods and retested (42, 45).
Electrophoresis mobility shift assay.
THP-1 cells (1 × 108) were grown in suspension at
37°C in 5% CO2-95% O2 with or without
pretreatment (30 min) with Mtb as described in Cell culture and
Mtb infection. Cells were washed with ice-cold PBS, nuclear
binding proteins were extracted as described (16), and protein concentrations were determined by the Bradford method (8). A double-stranded AP-1 consensus
oligonucleotide (5'-CGCTTGATGAGTCAGCCGGAA) was radiolabeled with
[-33P]ATP by using T4 polynucleotide kinase enzyme.
Nuclear proteins (5 µg) were incubated with the radiolabeled AP-1
oligonucleotide (3-600,000
counts · min
1 · ng
1) for 30 min at room temperature as described previously (45). For
competition reactions, a 50-fold molar excess of unlabeled double-stranded AP-1 oligonucleotide or a mutated AP-1 oligonucleotide with a two-base mismatch (5'-CGCTTGATGAGTTGGCCGGAA) was
added to the reaction. DNA-protein complexes were separated on
6% native PAGE gels (20:1 acrylamide-to-bis-acrylamide ratio) in
low-ionic strength buffer (22.25 mM Tris borate, 22.25 mM boric acid,
and 500 mM EDTA) for 2-3 h at 4°C at 10 V/cm. Gels were fixed in
a 10% acetic acid and 10% methanol solution for 10 min, dried
under vacuum, and exposed to X-ray film. The radiolabeled
DNA-protein complexes were extracted from gels and quantified by
scintillation counter.
DNA transfection of THP-1 cells. Electroporation of competent THP-1 human monocytes with a double-stranded 21-bp AP-1 consensus oligonucleotide (5'-CGCTTGATGAGTCAGCCGGAA) was done as previously described (48). Briefly, THP-1 cells were washed twice with cold PBS and added to serum-free media supplemented with 10 mM dextrose and 0.1 mM dithiothreitol to a final concentration of 6 × 107 cells/ml. THP-1 cells (5 × 106 cells) were added to electroporation cuvettes (0.4-cm gap) with or without 20 µg of the oligonucleotide and electroporated at 400 V and 1,075 µF (Gene Pulser II Electroporation System, Bio-Rad, Hercules, CA). The cells were then resuspended in RPMI 1640 media for 30 min, centrifuged, transferred to 75-mm2 tissue culture flasks containing fresh media, and incubated at 37°C and 5% CO2-95% O2. The cells were infected with Mtb bacilli at an MOI of 10 for 24 h, and gelatinolytic activity in the culture supernatant and cell extracts was determined by zymography.
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RESULTS |
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Mtb induces the expression of MMP-9 in THP-1 cells.
Human monocytic THP-1 cells were exposed to Mtb bacilli, after which
the culture supernatants and detergent cell lysates were prepared and
analyzed by gel zymography for the presence of MMP-9 activity and total
RNA was isolated for analysis of MMP-9 mRNA expression by RT-PCR.
Exposure of THP-1 cells to attenuated (H37Ra) and virulent (Erdman) Mtb
strains for 24 h resulted in abundant secretion of MMP-9 activity
(92-kDa band) into the culture media. Consistent with the secreted
nature of MMP-9, little activity was detected in the detergent cell
extract (Fig. 1A). To estimate the amount of MMP-9, secreted, purified MMP-9 (0.4 ng to 0.1 µg in
2-fold increments) was included in the gel zymograms and analyzed by
densitometry scanning, and a concentration curve was prepared (data not
shown). By linear regression analysis (R = 0.9), we estimated that the stimulation of THP-1 cells by Mtb bacilli for 24 h under the conditions described resulted in the secretion of
0.075-0.1 ng MMP-9/µg protein secreted into the culture
media.
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Induction of MMP-9 by Mtb is modulated by cytoskeletal arrangement.
The above results showing that Mtb bacilli induce the expression and
secretion of MMP-9 in THP-1 cells prompted us to investigate the
cellular mechanisms involved. We began by testing the role of the
cytoskeleton. To this end, THP-1 cells were treated with agents that
inhibit microtubule assembly, such as colchicine and taxol, and the
inhibitor of actin microfilament polymerization, cytochalasin D. THP-1
cells were treated with these agents for 30 min before infection with
Mtb Erdman bacilli and were harvested after 24 h for analysis of
MMP-9 activity by gelatin zymography. Figure
2 shows that infection of THP-1 cells
resulted in high levels of MMP-9 activity in culture supernatants,
whereas pretreatment of cells with colchicine and taxol resulted in
total inhibition of Mtb-induced MMP-9 activity. Analysis of cell pellet
detergent extracts showed no MMP-9 activity either (data not shown),
indicating that these agents did not affect MMP secretion.
Interestingly, treatment of THP-1 cells with cytochalasin D did not
inhibit MMP-9 activity but resulted in its induction. Also, a onefold
increase in MMP-9 activity was seen after infection of the cytochalasin D-treated cells, as determined by densitometry scanning of the zymogram
(data not shown). In all experiments, we demonstrated that the doses of
experimental reagents used did not affect viability, which remained
>99%. Together, these results suggest that the organization of
microtubules, but not microfilament assembly, is important for
induction and/or secretion of MMP-9 in THP-1 cells infected with Mtb.
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Induction of MMP-9 by Mtb requires protein kinase activation.
We next examined the role of protein kinases in the induction of MMP-9
expression in THP-1 cells after infection with Mtb bacilli. For this,
THP-1 cells were treated before infection with calphostin C, an
inhibitor of protein kinase C, or PD-98059, a MEK1 inhibitor that
blocks the mitogen-activated protein kinase pathway. As shown in Fig.
3, the treatment of THP-1 cells with light-activated calphostin C, but not inactive calphostin C, inhibited the induction of MMP-9 activity by Mtb H37Ra and Erdman bacilli. The
treatment of THP-1 cells with the MEK-1 inhibitor also resulted in
total inhibition of MMP-9 induction by Mtb Erdman. These results suggest that receptor-mediated signaling pathways involving protein kinases are important for MMP-9 induction by Mtb.
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Role of AP-1 in Mtb-induction of MMP-9.
To further investigate the cellular mechanisms involved in the
induction of MMP-9 expression by Mtb, we tested the role of the
transcription factor AP-1. There are two AP-1 binding motifs (TGAGTCA)
located upstream of the human MMP-9 gene (GenBank accession number
S83357) that have been implicated in the regulation of MMP-9 gene
expression (24). THP-1 cells were infected with Mtb Erdman for 24 h, and nuclear binding proteins were
isolated and tested for binding to a 21-bp radiolabeled double-stranded AP-1 consensus oligonucleotide. As shown in Fig.
4A, unstimulated THP-1 cells
contained AP-1. The infection of cells induced at least a onefold
increase in AP-1 binding to DNA. Addition of excess (50×) unlabeled
AP-1 oligonucleotide resulted in inhibition of AP-1 binding to the
labeled oligonucleotide. As expected, the addition of a mutated
unlabeled AP-1 oligonucleotide (2-base mismatch) did not inhibit AP-1
binding to the labeled oligonucleotide, demonstrating that binding of
transcription factor AP-1 to the oligonucleotide in our assay was
highly specific.
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Mannose receptor ligands induce MMP-9 activity in THP-1 cells.
The above data suggest that intracellular signals involved in the
induction of MMP-9 by Mtb are elicited cell surface receptors in THP-1
cells. Therefore, we decided to explore the receptors in THP-1 cells,
in particular those with specificity toward polysaccharide molecules
such as the mannose and the CR3 receptor, also known as the -glucan
receptor (12, 49, 58). We first tested various polysaccharide ligands for induction of MMP-9 activity in THP-1 cells.
THP-1 cells were incubated with the soluble
1,3-glucan polysaccharide laminarin, the soluble
-mannose polysaccharide mannan, the polysaccharide of
-mannose and
-glucose zymosan (a
particulate ligand of the mannose receptor), and mycobacterial LAM, followed by detection of MMP-9 activity by gel
zymography. Figure 5,
top, shows that incubation of THP-1 cells with increasing concentrations of mannan (100 µg to 1 mg/ml) results in a
dose-dependent increase in MMP-9 activity. In
contrast, incubation of THP-1 cells with laminarin at similar
concentrations did not result in increased MMP-9 activity.
Zymosan had a potent stimulatory effect that was noted with as little
as 25 µg/ml, whereas LAM was the most potent inducer.
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DISCUSSION |
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The mechanisms that regulate the inflammatory and tissue remodeling events that lead to the formation of caseating granulomas, tissue liquefaction, and cavity formation in the lungs of Mtb-infected animals are not fully understood. However, it is widely accepted that cell-mediated immune responses elicited in the host by mycobacteria are responsible for much of the tissue destruction observed (Ref. 70; reviewed in Refs. 13, 69). The release of cytokines by T cells and other effector host cells (e.g., monocyte and macrophage) and the production of components such as hydrogen peroxide, toxic fatty acids, cationic protein, lipases, phospholipases, nucleases, and antigen-antibody complexes have all been implicated in the tissue destruction process observed in tuberculosis (21, 22, 35, 40, 47, 59, 61-64). MMPs have also been implicated in the host response to mycobacteria (10, 46, 44). However, the exact role these molecules play in the pathophysiology of this illness and the factors that control their expression and function during Mtb infection are unknown.
MMPs are a group of matrix-degrading endopeptidases that have been implicated in tissue remodeling triggered by diverse forms of experimental and clinical lung disease (6, 38). Previously, we showed that the infection of C57BL6 mice with Mtb Erdman results in the induction of these gelatinases in lung (46). Chang et al. (10) reported that the infection of monocytic cells with Mtb elicited the production of MMP-9-related gelatinolytic activity. We confirmed this finding and demonstrated the induction of MMP-9 in transformed cells of the monocyte and macrophage lineage (i.e., U937 and THP-1) exposed to formalin-fixed killed Mtb bacilli, bacilli-free long-term culture filtrates, and purified cell wall components of mycobacteria (e.g., LAM) from attenuated and virulent Mtb strains (46). Using gel-filtration chromatography, we also isolated two protein fractions (17 and 156 kDa) that induced MMP-9 in THP-1 cells (46). These data suggest that MMP-9 induction is elicited by the direct interaction between cell wall components of Mtb bacilli and human monocytes and macrophages and that MMP-9 could play important roles in the pathogenesis of tuberculosis.
The present study explores the cellular mechanisms elicited in host cells that mediate the induction of MMP-9 by Mtb. To this end, we exposed THP-1 cells to virulent (Erdman) and attenuated (H37Ra) Mtb strains and demonstrated an increase in MMP-9-mediated gelatinolytic activity. Both strains induced gelatinolytic activity, suggesting that the ability of Mtb to induce MMP-9 activity in vitro is not necessarily related to its virulence. The examination of steady-state levels of mRNA coding for MMP-9 and its inhibitor TIMP-1 showed an induction of MMP-9 mRNA in response to Mtb but no significant changes in TIMP-1. This suggests that the induction of MMP-9 activity by Mtb is dependent on increased production of MMP-9 protein and not on the relative decrease in the expression of its inhibitor.
Further studies revealed that MMP-9 induction by Mtb was modulated by the organizational state of the cytoskeleton. The disruption of microtubules by colchicine and taxol inhibited MMP-9-related gelatinolytic activity, whereas depolymerization of actin microfilaments with cytochalasin D induced it. In a similar study, others found that colchicine treatment of human peripheral blood monocytes resulted in decreased secretion of MMP-9 (65). This effect might be dependent on the cell type tested because cytochalasin D inhibited MMP-9 in some malignant tumor cells (11, 32) and stimulated MMP-2 in others (20). Altogether, these results indicate that alterations in cytoskeletal structures in THP-1 cells exert a profound influence on the induction of MMP-9 by Mtb. How disruption of the cytoskeleton affects Mtb-induction of MMP-9 remains unclear. One explanation is that disruption of the cytoskeleton inhibits cellular phagocytosis. However, we found that phagocytosis of polystyrene beads alone does not induce MMP-9 in THP-1 cells, suggesting that phagocytosis in and of itself does not induce MMP-9. Another possibility is that cytoskeletal derangements might affect the secretion of MMP-9, resulting in the intracellular accumulation of MMP-9 in its inactive form, pro-MMP-9. However, analysis of the cell pellet extracts of Mtb-infected THP-1 cells did not show MMP-9 activity even after treatment with 4-aminophenylmercuric acid, an activator of MMP-9 (not shown). Effects of anticytoskeleton agents on the "stiffness or plasticity" of the cells is also possible, but the role of this process in MMP-9 expression has not been studied. Finally, another possibility is that MMP-9 expression in monocytes and macrophages is modulated by mechanotransduction, the intracellular events that link physical forces applied to cell membranes to intracellular signals that lead to altered gene expression (25). We favor this explanation particularly in view of data showing that cytoskeletal disruption is associated with the activation of kinases such as protein kinase C, protein kinase A, and mitogen-activated protein kinases and with the induction of transcription factors (1, 31, 51).
Consistent with the idea that Mtb induction of MMP-9 expression is dependent on the activation of intracellular protein kinases, we found that specific inhibitors of protein kinase C and mitogen-activated protein kinase pathways (calphostin C and PD-98058, respectively) inhibited MMP-9 expression. Similarly, others have shown that the exposure of cells to phorbol 12-myristate 13-acetate results in induction of MMP-9 (66) and that inhibitors of protein kinases result in inhibition (55).
We also examined the role of the transcription factor AP-1 in our system. In contrast to another gelatinase, MMP-2, the promoter for MMP-9 has a classic TATA box and AP-1 promoter elements that seem to be involved in transcriptional regulation (24). AP-1 elements bind the transcription factor AP-1, which consists of a complex of proteins, including members of the c-Fos and c-Jun family (3). Our data support a role for AP-1 in mediating MMP-9 induction by Mtb. We showed by electrophoresis motility shift assay that the exposure of THP-1 cells to Mtb bacilli resulted in increased AP-1 binding to DNA. Also, we showed that the induction of MMP-9 by Mtb was abolished by the transfection of cells with a competing AP-1 oligonucleotide, shown to be inhibitory for AP-1-dependent gene expression in previous studies (48). We believe this finding is important because AP-1 seems to be involved in the expression of other MMP genes, such as those encoding for collagenases and stromelysins (Ref. 24; reviewed in Refs. 33 and 38). Therefore, signaling events triggered by distinct forms of lung injury (including Mtb infection) might coincide at the level of AP-1, thereby stimulating the expression of MMPs capable of promoting tissue destruction.
Together, the above findings suggest that the interaction between
components of the cell wall of Mtb bacilli and cell surface molecules
on THP-1 cells elicit intracellular signals responsible for inducing
MMP-9 expression. Consequently, the last part of our work was directed
at identifying the receptor molecules responsible for these signals.
The observation that THP-1 monocytic cells respond to LAM by producing
MMP-9, whereas U937 cells do not (46), prompted us to
examine the role of polysaccharide-specific cell surface receptors such
as the mannose and the CR3 or -glucan (also know as Mac-1, integrin
CD11b) receptor. We showed that THP-1 cells treated with the soluble
ligand of the mannose receptor mannan, but not the
-glucan receptor
ligand laminarin, resulted in a dose-dependent increase in MMP-9
activity. We also found that the mycobacterial glycolipid LAM was the
most effective inducer of MMP-9 activity, followed by the particulate
ligand zymosan. In addition, exposure of cells to these polysaccharides
did not inhibit the subsequent induction of MMP-9 after infection with Mtb bacilli. The finding that large (LAM) and particulate (zymosan) molecules containing mannose effectively induce high levels of MMP-9
suggests a role for mannose receptors. In further support of a role for
these receptors in our system, we showed that a monoclonal antibody to
the human mannose receptor (anti-hMR), but not the CR3
-glucan
receptor (anti-CD11b), decreased the induction of MMP-9 in THP-1 cells
after infection with Mtb. These studies suggest that the induction of
MMP-9 in THP-1 monocytes by Mtb bacilli is partly mediated by the
binding of mannosylated glycans of the cell envelope of Mtb to mannose
receptors in THP-1 cells. However, because only partial inhibition of
MMP-9 activity was obtained, it is possible that other types of
receptors are also involved.
Mannose receptors are present in monocyte-derived macrophages (53, 56) and are involved in the nonopsonic phagocytosis of both pathogenic and nonpathogenic mycobacteria (18, 50). These receptors appear to represent a safe route of entry of Mtb into macrophages (4). The phagocytosis of LAM-coated microspheres by human monocyte-derived macrophages is believed to occur via mannose receptors and to be energy-, cytoskeleton-, and calcium-dependent and inhibitable by mannan (27). In murine and rabbit macrophages, ligand binding to mannose receptors triggers the secretion of lysosomal enzymes (7, 41), the production of superoxide (5, 28), and the secretion of cytokines (54, 68). Reminiscent to our observations with protein kinase inhibitors, others have shown that protein phosphorylation by tyrosine kinases is involved in mannose receptor signal transduction (30, 36). Further studies are needed to delineate the exact role of mannose receptors in the induction of MMP-9 by Mtb.
In summary, the present study provides evidence for Mtb-induced intracellular signaling capable of regulating the expression of the gelatinase MMP-9. This effect required receptor-mediated activation of protein kinases and the induction of the transcription factor AP-1, which in turn binds to the promoter of the MMP-9 gene and drives mRNA expression. These pathways seem to be sensitive to the state of cytoskeletal organization of the cell. Overall, our work strengthens the idea that MMPs are directly involved in the pathobiology of tuberculosis infection. However, it should be noted that MMP-9 levels are increased in lung diseases where tissue cavitation is not characteristic. This does not exclude a role for MMP-9 in Mtb-induced lung destruction. Instead, it suggests that the role of MMP-9 in tuberculosis might be more complex than previously considered. The relative expression of inhibitors and extracellular MMP activators is just one aspect that can determine the degree of tissue destruction observed. Furthermore, MMP-9 might play immunomodulatory roles not necessarily related to connective tissue degradation.
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ACKNOWLEDGEMENTS |
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This work was supported by National Institute of Allergy and Infectious Diseases Grant 1 RO1 AI-37937 (to J. Roman), an Established Investigator Award from the American Heart Association (to J. Roman), a Minority Supplement Award (to C. A. Rivera-Marrero), and a Veterans Affairs Merit Review Award (to C. A. Rivera-Marrero).
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FOOTNOTES |
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Address for reprint requests and other correspondence: J. Roman, Atlanta VA Medical Center, Pulmonary and Critical Care Section, Rm. 12C191, 1670 Clairmont Rd., Decatur, GA 30033 (E-mail: jesse.roman{at}med.va.gov).
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.
10.1152/ajplung.00175.2001
Received 17 May 2001; accepted in final form 26 October 2001.
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