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Am J Physiol Heart Circ Physiol 288: H293-H301, 2005. First published September 9, 2004; doi:10.1152/ajpheart.00165.2004
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TNF-{alpha} induces proliferation or apoptosis in human saphenous vein smooth muscle cells depending on phenotype

Zhongbiao Wang,1 Pulipaka J. Rao,2 Manuel R. Castresana,3 and Walter H. Newman1,3,4

1Division of Basic Medical Science, and Departments of 2Psychiatry and Behavioral Science, 3Anesthesiology, and 4Surgery, Mercer University School of Medicine, and Medical Center of Central Georgia, Macon, Georgia

Submitted 17 February 2004 ; accepted in final form 1 September 2004


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Tumor necrosis factor (TNF)-{alpha} is implicated in development of restenotic and atherosclerotic vascular lesions, which are pathological processes involving both proliferation and apoptosis of vascular smooth muscle cells (VSMCs). Human VSMCs were recently found to contain heterogeneous subpopulations. We therefore examined whether TNF has different effects on distinct subpopulations of VSMCs. With the use of cloning techniques, two stable subpopulations of VSMCs were isolated from human saphenous vein: spindle- and epithelioid-shaped smooth muscle cells (Sp- and Ep-SMCs, respectively). We found that TNF stimulated growth in Sp-SMCs but had a toxic effect on Ep-SMCs. TNF did not induce apoptosis in Sp-SMCs as determined by nuclear staining and cellular DNA electrophoresis. In contrast, the reduction of viability in Ep-SMCs was associated with induction of apoptosis as characterized by cellular DNA fragmentation and nuclear condensation. Higher levels of the TNF-R1 receptor subtype were detected in membrane preparations from Ep-SMCs than in membranes from Sp-SMCs. Activation of caspase-3 was also selectively induced in Ep-SMCs but not in Sp-SMCs. Cycloheximide, an inhibitor of protein synthesis, enhanced the toxicity of TNF in Ep-SMCs. This effect of cycloheximide was not seen in Sp-SMCs. The data presented here demonstrate for the first time that TNF either promotes growth or induces apoptosis in human VSMCs depending on phenotype.

tumor necrosis factor-{alpha}; spindle; epithelioid; cycloheximide; atherosclerosis; stenosis

TEN YEARS AFTER CORONARY BYPASS surgery, 40% of vein grafts are occluded, and half of the remaining patent grafts show significant stenosis (29). This disease process is associated with accumulation of vascular smooth muscle cells (VSMCs) within a hyperplastic neointima and eventual development of accelerated atherosclerosis. The neointima thickens the vessel wall, narrows the lumen, and represents a net increase in a population of vascular cells undergoing both proliferation and apoptosis (13, 14, 20, 23, 27). Additionally, formation of the neointima is characterized by an inflammatory process with the expression of numerous inflammatory cytokines including tumor necrosis factor-{alpha} (TNF) by the expanding population of VSMCs (8, 21, 26, 36, 37, 42). TNF is expressed by VSMCs within atherosclerotic plaques, in the neointima of balloon-injured arteries, and in diseased vein grafts (8, 7, 21, 35, 37, 42). Furthermore, in a mouse model of arterial injury, mice lacking functional TNF developed 14-fold less neointima than wild-type mice (35). Therefore, this multifunctional cytokine may play a significant role in the formation and development of neointima.

By signaling through two cell membrane receptors (p55 and p75, also known as TNF-R1 and -R2), TNF can induce mitogenic, antiapoptotic, and proapoptotic effects through transcriptional and translational mechanisms that require new protein synthesis. Both TNF-R1 and -R2 share homology with the death receptor Fas (44, 45). Like Fas, TNF-R1 has a death domain that induces signaling that activates caspase, whereas TNF-R2 lacks the death domain (45). The combination of TNF with its receptors induces multiple signals that result in effects on cells that are dependent on cell type. In studies of the activity of TNF on VSMCs, this cytokine has been consistently shown to be a chemoattractant; however, the effects on proliferation of VSMCs are conflicting (11, 12, 15, 19, 21, 22, 28, 32, 38, 39, 49, 53). Several investigations reported that TNF itself had no effect on VSMC proliferation (19, 28). In contrast, there are other reports that TNF induces proliferation of VSMCs through NF-{kappa}B-directed transcription mechanisms (38, 39). Similarly, investigations on the induction of apoptosis or the inhibition of proliferation by TNF are not consistent. Some investigators found no proapoptotic activity for TNF, whereas others found that under certain conditions, TNF was proapoptotic and activated apoptosis-related caspase-3 (11, 12, 15, 22, 32, 53). Thus it seems possible that TNF can induce both proliferation and apoptosis in VSMCs and regulate vascular intimal cellularity by a balance of these two effects.

Based on results from clonal studies (2, 18, 25, 40, 53), VSMCs are heterogeneous and manifest at least two morphologically distinct phenotypes: spindle- and epithelioid-shaped VSMCs (Sp- and Ep-SMCs, respectively). These subtypes are associated with different proliferative, apoptotic, and chemotactic behaviors in response to diverse stimuli (2, 18, 25). For example, ANG II, which is generally thought to be a mitogenic agent, induced apoptosis in rat aorta-derived Ep-SMCs but not in Sp-SMCs (2). In Ep-SMCs derived from human internal thoracic artery, PDGF was mitogenic, but it was not mitogenic in Sp-SMCs (25). Therefore, it seems possible that these variations in the effects of TNF on VSMCs originate from different responses of VSMC subtypes. In the present study, we tested this hypothesis. We cloned Ep- and Sp-SMCs from human saphenous vein. In these cells, we found that TNF induced proliferation in Sp-SMCs and was proapoptotic in Ep-SMCs. TNF-induced apoptosis in Ep-SMCs was associated with higher TNF-R1 levels than in Sp-SMCs and greater caspase-3 activation, which was not seen in Sp-SMCs. The apoptotic effect of TNF in Ep-SMCs was potentiated by inclusion of the protein synthesis inhibitor cycloheximide (CHX).


   MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Materials. Unless otherwise stated, all materials were from Sigma (St. Louis, MO). Recombinant human TNF was from PeproTech (Rocky Hill, NJ). Neutralizing antibodies to TNF were purchased from R&D Systems (Minneapolis, MN). Monoclonal anti-p55 TNF receptor subtype (TNF-R1) antibody was from Santa Cruz Biotechnology (Santa Cruz, CA). The protein assay kit was from Bio-Rad Laboratories (Hercules, CA). Cell culture reagents including medium and serum were purchased from GIBCO (Grand Island, NY) and Atlanta Biologicals (Norcross, GA). The enhanced chemiluminescence Western blotting analysis system was from Amersham (Piscataway, NJ). The caspase-3 substrate Ac-DEVD-p-nitroaniline (Ac-Asp-Glu-Val-Asp-pNA) was from Alexis.

Culture of VSMCs. With Institutional Review Board approval, VSMCs were isolated by a medial explant method from segments of saphenous veins obtained from three male patients who underwent coronary artery bypass surgery. Primary VSMCs were cultured and identified as previously described (31, 50). The primary cultures were homogeneously immunostained with anti-{alpha}-smooth muscle-actin ({alpha}-SM actin) antibodies but lacked immunoreactivity for the endothelial cell marker von Willebrand factor.

Cell cloning. Before cloning was started, primary cultures of VSMCs at near confluence were incubated in 20% fetal bovine serum (FBS)-Dulbecco's modified Eagle's medium (DMEM) for 2 days. This conditioned medium was collected and centrifuged at 200 g for 10 min and then filtered through 0.45-µm filters. The conditioned medium was stored at –80°C and was thawed just before use. Cloning medium was prepared by mixing the conditioned medium with 20% FBS-DMEM in a 1:1 proportion. Single cell suspension was prepared from the primary VSMC cultures and then plated at a density of ~2 cells/cm2 in 0.2% gelatin-coated, 100-mm petri dishes in the cloning medium. Single cell-derived individual colonies with uniform appearance were surrounded by cloning rings, released with 0.25% trypsin-0.02% EDTA, and expanded in 10% calf serum-DMEM. Multiple clones were generated and expanded using these procedures. Two types of cells with distinct morphology, Sp- and Ep-SMCs, were obtained. In all of the subsequent experiments, clones at passages 10–12 were studied under identical conditions.

Western blot analysis. Western blot analysis was performed as described previously (48). To analyze expression of {alpha}-SM actin, calponin, smooth muscle-myosin heavy chain (SM-MHC), and cytokeratin-8 (CK-8), the cells were lysed in buffer that contained 50 mM Tris·HCl (pH 8.0), 150 mM NaCl, 0.1% SDS, 100 µg/ml PMSF, 1 µg/ml aprotinin, and 1% Nonidet P-40 (NP-40). For detection of TNF-R1, cell membranes were isolated essentially as described by Zhang and Morrison (54). Briefly, the cells were scraped from plates and collected by centrifugation in PBS. Plasma membranes were isolated by incubating the cells at 4°C in hypoosmotic buffer that contained 10 mM Tris·HCl (pH 7.4) and 1 mM PMSF for 45 min followed by homogenization with Dounce homogenizer. After centrifugation at 750 g for 5 min, the resulting supernatant was recentrifuged at 15,000 g for 20 min, and the pellet containing crude plasma membrane suspension was resuspended in 25 mM Tris·HCl (pH 7.4), 25 mM MgCl2, and 1 mM PMSF. The membrane suspension was briefly sonicated three times for 10 s on ice. Protein concentrations of the whole cell lysates and the plasma membrane suspension were determined with Bio-Rad DC protein assay kit. After suspensions were boiled for 5 min, aliquots of the protein samples with equal amounts of protein were resolved by SDS-PAGE and transferred to nitrocellulose membrane (Bio-Rad) in 25 mM Tris·HCl, 20% methanol, and 192 mM glycine. The equivalence of sample loading was confirmed by Ponceau S staining of the membranes and Coomassie brilliant blue staining of gels with the same loading of protein samples in parallel experiments. The primary antibody used in immunoblotting was a monoclonal antibody to the corresponding antigen. Detection of antibody binding was performed using Amersham chemiluminescence reagents employing the protocols recommended by the manufacturer and exposing the blots to X-ray film to visualize protein bands. Prestained protein markers were used for molecular mass determinations.

Assay for cell viability. A colorimetric assay was used to analyze proliferative or cytotoxic effects of TNF on the cloned VSMCs. This assay detects cell viability by assessing the mitochondrial-dependent reduction of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrozolium bromide (MTT) to formazan, is dependent on the number of living cells, and has been widely applied in cell proliferation and cytotoxicity experiments (9). Preliminary experiments were performed to adjust cell plating conditions such that the cloned VSMCs reached ~40–50% confluence before TNF stimulation and did not reach 100% confluence at the end of these experiments. This was done to prevent the influence of cell contact on the experiments. Briefly, Sp- and Ep-SMCs at a density of 3 x 103 cells/well were seeded into 96-well plates in DMEM supplemented with 10% calf serum and were cultured for 24 h. The cells were then made quiescent by incubation in DMEM that contained 0.2% calf serum for an additional 2 days followed by addition of TNF at 1, 10, or 100 ng/ml in the absence or presence of 1 µg/ml CHX. Cells not treated with TNF served as controls. At the end of each experiment, cells were incubated with 0.5 mg/ml MTT dissolved in phenol red-free DMEM for 4 h. Formazan was extracted with DMSO and quantitated spectrophotometrically using a microplate reader. Viability of treated cells was expressed relative to control cells (relative viability). An increase in relative viability represents cell proliferation, and a decrease indicates cytotoxicity.

Apoptosis assay. Sp- and Ep-SMCs at a density of 9 x 104 cells/well were cultured in six-well plates in DMEM supplemented with 10% calf serum for 24 h. The cells were then made quiescent by incubation in DMEM that contained 0.2% calf serum for an additional 2 days, which was followed by addition of 10 ng/ml TNF. Apoptosis was analyzed 48 h later via both genomic DNA electrophoresis and fluorescence microscopy essentially as previously described (48).

For electrophoretic analysis of genomic DNA, adherent (harvested by scraping) and nonadherent cells were pooled and lysed in 50 mM Tris·HCl (pH 7.5), 20 mM EDTA, and 1% NP-40. After centrifugation for 5 min at 800 g and 4°C, the supernatant was collected, and extraction of the pellets was repeated twice. Pooled supernatants were precipitated with 7.5% polyethylene glycol 8,000 and 1 M NaCl for 15 min at 4°C. After centrifugation at 16,000 g for 20 min at room temperature, the supernatants were brought to 1% SDS, digested with 100 µg/ml DNase-free RNase A at 50°C for 1 h, digested with 500 µg/ml proteinase K at 37°C for 16 h, and then precipitated with isopropanol for 30 min at –20°C. After centrifugation, each pellet was dissolved in Tris·HCl-EDTA (pH 7.6). Equal amounts of the extracted genomic DNA were separated in 1% agarose gel by electrophoresis and stained with 0.5 µg/ml ethidium bromide. Ladder formation of oligonucleosomal DNA was detected under UV light.

For microscopic assay of apoptosis, after treatment, adherent (harvested by trypsinization) and nonadherent cells were pooled. Cells were pelleted by centrifugation, washed once with PBS, fixed by incubation in 4% paraformaldehyde for 30 min at room temperature, and then washed again with PBS to remove the fixative. The fixed cells were resuspended in PBS that contained 5 µg/ml Hoescht 33258 and incubated at room temperature for 15 min in the dark. Aliquots of cells were placed on glass slides and examined for cells with apoptotic morphology (nuclear condensation and chromatin fragmentation). To quantify apoptosis, 300 nuclei from random microscopic fields were analyzed. Data are presented as the mean percentages of apoptotic cells.

Caspase-3 activity assay. Caspase-3 activity was detected by a spectrophotometric procedure as previously described (43). Cells were plated and growth arrested as described (see Apoptosis assay). The cells were then treated with 10 ng/ml TNF for 24 h. After experimental treatments, cells in six-well plates were lysed in 0.5 ml of lysis buffer that contained 0.5% Triton X-100 or 0.25% NP-40, 2 mM EDTA, 1 mM PMSF, and 10 mM DTT. The cell lysates were collected and centrifuged in a bench-top centrifuge at 10,000 g for 5 min, and protein concentrations were determined by using a Bio-Rad protein assay dye reagent. Lysates were stored at –80°C until assayed. For detection of caspase-3 activity, aliquots of lysates containing an equal amount of protein were mixed with equal volumes of ICE buffer {200 mM HEPES-KOH (pH 7.5), 20% sucrose, 10 mM DTT, and 0.2% 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonate} that contained 4 µM Ac-DEVD-p-nitroaniline (the caspase-3 substrate) in Eppendorf tubes, were mixed, and were transferred to 96-well plates and incubated for 4 h at 37°C. The colorimetric release of p-nitroaniline from the Ac-DEVD-p-nitroaniline substrate was monitored at 405 nm.

Statistical analysis. Results are expressed as means ± SE for a minimum of three experiments using cloned cells derived from the veins of three patients. Paired data were evaluated by Student's t-test. Multiple comparisons were evaluated by ANOVA. P values <0.05 were considered to be statistically significant.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
VSMC phenotypes. We first used an explant technique to isolate primary VSMCs from human saphenous veins. After the primary cultures were identified as smooth muscle by {alpha}-actin staining, the cultures were cloned using a ring technique. Multiple clones were generated from each primary culture. As shown in Fig. 1A, these clones could be divided into two categories based on distinctly different morphology: Sp- and Ep-SMCs. Sp-SMCs grew with overlapping cells, whereas Ep-SMCs were smaller than Sp-SMCs and appeared to be contact inhibited. These two distinct subpopulations of SMCs were characterized by expression of smooth muscle proteins associated with various stages of differentiation (1, 16, 17, 24). These proteins include {alpha}-SM actin, calponin, SM-MHC, and CK-8. As shown in the Western blot in Fig. 1B, both clones expressed {alpha}-SM actin (43 kDa), calponin (34 kDa), and SM-MHC (200 kDa). However, expression of calponin and SM-MHC was more pronounced in Sp-SMCs than in Ep-SMCs. CK-8 (52 kDa) was detected only in Ep-SMCs. The morphology of each clone has been maintained over time (>22 mo at this writing) and with freezing-thawing procedures. Culturing the cells with conditioned medium or gelatin did not alter the morphology. Recloning of selected clones yielded cells with identical morphology. Thus these clones appear stable.



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  		Fig. 1. Two subpopulations of vascular smooth muscle cells (VSMCs) isolated from human saphenous vein. A: clones of VSMCs could be divided into two categories based on their distinctly different morphology: spindle and epithelioid shaped smooth muscle cells (Sp- and Ep-SMCs, respectively). Magnification, x100. B: both clones expressed differentiation markers of smooth muscle cells including {alpha}-smooth muscle ({alpha}-SM) actin, calponin, smooth muscle myosin heavy chain (SM-MHC), and cytokeratin-8 (CK-8) as determined by Western blots. These results are typical of three experiments. Clones were generated from primary cultures of VSMCs isolated from human saphenous veins. Multiple clones were generated from the primary culture with a cloning ring technique. To analyze expression of smooth muscle differentiation proteins, the cells were lysed. Equal amounts of cell lysates were resolved by SDS-PAGE. Separated proteins were transferred onto nitrocellulose membranes and probed with anti-{alpha}-SM actin, anti-calponin, anti-SM-MHC, and anti-CK-8 antibodies, respectively. Antigen-antibody bindings were visualized using enhanced chemiluminescence assay.

 
TNF either induced proliferation or reduced cell viability depending on phenotype. To examine the effects of TNF on viability of these two different subpopulations of VSMCs, MTT assay, which is a commonly used technique to determine cell viability, was performed. As shown in the time-course study (Fig. 2, A and B), in the absence of TNF, both subpopulations of VSMCs could be maintained for up to 3 days in the 0.2% calf serum without a significant change of cell viability. TNF (10 ng/ml) stimulated proliferation of Sp-SMCs as assessed by an increment in viability. Significant proliferation of Sp-SMCs was detected 48 h postexposure to TNF and was maintained at 72 h. In contrast, TNF at 10 ng/ml was toxic to Ep-SMCs, because cell viability was reduced in a time-dependent fashion. The toxic effect on Ep-SMCs was initially detected after 24 h poststimulation with TNF and increased with time up to 72 h. Experiments in which cells were counted after staining with Trypan blue yielded similar results to those using MTT assay (data not shown). The effect of TNF in both clones was blocked by preincubation with anti-TNF antibody, which further confirmed the ligand-induced effects (data not shown). We also determined the concentration-dependent effects of TNF on both subsets of VSMCs after 48 h of stimulation. As shown in Fig. 3A, incubation with TNF at concentrations of 1, 10, and 100 ng/ml induced proliferation in Sp-SMCs. Maximal stimulation was reached by incubation with 10 ng/ml TNF. As seen in Fig. 3B, TNF caused cytotoxicity in Ep-SMCs in a concentration-dependent manner.



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  		Fig. 2. Tumor necrosis factor-{alpha} (TNF) induced proliferation in Sp-SMCs but was cytotoxic in Ep-SMCs. A: in Sp-SMCs, 10 ng/ml TNF (TNF 10) significantly increased proliferation at 2 and 3 days. B: in contrast with A, TNF was cytotoxic in Ep-SMCs and significantly reduced viability in a time-dependent manner. Sp- and Ep-SMCs at a density of 3 x 103 cells/well in 96-well plates were growth arrested in 0.2% calf serum-DMEM for 48 h before addition of 10 ng/ml TNF. Cell viability was measured using 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrozolium bromide (MTT) assay immediately before (time 0) and after 1, 2, or 3 days of incubation with the cytokine. See text for detailed discussion of MTT assay. Values are means ± SE for three experiments conducted in cells isolated from saphenous veins from three patients. Not significant (NS), P > 0.05 vs. control (Ctrl) viability of each clone before TNF treatment; *P < 0.05 vs. corresponding control at each time point.

 


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  		Fig. 3. Concentration effects of TNF on viability in the two clones of VSMCs. A: concentration-dependent effects of TNF in increasing proliferation in Sp-SMCs. B: in Ep-SMCs, TNF was cytotoxic and reduced viability in a concentration-dependent manner. Sp- and Ep-SMCs at a density of 3 x 103 cells/well in 96-well plates were growth arrested in 0.2% calf serum-DMEM for 48 h before incubation in the absence or presence of TNF (1, 10, and 100 ng/ml). Cell viability was measured 48 h later using MTT assay. Cells in culture medium without TNF treatment served as controls. Values are means ± SE for three experiments; *P < 0.05 vs. control.

 
Because 10 ng/ml TNF significantly induced either proliferation or cytotoxicity depending on phenotype, this concentration of the cytokine was used in the subsequent experiments unless otherwise stated.

TNF induced phenotype-dependent apoptosis. Because previous studies have shown that TNF could be proapoptotic in VSMCs (11, 15, 32), we determined the effects of the cytokine on apoptosis in both subsets of VSMCs. We first analyzed genomic DNA integrity by agarose electrophoresis. Both Sp- and Ep-SMCs were stimulated with 10 ng/ml TNF, and genomic DNA was extracted 48 h later. As seen in Fig. 4A, DNA isolated from both untreated and TNF-treated Sp-SMCs was intact and migrated as high-molecular-weight DNA. In parallel experiments, DNA extracted from untreated Ep-SMCs was also intact. In contrast, the DNA isolated from TNF-treated Ep-SMCs was fragmented, which is a marker for cell apoptosis. To quantify the apoptotic incidence, we used a fluorescence microscopic assay of Hoescht 33258-stained cells. As shown in Fig. 4B, treatment with TNF in the Ep-SMC clones produced an apoptotic incidence of ~36%, which is significantly higher than the spontaneous incidence of ~3.5% in these cells. There was not an increase of apoptotic incidence in TNF-treated Sp-SMCs over control Sp-SMCs. These results indicated that TNF induced apoptosis in VSMCs depending on phenotype, and the reduced viability of the Ep-VSMCs was largely through induction of apoptosis.



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  		Fig. 4. Phenotype-dependent apoptosis induced by TNF in VSMCs. A: TNF (10 ng/ml) did not markedly influence the integrity of cellular DNA isolated from Sp-SMCs. B: in contrast, treatment with TNF caused fragmentation of cellular DNA in Ep-SMCs, a pattern consistent with apoptotic cell death. C: treatment with TNF did not increase the apoptotic incidence in Sp-SMCs. D: TNF treatment induced apoptosis in Ep-SMCs. Growth-arrested cells (Sp- and Ep-SMCs) were incubated in the absence (Ctrl) or presence of 10 ng/ml TNF. Cells were harvested 48 h later, and cellular DNA was extracted or cells were stained with Hoechst 33258. Cellular DNA was analyzed by 1% agarose gel electrophoresis for assessment of DNA integrity. Hoechst 33258-stained cells were examined for apoptotic characteristics (nuclear margination and chromatin condensation) using a fluorescence microscope. Apoptotic incidence was calculated. Results in A and B are representative of three individual experiments. Values in C and D are expressed as means ± SE for three experiments; *P < 0.05 vs. control. M.W., molecular weight.

 
Phenotype-dependent TNF-R1 expression and activation of caspase-3. Because signaling through TNF-R1 activates caspase-3 and this caspase plays a role in the execution phase of apoptosis (11, 32, 45), we determined the level of this receptor and caspase-3 activity in the two phenotypes. As seen in Fig. 5A, Western blot analysis shows a higher level of TNF-R1 expressed in membranes of Ep-SMCs than Sp-SMCs. We also measured activity of this caspase in untreated and TNF-treated clones. As shown in Fig. 5B, TNF (10 ng/ml for 24 h) did not increase activity of caspase-3 in Sp-SMCs. In contrast, treatment with TNF significantly increased the activity of caspase-3 in Ep-SMC clones. These results are consistent with TNF-selective induction of apoptosis in one subset of VSMCs.



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  		Fig. 5. Expression of receptor subtype TNF-R1 and activation of caspase-3 in VSMC subtypes. A: a higher constitutive expression of TNF-R1 was observed in cell membranes isolated from Ep-SMCs than from Sp-SMCs. This result is typical of three experiments. B: TNF (10 ng/ml) had no effect on caspase-3 activity in Sp-SMCs. C: in contrast, TNF induced increased caspase-3 activity in Ep-SMCs. Sp- and Ep-SMCs were plated and growth arrested as described for Fig. 4. Cells were either untreated (Ctrl) or treated with 10 ng/ml TNF for analysis of TNF-R1 expression and caspase-3 activity. Cell membrane proteins were prepared from untreated Sp- and Ep-SMCs. Equal amounts of membrane proteins were resolved by SDS-PAGE. Separated proteins were transferred onto nitrocellulose membranes and probed with anti-TNF-R1 antibodies. Antigen-antibody bindings were visualized using enhanced chemiluminescence assay. For determination of caspase-3 activity, untreated cells and cells treated for 24 h with TNF were lysed. Equivalent amounts of the lysates were used to measure caspase-3 activity (see MATERIALS AND METHODS). Values are means ± SE for three experiments; *P < 0.05 vs. control.

 
Augmentation of TNF effect by CHX was phenotype dependent. It is known that de novo protein synthesis protects some types of cells from the toxic effects of TNF (33, 46). We therefore examined the effects of TNF on viability of the two subpopulations of VSMCs in the presence of CHX, which is an inhibitor of protein synthesis. As seen in Fig. 6A, stimulation with 10 ng/ml TNF for 24 h did not significantly influence viability of Sp-SMCs. Incubation with 1 µg/ml CHX alone reduced viability of untreated cells. The combination of CHX and TNF did not further reduce viability of Sp-SMCs. Figure 6B shows that 24-h exposure of Ep-SMCs to 10 ng/ml TNF alone significantly reduced the cell viability by ~23%. Incubation with 1 µg/ml CHX for 24 h also reduced viability by ~12%. In the presence of CHX, TNF reduced viability of Ep-SMCs by ~83%, a value significantly greater (P < 0.05) than for TNF or CHX alone. These results suggest that de novo protein synthesis played a preventive role in antagonizing the toxic signal induced by TNF in the Ep-SMCs.



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  		Fig. 6. Effects of TNF on viability of smooth muscle cell clones in presence of the protein synthesis inhibitor cycloheximide (CHX). A: CHX (1 µg/ml) reduced viability in Sp-SMCs. This reduced viability was not potentiated by the addition of 10 ng/ml TNF. B: effects of CHX on viability were significantly potentiated by the addition of TNF in Ep-SMCs. Sp- and Ep-SMCs at a density of 3 x 103 cells/well in 96-well plates were growth arrested in 0.2% calf serum-DMEM for 48 h. Cells were incubated in the absence (Ctrl) or presence of 1 µg/ml CHX for 2 h before addition of 10 ng/ml TNF. Cell viability was measured 24 h later by MTT assay (see MATERIALS AND METHODS). Cells in culture medium without TNF and/or CHX treatment (Ctrl) reflect 100% cell viability. Other results are expressed as viability relative to control viability. Values are means ± SE for three experiments. NS; P > 0.05; *P < 0.05.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
VSMCs in intact arterial tissue and in cultured cells derived from arteries are heterogeneous in terms of morphology and biological function (2, 18, 25, 40, 53). Several lines of evidence have shown that VSMCs in arterial media and primary cultures derived from arteries contain at least two subsets of VSMCs including the highly differentiated Sp-SMCs and the poorly differentiated Ep-SMCs (2, 18, 25, 40, 53). Sp-SMCs appear to predominate in the arterial media and primary culture, whereas Ep-SMCs are the most common in intimal thickening after vascular injury (2, 18, 25). In the present study, we first isolated VSMCs from human saphenous veins via a commonly used medial explant technique. From these primary cultures of VSMCs, two distinctly different subsets of VSMCs were generated by clonal analysis: Sp- and Ep-SMCs. To further confirm the smooth muscle origin of these clones and compare their differentiation states, we used Western blot analysis to examine smooth muscle differentiation proteins expressed in various developmental stages including {alpha}-SM actin, calponin, SM-MHC, and CK-8. Both Sp- and Ep-SMCs had similar levels of {alpha}-SM actin, which is a smooth muscle differentiation protein expressed in all stages of smooth muscle development (16, 17, 24). Unlike {alpha}-SM actin, calponin and SM-MHC are exclusively expressed in late stages of smooth muscle differentiation (16, 24). Our Western blot analysis showed that calponin and SM-MHC were more pronouncedly expressed in Sp-SMCs. CK-8 is a marker of SMC phenotype at early stages of differentiation (1, 17). Expression of CK-8 was detected in Ep-SMCs but not in Sp-SMCs. Therefore, these results indicate that Sp-SMCs are a more mature phenotype than Ep-SMCs. In terms of morphology, growth pattern, and expression of the smooth muscle-specific differentiation proteins, these human vein-derived VSMC subtypes are similar to different subpopulations of VSMCs cultured from animal peripheral arteries and one segment of human internal thoracic artery (2, 18, 25, 40). The two distinct clones that we isolated from saphenous vein maintained their morphology through many passages and freeze-thaw cycles, thereby indicating that the clones are stable.

The presence of these two subtypes of VSMCs in primary cell cultures could contribute to the conflicting results reported in the literature. For example, it is well known that ANG II elicits both antiapoptotic and proapoptotic signals in VSMCs (52). ANG II has a potent antiapoptotic effect and weak proliferative activity in primary VSMC cultures isolated from rat aorta (4, 34). Conversely, ANG II can induce apoptosis in vivo (10, 30). In a recent study, Bascands et al. (2) used Sp- and Ep-SMCs cloned from rat aorta VSMCs and found that ANG II elicited apoptosis in Ep-SMCs but not Sp-SMCs. Therefore, it seems possible that the effect of ANG II would depend on the predominance of the phenotype in the experimental situation. Furthermore, as described in the opening of this report, previous investigations using primary VSMC cultures have shown conflicting effects of TNF (11, 12, 15, 19, 22, 28, 32, 38, 39, 53). Here we addressed the question of whether the diversity in the biological effects of TNF could originate from variations in the different VSMC subtypes. Our data show that TNF induces human vein-derived SMC proliferation or apoptosis depending on the phenotype. We found that TNF stimulated proliferation of Sp-SMCs and induced cytotoxic effects on Ep-SMCs. These results were confirmed by fluorescence microscopy and genomic DNA gel electrophoresis, which showed that in the Ep-SMCs, TNF induced apoptosis as characterized by DNA fragmentation and nuclear condensation. It is well known that TNF elicits diverse signals including proliferative, antiapoptotic, and proapoptotic signals (3, 33, 46, 47, 51). The final consequence of the cytokine interacting with the cell is determined by cell type (3, 33, 46, 47, 51). For instance, TNF is proapoptotic in some types of tumor cells and endothelial cells, whereas it is mitogenic in others such as fibroblasts (3, 33, 46, 47, 51). The present work shows for the first time that TNF induces diverse effects on VSMCs depending on subtype.

The explanation for this phenotype-dependent effect of TNF on cell survival is not totally defined in the present study. One speculation is that mitogenic signals override death signals in Sp-SMCs, whereas TNF-induced proapoptotic signals prevail over survival and growth signals in Ep-SMCs. Our findings that TNF activated caspase-3 [which is an effector caspase that plays a central role in apoptosis in many cell types including VSMCs (11, 32, 47)] only in Ep-SMCs and that Ep-SMCs expressed a higher level of TNF-R1 (the receptor linked to caspase activation) favors this assumption. Intriguingly, the death receptor TNF-R1 transduces not only death signals associated with caspase activation but also signals for survival and proliferation (5, 6, 41, 44, 45). It is the balance among these signals that determines whether a cell will finally die or proliferate in response to TNF (5). Although the molecular mechanisms by which TNF-R1 mediates these signals remain poorly understood, it is known that transcription and translation mechanisms mediated by factors such as NF-{kappa}B are important for production of the survival and proliferation signals (3, 5, 6, 11, 33, 38, 39, 41, 46, 47, 51). In TNF-sensitive tumor cells and endothelial cells, inhibition of de novo transcription and translation by actinomycin D and CHX potentiates the proapoptotic activity of TNF (33, 46). In human aorta-derived primary VSMCs, Selzman et al. (38, 39) reported that TNF-induced cell proliferation was reduced by 60% when NF-{kappa}B activation was inhibited by liposomal delivery of recombinant inhibitory protein (I-{kappa}B)-{alpha} protein. In the same type of cells, a study by Obara et al. (32) showed that the cytokine became proapoptotic in the presence of NF-{kappa}B inhibition caused by expression of truncated I-{kappa}B. These studies indicate that NF-{kappa}B-mediated transcription and translation transduce either mitogenic or antiapoptotic signals or both. In the present study, we observed the effects of TNF on VSMC clones in the presence of CHX. We found that incubation with the protein synthesis inhibitor alone for 24 h reduced cell viability in both clones. Additionally, we found that in the presence of CHX, TNF cytotoxic activity was significantly augmented in Ep-SMCs. This result suggests that in Ep-SMCs, TNF induces both proapoptotic and antiapoptotic signals and that antiapoptotic signals required de novo protein synthesis. In other cell types, it has been documented that transcription- and translation-mediated proliferative and antiapoptotic signals override proapoptotic signals and prevent TNF-induced cell death (33, 46, 47, 51). In the Sp-SMC clones, CHX reduced viability in untreated cells and TNF-stimulated cells equally, i.e., TNF in combination with CHX did not further reduce Sp-SMC viability. These results imply that in this clone, de novo protein synthesis was not required to antagonize the toxic signal generated by TNF, and instead, constitutive proteins antagonize the death signal elicited by TNF binding. Thus the findings in the present study may at least partly explain conflicting observations of the biological activities of TNF in primary cultures of VSMCs.

In a recent review, Geng and Libby (14) discussed in detail the evidence that supports an important role for apoptosis in stability of an atherosclerotic plaque. Apoptosis occurs in mature plaques leading to the death of VSMCs and loss of cellular matrix proteins thus weakening the plaque by reducing the strength of the fibrous cap. Loss of integrity of the cap promotes thrombogenesis in the affected vessel and potentially leads to an acute cardiac event. The authors point out that a complex interplay of factors such as cytokines like TNF found in the lesions, growth factors, modified lipids, and many others determines the balance between proliferation and apoptotic death of VSMCs in the lesion. Our data suggest that the phenotype of the VSMCs that predominates in the lesion could also critically contribute to this balance. If the Ep-SMC subtype, which responds to TNF with apoptosis, is the principal VSMC in the atheroma, this certainly would be another factor to consider in the pathology of atherosclerosis.

This study demonstrates that TNF can induce either proliferation or apoptosis of cultured human saphenous vein-derived VSMCs depending on phenotype. There are several limitations of this study that merit consideration particularly when the information is applied to the role of TNF in atherosclerosis. First, this study was conducted on venous VSMCs. The biology of venous VSMCs may extrapolate to vein-graft atherosclerosis and other venous occlusive disorders but may not directly relate to arterial atherosclerosis and restenosis. Studies of phenotypes of VSMCs derived from human artery are needed. Second, this study was performed on cloned VSMCs at relatively high passages (passages 10–12), which was necessary to grow enough cells from a single cell for study. VSMCs at relatively high passages might lose some of their original properties. Third, the concentrations of TNF in vascular diseases are to the best of our knowledge totally unknown. In this study, we only used TNF at concentrations commonly used in in vitro studies.

In conclusion, we believe the information presented here helps define the role of TNF in atherosclerosis and particularly in atherosclerosis developed within saphenous vein grafts for several reasons. TNF is expressed in diseased vein grafts at a level markedly higher than in atherosclerotic coronary arteries (7). Apoptosis of VSMCs is significant in vein-graft atherosclerosis and is found in both initial and advanced stages (23, 27). If TNF plays a significant role in the development of atherosclerotic or restenotic lesions, the results presented here could have important implications for understanding the complex control of cellularity in vascular proliferative processes.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported in part by a grant from the MedCen Foundation and the Clinical Research Center of the Medical Center of Central Georgia.


   ACKNOWLEDGMENTS
 
The authors thank Benjamin P. Mitchell and Jing Kang for assistance in this study.

Present address for M. R. Castresana: Dept. of Anesthesiology and Perioperative Medicine, Medical College of Georgia, Augusta, GA 30912.


   FOOTNOTES
 

Address for reprint requests and other correspondence: Z. Wang, Division of Basic Medical Science, Mercer Univ. School of Medicine, 1550 College St., Macon, GA 31207 (E-mail: wang_z{at}mercer.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
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

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