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Am J Physiol Heart Circ Physiol 288: H1080-H1087, 2005. First published October 14, 2004; doi:10.1152/ajpheart.00860.2004
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Abnormal cardiac wall motion and early matrix metalloproteinase activity

Ricardo A. García, Kristian L. Brown, Richard S. Pavelec, Katrina V. Go, James W. Covell, and Francisco J. Villarreal

Department of Medicine, University of California at San Diego, La Jolla, California

Submitted 23 August 2004 ; accepted in final form 8 October 2004


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Activation of matrix metalloproteinases (MMPs) in the heart is known to facilitate cardiac remodeling and progression to failure. We hypothesized that regional dyskinetic wall motion of the left ventricle would stimulate activation of MMPs. Abnormal wall motion at a target site on the anterior lateral wall of the left ventricle was induced by pacing atrial and ventricular sites of five open-chest anesthetized dogs. Changes in shortening at the left ventricular (LV) pacing site and at a remote site at the anterior base of the left ventricle were monitored with piezoelectric crystals. Simultaneous atrial and ventricular pacing resulted in abnormal motion at the LV pacing site, yielding early shortening and late systolic lengthening, whereas the shortening pattern at the remote site remained unaffected. Assessment of global myocardial MMP activity showed a sevenfold increase in substrate cleavage (P < 0.02) at the LV pacing site relative to the remote site. Gelatin zymography revealed increases in 92-kDa MMP-9 activity and 86-kDa MMP-9 activity at the LV pacing site relative to the remote site, whereas MMP-2 activity was unaffected. Abnormal wall motion was associated with increases in collagen degradation (~2-fold; P < 0.03), plasmin activity (~1.5-fold; P < 0.05), nitrotyrosine levels (~20-fold; P = 0.05), and inflammatory infiltrate (~2-fold; P < 0.02) relative to the remote site. Results indicate that regional dyskinesis induced by epicardial activation is sufficient to stimulate significant MMP activity in the heart, suggesting that abnormal wall motion is a stimulus for MMP activation.

dyskinesis; stretch; remodeling; collagen; metalloproteinases

ACTIVATION OF MATRIX METALLOPROTEINASES (MMPs) is known to contribute to the degradation of extracellular matrix (ECM), myocardial wall thinning, and left ventricular (LV) dilatation (23). MMPs are a family of enzymes that mediate ECM breakdown and are thought to be involved in the development of pathological LV remodeling (29). Despite the prevalence of reports detailing the activities of MMPs in cardiovascular disease, the exact mechanism(s) by which these enzymes are activated remains unclear.

Activation of MMPs can occur in the setting of myocardial ischemia/infarction (9). Ischemic insult to the myocardium gives rise to increased levels of oxygen radicals, inflammation, and abnormal motion of the dysfunctional myocardium by passive systolic stretch (i.e., dyskinesis) (1, 10, 11). Activation of MMPs by oxygen radicals has been implicated in myocardial contractile dysfunction (10), and activation of MMPs by inflammatory infiltrates is linked to the development of ventricular remodeling (1). Serine proteases such as plasmin have also been suggested as important activators of MMPs during postinfarction ECM remodeling (5).

In vitro studies using cardiac fibroblasts (27) and vascular smooth muscle cells (13) have shown that cyclic stretching stimulates increases in MMP activity. However, nothing is known about the effects of abnormal LV wall motion on in vivo activation of MMPs. In the present study, epicardial ventricular pacing was used to induce abnormal wall motion. Epicardial ventricular pacing is associated with late systolic lengthening (i.e., dyskinesis). Unlike dyskinesis induced by ischemia, this dyskinesis is induced in the absence of myocardial underperfusion. The data provided herein demonstrate that short-term (<4 h) dyskinesis is sufficient to selectively stimulate high levels of MMP-9 activity at the pacing site. Furthermore, we provide evidence for possible roles for plasmin, protein nitrosylation, and inflammatory infiltrates as potential activators of myocardial MMPs.


   MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
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Animal preparation. Animal studies were performed by guidelines described by the American Association for Accreditation of Laboratory Animal Care, and protocols were approved by the University of California-San Diego Animal Subjects Committee. Nine mongrel dogs (eight male; one female) weighing between 15 and 28 kg were sedated with thiopental sodium (25–50 mg/kg iv). Subjects were ventilated using a Harvard respirator, anesthetized by mask with 5% isofluorane, and maintained with 1–2% isofluorane for the rest of the study. The femoral veins were catheterized and used as infusion lines. The heart was exposed via a medial sternotomy and supported in a pericardial cradle. Arterial pressure was measured with a fluid-filled catheter placed into the aortic arch via the brachiocephalic or subclavian artery, and LV pressure was measured with a second matched fluid-filled catheter introduced through the apex of the heart. One pair of sonomicrometer crystals was placed near the center of the anterior LV wall [LV pacing site (LVP)], and a second pair was placed near the base of the anterior wall [LV remote site (LVR); Fig. 1]. Crystals were inserted into the midwall of the myocardium, perpendicular to the longitudinal axis of the heart. Epicardial pacing electrodes were sutured to the left or right atrium, and a second set was sutured between the crystal pair at the LVP site (Fig. 1). Arterial pressure, LV pressure, rate of change of LV pressure (LV dP/dt), sonomicrometer segment lengths, and electrical activity were recorded.



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  		Fig. 1. Pacing preparation. Sonomicrometer crystals were placed at the left ventricular pacing site (LVP) and at a left ventricular remote site (LVR) at the base of the heart. Epicardial pacing wires were sutured to the atrium and to the LVP site. Two connected stimulators were used for pacing, and LV pressure was measured with a fluid-filled catheter. LAD, left anterior descending coronary artery; CRX, circumflex coronary artery.

 
Six dogs were subjected to electrical pacing of the heart. In one animal, post hoc review showed that the left ventricle was not captured after 60 min of atrioventricular (AV) pacing: this animal was excluded from analyses. Three dogs served as shams that were instrumented but not paced. Zetabradine (0.5 mg/kg) was administered to dogs with heart rates >110 beats/min to capture the ventricle at 100–140 beats/min. Control measurements were obtained during atrial electrical stimulation. Induction of dyskinesis at the LVP site was accomplished by a second stimulus 0–80 ms after the atrial stimulus directed to LV electrodes. Dyskinesis was verified by the contraction pattern observed at the LVP site (Fig. 2B). Regional dyskinesis was maintained for 3.5–4 h. The ventricular stimulator was turned off, and a final reading of physiological parameters with atrial pacing was taken. Animals were euthanized with pentobarbital sodium (200 mg/kg), hearts were excised, and tissue was removed, frozen on dry ice, and stored at –80°C.



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  		Fig. 2. Segment motion and percent changes in segment length at LVR and LVP sites. Shown are representative recordings of segment motion from a single animal during atrial and atrioventricular (AV) pacing. A: LVR site with atrial pacing only (top) and with AV pacing (bottom). The solid tracing shows segment length during contraction. End diastole (ED) and end systole (ES) are shown (dashed vertical lines) with respect to LV pressure (dashed line). Both traces show normal phasic movement of the sonomicrometer crystals before and during AV pacing. B: LVP site with atrial pacing only (top) and with AV pacing (bottom). With AV pacing, LVP site shows early shortening that precedes ventricular ED (*) and early systolic bulging ({dagger}). Note change in segment length pattern from normal phasic movement (top) to abnormal movement (bottom). C: average percent changes in segment length at the LVR ({bullet}) and LVP ({blacksquare}) sites. Negative values reflect segment shortening, and positive values indicate lengthening. Percent change = [(ES – ED)/ED x 100]. Values are means ± SE of 5 animals per time point (except n = 4 animals at 150 min).

 
MMP and plasmin activity. MMP activity was measured with Omni-MMP fluorogenic substrate (P126, BioMol Research). Heart samples were taken from the right ventricle (RV) and from paced and nonpaced regions of the left ventricle adjacent to the crystal insertion sites; tissues at the crystal insertion sites were not used. Tissue (~50 mg) was homogenized in ice-cold buffer (50 mM Tris, pH 7.8, 150 mM NaCl, 5mM CaCl2, and 0.2 mM NaN3). Samples containing 50 µg of protein and 10 µM substrate were mixed in buffer. Selected samples were supplemented with the MMP inhibitor phenanthroline (1 mM) or the serine protease inhibitor aprotinin (50 kallikrein inhibitory units/ml). Kinetic fluorescence measurements were performed with a microplate reader (excitation of 340 nm, emission of 405 nm) at room temperature. Fluorescence emission was normalized to moles of cleaved peptide using the analogous unquenched fluorescent peptide of known concentration (P127, BioMol Research). Substrate cleavage rates were determined from the linear regions of the kinetic curves. Data are reported as percent ratios of rate of substrate cleavage with respect to the LVR site.

Plasmin activity was measured with N-(p-tosyl)-Gly-Pro-Lys 4-nitroanilide acetate salt, a plasmin-specific chromogenic substrate (T6140, Sigma). Tissues (~50 mg) were homogenized in 50 mM Tris, pH 7.7, 150 mM NaCl, and 2.5 mM 6-amino-n-hexanoic acid. Samples (100 µg) were incubated with substrate (700 µM), and activity was measured by absorbance (405 nm) at room temperature. Selected samples were supplemented with the serine protease inhibitor aprotinin (50 kallikrein inhibitory units/ml).

Gelatin zymography. Heart samples (~50 mg) were homogenized in 10 mM HEPES, pH 7.5, 150 mM NaCl, 0.2 mM EDTA, 25% glycerol, 100 µg/ml phenylmethylsulfonyl fluoride, and 0.2 kallikrein inhibitory units/ml aprotinin. Samples (10 µg of protein) were analyzed by SDS-PAGE as described (30). MMP-2 and MMP-9 protein standards were from Chemicon.

Immunoblotting. Protein levels were as assessed by immunoblotting (28) on polyvinylidene difluoride membranes with an anti-MMP-9 antibody (PC309, Calbiochem).

Myeloperoxidase activity. Myeloperoxidase (MPO) assay was carried out as previously described (20) with modifications. Tissue was homogenized in ice-cold buffer (50 mM KH2PO4 and 0.5% hexadecyltrimethylammonium bromide, pH 6.0) as described above. Homogenates were incubated on ice for 30 min and centrifuged at 4°C, and the supernatant was removed. Substrate solution (2 mg/ml tetramethylbenzidine dissolved in dimethyl sulfoxide) was diluted 1:10 in reaction buffer (50 mM KH2PO4, pH 6.0, supplemented with 4 µl of 30% hydrogen peroxide per 10 ml). Homogenates were diluted 1:10 in reaction buffer (100 µl final volume) and placed into a 96-well microplate. Substrate solution (100 µl) was added to the wells, and kinetic absorbance measurements of MPO activity were immediately monitored at 655 nm (readings every 40 s for ~20 min). Substrate cleavage rates were determined from the linear regions of the kinetic curves. Data were normalized to protein concentrations determined using the BCA protein assay kit (Pierce, Rockford, IL).

Nitrotyrosine analyses. Tissue samples were homogenized as described above, and gelatinases were affinity purified with gelatin-agarose resin (Sigma). Briefly, gelatin-agarose resin was washed with 10 bed volumes of buffer (50 mM Tris·HCl, pH 7.5, 400 mM NaCl, and 10 mM CaCl2). Supernatants from homogenized samples were incubated with resin for 30 min, washed with 4 bed volumes of buffer, and dissociated from resin with 2% dimethyl sulfoxide (21). Samples were analyzed for nitrotyrosine by slot blot (3) and Western blot analyses using a mouse anti-nitrotyrosine antibody (189542, Cayman Chemicals).

Hydroxyproline analysis. Hydroxyproline experiments were performed by the method of Edwards and O'Brien (8), with minor modifications. Transmural tissue samples, typically 300 mg wet wt, were trimmed of their inner and outer 1-mm layers, placed into 20-ml uncapped glass scintillation vials, and dried at 50°C for ~6–8 h. The dry weight of each sample was measured, 6 ml of 6 M HCl were added to each sample, and the vials were sealed with plastic caps. Samples were hydrolyzed for 36 h at 80°C with occasional mixing. The vials were uncapped, and the liquid was allowed to evaporate overnight at 80°C. The dried hydrolyzate film was suspended in 2 ml of deionized water and stored at –20°C. Hydroxyproline standards (8) were prepared in assay buffer (172 mM citric acid, 139 mM glacial acetic acid, 975 mM sodium acetate, 570 mM sodium hydroxide, 0.1% toluene, and 20% isopropanol, pH 6.5). About 10 µl of hydrolyzate were mixed with 190 µl of assay buffer, 100 µl of a hydroxyproline standard, and 150 µl of freshly prepared chloramine T reagent (0.141 g of chloramine T hydrate in 10 ml of deionized water) in plastic 1-ml microcentrifuge tubes. The samples were incubated at room temperature for 20–25 min. Ehrlich's reagent (1 M p-dimethylaminobenzaldehyde, 60% isopropanol, and 18.2% perchloric acid) was prepared within 5 min of use, 150 µl were added to each sample, and the tubes were incubated at 60°C for 15 min. The absorbance of each sample was read at 550 nm, and concentrations of hydroxyproline were determined from a standard addition curve (8). Collagen content was calculated with the assumption that the average hydroxyproline content of collagen is 10%, and thus 1 µmol of hydroxyproline is equivalent to 1 mg of collagen (25). Collagen content in each tissue sample was expressed as the percentage of the total dry weight of sample.

Collagen dansylation. Myocardial collagen was purified (22) and labeled with the fluorescent molecule dansyl ("dansylation") (12). Purified collagen was suspended in 0.5 M sodium bicarbonate. A volume of 0.5 ml of dansyl chloride (20 mg/ml in acetone) was added to collagen samples and incubated for 18 h in the dark at room temperature. Dansylated collagen was pelleted by centrifugation, supernatants were removed, and pellets were washed with three volumes of acetone. Samples were mixed in buffer (10 mM Tris, pH 7.5, 2% Triton X-100), and dansyl fluorescence was measured using an excitation wavelength of 360 nm and monitored at 516 nm. Fluorescence emission was normalized to collagen content as determined by hydroxyproline analyses and expressed as fluorescence emission per milligram of collagen (4).

Data analyses and statistics. Standardized measurements at end diastole were taken at the time of the initial rise in +dP/dt (where P is pressure and t is time) and LV pressure, and measurements at end systole were taken at the time of the dicrotic notch from arterial pressure. Sonomicrometer segment lengths at LVP and LVR sites were measured at times corresponding to end diastole and end systole. Zymographs and immunoblots were analyzed by densitometry. Bands of MMP-9 from tissue samples were normalized to the intensity of internal MMP-2 and MMP-9 standards (Chemicon). Statistical analyses were performed with either a Student's t-test or repeated-measures ANOVA. Results were considered statistically significant at P ≤ 0.05. All data are expressed as means ± SE.


   RESULTS
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 MATERIALS AND METHODS
 RESULTS
DISCUSSION
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Hemodynamics and abnormal wall motion. Averaged hemodynamics measurements are shown in Table 1. Comparisons of data measured during atrial pacing and after start of AV pacing did not reveal significant changes in heart rate, LV systolic pressure, or LV end-diastolic pressure. By contrast, peak +dP/dt was significantly depressed on initiation of AV pacing (P < 0.05, paired t-test). No significant changes occurred in LV pressure, heart rate, or peak ±dP/dt of shams (data not shown).


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  		Table 1. Hemodynamic measurements

 
As shown in Fig. 2, atrial pacing produced a normal phasic pattern of segment shortening during ejection and lengthening during diastole at both sites (Fig. 2, A and B, top). AV pacing, however, yielded dyskinetic segment motion at the LVP site (Fig. 2B, bottom). As shown in Fig. 2B, ventricular activation resulted in early regional shortening at the LVP site just before end diastole (denoted by "*") and was followed by early segment lengthening (or bulging) at the onset of systole (denoted by "{dagger}") followed by a second segment lengthening at end systole. The observed pattern of early ventricular activation followed by systolic segment lengthening has been observed in other comparable experimental ventricular pacing preparations (2) (19). Figure 2C shows that, at the LVR site, myocardial segments shorten (–9%) with atrial pacing. Upon AV pacing, the degree of shortening changes from –9% to –5%. Recovery to approximately –9% shortening occurred on return to atrial pacing at the end of the study period (Fig. 2C). By contrast, the LVP site initially shows normal shortening with atrial pacing (–11%) but becomes positive on AV pacing reflecting segment lengthening. Recovery to approximately –10% shortening occurs on return to atrial pacing, demonstrating that the effects of AV pacing at the site of abnormal wall motion are reversible. Myocardial segment shortening in shams (from –9% to –13% at mock LVP site and from –10% to –14% at LVR site) did not exhibit significant changes from initial to final measurements (P > 0.5, paired t-test).

MMP and plasmin activity. Representative gelatin zymography analysis of MMP activity is shown in Fig. 3A. The LVR site shows a moderate level of MMP-9 and MMP-2 activity of similar intensity whose molecular weights correspond to the prodomain forms of each enzyme. At the LVP site, there is a robust level of activity of 92-kDa MMP-9 that is ~40-fold greater than MMP-9 activity levels observed at the LVR site. A band directly below the 92-kDa MMP-9 band is visible in the LVP lane having a molecular mass of ~86 kDa, which is consistent with the active form of MMP-9. The 86-kDa MMP-9 activity at the LVP site is ~10-fold greater than 92 kDa MMP-9 activity levels at the LVR site (P < 0.01) and ~100-fold greater than 86-kDa MMP-9 levels at the LVR site (P < 0.001). The percent ratio of 86-kDa MMP-9 to 92-kDa MMP-9 is ~27%, as determined by densitometry. Ratios of 86- to 92-kDa MMP-9 measured from zymograms developed at different time points ensured that zymographic activity was in the linear range. MMP activity from the RV is of moderate intensity and resembles basal activity levels found at the LVR site. No significant differences in MMP-2 between LVR and LVP were measured (P > 0.2). Average MMP-9 activity for all pacing subjects is summarized in Fig. 3B.



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  		Fig. 3. Gelatin zymography showing activation of matrix metalloproteinase (MMP)-9 by abnormal wall motion. A: gelatin zymography bands corresponding to 92-kDa MMP-9, 86-kDa MMP-9, and 72-kDa MMP-2. MMP-2 and MMP-9 standards (Std) are shown in the left lane. Homogenate from the LVR site shows bands of moderate intensity corresponding to MMP-9 and MMP-2. Increased levels of MMP-9 are apparent at the LVP site. The 86-kDa band is ~27% as intense as the 92-kDa MMP-9 band. The right ventricle (RV) shows bands for 92-kDa MMP-9 and 72-kDa MMP-2 of similar intensity as those found in the LVR lane. B: bar graph showing averages of band intensity of MMP-9 activity.

 
A representative kinetic plot (Fig. 4A, inset) shows the progress of substrate cleavage with samples obtained from LVP, LVR, and RV sites. These data reveal that MMP activity at the LVP site is ~7-fold greater than activity levels at the LVR site and ~11-fold greater than activity levels at the RV. To establish that substrate cleavage was specifically mediated by MMPs, and not by other endogenous proteases within the tissue, parallel reactions were conducted with the MMP inhibitor phenanthroline. Phenanthroline completely inhibits cleavage of substrate (Fig. 4A, inset). Fluorescence data obtained with the omni-MMP substrate were converted to moles of cleaved fluorescent product. Figure 4A shows the ratios of percent substrate cleavage normalized to LVR regions in paced and sham animals. MMP activity from the LVP region shows a sevenfold higher activity (P < 0.02) than shams (LVP/LVR, 564 ± 122% vs. mock LVP/LVR, 85 ± 5%). Relative activity levels in the LVP region were also significantly higher (P < 0.04) than the RV of paced and sham subjects (143 ± 53 vs. 86 ± 16%).



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  		Fig. 4. Relative rate of substrate cleavage by MMPs and plasmin. A: tissue homogenates from LVP, LVR, and RV sites were exposed to a fluorogenic substrate specific for MMPs. Rate of substrate cleavage was determined for each sample and is reported as a ratio of basal MMP activity found at the LVR site. LVP shows ~7-fold more activity than LVR. Inset: representative fluorescence assay with tissue from the LVP ({blacktriangleup}), LVR ({triangleup}), and RV ({circ}) sites that were exposed to a fluorogenic substrate specific for MMPs. MMP activity was specifically blocked by 1 mM phenanthroline ({blacklozenge}). B: LVP site of paced dogs shows a 1.5-fold increase in plasmin activity than activity levels found at the mock "LVP" site of shams (P < 0.05) using a chromogenic substrate for plasmin. Rate of plasmin-mediated substrate cleavage was determined for each sample and is reported as a ratio of basal plasmin activity found at the LVR site.

 
Proteolytic activation of MMPs has been correlated with increased activity of serine proteases such as plasmin (5). Figure 4B shows comparisons of plasmin activity between LVP and mock LVP sites relative to LVR regions. Data reveal that plasmin activity from the LVP site of paced dogs is ~1.5-fold greater than activity from the mock LVP site of shams (164 ± 25% for LVP vs. 115 ± 9% for mock LVP; P < 0.05).

Immunoblotting for MMP-9. Representative immunoblots against MMP-9 are shown in Fig. 5A, with values for all subjects presented in Fig. 5B. MMP-9 protein levels from the LVP site are ~1.5-fold greater than the LVR site (400 ± 22 arbitrary units for LVP and 273 ± 30 arbitrary units for LVR; P < 0.04). No statistical difference was observed between mock LVP sites and LVR sites of shams (286 ± 9 arbitrary units for mock LVP and 298 ± 11 arbitrary units for LVR; P > 0.3).



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  		Fig. 5. Representative immunoblots for MMP-9. A: immunoreactive bands for MMP-9 in LVR and LVP samples. B: bar graph showing averages of MMP-9 band intensities. LVP site shows ~1.5-fold increase in protein levels relative to LVR (P < 0.04). Mock LVP and LVR sites of shams were not statistically different (P > 0.3).

 
Collagen cleavage assessment via amino-terminal dansylation. Figure 6 shows the extent of dansyl labeling ("dansylation") of purified collagen samples. The highest level of fluorescence emission occurs with collagen from the LVP site (24,113 ± 4,094 relative fluorescence units/mg), whereas lower levels are found in the LVR site and in both sites of shams (Fig. 6: 11,454–12,834 relative fluorescence units/mg). Fluorescence data were normalized to collagen levels measured by hydroxyproline analyses [3.8 ± 0.7 for LVP vs. 3.0 ± 0.4% collagen (dry wt) for LVR; P > 0.2]. Dansyl fluorescence increased twofold at the LVP site relative to the other regions (P < 0.03), indicating increased collagen degradation.



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  		Fig. 6. Collagen dansylation. Exposed amino-terminal ends of collagen were labeled with dansyl. Increased dansylation was observed with tissue from the site of abnormal wall motion (P < 0.02; n = 5). Fluorescence (relative fluorescence units, RFU) was normalized to collagen levels at each site.

 
MPO activity. LV homogenates were analyzed for inflammatory infiltrates by measuring MPO activity (18, 20). MPO activity from the LVP site shows a twofold increase in activity relative to the LVR region (16.1 ± 1.6 for LVR vs. 33.1 ± 6.1 arbitrary units x 10–3/min for LVP; P < 0.02; Fig. 7), whereas MPO activities in tissue from shams were comparable (21.8 ± 5.2 for LVR vs. 23.8 ± 8.4 arbitrary units x 10–3/min for mock LVP).



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  		Fig. 7. Myeloperoxidase (MPO) activity. Increased MPO activity was measured at the LVP site vs. the LVR site of paced animals (P < 0.02). No significant differences were measured between LVR and mock LVP sites of Shams (P > 0.6).

 
Nitrotyrosine levels. Tissue homogenates were enriched for gelatinases by affinity purification with gelatin agarose (21). Purified samples were analyzed by slot blot immunostaining for nitrotyrosine, as protein nitrosylation is known to activate latent MMPs (14). Figure 8A shows comparisons of nitrotyrosine levels between LVP and LVR sites from a single animal with averaged values for all paced dogs in Fig. 8B. Nitrotyrosine levels at the LVP site shows an average 20-fold increase in nitrotyrosine relative to the LVR site (125 ± 53 for LVP and 6 ± 1 arbitrary units for LVR; P = 0.05). Western blot analyses against nitrotyrosine reveal bands that approximate 92 and 86 kDa MMP-9 (Fig. 8C). Reblotting against MMP-9 verified that the amounts of MMP-9 in each lane were equivalent (Fig. 8D).



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  		Fig. 8. Nitrotyrosine levels. A: slot blot for 1 animal showing increased nitrotyrosine at the LVP site of paced animals (P = 0.05; n = 5). B: bar graph showing averages of nitrotyrosine band intensities from slot blots. The LVP site showed a 20-fold increase in protein levels relative to LVR. C: Western blot against nitrotyrosine showing bands corresponding to 92- and 86-kDa MMP-9. D: reblot against MMP-9 shows that MMP-9 protein levels are equivalent in each lane. Positions of molecular mass markers are shown at right.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Results demonstrate that short-term abnormal wall motion induced by epicardial ventricular activation is associated with enhanced MMP activity. Significant changes in activity were observed with MMP-9. Increases in MMP activity were concomitant with enhanced collagen degradation. These data are consistent with results from in vitro studies that correlate stretching of myocardial fibroblasts and vascular smooth muscle cells with increases in MMP levels and activity (13, 27). However, several alternate explanations for MMP activation described in this report must be considered including 1) regional variations in LV MMP activity and 2) potential for ischemia during ventricular pacing.

Regional variation in LV MMP levels could account for the observed differences in MMP activity. To our knowledge, no detailed studies have been reported that describe normal regional levels of MMPs present in the left ventricle. Wilson et al. (29) described the regional LV distribution of MMPs after myocardial infarction. In their study, no significant differences in regional MMP levels of normal LV samples were reported. In our studies, no regional differences (apex to base) in constitutive MMP activity within the LV were observed with sham subjects by zymography or by substrate cleavage analyses (data not shown). These observations indicate that increases in MMP activity at the LVP are not due to regional differences in MMP levels.

Systolic stretch of myocardium has been shown to increase oxygen demand at the site of regional dyskinesis (11). This point is critical because early ventricular activation by epicardial stimulation can reduce regional oxygen uptake at sites of LVP (7). This brings into question whether systolic stretching at the LVP site induces some degree of ischemia and/or oxidative stress and thus stimulates MMP-9 activity: increases in MMP activity are known to result from oxidative damage (6, 14). Figure 2C shows that myocardial LVP segments initially shorten by –11 ± 2% with atrial pacing. On cessation of ventricular pacing, shortening instantly returns to –10 ± 3% with atrial pacing. These data reveal that ventricular pacing does not result in significant changes in myocardial shortening from initial to final readings with atrial pacing (P > 0.7, paired t-test), thus demonstrating that ventricular pacing does not impair subsequent contractile function (i.e., no stunning). However, assessments of nitrotyrosine, a marker for oxidative stress, revealed increases in nitrotyrosine levels at the site of dyskinesis (P = 0.04). The increase in nitrotyrosine was not influenced by changes in pacing rate, voltage applied to the pacing site, or pacing duration (data not shown), making it unlikely that pacing, per se, stimulated protein nitrosylation. Because the increase in nitrotyrosine does not occur concomitant with myocardial contractile dysfunction or as a function of pacing parameters, even after at least 3.5 h of epicardial pacing, it is reasonable to infer that activation of MMPs at the LVP site by ischemia seems unlikely. The increase in nitrotyrosine, however, may be associated with activation of MMPs (6, 14).

Our observation that abnormal wall motion is associated with MMP activation is supported by studies that correlate mechanical stimulation of isolated cells with increases in MMP activity (13, 27). Tyagi et al. (27) demonstrated that MMP activity levels from infarcted human heart fibroblasts are identical to MMP activity levels achieved after 24 h of cyclic stretching of fibroblasts from normal tissue, suggesting that activation during ischemia may be due in part to deformation of the myocardium. In our studies, the magnitude of systolic bulging achieved during AV pacing at the LVP site ranged from 89% to 97% of the segment length at end diastole during atrial pacing. These data imply that the magnitude of myocardial stretching is not as important as is the timing of stretching (systolic bulging during dyskinesis) for MMP-9 activation. It is possible that the frequency of myocardial segment shortening and lengthening that occurs during a single beat (~3 cycles, Fig. 2B) might contribute to MMP activation at the site of abnormal wall motion. In addition, MMP activity was shown to be independent of pacing rate as comparisons of substrate cleavage data from subjects paced at different heart rates did not reveal a correlation between pacing rate and the degree of MMP activity observed at the LVP or LVR sites (data not shown).

The preponderance of 92-kDa MMP-9 in the zymograms suggests that abnormal wall motion stimulates de novo MMP-9 expression and/or triggers localization of enzyme to the dyskinetic site from preexisting reserves. Western blot analyses of MMP-9 showed moderate increases in MMP-9 levels at the LVP site (~1.5-fold), suggesting that zymographic activity of 92-kDa MMP-9 increases in part by an alternative manner. Recent evidence suggests that reserves of MMP-9 are stored in cytoplasmic secretory granules in endothelial cells (26). Shedding of MMP-2 and MMP-9 from endothelial cells occurs during angiogenesis (26). Secretory granule release occurs rapidly (within hours), and maximal release occurs by 4 h (26). Our studies are consistent with the time frame of MMP-9 release by shedding. More extensive studies to probe this issue may be warranted.

To investigate biochemical mechanisms of MMP activation, we examined possible involvement of serine proteases, inflammatory infiltrates, and protein nitrosylation. Activation of pro-MMPs has been correlated with activity of the serine protease plasmin (5). Plasmin activity was significantly increased by ~1.5-fold in the LVP site of paced animals (Fig. 4B). It is possible that increased plasmin activity at the LVP site might account for the enhanced presence of 86-kDa MMP-9 activity detected by zymography via proteolytic processing of 92-kDa MMP-9. Infiltration of inflammatory cells such as neutrophils into myocardium has been shown to activate MMP-9 in a canine ischemia-reperfusion model, where MMP-9 activity increased during the first hours of reperfusion (17). Increased MPO activity was observed at the LVP site relative to the LVR site of paced dogs (Fig. 7). The 2-fold increase in MPO activity at the LVP site was similar in magnitude to the increase in MMP-9 protein levels observed by immunoblotting (~1.5-fold; Fig. 5), suggesting that protein expression levels do not rise during induced dyskinesis; rather, they increase as a consequence of inflammatory cell infiltration. Protein nitrosylation of latent MMPs is know to stimulate MMP activity without proteolytic removal of the enzyme's prodomain (6, 14). Nitrotyrosine and MMP-9 immunoblotting of tissue homogenates affinity-purified for gelatinases showed bands close to 92 and 86 kDa (Fig. 8, C and D) in the LVP region. These data indicate that tyrosine residues of MMP-9 are specifically nitrosylated. Gu et al. (14) demonstrated that thiol-nitrosylation of pro-MMP-9 in vitro directly activates the enzyme. Although the exact mechanism by which nitrotyrosine levels increase is unclear, specific nitrosylation of 92-kDa MMP-9 may be involved in activation of its catalytic activity. Overall, our data imply that biochemical mediators such as plasmin, protein nitrosylation, and inflammatory infiltrates may be involved in activating MMPs within 4 h after the onset of abnormal wall motion.

Regional damage to collagen resulting from MMP-mediated proteolysis can yield a loss of ECM superstructure that leads to myocyte slippage and LV dilatation (15). Our data reveal that short-term dyskinesis stimulates a twofold increase in regional cleavage of collagen (Fig. 6). This increase in collagen degradation, however, does not result in significant changes in collagen content, as revealed by hydroxyproline analyses. The cessation of LVP allowed LV systolic function to return to normal. This observation indicates that the changes in ECM during this duration of pacing were not associated with changes in LV function. Under conditions of chronic rapid atrial pacing, Spinale et al. (24) found that LV collagen content dropped by ~16% after 3 wk of pacing tachycardia and was accompanied by LV systolic and diastolic dysfunction. The decrease in collagen was concomitant with the disruption of collagen fibrils, or "struts," that connect adjacent myocytes (24). The recovery of contractile function that we observed at the LVP site on termination of ventricular pacing suggests that myocardial ECM remained functionally intact after 4 h of pacing-induced abnormal wall motion, despite increases in MMP activity. Longer durations of ventricular pacing, however, could lead to further changes in ECM structure via MMP-9 activation and eventually could lead to LV dysfunction. Although not directly determined, increased MMP-9 activity by abnormal wall motion and the concomitant increase in collagen degradation suggest coordinate increases in collagenase activity. Early activation of the LV myocardium by ventricular pacing can also prestretch passive myocardium of later-activated regions distal to the ventricular pacing site (19). Thus other regions of the ventricle may also be subjected to stimulation of MMP activity by passive deformation of myocardium.

Regulation of MMP activity has been linked to the inhibitory action of tissue inhibitor of metalloproteinases (TIMPs) (23). Alterations in the balance of TIMPs/MMPs that favor increased MMP activity have been shown to occur in the failing human heart (16). Our data indicate that increased activation of MMPs at the site of abnormal wall motion is not ostensibly affected by endogenous TIMPs present at the site. However, it is possible that longer durations of induced dyskinesis might allow compensatory regulation of MMP activity by increased TIMP expression.

In conclusion, our data demonstrate that induction of short-term abnormal wall motion can stimulate significant levels of MMP activity in the heart. These results may provide an early view into the process of collagen degradation caused by abnormal wall motion that precedes pathological ECM remodeling.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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REFERENCES
 
This work was supported by National Heart, Lung, and Blood Institute Grant 1-R01-HL-143617 (to F. Villarreal). Support for R. Garcia was provided by National Heart, Lung, and Blood Institute, Research Training in Cardiovascular Physiology and Pharmacology at University of California-San Diego, Grant T32 HL-07444.


   ACKNOWLEDGMENTS
 
We are grateful to Rachel Alexander, Hiroshi Ashikaga, Oleg Ilchekes, and Kelly McCann for help with animal preparations and to Shirley Reynolds for early efforts with zymography experiments. We are also thankful to Professor Jeffrey Omens for many thoughtful suggestions given throughout the study.


   FOOTNOTES
 

Address for reprint requests and other correspondence: F. J. Villarreal, Dept. of Medicine, Univ. of California-San Diego, 9500 Gilman Dr., BSB 0613J, La Jolla, CA 92093 (E-mail: fvillarr{at}ucsd.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.


   REFERENCES
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Armstrong DG and Jude EB. The role of matrix metalloproteinases in wound healing. J Am Podiatr Med Assoc 92: 12–18, 2002.[Abstract/Free Full Text]
  2. Badke FR, Boinay P, and Covell JW. Effects of ventricular pacing on regional left ventricular performance in the dog. Am J Physiol Heart Circ Physiol 238: H858–H867, 1980.[Abstract/Free Full Text]
  3. Carnes CA, Chung MK, Nakayama T, Nakayama H, Baliga RS, Piao S, Kanderian A, Pavia S, Hamlin RL, McCarthy PM, Bauer JA, and VanWagoner DR. Ascorbate attenuates atrial pacing-induced peroxynitrite formation and electrical remodeling and decreases the incidence of postoperative atrial fibrillation. Circ Res 89: E32–E38, 2001.[Abstract/Free Full Text]
  4. Charney RH, Takahashi S, Zhao M, Sonnenblick EH, and Eng C. Collagen loss in the stunned myocardium. Circulation 85: 1483–1490, 1992.[Abstract/Free Full Text]
  5. Cleutjens JPM and Creemers EEJM. Integration of concepts: cardiac extracellular matrix remodeling after myocardial infarction. J Card Fail 8: S344–348, 2002.[CrossRef][ISI][Medline]
  6. Cox MJ, Hawkins UA, Hoit BD, and Tyagi SC. Attenuation of oxidative stress and remodeling by cardiac inhibitor of metalloproteinase protein transfer. Circulation 109: 2123–2128, 2004.[Abstract/Free Full Text]
  7. Delhaas T, Arts T, Prinzen FW, and Reneman RS. Regional fibre stress-fibre strain area as an estimate of regional blood flow and oxygen demand in the canine heart. J Physiol 477: 481–496, 1994.[ISI][Medline]
  8. Edwards CA and O'Brien WD. Modified assay for determination of hydroxyproline in a tissue hydrolyzate. Clin Chim Acta 104: 161–167, 1980.[CrossRef][ISI][Medline]
  9. Etoh T, Joffs C, Deschamps AM, Davis J, Dowdy K, Hendrick J, Baicu S, Mukherjee R, Manhaini M, and Spinale FG. Myocardial and interstitial matrix metalloproteinase activity after acute myocardial infarction in pigs. Am J Physiol Heart Circ Physiol 281: H987–H994, 2001.[Abstract/Free Full Text]
  10. Gao CQ, Sawicki G, Suarez-Pinzon WL, Csont T, Wozniak M, Ferdinandy P, and Schulz R. Matrix metalloproteinase-2 mediates cytokine-induced myocardial contractile dysfunction. Cardiovasc Res 57: 426–433, 2003.[CrossRef][ISI][Medline]
  11. Gayheart PA, Vinten-Johansen J, Johnston WE, Hester TO, and Cordell AR. Oxygen requirements of the dyskinetic myocardial segment. Am J Physiol Heart Circ Physiol 257: H1184–H1191, 1989.[Abstract/Free Full Text]
  12. Gray WR. Dansyl chloride procedure. Methods Enzymol XI: 139–151, 1967.
  13. Grote K, Flach I, Luchtefeld M, Akin E, Holland SM, Drexler H, and Schieffer B. Mechanical stretch enhances mRNA expression and proenzyme release of matrix metalloproteinase-2 (MMP-2) via NAD(P)H oxidase-derived reactive oxygen species. Circ Res 92: 1–7, 2003.[Free Full Text]
  14. Gu Z, Kaul M, Yan B, Kridel SJ, Cui J, Strongin A, Smith JW, Liddington RC, and Lipton SA. S-nitrosylation of matrix metalloproteinases: signaling pathway to neuronal cell death. Science 297: 1186–1190, 2002.[Abstract/Free Full Text]
  15. Lee RT. Matrix metalloproteinase inhibition and the prevention of heart failure. Trends Cardiovasc Med 11: 202–205, 2001.[CrossRef][ISI][Medline]
  16. Li YY, Feldman AM, Sun Y, and McTiernan CF. Differential expression of tissue inhibitors of metalloproteinases in the failing human heart. Circulation 98: 1728–1734, 1998.[Abstract/Free Full Text]
  17. Lindsey M, Wedin K, Brown MD, Keller C, Evans AJ, Smolen J, Burns AR, Rossen RD, Michael L, and Entman M. Matrix-dependent mechanism of neutrophil-mediated release and activation of matrix metalloproteinase 9 in myocardial ischemia/reperfusion. Circulation 103: 2181–2187, 2001.[Abstract/Free Full Text]
  18. Mullane KM, Kraemer R, and Smith B. Myeloperoxidase activity as a quantitative assessment of neutrophil infiltration into ischemic myocardium. J Pharmacol Methods 14: 157–167, 1985.[CrossRef][ISI][Medline]
  19. Prinzen FW, Hunter WC, Wyman BT, and McVeigh ER. Mapping of regional myocardial strain and work during ventricular pacing: experimental study using magnetic resonance imaging tagging. J Am Coll Cardiol 33: 1735–1742, 1999.[Abstract/Free Full Text]
  20. Romanic AM, Harrison SM, Bao W, Burns-Kurtis CL, Pickering S, Gu J, Grau E, Mao J, Sathe GM, Ohlstein EH, and Yue TL. Myocardial protection from ischemia/reperfusion injury by targeted deletion of matrix metalloproteinase-9. Cardiovasc Res 54: 549–558, 2002.[Abstract/Free Full Text]
  21. Sadatmansoori S, MacDougall J, Khademi S, Cooke LS, Guarino L, Meyer EF, and Forough R. Construction, expression, and characterization of a baculovirally expressed catalytic domain of human matrix metalloproteinase-9. Protein Expr Purif 23: 447–452, 2001.[CrossRef][ISI][Medline]
  22. Schofield JD, Freeman IL, and Jackson DS. The isolation, and amino acid and carbohydrate composition, of polymeric collagens prepared from various human tissues. Biochem J 124: 467–473, 1971.[ISI][Medline]
  23. Spinale FG. Matrix metalloproteinases: regulation and dysregulation in the failing heart. Circ Res 90: 520–530, 2002.[Abstract/Free Full Text]
  24. Spinale FG, Tomita M, Zellner JL, Cook JC, Crawford FA, and Zile MR. Collagen remodeling and ventricular function. Am J Physiol Heart Circ Physiol 261: H308–H318, 1991.[Abstract/Free Full Text]
  25. Takahashi S, Zhao M, and Eng C. Isolation and characterization of insoluble collagen of dog hearts. Protein Expr Purif 2: 304–312, 1991.[CrossRef][Medline]
  26. Taraboletti G, D'Ascenzo S, Borsotti P, Giavazzi R, Pavan A, and Dolo V. Shedding of the matrix metalloproteinases MMP-2, MMP-9, and MT1-MMP as membrane vesicle-associated components by endothelial cells. Am J Pathol 160: 673–680, 2002.[Abstract/Free Full Text]
  27. Tyagi SC, Kumar S, Voelker DJ, Reddy HK, Janicki JS, and Curtis JJ. Differential gene expression of extracellular matrix components in dilated cardiomyopathy. J Cell Biochem 63: 185–198, 1996.[ISI][Medline]
  28. Walther T, Schubert A, Falk V, Binner C, Kanev A, Bleiziffer S, Walther C, Doll N, Autschbach R, and Mohr FW. Regression of left ventricular hypertrophy after surgical therapy for aortic stenosis is associated with changes in extracellular matrix gene expression. Circulation 104: I54–58, 2001.
  29. Wilson EM, Moainie SL, Baskin JM, Lowry AS, Deschamps AM, Mukherjee R, Guy TS, John-Sutton MGS, Gorman JH, Edmunds LH, Gorman RC, and Spinale FG. Region- and type-specific induction of matrix metalloproteinases in post-myocardial infarction remodeling. Circulation 107: 2857–2863, 2003.[Abstract/Free Full Text]
  30. Yang JH, Briggs WH, Libby P, and Lee RT. Small mechanical strains selectively suppress matrix metalloproteinase-1 expression by human vascular smooth muscle cells. J Biol Chem 273: 6550–6555, 1998.[Abstract/Free Full Text]



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