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Am J Physiol Heart Circ Physiol 287: H1730-H1739, 2004. First published May 27, 2004; doi:10.1152/ajpheart.00098.2004
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Increased cell death in osteopontin-deficient cardiac fibroblasts occurs by a caspase-3-independent pathway

Ron Zohar,1,3 Baoqian Zhu,1,3 Peter Liu,2 Jaro Sodek,1,3 and C. A. McCulloch1,3

1Group in Matrix Dynamics, Faculty of Dentistry, 2Heart and Stroke/Richard Lewar Centre of Excellence, and 3Canadian Institutes of Health Research, University of Toronto, Toronto, Ontario M5G 1G6, Canada

Submitted 4 February 2004 ; accepted in final form 18 May 2004


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Reperfusion-induced oxidative injury to the myocardium promotes activation and proliferation of cardiac fibroblasts and repair by scar formation. Osteopontin (OPN) is a proinflammatory cytokine that is upregulated after reperfusion. To determine whether OPN enhances fibroblast survival after exposure to oxidants, cardiac fibroblasts from wild-type (WT) or OPN-null (OPN–/–) mice were treated in vitro with H2O2 to model reperfusion injury. Within 1 h, membrane permeability to propidium iodide (PI) was increased from 5 to 60% in OPN–/– cells but was increased to only 20% in WT cells. In contrast, after 1–8 h of treatment with H2O2, the percent of terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling (TUNEL)-stained cells was more than twofold higher in WT than OPN–/– cells. Electron microscopy of WT cells treated with H2O2 showed chromatin condensation, nuclear fragmentation, and cytoplasmic and nuclear shrinkage, which are consistent with apoptosis. In contrast, H2O2-treated OPN–/– cardiac fibroblasts exhibited cell and nuclear swelling and membrane disruption that are indicative of cell necrosis. Treatment of OPN–/– and WT cells with a cell-permeable caspase-3 inhibitor reduced the percentage of TUNEL staining by more than fourfold in WT cells but decreased staining in OPN–/– cells by ~30%. Although the percentage of PI-permeable WT cells was reduced threefold, the percent of PI-permeable OPN–/– cells was not altered. Restoration of OPN expression in OPN–/– fibroblasts reduced the percentage of PI-permeable cells but not TUNEL staining after H2O2 treatment. Thus H2O2-induced cell death in OPN-deficient cardiac fibroblasts is mediated by a caspase-3-independent, necrotic pathway. We suggest that the increased expression of OPN in the myocardium after reperfusion may promote fibrosis by protecting cardiac fibroblasts from cell death.

necrosis; reperfusion; myocardium; nuclear fragmentation

AFTER MYOCARDIAL INFARCTION, ischemic damage to cardiac tissues causes tissue necrosis and an associated inflammatory response that leads to disorganization of myocardial architecture (1, 14). The death of cardiomyocytes is an integral part of pathological tissue destruction after infarction, but the viability of endothelial cells and fibroblasts is protected, thereby enabling the formation of a reparative matrix. Although matrix production by fibroblasts is important for maintaining the integrity of cardiac tissues, the persistence of activated fibroblasts (5, 10, 15) leads to the formation of a stiff, noncompliant scar tissue that compromises cardiac function. Accordingly, present therapeutic approaches are aimed at minimizing fibrosis during reperfusion (22).

During the reperfusion stage after myocardial infarction, there is increased production of oxidants and cytokines including IL-10, transforming growth factor-{beta}1, and osteopontin (OPN) that stimulate fibroblast proliferation and matrix formation. In postischemic hearts, OPN expression is increased (51, 54, 58); this enhanced expression is considered an important mediator of ANG II-mediated regulation of cardiac fibroblast behavior including increased collagen synthesis (3, 29, 58). Although OPN is important for the generation of immune responses in several inflammatory diseases (4, 8, 41), it is also required for the survival (12, 34, 49, 56) of endothelial, smooth muscle, kidney, and tumor cells (12). Because the development of cardiac fibrosis is due in part to the persistence of activated cardiac fibroblasts in the damaged myocardium, we investigated the impact of OPN on cardiac fibroblast deletion by necrosis and apoptosis after oxidative injury in vitro.


   MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
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Experimental models. Animal experiments were conducted according to guidelines established by the Animal Care Committee of the University of Toronto. Cells were obtained from OPN-null (OPN–/–; Ref. 46) and control wild-type (WT) mice, strain B6129SF2/J, provided by S. R. Rittling and D. T. Denhardt (Rutgers University, Piscataway, NJ). Ischemia-reperfusion injury in vivo was induced in 12-wk-old male mice by experimental myocardial infarction mediated by ligation of left coronary arteries as previously described (52). Functional cardiac evaluation was performed using transthoracic 2-D and M-mode echocardiographic examination with an Acuson Sequoia C256 system equipped with a 15-MHz linear transducer (15L8, version 4.0, Acuson; Mountain View, CA). For studies that involved immunostaining, mice were perfusion fixed with 10% buffered formalin, and the left ventricles were harvested. Paraffin sections were immunostained for OPN (see below).

For studies of cell injury in vitro, cardiac fibroblasts were prepared as previously described (16, 55). Briefly, hearts were removed under sterile conditions, and the ventricular tissue was excised, minced, and digested with 0.3% collagenase that contained (in wt/vol) 1.8% sorbitol, 0.05% DNase, 6.25 U/ml elastase, and 0.05% trypsin in Krebs buffer. Cells were plated for 1 h, and nonadherent cells were washed away. Cardiac fibroblasts attached and proliferated much more rapidly than cardiac myocytes; this produced virtually pure fibroblast cultures after the first passage, which was confirmed by immunostaining for collagen and vimentin. Cells were detached with 0.01% trypsin (GIBCO; Burlington, ON, Canada) for passaging, and culture studies were performed at passages 1 and 2. Fibroblastic cell lines from WT and OPN–/– mouse fetuses and clone 135–3T3 were provided by S. R. Rittling and D. T. Denhardt. A stable cell line was subcloned (clone 135–3T3/hOPNmet) from OPN–/– cells after transfection with a human OPN expression vector driven by a metallothionein promoter (pNMH-OP10). Cells were grown in {alpha}-MEM, (high-growth DMEM for cardiac fibroblasts) that contained 10% heat-inactivated FBS and antibiotics (in µg/ml: 100 penicillin G, 50 gentamycin sulfate, and 0.3 fungizone). Cultures were incubated at 37°C in a humidified atmosphere that contained 5% CO2.

Induction of cell death. Cells were analyzed primarily after incubation with 0.5 mM H2O2 in time-course experiments. In some experiments, cell death was induced by incubating primary cardiac mouse fibroblasts with staurosporine (1 µM), actinomycin D (500 ng/ml), thapsigargin (1 µM), or cycloheximide (100 µg/ml) for 24 h. To study the importance of active caspase-3 in the death of OPN–/– cardiac fibroblasts, cultures were treated for 3 h before apoptotic induction with Ac-AAVALLPAVLLALLAP-DEVD-CHO (1 µM), which is a cell-permeable, irreversible caspase-3 inhibitor (catalog no. 235423; Calbiochem; San Diego, CA).

Transmission electron microscopy. Cultured cells were fixed in Karnovsky's solution (2% paraformaldehyde and 2.5% glutaraldehyde in 0.1 M sodium cacodylate at pH 7.3) at 4°C for 4 h, washed three times in 0.1 M sodium cacodylate buffer, postfixed in 2% OsO4 in 0.1 M sodium cacodylate for 90 min at room temperature (21°C), and again washed three times in 0.1 M sodium cacodylate buffer. Samples were embedded in Epon 812 resin. Thin sections were placed on nickel grids, stained with uranyl acetate and lead citrate, and examined with an electron microscope (Hitachi) to detect morphological alterations of cell membranes, cytoplasmic organelles, and nuclei that are suggestive of apoptosis or necrosis.

Immunostaining. The presence of OPN in ventricles was examined by immunoperoxidase staining of 5-µm paraffin sections from WT mice subjected to experimental myocardial infarction. OPN was localized with a rabbit OPN antibody (LF123, provided by L. W. Fisher, National Institute of Dental and Craniofacial Research, Bethesda, MD). To detect apoptosis in vivo, we used terminal deoxynucleotidyl transferase-mediated dUTP nick end-labeling (TUNEL; Roche Diagnostics) staining of postinfarction specimens of myocardium. Paraffin sections (5 µm) were stained with a CardioTACS Kit (Trevigen; Gaithersburg, MD) according to the supplier's protocol.

For culture experiments, cells were fixed with 2% paraformaldehyde in Ca2+- and Mg2+-free PBS for 30 min, washed three times with BSA buffer, permeabilized with 0.1% Triton X-100 in PBS for 30 min at room temperature, and stained for OPN. Antibody binding was detected with a Texas red-conjugated goat anti-rabbit IgG (heavy and light chains; Vector Laboratories; Burlingame, CA; Ref. 31). Immunofluorescence was examined via confocal microscopy.

Cells were stained with propidium iodide (PI; 20 µg/ml for 5 min; Molecular Probes; Eugene, OR) to label cells with permeable cell membranes, which are seen in dying cells. For detection of apoptosis, cells were fixed with paraformaldehyde (4% in PBS, pH 7.4, for 1 h) and permeabilized with Triton X-100 (in 0.1% sodium citrate buffer for 2 min at 4°C). Endonuclease-fragmented DNA was detected with TUNEL staining. Cells were counterstained with 4',6-diamidino-2-phenylindole dihydrochloride (DAPI) to facilitate estimates of total cell numbers. Cells were stained either as spread cells grown on Lab-Tek eight-well chamber slides or as single cell suspensions for cytospin analyses (59). Cells exhibiting condensed nuclei (as indicated by DAPI staining) or fragmented nuclei and chromatin (via DAPI and TUNEL staining) were counted in three fields on replicate dishes, and the percentages of apoptotic cells were calculated. To distinguish between apoptosis and other forms of cell death (i.e., necrosis), the total number of PI-positive cells was enumerated separately from cells that were both PI and TUNEL positive (defined here as apoptotic; Ref. 24). In some experiments, TUNEL- and PI-stained cell suspensions were analyzed with either a FACSTAR Plus flow cytometer (Becton-Dickinson) or an Altra flow cytometer (Beckman Coulter).

Apoptosis was examined further by staining for active caspase-3. Fixed cells (2% paraformaldehyde in Ca2+- and Mg2+-free PBS for 30 min at 4°C) were permeabilized with 0.3% Triton X-100. Washing and antibody dilution were in 0.25% BSA in PBS (Ca2+ and Mg2+ free) except as outlined below. Cells were stained with rabbit anti-human/mouse caspase-3 (0.3 µg/ml; R&D Systems; Minneapolis, MN). Binding was detected with FITC or Alexa-conjugated goat anti-rabbit F(ab)2 fragments diluted 1:100 in the same BSA solution and incubated for 1 h at 4°C.

Mitochondrial and cell membrane potentials were analyzed to detect earlier changes in the cell death process. For measurement of mitochondrial membrane potential, single-cell suspensions were stained with 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide (JC-1, 0.5 µM for 10 min; Molecular Probes). With excitation at 488 nm, the relative intensity of red and green fluorescence was analyzed by flow cytometry (n = 10,000 cells). The formation of "J aggregates" with enhanced red fluorescence provides an estimate of the mitochondrial membrane potential (28, 37). In some experiments, the mitochondrial membrane potential was dissipated with carbonyl cyanide m-chlorophenylhydrazone (CCCP, 1 µmol; Sigma-Aldrich) to provide controls.

To measure cell membrane potential, time-mode, flow-cytometric analyses of OPN–/– and WT cardiac fibroblasts were stained with bis-(1,3-dibutylbarbituric acid)trimethine oxonol [DiBAC4(3), 10 µM in ethanol; Molecular Probes] for 30 min. Background fluorescence was determined in cell suspensions stained with DiBAC4(3) and compared with cells previously incubated with 0.5 mM H2O2. Increased DiBAC4(3) fluorescence intensity, which is indicative of membrane depolarization associated with cell death, was measured at 15-min intervals at 520 nm. As a positive control, OPN–/– cell suspensions were incubated with 100 mM KCl to depolarize cells (24).

Statistical analysis. For all quantitative data, means and standard errors of the means were computed. For multiple comparisons, ANOVA was performed and subsequent post hoc comparisons were done with Tukey's test. For all experiments, at least three replicates were included, and experiments were repeated at least three times. The counting of cells and nuclei on stained slides was performed in three fields that exhibited uniform distributions of cells.


   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
OPN in cardiac fibroblasts. We examined OPN–/– and WT mice to determine whether the lack of OPN expression affected the growth of cardiac tissues. Hearts from WT and OPN–/– mice were weighed before subsequent experimentation, and the weights were expressed as percentages of individual mouse body weights. When adjusted for individual body weights, the OPN–/– hearts were ~10% smaller as indicated by a weight ratio of 0.50 ± 0.04% compared with 0.56 ± 0.03% for the WT mice (P < 0.1). We examined the functional implications of OPN on post-myocardial infarction cardiac function.

Echocardiographic assessments of 9-wk-old WT and OPN–/– mice were performed 4 wk after infarction. There were no significant differences in heart rate (OPN–/–, 480 ± 60; WT, 460 ± 35 beats/min; P > 0.2), anterior wall thickness (OPN–/–, 0.134 ± 0.012 cm; WT, 0.134 ± 0.032 cm; P > 0.2), or left ventricular end-diastolic dimension (OPN–/–, 0.418 ± 0.034 cm; WT, 0.374 ± 0.064 cm; P > 0.2) between the OPN–/– and WT mice. However, the percent ejection fraction was significantly reduced in the OPN mice (WT, 48 ± 8%; OPN–/–, 30 ± 8%; P < 0.05), which indicates that OPN affected cardiac function. Furthermore, ex vivo cardiac function analyses using Langendorff preparations showed reduced contractile ability in OPN–/– mice compared with WT (+dP/dt mmHg/s: OPN–/–, 3,000 ± 600; WT, 3,567 ± 681; P < 0.05).

We determined whether OPN expression in vivo was increased in mouse ventricles after myocardial infarction followed by reperfusion injury. Immunoperoxidase staining of mouse ventricles showed very little OPN staining in 6-wk-old WT control mice (Fig. 1A). Within 12 days after injury, there was a marked increase in OPN staining of nonmyocytic cells, which at this time after injury were most likely fibroblasts (Fig. 1A). The staining was primarily cytoplasmic with weak staining of the surrounding extracellular matrix; this is consistent with OPN being synthesized and secreted as a cytokine and also with its possible retention within the cells (60). These findings suggested that cardiac injury promotes OPN expression in ventricular fibroblasts of WT mice.



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  		Fig. 1. Immunostaining of osteopontin (OPN) in cardiac tissues. A: immunolocalization of OPN in mouse hearts. Control ventricle (left) shows minimal OPN staining, whereas at 12 days postinjury (right), there is strong OPN staining in cells (black arrows) at the periphery of the infarcted myocardium. MI, myocardial infarction. B: at 12 days postischemia, ventricle walls exhibit numerous terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling (TUNEL)-positive nuclei (stained blue using TACS Blue) in both cardiomyocyte and noncardiomyocyte cells. Black bar, 50 µm. C: staining for propidium iodide (PI) and TUNEL in wild-type (WT, top) and OPN-null (OPN–/–, bottom) cardiac fibroblasts. Cells grown in 8-well chamber slides were treated with H2O2, stained with PI, fixed, permeabilized, and stained for TUNEL. Nuclear staining with 4',6-diamidino-2-phenylindole dihydrochloride (DAPI) was used to determine the total number of cells in a field. Only a few nuclei in the OPN–/– cardiac fibroblast cultures show both TUNEL and PI labeling despite the large number of PI-positive nuclei, whereas almost all of the WT cells stained for PI also stained for TUNEL. White bar, 20 µm.

 
Cell death. We determined whether myocardial infarction and postinfarction reperfusion are associated with increased cell death by apoptosis. In sections of mouse ventricle stained for TUNEL, we noted large numbers of TUNEL-positive cells after experimental infarction (Fig. 1B). We next examined the response of cultured mouse cardiac fibroblasts to oxidative injury that models postinfarction reperfusion. Cells were treated with 0.5 mM H2O2, stained with PI, fixed, stained with TUNEL reagent, and counterstained with DAPI (Fig. 1, C and D). Most of the nuclei of the WT cells were positive for PI and TUNEL after 4 h as shown by colocalization (yellow) of the red (PI) and green (TUNEL) fluorescence (Fig. 1C), whereas few OPN–/– cells were positive for PI and TUNEL (Fig. 1D). As the TUNEL assay is not absolutely diagnostic of apoptosis because of the possibility of detecting double-stranded RNA and other artifacts, we used distinct morphological features of necrosis and apoptosis (20, 25) to examine H2O2-induced death in more detail. The ultrastructural features of untreated cardiac fibroblasts (Fig. 2A) were compared with cells treated with 0.5 mM H2O2 (Fig. 2B). Chromatin condensation was observed in WT cells, whereas cardiac fibroblasts from OPN–/– mice exhibited signs of cellular and nuclear degeneration. However, although the integrity of the cellular and nuclear membranes of WT cells was preserved, the membranes in most of the OPN–/– cells were ruptured, and there was loss of cytoplasmic structure. These changes were consistent with the increased permeability to PI in OPN–/– fibroblasts (see Fig. 1, C and D) and the alterations of nuclear morphology shown by DAPI staining described below. Taken together with the absence of TUNEL staining, which is associated with endonuclease activity and the generation of DNA strand breaks, the morphological changes in the majority of the OPN–/– cells were indicative of nonapoptotic cell death.



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  		Fig. 2. Morphological characterization of cell death. A and B: transmission electron microscopy of cytoplasm and nuclear morphology in mouse cardiac fibroblasts 2 h after treatment with H2O2. Compared with untreated cells (A), H2O2-treated cells show loss of electron density in the cytoplasm of both WT and OPN–/– cells, whereas damage to the nuclear membrane was also evident in the OPN–/– cells (B). C–F: staining of nuclei with DAPI at different time points after stimulating cell death by H2O2. Compared with normal nuclear morphologies in control cultures (C), WT nuclei treated for 1 h show chromatin condensation, whereas OPN–/– nuclei are oncotic with no chromatin changes (D). After 2 h, WT cells show nuclear fragmentation, whereas damage to the nuclear membrane (arrow) is significant in many of the OPN–/– nuclei (E). After 8 h, WT nuclei are pyknotic, but the nuclear membranes remain intact, whereas chromatin in the OPN–/– cells is fragmented (F). Nuclei were photographed using a x40 magnification objective, 1.3-numerical aperture, oil-immersion lens.

 
Fluorescence microscopy of the DAPI-stained nuclei of the cardiac fibroblasts was used to identify progressive cell death-related changes at early and late stages of H2O2 treatment (Fig. 2, C–F). In untreated control cultures, the DAPI-stained nuclei were intact and of normal size and appearance in both WT and OPN–/– cells (Fig. 2C). After 15 min of H2O2 treatment, nuclear condensation was evident in some WT cells, whereas nuclear swelling was observed in most of the OPN–/– cells (Fig. 2D). After 2 h, nuclear fragmentation could be seen in nuclei of WT cells, whereas OPN–/– cells displayed extensive damage to the nuclear membrane (Fig. 2E). After 6–8 h, the WT nuclei were shrunken, and only nuclear debris could be detected in the OPN–/– cultures (Fig. 2F).

Quantification of cell death. Cell death in cultures of murine cardiac fibroblasts treated with H2O2 was examined over time using cell detachment, permeability to PI, and TUNEL staining as outcome measures (Fig. 3). The number of DAPI-stained OPN–/– and WT cells that remained attached to the culture dishes was reduced progressively over the first 2 h of H2O2 treatment so that after 8 h, only ~50% of the cells remained attached (Fig. 3A; P < 0.01). However, compared with WT cells, higher percentages of OPN–/– cells exhibited PI-positive nuclei (P < 0.01), and this percentage increased from 22% in untreated cells to >90% after 8 h. In contrast, <5% of untreated WT cells were PI positive, and the percentage of PI-positive cells increased more slowly than the OPN–/– cells, exceeding 80% only after 8 h. Thus at 2 h, only 50% of the attached WT cells showed PI staining, whereas most of the OPN–/– cells were permeable to PI (Fig. 3B; P < 0.01). Despite the higher death rates in the OPN–/– cells, WT cells exhibited more than twofold higher percentages of PI- and TUNEL-positive cells after 8 h of H2O2 treatment (Fig. 3C; P < 0.01). Notably, after 2 h of treatment, almost all of the WT cells that were PI positive were also TUNEL positive, whereas only 15% of the PI-positive OPN–/– cells were TUNEL positive. We also examined the percentages of PI- and TUNEL-positive cells in the detached cell populations (floaters; Fig. 3D). There were >10-fold more TUNEL-positive WT than OPN–/– cells (P < 0.001), and the relative number of TUNEL-positive OPN–/– cells was very low. Collectively, these data suggested that cell death in the majority of the OPN–/– cells was not via an apoptotic pathway.



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  		Fig. 3. Time-course analysis of cardiac fibroblast death. Permeability to PI and TUNEL staining was determined in mouse cardiac fibroblasts as a percentage of total DAPI-stained nuclei over an 8-h period after stimulation with H2O2 for 30 min. Results are expressed as means ± SE. A: measurements of cell detachment, determined by counting the number of DAPI-stained nuclei per field, were not significantly different between OPN–/– and WT cells (P > 0.2). B: analysis of PI-stained cells remaining attached over 4 h showed that OPN–/– cardiac fibroblasts exhibited increased permeability more rapidly than WT cells (% PI). After 8 h, the percentages of PI-stained cells were not significantly different between WT and OPN–/– cells. C: analysis of apoptotic cells (TUNEL+PI/DAPI cells) over the time course showed a greater than twofold higher percentage of apoptotic WT cells compared with OPN–/– cardiac fibroblasts. D: analyses of detached cells 4 h after H2O2 stimulation showed that most of the OPN–/– and WT cardiac fibroblasts were permeable to PI, whereas only the WT cells exhibited significant numbers of TUNEL-positive cells (>10-fold more than the OPN–/– detached cells).

 
Caspase-3. Because active caspase-3 is an indicator of apoptotic cell death and provides another, separate indicator of apoptosis, cardiac fibroblasts were examined for caspase-3 activity 8 h after treatment with H2O2 (Fig. 4). Abundant cytoplasmic staining for active caspase-3 was evident in most of the WT cells but in only a few of the OPN–/– cells (Fig. 4A). Quantification of active caspase-3-positive cells after 30 min and 8 h of H2O2 treatment showed (Fig. 4B) that OPN–/– cardiac fibroblasts were fourfold less likely to exhibit staining than WT cultures (P < 0.001). At 8 h, only ~20% of the attached OPN–/– cells showed active caspase-3 staining.



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  		Fig. 4. Involvement of active caspase-3. A: immunofluorescence staining for active caspase-3 (C3) in mouse cardiac fibroblasts treated with H2O2 for 8 h. Cells were permeabilized and immunostained for active caspase-3. Most of the WT cells but only a few OPN–/– cells exhibited caspase-3 staining. B: quantification of the percentage of mouse cardiac fibroblasts stained for active caspase-3 after stimulation with H2O2 was determined after 30 min and after 8 h. At both time points, OPN–/– cells showed fourfold fewer cells with active caspase-3 staining than WT cells. Results are expressed as means ± SE. C: treatment with a caspase-3 inhibitor reduced TUNEL staining more than fourfold and membrane permeability (PI, open bars) more than twofold in WT cardiac fibroblasts. In contrast, apoptosis was reduced <60% for OPN–/– cells with no effect on cell permeability (PI, hatched bars).

 
Cells were also incubated with a caspase-3 inhibitor 3 h before treatment with H2O2. After H2O2 treatment, the caspase-3 inhibitor reduced the percentage of TUNEL-positive cells in WT cultures by more than fourfold, whereas there was a decrease of only ~30% in OPN–/– cultures (Fig. 4C; P < 0.01). Cell membrane permeability, examined by PI staining, was reduced by 50% in WT cells treated with H2O2 (Fig. 4C; P < 0.01), whereas OPN–/– cells did not show any change in permeability in response to the caspase-3 inhibitor.

Membrane permeability. The possible loss of membrane permeability suggested by the PI staining was measured in vital OPN–/– cells during the first 30 min of H2O2 treatment by flow-cytometric analyses of DiBAC4(3) fluorescence (Fig. 5, A and B). Incubation of cell suspensions with 100 mM KCl was used as a positive control to depolarize cells and cause increased fluorescence of DIBAC4(3) (24). There was a time-dependent increase in DiBAC4(3) fluorescence in most of the OPN–/– cells 30 min after treatment with H2O2.



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  		Fig. 5. Cellular and mitochondrial membrane depolarization. A and B: loss of membrane permeability observed in OPN–/– cardiac fibroblasts by PI staining was confirmed by time-mode analyses of membrane depolarization. After 30 min of H2O2 stimulation (A), OPN–/– cardiac fibroblasts were incubated with bis-(1,3-dibutylbarbituric acid)trimethine oxonol [DiBAC4(3)], and the fluorescence of single cell suspensions were measured (excitation, 488 nm; emission, 520 nm) at time 0 (green tracing), after 15 min (pink tracing), and after 30 min (blue tracing). Notably, a second attempt to depolarize the cells with a second dose of H2O2 was unsuccessful, which indicates leakage of DiBAC4(3) through damaged membranes of dead cells. Incubation of cells in 100 mM KCl for 30 min served as a positive control for membrane depolarization (B). C–F: bivariate plots show flow-cytometric analyses of mitochondrial membrane potential in cardiac fibroblasts stained with 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide (JC-1). Cells were monitored for changes in red fluorescence (y-axis; red arrow), which indicate normally functioning mitochondria, compared with increases in green fluorescence and reduced red fluorescence, which indicate mitochondrial depolarization, a component of the cell death process (x-axis; green arrow). WT and OPN–/– cardiac fibroblasts were incubated in the absence (C) or presence (D) of cyanide [5 µmol/µl carbonyl cyanide m-chlorophenylhydrazone (CCCP) for 10 min], which dissipates mitochondrial membrane potential and serves as a positive control. Cyanide-treated cells exhibited mainly reduced red fluorescence and increased green fluorescence in both the WT and OPN–/– cells. Similar results were observed when both cell types were stimulated with H2O2 (E). Shift to green fluorescence was more marked in OPN–/– cells. Pretreatment of cells by addition of 5 µg/ml of purified soluble OPN showed a protective effect against mitochondrial membrane depolarization induced by oxidative injury. Effect of OPN was more pronounced in the OPN–/– group as shown by higher number of cells with increased red fluorescence (F).

 
Mitochondrial membrane potential. Because mitochondrial membrane potential provides an earlier marker for cell death than TUNEL or PI staining (28), we used the dual-emission dye JC-1 to analyze the effects of H2O2 on OPN–/– and WT cells (Fig. 5, C–F). Low mitochondrial potential, as estimated by increased green fluorescence and reduced red fluorescence, was observed after incubation with CCCP (1 µmol), which was used as a positive control for both types of cells (Fig. 5D). H2O2 caused an increase of green fluorescence and a reduction of red fluorescence in most of the OPN–/– and WT cells (Fig. 5E). Addition of 5 µg/ml OPN to the medium reduced cell death as shown by the higher number of cells that exhibited red fluorescence (Fig. 5F). The effect of the exogenous OPN was more clearly evident in the OPN–/– cells, although some effect was also apparent in the WT cells.

Specificity of apoptotic induction. To determine whether the induction of nonapoptotic cell death in OPN–/– cells was restricted to H2O2, which induces death by oxidative injury, we analyzed the effects of other agents that are known to induce apoptotic cell death. Staurosporine, a protein kinase C inhibitor (57), thapsigargin, a Ca2+-ATPase inhibitor (11), actinomycin D, which induces apoptosis through cytoplasmic translocation and cleavage of nuclear protein RNA helicase and thus interferes with transcription (42), and cycloheximide, which inhibits protein synthesis and activates caspase-dependent death pathways (36) were used to treat mouse cardiac fibroblasts. All of these reagents increased the PI permeability of WT and OPN–/– mouse cardiac fibroblasts to a similar extent (Fig. 6A; P < 0.01). However, when these cells were examined for DNA strand breakage by TUNEL staining (Fig. 6B; P < 0.01), only a small proportion of OPN–/– cells were TUNEL positive, whereas a high percentage of WT cells were TUNEL positive. We also examined the percentage of detached cells that were TUNEL positive (Fig. 6C). These data showed that detached OPN–/– cells stained with TUNEL were much less prevalent than WT cells particularly in cells treated with either thapsigargin, staurosporine, or cycloheximide (differences between WT and OPN–/– cells were all significantly different, P < 0.02, for each treatment).



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  		Fig. 6. Effects of several different apoptotic agents on cell death in WT and OPN–/– cardiac fibroblasts were determined. A: PI used alone. B: PI and TUNEL staining. Data for staurosporine (1 µM), actinomycin D (500 ng/ml), thapsigargin (1 µM) and cycloheximide (100 µg/ml) are compared with H2O2. OPN–/– cardiac fibroblasts exhibited small but consistently higher percentages of PI permeability than WT cells. However, compared with WT cells, only a small proportion of OPN–/– cells were TUNEL positive. There were no TUNEL-positive OPN–/– cells in the cycloheximide-treated group. Data are percentage means ± SE. C: detached cells were examined with TUNEL staining. WT cells exhibited at least 20% of TUNEL-positive cells for all death stimulants, whereas OPN–/– cardiac fibroblasts showed <20% TUNEL-positive cells and only after thapsigargin, staurosporine, or actinomycin-D treatment.

 
Effect of restoring OPN on induction of cell death. The ability of exogenous OPN to prevent cell death varies depending on the mode of presentation (18, 39, 45, 49) and cell type (56). Our preliminary data using soluble or coated OPN confirmed these previous reports. To minimize the variations associated with addition of exogenous OPN, we examined cell death in an OPN–/– mouse fibroblast cell line that was stably transfected with a Zn2+-activated OPN expression vector to restore endogenous OPN expression. When OPN–/– cells were stimulated with Zn2+ 48 h before oxidative injury, the percentage of PI-stained cells was reduced to control levels (P < 0.01), whereas the percentage of TUNEL- and PI-stained cells was not significantly different than controls subjected to oxidative injury but without Zn2+-activated OPN expression (Fig. 7; P > 0.2). As expected, the percentage of PI-stained, nontransfected WT cells was not affected by treatment with Zn2+ nor did Zn2+ affect the percentage of TUNEL- and PI-stained cells (data not shown).



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  		Fig. 7. Rescue of cell death by OPN. Ability of endogenously expressed OPN to prevent cell death in a mouse OPN–/– fibroblast line treated with H2O2 was analyzed by fluorescence microscopy of cytospin preparations. A: cells were stained with PI. B: cells were stained with TUNEL, PI, and DAPI. Cells were stably transfected with an expression vector that contained OPN cDNA driven by a Zn-activated metallothionein promoter. Cells were stimulated with Zn2+ for 4 h to induce OPN expression, and 48 h later, cell death was induced with H2O2 followed by 4 h of recovery in serum-free conditions. After recovery, cells were trypsinized, counted, and analyzed by PI and TUNEL staining using fluorescence microscopy of cytospin preparations. Stimulation of OPN expression exerts little effect on TUNEL staining but rescues permeability to PI.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The development of fibrosis in response to postischemic reperfusion injury compromises cardiac function. Although the molecular mechanisms that lead to cardiac fibrosis and scar formation are still obscure, in most cases the persistence of fibroblasts is an impediment to the healing of scar tissue (13). Here cells from mice with a targeted disruption of the OPN gene were used to evaluate the importance of OPN in the survival of cardiac fibroblasts subjected to oxidative injury. Under these conditions, WT cardiac fibroblasts undergo apoptosis characterized by the formation of apoptotic bodies and the expression of active caspase-3. In contrast, in the absence of OPN expression, oxidative injury primarily induces a nonapoptotic death pathway with morphological and biochemical characteristics consistent with necrosis (23, 43, 50). Zn2+-activated OPN expression in OPN–/– cells reduced PI staining to control levels. Collectively, these studies indicate that OPN promotes the survival of cardiac fibroblasts, cells that contribute to the formation of fibrotic tissue in reperfusion injury.

Cell death is an important part of developmental and wound-healing processes (30) and occurs in a cell type-specific manner in response to signals in the local environment (2, 9, 17). These signals can be simulated in vitro with different cell death-inducing agents. Here we used H2O2 to model the generation of free radicals produced by inflammatory cells that can induce cell death. Using PI staining to measure cell permeability, OPN–/– cardiac fibroblasts showed marked increases of permeability that were accompanied by irreversible cell membrane depolarization in most cells. In contrast with dying WT cells in which PI-stained cells were also positive for TUNEL and active caspase-3 (indicative of apoptosis), most of the OPN–/– cells that were PI positive were not stained for both TUNEL and active caspase-3. Furthermore, electron and fluorescence microscopy of WT cardiac fibroblasts induced to undergo cell death revealed the classical morphological signs of apoptosis such as chromatin condensation, nuclear fragmentation, and cell shrinkage. In contrast, the majority of the OPN–/– cardiac fibroblasts exhibited swelling, rupture of the cellular and nuclear membranes, and cellular disintegration. These morphological characteristics indicate that WT cells undergo apoptosis, whereas most of the OPN–/– cells, under the same conditions, exhibit features of necrosis (23, 38, 50). This is further supported by the inability of the caspase-3 inhibitor to significantly reduce the nonapoptotic cell death while it reduced apoptosis in the OPN–/– and WT cardiac fibroblasts. Notably, in contrast with apoptosis, in which nucleoproteins generated by enzymatic internucleosomal splicing are packaged into apoptotic bodies and the cells are cleared by phagocytic cells, the release of autoantigens in necrotic cell death can trigger and sustain inflammatory processes (43).

Irreversible depolarization of cell membranes and reduction of mitochondrial membrane potential indicate damage to cells that is associated with cell death (6, 24, 32). The speed of these events in OPN–/– cardiac fibroblasts emphasizes the role of OPN expression in these events and also confirms the increased plasma-membrane permeability observed with PI. However, these alterations are not exclusive to apoptosis. Release of mitochondrial proteins such as cytochrome c, which occurs in apoptosis as a reaction to oxygen radicals (7), can also occur in response to other cell death pathways. With the recognition of new forms of cell death that occur in cardiac ischemia-reperfusion (21), many cellular events could be the result of both apoptosis and necrosis, or "necrapoptosis" (27). Thus an increase in mitochondrial potential could lead to either apoptosis or necrosis (43).

We found sparse staining for active caspase-3 in dying cardiac OPN–/– fibroblasts. Active caspase-3 is considered a valid indicator of apoptosis (19, 44, 53). Inhibition of caspase-3 activity directs cell death through a necrotic instead of an apoptotic pathway (48). Thus low levels of active caspase-3 in the OPN–/– cardiac fibroblasts suggest nonapoptotic death possibly by necrosis. The observation that the caspase-3 inhibitor did not alter cell death may correlate OPN expression with caspase-3 activity.

OPN has been implicated as a survival factor in diverse cell types including endothelial cells (26, 45, 49), vascular smooth muscle cells (56), renal epithelial cells (40, 47), melanoma (18), B cells (35), and breast cancer cells (39). However, there are cell-specific differences in the ability of exogenous OPN to prevent cell death. Native OPN in solution inhibits apoptosis of human umbilical vein endothelial cells (26) but not vascular smooth muscle cells (56). To overcome the complications associated with the addition of exogenous OPN, we endogenously expressed OPN in transfected OPN–/– cells. With this approach, endogenous OPN strongly reduced PI staining but not TUNEL staining. Thus in fibroblasts, endogenous OPN expression may be important in preventing cell necrosis. Notably, previous studies have indicated that some OPN may be retained and function inside fibroblasts (60). This intracellular OPN could be important in preventing cell necrosis and may account for some of the cellular staining seen in Fig. 1. However, more definitive experiments are needed to determine the relative contributions of secreted and intracellular OPNs in cell survival.

Our results suggest that cardiac fibroblasts that respond to reperfusion injury in vivo survive the stimulus of oxidative burst by expression of OPN possibly including an intracellular OPN isoform (24). Moreover, the ability of OPN to protect cardiac fibroblasts from cell death may explain the impaired connective-tissue wound healing that is observed in OPN–/– mice. Thus OPN plays a pivotal role in tissue repair and in the subsequent fibrosis and cardiac muscle inflammation (33, 54) that typically occur after myocardial injury.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was funded by Grant T4819 from the Ontario Heart and Stroke Foundation (to C. A. McCulloch) and Grants MOP-36333 (to R. Zohar) and MOP-36332 (to J. Sodek and C. A. McCulloch) from the Canadian Institutes of Health Research.


   ACKNOWLEDGMENTS
 
We thank Drs. D. T. Denhardt and S. R. Rittling for their generosity in providing the OPN–/– mice and cell lines, Dr. L. W. Fisher for the LF123 antiserum, Dr. F. Dawood for performing the ischemia-reperfusion model and analyses, and Dr. Y. Gao for help with histology.


   FOOTNOTES
 

Address for reprint requests and other correspondence: R. Zohar, Faculty of Dentistry, Univ. of Toronto, 124 Edward St., Rm. 464A, Toronto, ON M5G 1G6, Canada (E-mail: ron.zohar{at}utoronto.ca)

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
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 DISCUSSION
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