Vascular remodeling is a key feature of many pathologic states, including atherosclerosis, restenosis, or hypertension (1). Vascular smooth muscle cells (VSMCs) strongly participate in determining the vessel structure as they may undergo different mechanisms like cell migration, cell growth (cell hyperplasia or cell hypertrophy), or cell death. Initially, cell death in the vasculature was thought to be the consequence of a toxic injury, due to factors like free radicals, resulting in VSMC necrosis (2). However, in the last few years, programmed cell death, or apoptosis, has been shown to play a predominant role in determining the vessel architecture (3). Indeed, there is increasing evidence showing that VSMC apoptosis is involved in the pathogenesis of vascular diseases, like hypertension (4), atherosclerosis, restenosis (5), or diabetic vasculopathy (6), which in turn raises the necessity of understanding the mechanisms underlying VSMC apoptosis.
A wide variety of drugs have been used to study apoptosis using VSMC cell cultures. Thapsigargin is an inhibitor of an adenosine triphosphate (ATP)-gated calcium pump controlling the influx of calcium from the cytosol to the endoplasmic reticulum (7). Thapsigargin has been widely used to induce apoptosis in different cell types (8-12). However, to our knowledge, there are no reports concerning the ability of thapsigargin to promote apoptosis in human VSMCs. The aim of the present work was to determine whether thapsigargin is able to induce apoptosis in human VSMCs and therefore if this drug can be used as a model of apoptosis in this cell type.
MATERIALS AND METHODS
VSMC cultures were obtained from the aortas of three organ donors (age, 37 ± 3 years). In brief, the aortic fragments were cleaned free of the adventitia layer, and the endothelium was removed by gently rubbing the lumen of the vessel. Afterward, the fragments were cut into small pieces and incubated for 90 min in Dulbecco's Modified Eagle Medium (DMEM) containing 0.1% bovine serum albumin and 4 mg/ml collagenase (Type II; Sigma, St. Louis, MO, U.S.A.). At the end of the incubation, the resulting cell suspension was washed twice with fresh DMEM, resuspended in DMEM containing 10% fetal calf serum (FCS), 100 μg/ml streptomycin, 100 U/ml penicillin, and 2.5 μg/ml amphotericin B (Sigma), and plated onto 25-cm2 culture flasks. Cells were characterized as smooth muscle based on (a), morphologic criteria: cells were spindle-shaped and showed a typical "hill and valley" pattern when postconfluent, and (b), indirect immunofluorescence staining for α-smooth muscle actin, as previously described by others (13). At confluence, cells were passaged using a solution containing 0.02% EDTA: 0.05% trypsin and split in a 1:2 ratio. In the present study, cells were used between passages two and six.
Chromatin binding dye staining
For experiments, HVSMCs were plated onto 24-well culture plates in DMEM containing 10% FCS. After cell attachment, medium was switched to DMEM containing 0.5% FCS. When confluent, HVSMCs were treated with different concentrations of thapsigargin for 60 min. Afterward, the medium was removed and replaced by fresh DMEM containing 0.5% FCS. At different time points, differential chromatin staining was performed, as previously described by others (14). In brief, chromatin binding dyes Hoescht 33342 (5 μM) and propidium iodide (1 μM; Molecular Probes, Leiden, The Netherlands) were added to cell cultures for 30 min. Hoescht 33342 is membrane permeant, whereas propidium iodide penetrates only into cells with disrupted plasma membrane. Afterward, HVSMCs were collected, centrifuged, and the cell pellet was resuspended in FCS-containing medium. Cell nuclei were observed using an epifluorescence microscope (Eclipse TE300; Nikon, Tokyo, Japan) at ×40 magnification. For every treatment, 1,000 nuclei from randomly selected microscopic fields were counted by a blinded observer.
Cell death detection ELISA
Cell death detection enzyme-linked immunosorbent assay (ELISA) was performed using a commercial kit that detects apoptosis by quantifying cytoplasmic histone-associated DNA fragments (Boehringer Mannheim, Mannheim, Germany). In brief, after submitting HVSMCs to the different treatments described earlier, cells were lysed, and the cytoplasmic fraction was transferred onto 96-well plates precoated with streptavidin. The cytoplasmic fractions were incubated with an immunoreagent mix containing anti-histone-biotin antibody and anti-DNA-peroxidase for 2 h at room temperature. Plates were then washed, the substrate for peroxidase was added, and absorbance was measured at 405 nm.
In situ nick-end labeling
The terminal deoxyribonucleotidyl transferase-mediated dUTP-digoxigenin nick-end labeling (TUNEL) assay was performed using a commercial kit (ApopTag; Oncor, Gaithersburg, MD, U.S.A.). In brief, samples were fixed with 4% paraformaldehyde, permeabilized with 2:1 ethanol/acetic acid, and incubated with equilibration buffer for 5 min at room temperature. Samples were then incubated with terminal deoxyribonucleotidyl transferase for 1 h at 37°C, rinsed with PBS, and incubated for further 10 min in stopping buffer. After rinsing with PBS, the fluorescein-labeled anti-digoxigenin antibody was applied for 30 min at room temperature and rinsed again with PBS. The samples were then mounted with mounting medium containing 1 μM 4′,6-diamidino-2-phenylindole (DAPI) and viewed under fluorescence microscopy. For quantifying the amount of TUNEL-positive cells, five independent fields per treatment were analyzed, and the percentage of labeled nuclei related to total nuclei number was calculated.
Culture plastic ware was obtained from Costar (Corning, NY, U.S.A.). DMEM, fetal calf serum, and trypsin-EDTA solutions were from Biological Industries (Beit Hamek, Israel). Thapsigargin was purchased from RBI (Natick, MA, U.S.A.). Thapsigargin was dissolved in dimethyl sulfoxide (DMSO) at a concentration of 1 mM, aliquoted, and stored at −70°C until used. Further dilutions were done in deionized water.
Data are expressed as mean ± SEM. The statistical analysis was evaluated by Student's t test or analysis of variance (ANOVA) for data points or curves, respectively. A value of p < 0.05 was taken as the statistical significance.
Treatment of HVSMCs with thapsigargin resulted in morphologic changes associated with apoptotic cell death. When observed under phase contrast microscopy, a large number of HVSMCs exhibited morphologic features characteristic of apoptosis, such as cell shrinkage and rounding (Fig. 1A). To observe nuclear chromatin morphology, HVSMCs were differentially stained with DNA-binding dyes and observed under epifluorescence microscopy. As shown in Fig. 1B, cells exhibiting apoptosis-related morphologic changes also exhibited nuclear chromatin condensation and fragmentation after staining with Hoescht 33342 when viewed under epifluorescence microscopy.
As a quantitative index of apoptosis, the number of cells showing nuclei with condensed or fragmented chromatin was determined. Propidium iodide-stained cells (∼3% of total cell number) were discarded, as they were considered toxic or necrotic cells. The basal percentage of apoptotic nuclei in HVSMCs cultures maintained in DMEM with 0.5% FCS without thapsigargin treatment was 0.19 ± 0.04% (results from eight independent experiments). When HVSMCs were treated with increasing concentrations of thapsigargin (100 nM-10 μM), the number of apoptotic nuclei increased in a concentration-dependent manner (Fig. 2A). The percentage of apoptotic nuclei was around sixfold, 15-, and 57-fold increase over basal for 100 nM, 1 μM, and 10 μM thapsigargin, respectively. DMSO by itself did not affect the number of apoptotic nuclei (data not shown).
When a time-course experiment was performed using 10 μM thapsigargin, a slight increase in the number of apoptotic nuclei was observed 3 h after the beginning of the thapsigargin pulse (Fig. 2B). However, it was between 3 and 6 h after thapsigargin treatment when a dramatic increase in the percentage of apoptotic nuclei was observed. Between 6 and 24 h, the number of apoptotic nuclei was moderately increased (Fig. 2B). The basal apoptotic index remained unmodified over the 24-h period (Fig. 2B).
Using another experimental approach to quantify thapsigargin-induced apoptosis, histone-associated DNA fragmentation was measured by ELISA. As shown in Fig. 3A, DNA fragmentation was also increased in a concentration-dependent manner when HVSMCs were treated with growing concentrations of thapsigargin (100 nM-10 μM). Regarding the time course of DNA fragmentation (Fig. 3B), again a clear increase in DNA fragmentation was observed around 6 h after the beginning of the exposure to thapsigargin, whereas in untreated cultures, the basal levels of DNA fragmentation remained unchanged along the 24-h period. DMSO by itself did not affect basal ELISA values (data not shown).
Finally, in situ nick-end labeling was also performed as a method that enables visualization of early apoptosis-associated DNA fragmentation. As shown in Fig. 4, cultures incubated with 10 μM thapsigargin exhibited a significantly higher number of TUNEL-positive nuclei 3 h after thapsigargin treatment compared with untreated cultures (23 ± 6% and 2.3 ± 1.3%, respectively; p < 0.05, results from three independent experiments).
VSMCs are determinant in designing the vessel structure through different phenomena, such as cell migration, cell growth, and cell death (necrosis or apoptosis) (3). Emerging evidence shows that apoptosis may play a pivotal role in determining the vessel architecture in many pathologic states, like hypertension (4), atherosclerosis and restenosis (5), or diabetic vasculopathy (6). Indeed, there is growing interest in studying the mechanisms underlying VSMC apoptosis.
Different agents have been used to stimulate apoptosis in VSMC cultures. The nature of these agents is highly heterogenous, including free radical generating systems (15), inhibitors of postranslational lipidic protein modification, like perillyl alcohol (16); pentoxifylline, a tumor necrosis factor (TNF)-α synthesis inhibitor (17); camptothecin, a chemotherapeutic agent (18); or several nitric oxide donors (14).
In the last few years, thapsigargin has been reported to promote apoptosis in many different cell types, such are rat mesangial cells (8), mouse osteoblasts (9), and hypothalamic neurons (10), or different human and mouse tumoral cell lines (11,12).
Concerning VSMCs, previous reports have shown that thapsigargin inhibits proliferation of rat aortic VSMCs in culture (19,20). Furthermore, when added for a short time to human saphenous vein segments, thapsigargin inhibits neointima formation by reducing both VSMC migration and proliferation, without affecting the rate of cell death (21). In the present work, however, we observed that a short treatment with thapsigargin promoted HVSMC death, which was not due to a necrotic response, as shown by the negative propidium iodide staining. On the contrary, HVSMCs died in an apoptotic pattern, which was particularly evident between 3 and 6 h after exposure to thapsigargin, as assessed by chromatin morphology, ELISA quantification of DNA fragmentation, and TUNEL. In fact, these different techniques are able to detect several stages of the apoptotic process, all of them being consistent with the proapoptotic capability of thapsigargin. Conversely, the lack of cell death-promoting effects of thapsigargin described in human saphenous vein segments is likely explained by the fact that those segments were exposed to a thapsigargin concentration of 10 nM(21), which is 10-fold lower than the minimal concentration we have shown to induce HVSMC apoptosis in the present work. It may therefore be concluded that, depending on the concentration used, thapsigargin may act either as a growth inhibitor or a as proapoptotic agent for cultured VSMCs, as has been observed with other drugs, like some nitric oxide donors (14).
At present the cellular pathways mediating thapsigargin-induced apoptosis are not well understood. However, in addition to calcium homeostasis disruption, different mechanisms have been proposed to be involved, like nitric oxide production (8,22), increased c-Jun NH(2)-terminal kinase activity (9,22), enhanced transcriptional activity of nuclear factor-κB (NF-κB) (9), or a decrease in the Bcl-2/Bax ratio (8). Whether these mechanisms are shared by the different cell types still remains to be determined.
In conclusion, here we report that thapsigargin acts as a proapoptotic agent in cultured VSMCs, providing a useful tool to study the mechanisms leading to apoptosis in this cell type.
Acknowledgment: The present work was supported by grants from CICYT (SAF 98-0010), CAM (04.8/0027/1998), FEDER (2FD97-0445-CO2), FISS 99/0246, and Bayer España. C. Peiró is the recipient of a postdoctoral fellowship from Comunidad Autónoma de Madrid.
1. Gibbons GH, Dzau VJ. The emerging concept of vascular remodeling. N Engl J Med
2. Guyton JR, Black BL, Seidel CL. Focal toxicity of oxysterols in vascular smooth muscle
cell culture: a model of the atherosclerotic core region. Am J Pathol
3. Hamet P, Moreau P, Dam TV, et al. The time window of apoptosis
: a new component in the therapeutic strategy for cardiovascular remodeling. J Hypertens Suppl
4. Díez J, Panizo A, Hernández M, et al. Is the regulation of apoptosis
altered in smooth muscle cells of adult spontaneously hypertensive rats? Hypertension
5. Isner JM, Kearney M, Bortman S, et al. Apoptosis
in human atherosclerosis and restenosis. Circulation
6. Fukumoto H, Naito Z, Asano G, et al. Immunohistochemical and morphometric evaluations of coronary atherosclerotic plaques associated with myocardial infarction and diabetes mellitus. J Atheroscl Thromb
7. Thastrup O, Cullen PJ, Drobak BK, et al. Thapsigargin
, a tumor promoter, discharges intracellular Ca2+
stores by specific inhibition of the endoplasmic reticulum Ca2(+)-ATPase. Proc Natl Acad Sci U S A
8. Rodríguez-López AM, Flores O, Martínez-Salgado C, et al. Increased apoptosis
susceptibility in mesangial cells from spontaneously hypertensive rats. Microvasc Res
9. Chae HJ, Chae SW, Weon KH, et al. Signal transduction of thapsigargin
in osteoblast. Bone
10. Wei H, Wei W, Bredesen DE, et al. Bcl-2 protects against apoptosis
in neuronal cell line caused by thapsigargin
-induced depletion of intracellular calcium stores. J Neurochem
11. Xue L-Y, Qiu Y, He J, et al. Etk/Bmx, a PH-domain containing tyrosine kinase, protects prostate cancer cells from apoptosis
induced by photodynamic therapy or thapsigargin
12. McCormick TS, McColl KS, Distelhorst CW. Mouse lymphoma cells destined to undergo apoptosis
in response to thapsigargin
treatment fail to generate a calcium-mediated grp78/grp94 stress response. J Biol Chem
13. Dubey RK, Roy A, Overbeck HW. Culture of renal arteriolar smooth muscle cells: mitogenic responses to angiotensin II. Circ Res
14. Pollman MJ, Yamada T, Horiuchi M, et al. Vasoactive substances regulate vascular smooth muscle
: countervailing influences of nitric oxide and angiotensin II. Circ Res
15. Li P-F, Dietz R, von Harsdorf R. Reactive oxygen species induce apoptosis
of vascular smooth muscle
cell. FEBS Lett
16. Unlu S, Mason CD, Schachter M, et al. Perillyl alcohol, an inhibitor of geranylgeranyl transferase, induces apoptosis
of immortalized human vascular smooth muscle
cells in vitro. J Cardiovasc Pharmacol
17. Hamet P, Richard L, Dam T-V, et al. Apoptosis
in target organs of hypertension. Hypertension
18. Min I, Stubbs MC, Strachan GD, et al. Cyclin A levels, the duration of S phase and sensitivity to a chemotherapeutic agent are altered in fibroblasts cultured on a fibronectin matrix. Int J Oncol
19. Kwan CY, Chaudhary CY, Zheng XF, et al. Effects of sarcoplasmic reticulum calcium pump inhibitors on vascular smooth muscle
20. Shukla N, Jeremy JY, Nicholl P, et al. Short-term exposure to low concentrations of thapsigargin
inhibits replication of cultured human vascular smooth muscle
cells. Br J Surg
21. George SJ, Johnson JL, Angelini GD, et al. Short-term exposure to thapsigargin
inhibits neointima formation in human saphenous vein. Arterioscler Thromb Vasc Biol
22. Srivastava RK, Sollott SJ, Khan L, et al. Bcl-2 and Bcl-X(L) block thapsigargin
-induced nitric oxide generation, c-Jun NH(2)-terminal kinase activity, and apoptosis
. Mol Cell Biol