Department of Medicine and Research, Northport Veterans Affairs Medical Center, Northport 11768; and The State University of New York at Stony Brook, Stony Brook, New York 11794
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ABSTRACT |
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High-volume mechanical ventilation leads to ventilator-induced lung injury. This type of lung injury is accompanied by an increased release and activation of matrix metalloproteinases (MMPs). To investigate the mechanism leading to the increased MMP release, we systematically studied the effect of mechanical stretch on human microvascular endothelial cells isolated from the lung. We exposed cells grown on collagen 1 BioFlex plates to sinusoidal cyclic stretch at 0.5 Hz using the Flexercell system with 17-18% elongation of cells. After 4 days of cell stretching, conditioned media and cell lysate were collected and analyzed by gelatin, casein, and reverse zymograms as well as Western blotting. RT-PCR of mRNA extracted from stretched cells was performed. Our results show that 1) cyclic stretch led to increased release and activation of MMP-2 and MMP-1; 2) the activation of MMP-2 was accompanied by an increase in membrane type-1 MMP (MT1-MMP) and inhibited by a hydroxamic acid-derived inhibitor of MMPs (Prinomastat, AG3340); and 3) the MMP-2 release and activation were preceded by an increase in production of extracellular MMP inducer (EMMPRIN). These results suggest that cyclic mechanical stretch leads to MMP-2 activation through an MT1-MMP mechanism. EMMPRIN may play an important role in the release and activation of MMPs during lung injury.
matrix metalloproteinase; ventilator-induced lung injury; extracellular matrix metalloproteinase inducer; human lung microvascular endothelial cell; mechanical ventilation
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INTRODUCTION |
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MECHANICAL
VENTILATION (MV) has become an indispensable therapeutic modality
for patients with respiratory failure. However, it has been recognized
that MV per se can lead to a number of serious complications. The most
serious of these complications is the newly described
ventilator-induced lung injury (VILI) (5). The concept has
emerged that stretching of the lung by MV with high tidal volumes may
lead to acute lung injury with severe damage to the alveolar-capillary
barrier and pulmonary edema (5, 6). This type of lung
injury is characterized by an increased endothelial and
epithelial barrier permeability in the lung. The mechanism of such an
increase in permeability is not well understood. It was recently
reported that MV might play a role in initiating and propagating an
inflammatory response in the lung by increasing the release of
cytokines (TNF- and IL-1
) from the lung (27).
Matrix metalloproteinases (MMPs) are a family of enzymes that degrade components of the extracellular matrix (ECM) (23, 24). The 72-kDa gelatinase A (MMP-2) is the most widely distributed of all the MMPs (2) and is expressed constitutively by a number of cells, including endothelial and epithelial cells. The 92-kDa gelatinase B (MMP-9) is produced by several types of inflammatory cells, including polymorphonuclear neutrophils and alveolar macrophages, as well as stimulated connective tissue cells. MMP-2 and MMP-9 play an important role in pericellular basement membrane turnover by degrading the main components of the basement membrane. The extracellular matrix metalloproteinase inducer (EMMPRIN) is a 58-kDa, membrane-bound protein that has been identified in both normal (4) and diseased human tissues (1, 14, 21). Exposure of human fibroblast to recombinant EMMPRIN caused induction of MMP-1, MMP-2, and MMP-3 (4, 14).
We recently reported that in a rat model of VILI there was an upregulation and increased release and activation of MMPs, especially MMP-2, MMP-9, and membrane type-1 MMP (MT1-MMP). This increase in MMP-2 and MT1-MMP was preceded by the upregulation of EMMPRIN. We further found that the synthetic MMP inhibitor Prinomastat (AG3340) protected rat lungs from VILI (9). One possible mechanism leading to MMP upregulation and activation may be the effect of cell stretching caused by MV. Pulmonary microvascular endothelial cells are the cells forming the alveolar-capillary barrier (25) and are most likely to be subjected to stretch stresses during MV (16). Therefore, we sought in this study to systematically examine the effect of cell stretching in vitro on the release and activation of MMPs from pulmonary microvascular endothelial cells.
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METHODS |
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Cell culture.
Human lung microvascular endothelial cells (HMVEC-L) were purchased
from Clonetics (San Diego, CA). HMVEC-L cells were grown in EGM-2 MV
media (human endothelial cell media supplemented with 5% fetal bovine
serum, 0.04% hydrocortisone, 0.4% human fibroblast growth factor-,
0.1% vascular endothelial growth factor, 0.1% insulin-like growth
factor, 0.1% ascorbic acid, 0.1% human epidermal growth factor, and
1% GA-1000) (Clonetics). The cells were grown in a humidified
atmosphere of 95% air-5% CO2 at 37°C and passaged every
5-7 days. Cells from the second to the fifth passage were employed
in this study.
Application of cyclical mechanical strain.
HMVEC-L (5 × 105 cells/ well) were propagated in
six-well plates coated with type 1 collagen (BioFlex collagen 1 culture
plate; Flexcell International, Hillsborough, NC). We exposed the cells grown to confluence on the flexible surface of the BioFlex plates to
cycles of stretch and relaxation using a computer-driven,
vacuum-operated, stress-providing instrument (Flexercell Strain Unit
FX-4000 Tension plus; Flexcell International). The vacuum induced
17-18.5% elongation in the diameter of the flexible surface. We
used a "Flexstop," which is a rubber stopper inserted into the
underside of the BioFlex culture plate wells of the control
nonstretched cells to prevent the vacuum-induced flexing of the BioFlex
growth surface. The cells were exposed to stretch for 1, 2, 3, and 4 days. In the 3- and 4-day stretch experiments, serum-enriched medium
was added for 2 days and then replaced with serum-free medium
(endothelial cell basic medium; Invitrogen) for the following two days.
In the 1- to 2-day stretch experiments, serum-free medium was added from the start. Cells were examined by phase-contrast microscopy and
trypan blue exclusion to verify cell attachment and viability after
mechanical stretch. The conditioned medium collected after 1, 2, 3, and
4 days of stretch was centrifuged at 1,500 g for 5 min to
remove particulate matter and nonadherent cells and then stored at
80°C until assayed. At the end of each experiment, either cell
lysate or total cellular RNA was obtained. We obtained cell lysate by
incubating the cells with lysis buffer containing 0.1% SDS, 0.5%
sodium deoxycholate, and 1% Nonidet P-40 in PBS containing protease
inhibitor (Sigma, St. Louis, MO) for 1 h at 4°C. The cell lysate
was then pelleted by centrifuging at 1,500 g for 15 min to
remove cellular debris and collagen. Total RNA from the cells was
isolated with Tri reagent (Molecular Research Center, Cincinnati, OH)
following the manufacturer's instructions. All stretch experiments
were repeated 3-5 times.
Lactate dehydrogenase cytotoxicity assay.
Conditioned media of HMVEC-L exposed to stretch and nonstretch
conditions were assayed for lactate dehydrogenase (LDH) release. This
test provides a method for quantitating cytotoxicity based on the
measurement of the activity of LDH released from damaged cells. The LDH
assay was performed according the manufacturer's instructions (LDH
Cytotoxicity Detection Kit; Biovision). Briefly, 100 µl of
each of the following were examined: 1) serum-free medium as
background control, 2) conditioned medium of HMVEC-L cells as low control, 3) conditioned medium of HMVEC-L cells
incubated with serum-free medium containing 1% Triton X-100 as high
control, and 4) test samples of conditioned medium of
HMVEC-L cells exposed to stretch and nonstretch conditions.
Each sample was then mixed with 100 µl of reaction mixture (catalyst
and dye solution) in a 96-well plate and incubated for 30 min at room
temperature in the dark. Absorbance was measured at 490 nm. The mean of
the background value was subtracted from all other values. Cytotoxicity
was calculated with the following equation
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Gelatin zymography. Conditioned media of both stretched and nonstretched cells were diluted 1:1 in nonreducing sample buffer and separated on 10% SDS-PAGE gels containing 0.1% gelatin (precast gels from Invitrogen) for 150 min at 125 V. SDS was removed by incubation with renaturing buffer (Triton X-100, 25% in water) for 30 min at room temperature. The gels were washed for 30 min in developing buffer (Tris base, Tris · HCl, NaCl, CaCl2, and Brij 35 in distilled water) and then incubated for 24 h at 37°C in developing buffer as previously described (7). Finally, gels were stained with Coomassie brilliant blue r-250. Zones of enzymatic activity were characterized by the absence of Coomassie blue staining. Gelatinolytic bands were quantified by gel scanning and densitometry with an Alpha Imager (Alpha Innotech, San Leandro, CA).
Treatment with the MMP inhibitor Prinomastat. Prinomastat (AG3340; Agouron Pharmaceuticals, Pfizer) is a potent inhibitor of MMP-2, MMP-9, MMP-13, and MT1-MMP activity (20). An aqueous stock solution was prepared by dissolving Prinomastat in DMSO and used at a final concentration of 300 ng/ml in sterile PBS. This nontoxic concentration of Prinomastat is sufficient to inhibit MMP-2 and MMP-9 activation (20). DMSO at a final concentration of 0.5% in PBS was used as a vehicle control.
Casein gel. Equal amounts of conditioned media from both stretched and nonstretched cells were mixed with an equal volume of nonreducing Laemmli sample buffer and electrophoresed in SDS-7.5% polyacrylamide gels containing 1 mg/ml casein. After electrophoresis, we cleared the gels of SDS by incubating them for 1 h with incubation buffer (50 mM Tris · HCl, 2.5% Triton X-100, 5 mM CaCl2, and 1 µM ZnCl2 in distilled water). Gels were then incubated overnight in developing buffer at 37°C. The gels were then stained with Coomassie brilliant blue. Caseinolytic bands were quantified by gel scanning and densitometry with an Alpha Imager.
Reverse zymography. Reverse zymogram was performed as described previously (8). Briefly, equal amounts of conditioned media of both stretched and nonstretched cells were mixed with an equal volume of nonreducing Laemmli sample buffer and electrophoresed in SDS-7.5% polyacrylamide gels impregnated with both 0.05% gelatin and excess activated MMP-2. Tissue inhibitors of metalloproteinase (TIMPs) were identified by Coomassie blue staining bands on a clear background. Semiquantitative analysis of the amount of TIMP was performed with gel scanning and densitometry with the Alpha Imager.
Immunoblotting. Immunoblotting for MT1-MMP and EMMPRIN of cell lysates of stretched and nonstretched cells was performed with affinity-purified rabbit antibodies directed against MT1-MMP (hinge region) (Chemicon International, Temecula, CA), a mouse monoclonal antibody against the hemopexin-like domain of MT1-MMP (Oncogene, Cambridge, MA), and a mouse monoclonal antibody directed against TIMP-2 and EMMPRIN (Santa Cruz Biotechnology), respectively. Cell lysates were run in 8-16% SDS-polyacrylamide gel (Invitrogen). Gels were blotted onto polyvinylidene fluoride membranes (Invitrogen). Membranes were blocked for 1 h in PBS containing 0.1% Tween 20 and 5% milk. Primary anti-MT1-MMP, -TIMP-2, and -EMMPRIN antibodies diluted to a final concentration of 20 ng/ml, 2 µg/ml, and 200 ng/ml, respectively, in blocking solution were added and incubated overnight at 4°C. Secondary goat anti-rabbit and sheep anti-mouse immunoglobulin horseradish peroxidase-conjugate (Amersham) were used at 1:5,000 dilution (7, 9). Enhanced chemiluminescence (ECL) detection was performed per manufacturer's instruction (Amersham).
Immunoblotting of cell conditioned media was performed using rabbit antibodies directed against human MMP-3 and MMP-1 (Biogenesis) or mouse antibodies directed against human TIMP-2 (Oncogene). Conditioned medium with sample buffer was sized fractionated in 8-16% SDS-polyacrylamide gel. Gels were blotted onto nitrocellulose membranes. Membranes were blocked for 1 h in PBS containing 0.1% Tween 20 and 5% milk and incubated overnight at 4°C with primary antibodies. The final concentrations of the primary antibodies employed were as follows: anti-MMP-1 (2.6 µg/ml), anti-MMP-3 (1.36 µg/ml), and anti-TIMP-2 antibodies (2 µg/ml). Secondary antibodies, goat anti-rabbit or sheep anti-mouse immunoglobulin, and horseradish peroxidase-conjugate (Amersham) were used at 1:5,000 dilution. ECL detection was performed per manufacturer's instruction. Bands were identified, analyzed and photographed using alpha imaging.RT-PCR for MMP-2, MMP-1, MT1-MMP, EMMPRIN, and TIMP-2.
After 4 days of mechanical stretch, total RNA was isolated from HMVEC-L
by lysis and extraction in Tri reagent (Molecular Research Center) per
manufacturer's instructions. We performed reverse transcription by
boiling 1 µg RNA with 1 µM oligo(dT12-18) (Amersham Pharmacia) for 1 min, followed by incubation for 1 h at
37°C with the following reagents: 1× PCR buffer II (Sigma), 2.5 mM MgCl2 (Sigma), 1 mM dNTPs (Epicentre, Madison, MI),
800 U/ml ribonuclease inhibitor (Sigma), and 20 units Moloney murine leukemia virus-RT. Reactions were terminated by incubation at 95°C
for 10 min. PCR was performed with 200 ng cDNA, 1× PCR buffer II, 1.5 mM MgCl2, 0.5 unit Tfl DNA polymerase (Epicentre), and the
following oligonucleotide primers used at 0.4 µM: MMP-2 (+), 5'
CGCCGTCGCCCATCATCAAGT 3'; MMP-2 (), 5'
TGGATTCGAGAAAACCGCAGTGG 3'; EMMPRIN (+), 5' GTGAAGGCTGTGAAGTCGTCA 3';
EMMPRIN (
), 5' TTCCGGCGCTTCTCGTAGATGAA 3'; MT1-MMP (+), 5'
ATTCGCAAGGCGTTCCGCGTGTG 3'; MT1-MMP (
), 5' TGATGGCCGAGGGGTCACTGGA 3';
TIMP-2 (+), 5'TGCAGCTGCTCCCCGGTGCAC 3'; TIMP-2 (
),
5'TTATGGGTCCTCGATGTCGAG 3'; MMP-1 (+), 5'GAAATCTTGCTCATGCTTTTCAACC 3';
MMP-1 (
), 5'AAGGTTAGCTTACTGTCACATGCTT 3'.
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RESULTS |
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MMP-2 is both induced and activated by mechanical stretch.
Both static and stretched cells produced a constitutive 72-kDa
gelatinolytic band consistent with MMP-2. The size of the 72-kDa band
increased progressively with time and was much greater in the
conditioned media from stretched cells compared with nonstretched cells. There was a time-dependent increase in 64- and 62-kDa
gelatinolytic bands in conditioned media of stretched cells
consistent with intermediate and active forms of MMP-2 (Fig.
1). The conditioned media of
stretched cells treated with concanavalin A (ConA) showed the same
activated gelatinolytic band (62 kDa) as conditioned media from
stretched cells without ConA treatment (data not shown). Both static
and cyclically stretched HMVEC-L cell cultures had >90% cell
viability at the end of the experiment, as assessed by trypan blue and
LDH cytotoxicity assay, indicating that growth on collagen 1-coated
BioFlex membrane and cell stretching did not interfere with essential
cell functions.
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MT1-MMP is upregulated and activated by stretch.
Immunoblotting of cell lysates using the polyclonal rabbit anti-MT1-MMP
antibodies (against the hinge region) detected a 63-kDa band
representing pro-MT1-MMP, a 54-kDa band representing activated MT1-MMP,
and a 45-kDa band representing a degradation product of MT1-MMP in
cells exposed to stretch (12) (Fig.
2A). The intensity of the
MT1-MMP bands increased progressively over 4 days in stretched cells
but not in nonstretched cells (Fig. 2B). When the monoclonal mouse anti-MT1-MMP antibody was used against the hemopexin-like domain,
only the active form of MT1-MMP at 54 kDa was most prominently displayed (18) (Fig. 2B). Immunoblotting of
cell lysates of cells exposed to stretch for 1, 3, and 4 days showed
progressive increase in activated MT1-MMP compared with cell lysates
from nonstretched cells (Fig. 2C).
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MMP inhibitor Prinomastat attenuates the activation of MMP-2 caused
by stretch.
Continuous treatment of stretched cells with Prinomastat (300 ng/ml)
blocked the cleavage of the 72-kDa pro-MMP-2 to the active 62-kDa form.
However, the MMP inhibitor had no effect on pro-MMP-2 secretion (Fig.
3). These results suggest that the
activation of MMP-2 by mechanical stretch may be MT1-MMP dependent
through a signal transduction pathway.
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TIMP-2 is upregulated by stretch.
Reverse zymography and immunoblotting of conditioned media showed a
single 21-kDa band representing TIMP-2 in the conditioned media from
stretched cells; this band became evident in the conditioned media only
after 4 days of stretch. No bands were evident in the conditioned media
collected on day 1, 2, or 3. No TIMP-2 bands were
evident in conditioned media from nonstretched cells (Fig. 4A). Immunoblotting of cell
lysates from stretched and nonstretched HMVEC-L for TIMP-2 using mouse
monoclonal anti-TIMP-2 showed a band at 57 kDa representing TIMP-2
[most possibly bound to active MT1-MMP (13) or
representing a multimer of TIMP-2]. These bands were present after one
day of stretch. Stretch increased the density of the bands at each time
point (Fig. 4B).
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MMP-1 but not MMP-3 is induced and activated by stretch.
Casein gel zymography revealed a band at 52 kDa, consistent with either
MMP-3 or MMP-1, which was more prominent in the conditioned media of
cells exposed to stretch than in the conditioned media of nonstretched
cells (data not shown). Immunoblotting of conditioned media using
antibodies directed against MMP-1 confirmed the result of the casein
gel and showed two bands at 52 and 43 kDa, consistent with pro-MMP-1
and its activation product MMP-1 (Fig.
5). No bands consistent with MMP-3 were
observed when the antibody directed against MMP-3 was used. In the
time-course experiments, immunoblotting of conditioned media of cells
exposed to stretch and nonstretch for 1, 2, 3, and 4 days showed a
progressive increase in MMP-1 in the conditioned media of stretched
cells, which was significantly more than what was observed with
nonstretched cells (data not shown).
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EMMPRIN is upregulated by stretch.
Immunoblotting of lysate of cells exposed to stretch and
nonstretch using the mouse antibodies directed against EMMPRIN
showed a 58-kDa band representing EMMPRIN; EMMPRIN was significantly increased in stretched cells compared with nonstretched cells. EMMPRIN
increased in a time-dependent fashion beginning on day 1 in
cell lysate from stretched cells compared with nonstretched cells (Fig.
6).
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MMPs, TIMP-2, and EMMPRIN mRNA are induced by mechanical stretch.
Utilizing RT-PCR on RNA extracted from stretched and nonstretched cells
at the end of 4 days, we found a marked induction of mRNA for MMP-1,
MMP-2, MT1-MMP, TIMP-2, and EMMPRIN in stretched cells compared with
nonstretched cells (Fig. 7). The
expression patterns of these genes was consistent with our findings at
the protein level in our experiments described above using gelatin zymography, casein gel zymography, reverse zymography, and
immunoblotting.
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DISCUSSION |
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We recently reported that in a rat model of VILI there was an upregulation and increased release and activation of MMPs, especially MMP-2, MMP-9, and MT1-MMP (9). One of the possible mechanisms leading to MMP upregulation in these experiments is the effect of cell stretching caused by MV. Pulmonary microvascular endothelial cells are likely to be subjected to stretch stresses during MV. Therefore, we sought in this study to systematically examine the effects of cell stretching in vitro on the release and activation of MMPs from HMVEC-L. We used a degree of stretch (17% elongation) that is well above the expected stretch with normal breathing (15, 29) in an attempt to mimic the type of stretch that may be seen with high-volume lung MV. Our results show that cyclic mechanical stretch of HMVEC-L leads to an increase in the release and activation of MMP-2. This increase was accompanied by an increase in MT1-MMP. The activation of pro-MMP-2 was inhibited by the MMP inhibitor Prinomastat (AG3340). We also found that cell stretching caused an increase in the release and activation of MMP-1. EMMPRIN synthesis was also increased by cell stretching. Together, our results suggest that cyclic mechanical stretch caused by MV can cause lung injury by upregulating and increasing the release and activation of MMPs, especially MMP-2, MT1-MMP, and MMP-1.
MMP-2 is not generally regulated at the level of transcription (22) but, rather, is constitutively expressed and controlled through a MT1-MMP-, TIMP-2-specific mechanism of activation (13, 26, 31). However, there are data to indicate that the basal expression of MMP-2, MT1-MMP, and TIMP-2 is coregulated (22), consistent with their cooperation in MMP-2 activation. Our data support these conclusions in showing that MMP-2 upregulation was accompanied by a similar increase in MT1-MMP and TIMP-2. Our results support the role of MT1-MMP in the activation of pro-MMP-2, especially the data demonstrating that MT1-MMP levels increased in a time course that paralleled the increased activation of pro-MMP-2. Furthermore, treatment with Prinomastat prevented MMP-2 activation. These data strongly suggest a direct relationship between MT1-MMP expression and MMP-2 activation, consistent with the currently accepted mechanism of MMP-2 activation (3, 31). The results presented here are consistent with those reported in cardiac fibroblasts showing increased production of MMP-2 and MT1-MMP in response to stretch (28). Mechanical strain has also been reported to increase MMP-2 production and activity in vascular smooth muscle cells (17) and chondrocytes (10). Recently it has been shown that cyclic strain upregulates the early growth response gene product-mediated expression of MT1-MMP in rat microvascular endothelial cells (30).
In our study, reverse zymograms and Western blots established that TIMP-2 levels did not increase in the conditioned media during the initial 3 days of stretch, but TIMP-2 increased in response to stretch on the 4th day. However, in cell lysate, TIMP-2, which appears to be present in complex [possibly bound to active MT1-MMP (13) or as a multimer of TIMP-2 (8)], was enhanced by stretch beginning on day 1. The increase in the amount of activated MMP-2, with no change in stoichiometric amounts of TIMP-2 in the conditioned media during the initial 3 days of stretch, favors an environment where MMP-2, MT1-MMP, and TIMP-2 function together, leading to the degradation of ECM. The appearance of excess TIMP-2 on the 4th day of stretch in conditioned media is consistent with an inhibitory effect of increased TIMP-2 production.
Mechanical stretch increased EMMPRIN gene expression and protein synthesis in endothelial cells. This increase started early in the course of stretch (day 1). EMMPRIN is known to induce MMP-1, MMP-2, and MMP-3 expression in human fibroblasts (4, 14). EMMPRIN also causes increased expression of MT1-MMP and MT2-MMP as well as increased production and activation of MMP-2 in brain-derived fibroblasts (19). Increased EMMPRIN levels have also been shown to be associated with increased production and activation of MMP-2 and MT1-MMP in dilated cardiomyopathy (21). Recently, we reported that EMMPRIN mRNA was increased in rat lung tissues subjected to VILI; this increased EMMPRIN mRNA preceded the increase in MMP-2 and MT1-MMP mRNA seen in this type of injury (9). Together, our results suggest that mechanical stretch causes an increased upregulation of EMMPRIN that in turn leads to the increased production of MMP-1, MMP-2, and MT1-MMP seen in our study.
In conclusion, our results suggest that mechanical stress in the form of cell stretching may be an important stimulus to increased production and activation of MMP-2 through MT1-MMP- and EMMPRIN-dependent mechanisms. The clinical use of pharmacological MMP inhibitors to prevent early activation of MMPs leading to VILI should be considered. A limitation to the use of MMP inhibitors in inflammation, however, is in the difficulty distinguishing between the negative and positive effects of activated MMPs in the tissue injury versus the repair process (11).
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ACKNOWLEDGEMENTS |
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We thank Dr. David Shalinsky (Agouron Pharmaceuticals, Pfizer) for generously providing Prinomastat.
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FOOTNOTES |
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This work was supported by funds from the Veterans Affairs (V. A.) Research Enhancement Award Program REAP, National Heart, Lung, and Blood Institute Grant HL-646340 (H. D. Foda), and a V. A. Merit Review grant (S. Zucker).
Address for reprint requests and other correspondence: H. D. Foda, Pulmonary and Critical Care Medicine, SUNY at Stony Brook, Health Science Center, Stony Brook, NY, 11794-8172 (E-mail: hfoda{at}mail.som.sunysb.edu).
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.
First published November 27, 2002;10.1152/ajplung.00290.2002
Received 22 August 2002; accepted in final form 25 November 2002.
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