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1 Division of Cardiology, Department of Medicine, and 2 Department of Biomedical Engineering, Emory University School of Medicine, Atlanta 30322; and 3 The Institute of Bioengineering and Bioscience, Georgia Institute of Technology, Atlanta, Georgia 30332
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ABSTRACT |
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 TOP
 ABSTRACT
 INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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Arteries remodel in response to environmental changes. We investigated whether mechanical strain modulates production of matrix metalloproteinase (MMP)-2 and -9 by cultured vascular smooth muscle cells (SMC). MMP-2 and MMP-9 expression were tested using human saphenous vein SMC cultured on silicone membranes at rest or subjected to physiological levels (5%) of stationary or cyclical (1 Hz) uniaxial strain. Compared with control, stationary strain significantly increased MMP-2 mRNA levels at all time points, whereas cyclic strain decreased it after 48 h. Both secreted and cell-associated pro-MMP-2 levels were increased by stationary strain at all times (P < 0.01), whereas cyclic strain decreased secreted levels after 48 h (P < 0.02). MMP-9 mRNA levels and pro-MMP-9 protein were increased after 48 h of stationary stretch (P < 0.01) compared with both no strain and cyclic strain. Our study indicates that vascular SMC show a selective response to different types of strain. We suggest that local increases in stationary mechanical strain resulting from stenting, hypertension, or atherosclerosis may lead to enhanced matrix degradation by SMC.
matrix metalloproteinase; mechanical stretch; vascular remodeling
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INTRODUCTION |
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TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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BLOOD VESSELS UNDERGO PHYSIOLOGICAL and pathological remodeling. This process necessitates the breakdown and reorganization of the extracellular matrix scaffold by vascular cells. Mounting evidence, obtained mainly from the study of remodeling during the development and complication of naturally occurring atherosclerotic lesions (5, 12, 17, 30), and that of vascular lesions triggered by interventions (2, 31, 35), implicates the contribution of specialized enzymes called matrix metalloproteinases (MMPs) (9). MMPs are secreted, both constitutively and inducibly, as inactive zymogens by human vascular (10, 15) and inflammatory (33) cells. In vitro and in vivo studies have shown that certain factors associated with vascular pathology, such as cytokines and mechanical injury, induce the expression and trigger the activation of MMPs (2, 10). MMPs have been found to be capable of degradation of collagen in the vulnerable shoulders of atherosclerotic plaques (30) and may be the cause of vascular tissue weakening and plaque destabilization. Because plaque failure was discovered to represent the major trigger for clinical events such as myocardial infarction and stroke (8), the potential action of MMPs has recently received a lot of attention. Similarly, increased expression of MMPs was demonstrated in several experimental models of arterial mechanical injury and graft failure (2, 31, 35), whereas MMP inhibitors decrease such lesions (1). Thus MMPs are believed to facilitate a variety of vascular pathologies.
One potentially relevant association is the observation that MMPs are overexpressed and activated in the vulnerable shoulders of atherosclerotic plaques (12), known to be exposed to the highest mechanical stress (6, 27). Colocalization of increased MMP expression and matrix degrading activity in the vulnerable shoulders (13) with areas of high tensile stress (21) may simply be an unfortunate coincidence, in which the weakest spot is subject to the highest tensile stress, but could also be indicative of a response to a specific mechanical environment. The cells and the extracellular matrix of the arterial vessel wall are likely responsive not only to biochemical but also to mechanical cues from the environment.
Hemodynamic forces have been demonstrated to play an important role in the development and evolution of arterial lesions in atherosclerosis and hypertension (29). Previous studies (6, 27) have shown that the distribution of forces that act on the vessel wall is affected by the development of an atherosclerotic lesion in the intimal layer of artery and that plaques tend to rupture in those areas exposed to the highest mechanical stresses. The same vulnerable areas present MMP overexpression and MMP activity (12), and colocalization of increased circumferential stress and overexpression of MMP-1 was demonstrated by Lee et al. (21). However, the cyclic deformation of human smooth muscle cells (SMC) in vitro was found to actually inhibit MMP-1 expression (34). Other work (20) has shown that axial strain in vivo upregulates MMP-2 in rabbit carotid arteries. We undertook the present study based on the same general assumption that mechanical stress could contribute to the increased MMP expression in atherosclerotic lesions, but we decided to investigate the possibility that this modulation depends on the nature of the mechanical stress. We thus compared the in vitro effects of cyclic versus stationary uniaxial strain on the production of MMP-2, the main MMP constitutively produced by human SMC, and of MMP-9, an MMP shown to be inducible by cytokines in human SMC in vitro (10).
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METHODS |
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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Cell culture and stretching experiments.
Experiments were performed with the use of cultured human SMC obtained
from explants of excess saphenous veins obtained after bypass surgery
(10). Primary SMC grown from vein explants were passaged
and used between passages 3 and 6. Culture purity
was verified with the use of anti-SMC 
-actin with immunostaining (Sigma A2547, 1:200). Each passage was monitored to ensure the lack of
any potential morphological or phenotypical changes or change in the
growth rate. SMC were grown to confluency in Dulbecco's modified Eagle
medium (DMEM) supplemented with 10% fetal calf serum (Cellgro/Fisher).
The silicon membranes were prepared beforehand by being washed with 10 N sulfuric acid for 1.5 h and then by being rinsed extensively
with autoclaved water. Each membrane was coated with a sterile solution
that was mixed with (1:1) 1 mg/ml collagen type I (Collaborative
Biomedical Products) and 100 µg/ml chitosan (USB) in 0.2 N acetic
acid through five spraying/drying cycles, UV sterilized (30 min),
rehydrated with warm (37°C) DMEM containing an antibiotic-antimycotic
mixture (100 U/ml penicillin; 100 mg/ml streptomycin), and supplemented
with 10% fetal calf serum. SMC were seeded on the coated membranes and
allowed to reach confluency (typically overnight). Membranes carrying
confluent cells were washed twice with Hanks' balanced salt solution
and then transferred into serum-free medium DMEM/F12 (1:1) supplemented with 1 mmol/l insulin and 5 mg/ml transferrin 24 h before the stretch experiments.
Sample collection and analysis. Culture medium was collected from the reservoir of the stretching device at the end of 24, 48, or 72 h. In some experiments, culture medium was concentrated by precipitation with trichloroacetic acid (TCA). The SMC were lysed using 10 mM phosphate buffer, pH 7.2, and 150 mM NaCl (PBS) containing 1% Triton X-100, 0.1% SDS, 0.5% sodium deoxycholate, and 0.2% sodium azide, as described previously (10). Protein concentration was measured using DC Protein Assay (Bio-Rad; Hercules, CA) as recommended for detergent-solubilized samples.
RNA extraction.
Total SMC RNA was extracted by using TRI reagent (Sigma) following the
manufacturer's instructions with slight modifications. These included
sample extraction on ice, longer centrifugation time (25 min), and
precipitation at 
20°C for 60 min. RNA was measured from the optical
absorbency at 260 nm.
Quantitative competitive RT-PCR. Quantitative competitive RT-PCR was performed essentially as described previously (32). This method uses the competitive amplification of mRNA sequences from the sample against a similar but shorter fragment of standard probe. Double-stranded DNA sequences that contained full-length inserts of MMP-2 or MMP-9 were generously provided by Dr. G. Wells (Neures).
To generate the MMP-2 standard from the MMP-2 cDNA, we used a sense primer corresponding to the T7 promoter and the reverse primer: 5'-ACATTGACCTTGGCACCGG
TGTCACTCCTGAGATCTGCAA-3' (where the
"" indicates a 21 base pair deleted sequence of the cDNA). For the MMP-9 standard, we used the same sense T7 promoter primer and the
reverse primer 5'-AAGATGCTGCTGTTCAGCGACGTGAAGGCGCAGATGGT-3'. The
double-stranded DNA PCR product was gel purified and used to generate a
340-nt synthetic RNA sequence for MMP-2 and a 302-nt synthetic RNA
sequence for MMP-9 with the use of T7 polymerase. These exogenous RNA
standards were identical to the endogenous cDNAs of MMP-2 and MMP-9 of
human SMC except that 21 base pairs near to the 3' regions had been
deleted. Exogenous standards were treated with RNase-free DNase (10 U/µl) for 15 min at 37°C and extracted using phenol-chloroform. To
determine the range of mRNA levels in the samples, we ran simultaneous
reactions containing 0.1µg of sample RNA and up to 100 pg of MMP
standard RNAs. Reverse transcription was performed using a commercially available kit (Gene Amp RNA PCR, Perkin-Elmer Applied Biosystems; Foster City, CA). PCR was then performed with AmpliTaq (Applied Biosystems), using the sense primer 5'-GCAAAGAACACAGCCTTCTC-3' and the
antisense primer 5'-ACATTGACCTTGGCACCGG-3' for MMP-2. For MMP-9,
we used the sense primer 5'-ACCTGGTTCAACTCACTCCG-3' and the
antisense primer 5'-AAGATGCTGCTGTTCAGCG-3'. These PCR reactions
resulted in amplification of a 257-bp sequence from the sample RNA, a
236-bp sequence from the exogenous standard of MMP-2, a 205-bp sequence
from the sample RNA, and a 184-bp sequence from the exogenous standard
of MMP-9. The PCR reactions were run on a 2.0% agarose gel and
visualized using ethidium bromide staining. The level of mRNA was
estimated on the basis of densitometric analysis of the signal for two
competing RNA products (Molecular Analyst, Bio-Rad). After an initial
approximation of the concentration, we selected several concentrations
closely distributed within that range to more precisely determine mRNA
abundance. The mRNA abundance (per µg total RNA) was calculated as
the amount of competitor required to produce a band of equal intensity
to the mRNA product. For assay of MMP-2 mRNA in nonstretch or cyclic
samples, the range used for template RNA was 0-0.4 pg per 0.1 µg
of sample RNA. For stationary conditions, the range was 0-20 pg.
For MMP-9 mRNA, the range was 0-0.5 pg for nonstretch, cyclic, or stationary.
Western blotting. Proteins were identified by immunoblotting of cell lysates and of cell culture media. Cell lysates were used directly, whereas culture media were concentrated by TCA precipitation. Equal amounts of protein (30 µg) from each sample were loaded on 10% SDS-PAGE mini gels. After electrophoresis, proteins were transferred onto nitrocellulose membrane with the use of a Trans-Blot system (Bio-Rad). Nonspecific binding was blocked with the use of 0.1% Tween and 5% dry milk in PBS. The blots were incubated with monoclonal anti-human MMP-2 (1 ng/µl Ab-3) (Oncogene), anti-human MMP-9 (2 ng/µl Ab-1), or polyclonal anti-human TIMP-2 (1:5,000 Ab-801) (Chemicon) or TIMP-1 (1:1,000, Ab-8122) in PBS containing 0.1% Tween and 5% milk. The blots were then washed and incubated in horseradish peroxidase-linked secondary antibodies (1:1,000, Amersham). Positive bands were revealed by using the ECL chemiluminiscence kit (Amersham) and quantified by laser densitometry and image analysis with Molecular Analyst software (Bio-Rad).
SDS-PAGE zymography. Proteins with gelatinolytic activity in cell lysates or culture media were identified by electrophoresis in the presence of SDS in 10% polyacrylamide gels containing 1 mg/ml gelatin, as described before (10). Equal amounts (30 µg) were loaded on each lane. Gels were stained with colloidal Coomassie blue (Sigma) and bands were quantified by laser densitometry and image analysis.
Statistical analysis. For each variable, data from at least four independent experiments were quantified and analyzed. Statistical analysis was performed with the use of ANOVA and Student's t-test. Comparisons were made as two-tailed Student's t-tests. P values <0.05 were considered to indicate statistical significance.
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RESULTS |
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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Under conventional static cell culture conditions, human SMC constitutively produced pro-MMP-2. The second gelatinase, pro-MMP-9, is also produced by human SMC after stimulation with cytokines (10). In the present study, we investigated the effect of stretch upon expression of these two SMC gelatinases. We compared SMC maintained in the classic, nonstrained, cell culture conditions with those subjected to static or cyclic uniaxial strain. We investigated mRNA and protein levels, including both cell-associated protein and protein secreted in the culture medium. The methods we used allowed detection of both pro-enzyme and activated forms of MMP-2 and MMP-9.
Expression of MMP-2.
Levels of MMP-2 mRNA, as assayed by RT-PCR, extracted from SMC
subjected to nonstretch and cyclic stretch did not vary significantly with time and were comparable at all time points (Fig.
1A). In contrast, stationary
stretch increased the MMP-2 mRNA level such that after 24 h this
was ~50-fold higher than the level of MMP-2 mRNA of both nonstretched
and cyclically stretched SMC (Fig. 1B).
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Expression of MMP-9. Levels of MMP-9 mRNA detected under the three different conditions were very low (0-4 pg/0.1 µg total RNA). We did not detect significant differences between different conditions at various time points (data not shown).
Levels of secreted pro-MMP-9 protein, measured by Western blotting (Fig. 5A), were close to the lower detection limit in most samples with the exception of samples obtained after stationary stretch at 48 and 72 h. These values were significantly higher that those detected in both other conditions (Fig. 5B). Levels of pro-MMP-9 secreted in the culture medium were also very low by SDS-PAGE zymography (data not shown).
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DISCUSSION |
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
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Vascular tissue remodeling (13) underlies the development, evolution, and complication of vascular lesions in atherosclerosis, hypertension, aneurysmal dilatation, graft failure, and after vascular interventions such as balloon angioplasty or stenting. The naturally occurring vascular pathological conditions are largely limited to the arterial side of circulation, which is exposed to pulsatile blood flow and frequent changes of the hemodynamic environment. These are not seen in the venous circulation, but occur in veins grafted into the arterial circulation, further supporting a role for physical forces in the natural history of arterial lesions (13, 24, 29). Similarly, aortic aneurysms occur in segments exposed to high pressure. Inside the vessel wall, the residing SMC are exposed to pressure, which manifests at the cellular level as mechanical stress. Mechanical stretching may occur chronically in hypertension, and prolonged stationary stretch results from the placement of a stent to open an occluded artery. Previous studies (21) showed that increased tensile stress colocalizes with increased MMP expression in the vulnerable shoulders of human atheroma. We have also found that increasing the ex vivo perfusion pressure enhances the expression and activity of MMPs within the wall of porcine arteries (7). We also showed that longitudinally stretched human saphenous veins had increased MMP expression and activity (23). These findings suggest that a combination of high mechanical stresses and areas with weakened matrix scaffold could endanger the mechanical strength of vascular tissue. At the same time, these observations have also raised the possibility that high mechanical stress may be a cause for increased matrix degradation via the action of MMPs. We therefore decided to test the hypothesis that uniaxial stress modulates the expression of the major SMC-derived MMPs, MMP-2, and MMP-9.
Previously reported (4) in vitro effects of mechanical strain on SMC include increased cell proliferation and deposition of a matrix necessary for organization and development of arteries, and later for vascular remodeling during atherogenesis and hypertension. This reaction was suspected as contributing to increased intimal thickening triggered by overstretching of arteries during balloon angioplasty. By using a similar simplified in vitro model, we now find that stationary strain of SMC increases their capacity to break down matrix. This could enhance arterial remodeling where SMC are exposed to such forces in situ and may enable compensatory remodeling and repair.
By examining MMP-2 and MMP-9, we found that compared with conventional static cell culture conditions, cyclic strain decreases the expression of MMP-2 in human SMC, as previously reported for MMP-1 (34). In contrast, stationary strain significantly increased the level of mRNA as well as latent and active MMP-2. Similarly, although much attenuated, static strain had enhancing effects upon expression of MMP-9. Thus we found that the in vitro response of SMC is dependent on the strain regimen. Furthermore, we found that stationary strain increased not only the expression of latent MMP-2, but also the level of cell-associated and secreted active MMP-2. This finding has important functional implications because degradation of matrix substrates by MMPs requires posttranslational processing of the latent forms secreted by cells. Conditions that trigger the activation of MMP zymogens are still poorly understood. A cell-associated activator of MMP-2 was described on the surface of transformed cells (28). This type of molecule is also expressed by activated SMC (26) and it may also be upregulated by mechanical strain.
Another potential mechanism for in situ MMP-2 activation may involve the generation of reactive oxygen species. We have shown that reactive oxygen radicals can activate in vitro latent forms of gelatinases secreted by cultured human SMC (25). Production of reactive oxygen species known to be present in atheroma is commonly attributed to inflammatory cells and may activate MMPs in the vulnerable shoulders of plaques (11); however, reactive oxygen species can also be produced by vascular cells, especially on stimulation (14). The fact that mechanical strain may act as a stimulus for vascular cells is supported by evidence that stretching of cultured endothelial cells leads to release of hydrogen peroxide (19). In addition, we showed that oxidative stress might contribute to the remodeling of human saphenous vein grafts experiencing arterial hemodynamic conditions through activation of latent MMPs (22). Thus an increased level of available latent MMP-2 in areas where SMC are subjected to stationary stretch will likely generate increased levels of enzymatically active MMP-2 in situ, through several potential pathways of activation that may be at work in the vessel wall.
Observed differences between the magnitude of effects at the level of mRNA and that of MMP-2 protein may be related to the translational and posttranslational distribution of mRNA. MMP-2 mRNA is translated into pro-MMP-2 protein, which is then distributed between cell-associated and secreted pro-MMP-2. This is in turn posttranslationally processed into the active forms of the enzyme and ultimately degraded by autolysis (3).
The results of our current in vitro study also support the notion that cyclic strain downregulates production of MMPs. This suggests that under normal hemodynamic conditions the matrix-degrading activity in healthy arteries is low. However, the increased stiffening due to the development of advanced calcified lesions in the arterial wall (18) and the increased oxidative stress impair the normal cyclic expansion and recoil of diseased arteries (16), while the wall is subjected to increased circumferential tensile stress (27). Similarly, diseased arteries are subjected to acute increases in stationary stretch during angioplasty, and chronic increases after stenting. We propose that increased stationary strain acts as a stimulus to increase production of matrix-degrading enzymes by the SMC, possibly as a natural reaction meant to allow the expansion of the arterial wall subjected to stretch. However, this reaction, triggered by vascular interventions or possibly mismatched compliance of an engineered vascular substitute to the adjacent tissue, will eventually facilitate development of lesions. In the shoulder areas of atherosclerotic plaques, where the stress is maximal (6), such process would contribute to the weakening of the plaque's fibrous cap and ultimately to its failure. Thus our in vitro results provide a likely mechanistic support to observations made in human and experimental vascular lesions, which suggest a relation between mechanical stretch and increased remodeling. A better understanding of the connection between mechanical stimulation and remodeling of vascular tissue in general, and of the matrix scaffold in special, should also aid in the design of cell seeded vascular grafts.
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ACKNOWLEDGEMENTS |
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The authors thank Dr. Karen Schentzer for valuable advice regarding the stretching device and Xiaoping Meng for assistance with the cell culture.
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FOOTNOTES |
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This study was supported in part by Georgia Tech-Emory Biotechnology Research Center, the Whitaker Foundation, and National Heart, Lung, and Blood Institute Grant R01 HL-64689-01A1.
Address for reprint requests and other correspondence: Z. S. Galis, Emory Univ. School of Medicine, Div. of Cardiology, 1639 Pierce Dr., WMB 319, Atlanta, GA 30322 (E-mail: zgalis{at}emory.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 January 23, 2003;10.1152/ajpheart.00494.2002
Received 13 June 2002; accepted in final form 19 January 2003.
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INTRODUCTION
METHODS
RESULTS
DISCUSSION
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 J D Raffetto and R A Khalil Mechanisms of varicose vein formation: valve dysfunction and wall dilation Phlebology, April 1, 2008; 23(2): 85 - 98. [Abstract] [Full Text] [PDF]
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 F. G. Spinale Myocardial Matrix Remodeling and the Matrix Metalloproteinases: Influence on Cardiac Form and Function Physiol Rev, October 1, 2007; 87(4): 1285 - 1342. [Abstract] [Full Text] [PDF]
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 T. L. Haas, J. L. Doyle, M. R. Distasi, L. E. Norton, K. M. Sheridan, and J. L. Unthank Involvement of MMPs in the outward remodeling of collateral mesenteric arteries Am J Physiol Heart Circ Physiol, October 1, 2007; 293(4): H2429 - H2437. [Abstract] [Full Text] [PDF]
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 K. M. Rice, D. H. Desai, D. L. Preston, P. S. Wehner, and E. R. Blough Vascular: Uniaxial stretch-induced regulation of mitogen-activated protein kinase, Akt and p70 S6 kinase in the ageing Fischer 344 x Brown Norway rat aorta Exp Physiol, September 1, 2007; 92(5): 963 - 970. [Abstract] [Full Text] [PDF]
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 M. Flamant, S. Placier, C. Dubroca, B. Esposito, I. Lopes, C. Chatziantoniou, A. Tedgui, J.-C. Dussaule, and S. Lehoux Role of Matrix Metalloproteinases in Early Hypertensive Vascular Remodeling Hypertension, July 1, 2007; 50(1): 212 - 218. [Abstract] [Full Text] [PDF]
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S. W. Watts, C. Rondelli, K. Thakali, X. Li, B. Uhal, M. H. Pervaiz, R. E. Watson, and G. D. Fink Morphological and biochemical characterization of remodeling in aorta and vena cava of DOCA-salt hypertensive rats Am J Physiol Heart Circ Physiol, May 1, 2007; 292(5): H2438 - H2448. [Abstract] [Full Text] [PDF]
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P. M. Cummins, N. von Offenberg Sweeney, M. T. Killeen, Y. A. Birney, E. M. Redmond, and P. A. Cahill Cyclic strain-mediated matrix metalloproteinase regulation within the vascular endothelium: a force to be reckoned with Am J Physiol Heart Circ Physiol, January 1, 2007; 292(1): H28 - H42. [Abstract] [Full Text] [PDF]
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 V. Gupta and K. J. Grande-Allen Effects of static and cyclic loading in regulating extracellular matrix synthesis by cardiovascular cells Cardiovasc Res, December 1, 2006; 72(3): 375 - 383. [Abstract] [Full Text] [PDF]
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C. M. Dollery and P. Libby Atherosclerosis and proteinase activation Cardiovasc Res, February 15, 2006; 69(3): 625 - 635. [Abstract] [Full Text] [PDF]
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 L. Pascarella, A. Penn, and G. W. Schmid-Schonbein Venous Hypertension and the Inflammatory Cascade: Major Manifestations and Trigger Mechanisms Angiology, November 1, 2005; 56(1_suppl): S3 - S10. [Abstract] [PDF]
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L. Pascarella, A. Penn, and G. W. Schmid-Schonbein Venous Hypertension and the Inflammatory Cascade: Major Manifestations and Trigger Mechanisms Angiology, November 1, 2005; 56(6_suppl): S3 - S10. [Abstract] [PDF]
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 A. Reich, N. Jaffe, A. Tong, I. Lavelin, O. Genina, M. Pines, D. Sklan, A. Nussinovitch, and E. Monsonego-Ornan Weight loading young chicks inhibits bone elongation and promotes growth plate ossification and vascularization J Appl Physiol, June 1, 2005; 98(6): 2381 - 2389. [Abstract] [Full Text] [PDF]
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 A. M. Deschamps, K. A. Apple, A. H. Leonardi, J. E. McLean, W. M. Yarbrough, R. E. Stroud, L. L. Clark, J. A. Sample, and F. G. Spinale Myocardial Interstitial Matrix Metalloproteinase Activity Is Altered by Mechanical Changes in LV Load: Interaction With the Angiotensin Type 1 Receptor Circ. Res., May 27, 2005; 96(10): 1110 - 1118. [Abstract] [Full Text] [PDF]
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 S. Lehoux and A. Tedgui Making Up and Breaking Up: The Tortuous Ways of the Vascular Wall Arterioscler Thromb Vasc Biol, May 1, 2005; 25(5): 892 - 894. [Full Text] [PDF]
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C. Bouvet, L.-A. Gilbert, D. Girardot, D. deBlois, and P. Moreau Different Involvement of Extracellular Matrix Components in Small and Large Arteries During Chronic NO Synthase Inhibition Hypertension, March 1, 2005; 45(3): 432 - 437. [Abstract] [Full Text] [PDF]
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 S. S. Signorelli, G. Malaponte, M. Libra, L. D. Pino, G. Celotta, V. Bevelacqua, M. Petrina, G. S Nicotra, M. Indelicato, P. M Navolanic, et al. Plasma levels and zymographic activities of matrix metalloproteinases 2 and 9 in type II diabetics with peripheral arterial disease Vascular Medicine, February 1, 2005; 10(1): 1 - 6. [Abstract] [PDF]
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N. von Offenberg Sweeney, P. M Cummins, Y. A Birney, J. P Cullen, E. M Redmond, and P. A Cahill Cyclic strain-mediated regulation of endothelial matrix metalloproteinase-2 expression and activity Cardiovasc Res, September 1, 2004; 63(4): 625 - 634. [Abstract] [Full Text] [PDF]
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