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Cardiovascular Research 2001 50(1):97-107; doi:10.1016/S0008-6363(01)00196-1
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Copyright © 2001, European Society of Cardiology

Loss of cyclin A and G1-cell cycle arrest are a prerequisite of ceramide-induced toxicity in human arterial endothelial cells

Ioakim Spyridopoulosa,*, Petra Mayera, Kerida S. Shooka, Dorothea I. Axela, Richard Viebahnb and Karl R. Karscha

aDepartment of Cardiology and Cardiovascular Research, Medizinische Klinik III, Otfried-Mueller-Str. 10, 72076 Tübingen, Germany
bDepartment of Surgery, University of Tübingen, Tübingen, Germany

* Corresponding author. Tel.: +49-7071-298-2711; fax: +49-7071-293-169 ioakim_s{at}hotmail.com

Received 31 August 2000; accepted 27 December 2000


   Abstract
Top
 Abstract
1 Introduction
 2 Methods
 3 Results
 4 Discussion
 References
 
Background: Ceramide is an important messenger of TNF- and lipid-induced apoptosis. We previously demonstrated the adverse effect of TNF in the process of reendothelialization as well as the dependence of its effect on cell-cycle regulation. The current study was designed to investigate the linkage between ceramide induced toxicity and growth arrest in human endothelial cells. Methods and results: Cultured human arterial endothelial cells (HAEC) served as an in-vitro model to test the cellular effects of C2-ceramide (C2). C2-induced cell death in HAECs occurred time- and dose-dependently. The LD50 in subconfluent cells was three times lower than in confluent cell layers (25 vs. 75 µM). C2 caused up to 70% inhibition of BrdU and [3H]thymidine incorporation at non-toxic concentrations as a result of G1 cell-cycle arrest. Downregulation of cyclin A and p21Cip1/Waf1 protein expression was observed independently of C2-toxicity, while expression of other cell-cycle regulatory genes was not affected. Inhibition of cyclin A protein expression by sequence-specific antisense-oligonucleotides was paralleled by significant growth-inhibition. The protein phosphatase inhibitor okadaic acid induced endothelial cell proliferation, which was completely abrogated by C2. In contrast, aphidicolin-synchronized endothelial cells demonstrated elevated cyclin A levels along with 30% higher BrdU-incorporation and 70% less C2-toxicity. G1-arrested cells, however, showed significantly enhanced C2-toxicity, lack of cyclin A expression and induction of uncleaved caspase-3 (CPP32). Conclusions: Ceramide abrogates endothelial cell proliferation independently of apoptosis or necrosis at low concentrations (≤10 µM) through loss of cyclin A expression with subsequent G1 cell-cycle arrest. Synchronization of HAECs in S-phase with aphidicolin overcomes C2-induced G1-arrest and partially blocks ceramide toxicity. These findings demonstrate the dependence of ceramide toxicity on cell cycle regulation, suggesting a strong bidirectional relationship between cell-cycle control and cell death in vessel biology.

KEYWORDS C2, C2-ceramide (D-erythro-N-acetylsphingosine); C6, C6-ceramide (D-erythro-N-hexanoylsphingosine); HAEC, human arterial endothelial cells; MTS, 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)2H-tetrazolium; OA, okadaic acid; PI, propidium iodide; PMA, phorbol 12-myristate 13-acetate; PP2A, protein phosphatase 2A; SMase, sphingomyelinase; TNF, tumor-necrosis-factor {alpha}


   1 Introduction
Top
 Abstract
 1 Introduction
2 Methods
 3 Results
 4 Discussion
 References
 
Apoptosis and cell growth are tightly connected in most cellular systems [1]. Induction of the tumor suppressor gene p53 on the one hand serves as a G1 phase checkpoint in the cell-cycle, prohibiting the cell from entering S-phase for replication of its DNA prior to mitosis [2,3]. On the other hand, overexpression of p53 induced by hypoxia, oxygen radicals or other noxious stimuli can simultaneously lead to initiation of programmed cell death [4]. Proliferation and suppression of endothelial cell death play an important role in vascular biology during angiogenesis [5–7] as well as in reendothelialization after barotraumatic vessel injury [8].

The enzyme sphingomyelinase (SMase) catalyzes the hydrolysis of sphingomyelin to ceramide and choline phosphate [9]. SMase reactions have been implicated in specific atherogenic processes [10,11], such as hydrolyzing subendothelial LDL. Cultured endothelial cells are an abundant source of SMase [12].

We have previously shown that neutralizing the physiologically synthesized cytokine tumor necrosis factor alpha (TNF) after experimental balloon angioplasty in a rat carotid artery model significantly accelerates reendothelialization of the traumatized vessel [13]. Using an in-vitro model of cultured human endothelial cells we also found that restoring cyclin A through overexpression of the transcription factor E2F in TNF-treated cells protected them from apoptosis. Since ceramide is an important messenger of cell death induced by cytokines [14], oxysterols [11] and oxygen radicals [15], this study aimed to further investigate the relationship between growth control and cell death in human endothelial cells.


   2 Methods
Top
 Abstract
 1 Introduction
 2 Methods
3 Results
 4 Discussion
 References
 
2.1 Endothelial cell isolation and culture
Human iliac and renal arteries were obtained from organ donors (approved by local ethics committee). Human arterial endothelial cells (HAECs) were isolated and passaged according to techniques described previously [16]. HAECs were subcultured onto collagen-coated plastic culture dishes (Iwaki Glass) and grown in an endothelial medium kit (EBM2, Clonetics) containing VEGF, hEGF, hFGF, R3-IGF-1, hydrocortisone, 2% FCS and 1% penicillin/streptomycin. Subcultured HAECs were characterized by immunocytochemical staining with polyclonal antibodies against von Willebrand factor (Boehringer Mannheim), and contamination with vascular smooth muscle cells was excluded by additional double staining using antibodies against smooth muscle {alpha}-actin. Monolayers showed characteristic cobblestone morphology. Cells were fed every 3rd day and used for experiments in low passages (2 through 4). DAPI staining (Boehringer Mannheim) was performed in order to exclude contamination with mycoplasma. Unless stated otherwise, all experiments were performed with subconfluent layers of endothelial cells.

2.2 Antibodies and chemicals
The following antibodies were used for Western blotting: anti-cyclin A pAb rabbit, Santa Cruz #sc-4072, anti-cyclin E (M-20) pAb rabbit, Santa Cruz #sc-481, anti-cdk2 (H-298) pAb rabbit, Santa Cruz #sc-748, anti-CPP32 pAb goat, R&D #AF-605-NA, anti-p21 (anti-Cip1) mAb mouse, Transduction #C24420, anti-p53 (DO-1) pAb rabbit, Santa Cruz #sc-126, anti β-actin mAb mouse, Sigma chemicals #A5441, anti-rabbit IgG-HRP, Santa Cruz #sc-2004, anti-mouse IgG-HRP, Santa Cruz #sc-2005, and anti-goat IgG-HRP, Santa Cruz #sc-2020. C2-Ceramide (D-erythro-N-acetylsphingosine, BioMol #SL-100) was dissolved in DMSO (10 mg per 580 µl DMSO resulting in a 12.5-mM stock solution) and stored in aliquots at –20°C. Dihydro-C2 (D-erythro-N-acetylsphinganine, BioMol #SL-101), C6-ceramide (D-erythro-N-hexanoylsphingosine, BioMol #SL-110) and dihydro-C6 (D-erythro-N-hexanoylsphinganine, BioMol #SL-111) were also dissolved at 12.5-mM stock solutions in DMSO. Okadaic acid and PMA were purchased from Sigma, while roscovitine, aphidicolin and nocodazole were purchased from BioMol.

2.3 [3H]Thymidine incorporation
To measure DNA synthesis, 15,000 endothelial cells per 35-mm dish were starved for 48 h in EBM2 with 0.5% FCS lacking growth factors. Growth-media and 3[H]thymidine (3 µCi/ml) were added for selected times up to 44 h. Adherent cultures were fixed with 1 ml of 10% TCA, solubilised in 0.25 N NaOH and then harvested. 3[H]Thymidine incorporation was determined by liquid scintillation counting. Each sample was done in triplicate, and the data are presented as mean values±S.E.M. of the replicates per assay.

2.4 BrdU-ELISA
BrdU ELISA was performed in 96-well plates. HAECs were seeded at subconfluent density, and conditions were added the next day. Then 10 µl of 10 mM BrdU labeling reagent (5'bromo-2'deoxyuridine) was added 42 h after the incubation with the indicated substance had been started. Another 6 h later, cells were fixed for 45 min at room temperature. Next 100 µl of anti-BrdU antibody in a 1:100 dilution were added per well and incubated for 90 min at 37°C. After three washing steps peroxidase-substrate solution (100 µl) was added to each well and the color reaction was stopped after 5 min with 25 µl 1 M H2SO4. Absorbance was measured at 450 nm. Each condition was tested in parallel in 18 wells, and the data are presented as mean values±S.E.M. of the replicates per assay.

2.5 MTS viability assay and measurement of cell death
CellTiter 96 AQueous non-radioactive cell viability assay (Promega) was used to assess cell viability and proliferation. The assay is composed of the tetrazolium compound MTS (3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)2H-tetrazolium) and an electron coupling reagent PMS (phenazine methosulfate). MTS is reduced by viable cells to formazan, which can be measured with a spectrophotometer by the amount of the increased 490-nm absorbance. Previous studies in other cell lines show clearly that the formazan product is time-dependent and proportional to the number of viable cells [17]. Endothelial cells were cultured in 100 µl of HAEC medium in 96-well flat bottomed, fibronectin coated culture plates (Becton Dickinson). Cultures were seeded at 6x103 cells/well for subconfluent conditions. After the indicated time of incubation with the appropriate medium, 20 µl of MTS/PMS (1:0.05) mixture was added per well and cells were incubated 2 h before measuring absorbance at 490 nm. Background absorbance from the control wells (same medium, no cells) was subtracted. Viability of control cells was set to 100% and cell death was measured as 100% — % viability of treated cells. Throughout this paper, each calculated value for viability or cell death, respectively, is the average of six wells per group.

2.6 Flow cytometry for measurement of cell-cycle distribution
Adherent cells were detached by trypsinization (0.05% (w/v) trypsin in 0.02% (w/v) EDTA in HBSS), incubated for 5 min at 37°C, and harvested. After washing once in PBS and slow centrifugation (400xg) the pellet was resuspended in ice cold ethanol (70%) and fixed overnight at 4°C. As previously described [18] the cell pellet was stained in PBS, pH 7.4, with addition of 0.1% (w/v) Triton X-100 (Sigma), 0.5 mM EDTA, pH 7.4, 0.05 mg/ml RNase A (50 U/l, Sigma) and 50 µg/ml propidium iodide (Boehringer Mannheim) at 4°C overnight. A final concentration of 106 cells/ml staining solution was achieved. DNA content was analyzed from 10,000 cells (events) per group within the fluorescence gate using a Beckton Dickinson Flow Cytometer in combination with the CytoFlow program version 2.2. CellFIT cell-cycle software from Becton Dickinson (Version 2.01.2) was used to determine the percentage of cells in G0/G1, S or G2/M phase.

2.7 Flow cytometry for measurement of propidium iodide uptake
Staining for propidium iodide uptake was performed as previously described [1]. In brief, pre-treated endothelial cells were washed twice with PBS and detached using trypsin/1 mM EDTA. Bovine serum albumin was added to neutralize trypsin and cells were centrifuged for 5 min at 1000xg. Following two more washing steps the cell pellet was dissolved in 100 µl incubation buffer (10 µl 10xbinding buffer, 10 µl propidium iodide and 80 µl dH2O). After 15-min incubation 400 µl 1xbinding buffer was added and PI-fluorescence was measured recording 20,000 events.

2.8 Whole cell extracts
Cells were washed three times in cold phosphate-buffered saline (PBS), and then lysed for 30 min at 4°C in lysis buffer containing 50 mM Tris–HCl (pH 8.0), 2 mM EDTA (pH 8.0), 150 mM NaCl, 0.5% Nonidet P-40 and the following protease inhibitors: 0.5 mM/l PMSF, 1 µg/ml aprotinin, 1 µg/ml leupeptin and 0.5 µg/ml pepstatin A. After centrifugation at high speed supernatant was collected and protein content of all samples was determined using the Bio-Rad protein assay with {gamma} globulin as a standard.

2.9 Western blot analysis
Electrophoresis was performed on 12% SDS–polyacrylamide gels loading 30 µg of protein per lane. After transfer to a 0.2-µm PVDF membrane (Bio-Rad) membranes were blocked in 10% (w/v) non-fat dry milk (in PBS, pH 7.5, and 0.1% Tween 20) for 1 h and incubated for 150 min with the monoclonal or polyclonal antibodies at 4°C. Each antibody incubation period was followed by 1 h of membrane washing in 0.1% Tween-20 in PBS.

Detection was carried out by use of a secondary horseradish peroxidase-linked anti-rabbit (1:7500 in PBS/1% Tween 20/2% non-fat milk) or anti-mouse (1:5000 in PBS/1% Tween 20/2% non-fat milk) antibody (Santa Cruz) and the Amersham enhanced chemiluminescence system (ECL kit).

2.10 Inhibition of cyclin A expression by antisense-oligonucleotides
Endothelial cells were cultured in 100 µl of HAEC medium in 96-well flat bottomed, fibronectin coated culture plates (Becton Dickinson). Cultures were seeded at 6x103 cells/well for subconfluent conditions. Antisense oligonucleotides (Biognostics, Heidelberg) were added at 5-µM concentration to HAECs for 48–72 h, depending on the experiment.

2.11 Statistical analysis
All values are expressed as mean±S.E.M., whether in numbers or charted. Differences between groups were assessed by unpaired t-test (2 groups) or one-way ANOVA (<2 groups). Subsequent multiple comparisons for ≥3 groups were performed only if one-way ANOVA reached statistical significance P<0.05, using the Student–Newman–Keul's test to compare all pairs or Dunnett's post test to compare each group against control. The level of statistical significance is indicated in each figure. Statistical calculations were carried out with GraphPad Prism version 3.00 for Windows (GraphPad Software, San Diego California USA, www.graphpad.com).


   3 Results
Top
 Abstract
 1 Introduction
 2 Methods
 3 Results
4 Discussion
 References
 
3.1 Role of cell density in ceramide-induced cell death
Ceramide toxicity was compared in subconfluent (6000 cells/well) versus confluent (40,000 cells/well) endothelial cell layers. Concentrations of 10 µM and less had no effect on cell viability, independent of cell density. Half-maximal cell death (LD50) was reached at 25 µM ceramide in subconfluent endothelial cells versus 75 µM in confluent cells (Fig. 1A). Ceramide induced cell death was specific, since introduction of a dihydro-group into C2- or C6-ceramide completely abrogated its toxic effect (Fig. 1B). Flowcytometric analysis of PI uptake is a marker for disruption of the cytoplasmic membrane, as seen in necrosis or later stages of apoptosis. While 10 µM of C2-ceramide did not lead to a significant increase, 20 µM and higher concentrations showed a more than 2-fold increase in PI uptake by HAECs (P<0.01) (Fig. 2A). To evaluate the morphology of cell nuclei under treatment with ceramide, we performed DAPI staining. Control cells showed oval shaped nuclei with small nucleoli (Fig. 2B) whereas C2-treated endothelial cells (30 µM) contained rounded, condensed nuclei with intense fluorescence (Fig. 2C).


Figure 1
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  		Fig. 1 Effect of cell density on ceramide toxicity. Confluent endothelial cells are more resistant to C2-ceramide. (A) MTS assay was performed with subconfluent (6000 cells/well) versus confluent (40,000 cells/well) HAECs in 96-well plates (FALCON, tissue culture treated) over 24 h. Cell death was calculated as described under methods from six wells per treatment group, and a representative of four experiments is shown. (B) C2-ceramide, dihydro-C2, C6-ceramide and dihydro-C6 were compared for their potential to induce cell death. MTS assay was performed with subconfluent HAECs in 96-well plates over 48 h in six wells per treatment group, and a representative of three experiments is shown. Data are presented as mean±S.E.M. *P<0.01 versus control; NS, not significant.

 

Figure 2
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  		Fig. 2 Morphology of ceramide-induced cell death. (A) Uptake of PI by necrotic endothelial cells was quantified by FACS analysis. Cells were treated with up to 25 µM C2-ceramide. Data are presented as mean±S.E.M. Some 20,000 events per group served as the basis for calculation (n = 2). Data are presented as mean±S.E.M., and a representative of two experiments is shown. (B–C) Fluorescence-microscopic image of control versus C2-ceramide (30 µM for 24 h) treated HAECs after staining with DAPI.

 
3.2 Ceramide inhibits growth of endothelial cells independent of cell death
To evaluate the effect of ceramide on endothelial cell growth, DNA synthesis was measured by two independent methods: BrdU-incorporation at 48 h was significantly inhibited at 6.25 µM (24%) and 12.5 µM (62%), while cell viability remained unchanged (Fig. 3A). Dihydro-C2 was used as a control for C2 specificity and did not have any anti-proliferative effect. [3H]Thymidine incorporation studies also showed significant inhibition of DNA synthesis as early as 12 h after ceramide treatment (Fig. 3B). Conversely, primary toxic concentrations of C2-ceramide (25 µM) also led to growth-inhibition measured after 48 h, even when applied only for up to 2 h while no toxicity could be shown in a parallel MTS assay from such short exposure (Fig. 4A).


Figure 3
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  		Fig. 3 Ceramide inhibits growth of endothelial cells in non-toxic concentrations. (A) BrdU assay (black bars) and MTS assay (grey bars) were performed on HAECs in parallel plates using equal ceramide concentrations. Ceramide was dissolved in DMSO as a 12.5-mM stock solution, and 0.2% DMSO served as vehicle control. MTS and BrdU assays were performed with subconfluent HAECs in 96-well plates over 48 h in six wells per treatment group, and a representative of three experiments is shown. (B) HAECs were serum starved for 48 h in six-well plates and then restimulated with growth media with or without ceramide at different concentrations. [3H]Thymidine was added at 1 µCi between 0 and 1 h (serum starved=0 h) or between 11 and 12 h. Cells were fixed and [3H]thymidine incorporation was measured with a scintillation counter (n = 3). A representative of two experiments is shown. Data are presented as mean±S.E.M. *P<0.05 versus control; §P<0.01 versus control; NS, not significant.

 

Figure 4
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  		Fig. 4 Ceramide-induced growth-arrest is due to G1 arrest. (A) BrdU assay (black bars) and MTS assay (grey bars) were performed on HAECs in parallel plates using 25 µM C2-ceramide (Co, control). At the indicated time points C2-ceramide containing medium was replaced with growth-medium. Assays were performed with subconfluent HAECs in 96-well plates over 48 h in six wells per treatment group, and a representative of three experiments is shown. (B) Flow cytometric analysis of cell-cycle distribution. HAECs were serum starved for 24 h (1% serum, no growth factor supplement) in 10-cm plates and then restimulated with growth media with or without 12.5 µM C2-ceramide (n = 2, with 20,000 events in each group). A representative of three experiments is shown. Data are presented as mean±S.E.M.

 
3.3 Loss of cyclin A leads to G1 cell-cycle arrest and subsequent growth-inhibition
We previously showed that TNF induced G1 cell-cycle arrest by downregulation of cyclin A. To evaluate whether ceramide induces growth arrest through similar mechanisms, we performed flow cytometric analysis of propidium-iodide stained cell samples. C2 treated HAECs demonstrated a significant increase in the G1 population after 12 h (87 vs. 50%, P<0.0001) compared to untreated control cells. S-phase was also repressed with addition of C2 (5 vs. 42%, P<0.0001), proving growth-inhibition due to G1 cell-cycle arrest with C2 treatment (Fig. 4B). Western blot analysis revealed downregulation of cyclin A and p21Cip1/Waf1 at 10-µM C2 concentration whereas other cell-cycle regulatory genes such as cyclin E, cdc2 or p53 remained unchanged at this concentration (Fig. 5A). To demonstrate that loss of cyclin A expression by itself is sufficient to cause growth arrest in human endothelial cells, we used antisense oligonucleotides with specific sequences against cyclin A mRNA or a scrambled sequence, respectively (Fig. 5B,C). BrdU-incorporation was reduced to 50% of control, while the scrambled sequence did not affect proliferation (P<0.001, Fig. 5D).


Figure 5
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  		Fig. 5 Cyclin A is required for cell-cycle progression of HAECs. (A) Western blot analysis of HAECs treated for 20 h with increasing concentrations of C2-ceramide. Co, control; Ve, vehicle (0.2% DMSO). (B) fluorescence micrograph from HAECs treated 24 h with 5 µM of cyclin A antisense oligonucleotides (FITC-conjugated). (C) Western blot showing expression of cyclin A in HAECs treated with growth medium (Cont), cyclin A antisense oligonucleotides (AS, 5 µM) or the scrambled sequence (CO-AS, 5 µM). Two representative blots are shown (n = 3). (D) Cyclin A antisense oligonucleotides (AS, 5 µM) or the scrambled sequence (CO, 5 µM) were added to the media for 54 h and BrdU added for another 18 h (n = 6). BrdU incorporation was measured at 72 h. A representative of three experiments is shown. Data are presented as mean±S.E.M.

 
3.4 Phosphatase PP2A is involved in ceramide-mediated G1-arrest
Phosphatase PP2A can be activated by induction of stress and hence cause growth arrest in other cell systems. First we verified the existence of endogenous PP2A and its catalytic subunit PP2Ac in HAECs by Western blotting (data not shown). We used okadaic acid (OA), a protein phosphatase inhibitor, which has been shown to specifically inhibit PP2A at 1-nM concentration, while inhibition of other phosphatases required higher concentrations. OA was found to be a strong inducer of cell proliferation with maximum increase in BrdU-incorporation (+43%) at 0.25 nM (Fig. 6A). Concentrations higher than 1 nM caused opposite effects, leading to growth inhibition, most likely due to inhibition of other protein phosphatases. To find out whether PP2A induction by C2 was responsible for growth inhibition, we used increasing amounts of subtoxic ceramide concentrations in combination with 0.25 nM OA. Addition of 10 µM C2 abrogated the proliferative effect of 0.25 nM OA (45 vs. 146% cell number after 72 h, P<0.0001) (Fig. 6B).


Figure 6
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  		Fig. 6 Influence of the phosphatase inhibitor okadaic acid on HAEC proliferation. (A) BrdU incorporation was measured at 48 h (n = 6). Negative percentage indicates growth inhibition, positive gain growth stimulation. Data are presented as mean±S.E.M. A representative of three experiments is shown. *P<0.05 versus control; §P<0.01 versus control; NS, not significant. (B) HAECs were treated simultaneously with 1 nM of okadaic acid and various C2 concentrations for 72 h. Relative cell number, as measured by the MTS assay, is indicated (control=100%). The assay was performed with subconfluent HAECs in 96-well plates in six wells per treatment group, and a representative of three experiments is shown. §P<0.01 versus cells treated with 1.0 nM okadaic acid.

 
3.5 Loss of cyclin A and G1-cell cycle arrest are a prerequisite of ceramide-induced toxicity
To investigate whether cyclin A was regulated throughout the cell-cycle in human endothelial cells, we synchronized cells in G1 phase with the phorbol ester PMA [19,20] (Fig. 7A–C) or the CDK-inhibitor roscovitine [21] (Fig. 7B–C). Both led to significant inhibition of BrdU-incorporation as well as cyclin A downregulation. Nocodazole arrested endothelial cells in G2/M phase, as we showed previously [13], and caused cyclin A downregulation (data not shown) accompanied by diminished BrdU-incorporation (Fig. 7B). Aphidicolin treated HAECs accumulated in S-phase [13,22] (Fig. 7A) and showed a 27% increase in BrdU-incorporation (Fig. 7B) together with significant upregulation of cyclin A protein expression (Fig. 7C), demonstrating a tight connection between cyclin A expression, S-phase and DNA-synthesis, respectively. Interestingly, the uncleaved 32-kDa fragment of caspase-3 (CPP32) was upregulated in G1- and G2-arrested HAECs when no cell death occurred (Fig. 7C). Finally, HAECs synchronized in G1-phase by roscovitine (Fig. 8A) and PMA (Fig. 8B) showed a 3- to 5-fold increase in cell death with C2-ceramide treatment (P<0.001). In contrast, S-phase synchronized HAECs were protected from C2-mediated cell death (6 vs. 17%, P<0.05) (Fig. 8C).


Figure 7
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  		Fig. 7 Relationship between cell-cycle position, BrdU-incorporation and cyclin A expression. (A) Flowcytometric analysis of propidium-iodide stained HAECs (20,000 events each). Cells were treated with growth medium (Control), 0.2 µg/ml aphidicolin, 100 nM PMA or 1 nM okadaic acid. S-phase population is either increased (aphidicolin, okadaic acid) or decreased (PMA) compared to control cells. (B) Influence of synchronization on HAEC proliferation, as determined by change in BrdU-incorporation relative to control (Ap, aphidicolin 0.2 µg/ml; Co, control medium with 2% FCS and without growth factor addition; No, nocodazole 0.02 µg/ml; PM, PMA 100 nM; Ro, roscovitine 2.5 µg/ml). A representative of three experiments is shown (six wells per treatment group). Data are presented as mean±S.E.M. *P<0.01 versus control, §P<0.001 versus control. (C) Western blot showing expression of cyclin A and uncleaved CPP32 in different phases of the endothelial cell cycle. (control lanes: growth media, PMA: 500, 100 and 10 nM, aphidicolin: 0.2, 0.1 and 0.02 µg/ml; Roscov: roscovitine 2.5 and 4 µg/ml). A representative blot is shown (n = 3).

 

Figure 8
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  		Fig. 8 Synchronization of HAECs in different phases of the cell-cycle alters susceptibility for ceramide. (A–C) HAECs were synchronized in G1 phase by 16-h pretreatment with 2.5 µg/ml roscovitine or 100 nM PMA, respectively, and in S-phase by 0.1 µg/ml aphidicolin. After medium exchange C2-ceramide was added at the indicated concentrations for 24 h and cell death was determined by the MTS assay. MTS assays were performed with subconfluent HAECs in 96-well plates. A representative of three experiments is shown (six wells per treatment group). Data are presented as mean±S.E.M., NS, not significant.

 

   4 Discussion
Top
 Abstract
 1 Introduction
 2 Methods
 3 Results
 4 Discussion
References
 
4.1 Ceramide-induced G1-arrest
Ceramide, a product of regulated sphingomyelin hydrolysis, is released within minutes following stimulation with tumor necrosis factor and has been implicated as a mediator of TNF-induced programmed cell death [15,23–25]. Beside its apoptotic effect, differences exist about the influence of ceramide on cell proliferation [10,26]. Though it is likely that apoptosis results in growth-arrest, i.e. through induction of p53 or p21Cip1/Waf1 [4,8] or simply destruction of the nuclear cell-cycle machinery, an independent effect of ceramide on cell cycle progression in vascular endothelial cells, however, has never been examined.

In this study we found that ceramide induces growth arrest in human endothelial cells independent of its toxic effect. Cell death occurred at concentrations of 15 µM ceramide and higher in subconfluent HAECs, while at subtoxic concentrations (≤12.5 µM) ceramide only caused G1 cell-cycle arrest subsequent to cyclin A downregulation.

Ceramide induced G1-arrest has been described by others in yeast, human leukemia cells (HL-60 and MOLT-4) and a Wi38 fibroblast cell-line, but not in human primary cells [26–28]. The mechanisms for ceramide mediated G1 arrest in these models relied on two distinct proteins: (a) in yeast, mutation of regulatory or catalytic subunits of a ceramide-activated protein phosphatase (CAPP) abrogates the antiproliferative effect of ceramide [28], and (b) in human leukemia cells ceramide blocked c-myc mRNA elongation as determined by nuclear run-on analysis [27]. Since Rudolph et al. identified a myc-responsive element in the cyclin A promoter [21], this could very well be one explanation for cyclin A downregulation as shown by our studies. In this context it is noteworthy that ceramide concentrations used in the other studies to induce G1-arrest in mammalian cell lines were also sufficient to induce apoptosis or cell death. This supports previous data showing a direct effect of TNF on the cyclin A promoter [29]. ‘Unspecific’ cleavage of other cell-cycle regulatory proteins, such as p21, under apoptotic C2-concentrations can be explained by modulation of caspase activity [30].

4.2 Role of protein phosphatase 2A
C2-ceramide as well as naturally occurring ceramide have been shown to activate a cytosolic protein phosphatase in vitro, an effect denied to other related sphingolipids [28]. This ceramide activated protein phosphatase (CAPP) seems to be a member of the 2A family of protein phosphatases (PP2A), all of which are heterotrimeric proteins comprised of a small catalytic subunit C, and two larger regulatory subunits, A and B [31]. Several PP2A species exist in the cell, and although CAPP has not yet been identified at the molecular level, we demonstrate that inhibiting PP2A with okadaic acid markedly increases endothelial cell proliferation. Interestingly, this phenomenon occurred only between 0.1- and 1.0-nM concentrations of okadaic acid while higher concentrations caused cell death. Okadaic acid is a polyketal fatty acid which can act as a strong tumor promoter [31,32]. It binds to and inhibits PP1c and PP2Ac with dissociation constants (Ki) of 147 and 0.032 nM, respectively. It can also inhibit nuclear telomerase activity necessary for S-phase progression with an ED50 of 0.2 nM, suggesting that the catalytic activity of PP2A is required in this process [33]. Therefore the effect of okadaic acid we saw in our experiments was specific for PP2A. Wolff et al. found that okadaic acid inhibited down-regulation of c-myc by TNF and C2-ceramide in a dose-dependent manner in HL-60 cells [27], though using concentrations between 10 and 100 nM. Since these are 10- to 100-fold higher concentrations as predicted from the kinetic data available on PP2Ac, it remains doubtful that C2 induced PP2Ac was responsible for c-myc down-regulation or simply other phosphatases, such as PP1. Our experiments find that phosphatase PP2A is involved in ceramide-mediated G1-cell cycle arrest.

4.3 Dependence of ceramide toxicity on cell cycle regulation
A potential link between cell proliferation and apoptosis has been suggested by the observation that several factors, including adenovirus E1A and members of the E2F gene family, induce both cell cycle progression and apoptosis [34]. In mice E2F can promote apoptosis and suppress proliferation [35]. In previous work we demonstrated that E2F rescues endothelial cells from TNF-induced apoptosis by restoration of cyclin A, thus overcoming G1-arrest [13]. Cyclin A is characterized by repression of its promoter during the G1 phase of the cell cycle and its induction at S-phase entry [36–38]. Furthermore, it can promote adhesion-dependent cell cycle progression [39]. Our experiments confirm the unique role of cyclin A in S-phase of endothelial cells, which notably are anchorage-dependent cells. In addition, S-phase cells were relatively protected from ceramide-induced cell death. Cyclin A seems therefore to be a possible contender for a cell-cycle dependent survival factor in endothelial cells. Our results demonstrated that (a) cyclin A is downregulated in G1 or G2 phase, (b) highly expressed in S-phase and (c) required for cell proliferation. Ceramide first leads to growth inhibition due to cyclin A downregulation. Further treatment or higher concentrations then cause toxic effects, resembling the morphology of necrosis and apoptosis in human endothelial cells. The toxic effect of ceramide can be enhanced by artificially keeping cyclin A levels low (PMA or roscovitine) and most importantly, toxicity can be abrogated by overcoming ceramide-induced G1-arrest through prior synchronization of cells in S-phase with aphidicolin. These findings strongly suggest that ceramide can only develop its toxicity when cells are already in G1-phase and cyclin A levels are suppressed. These findings suggest a strong bidirectional relationship between cell-cycle control and cell death in vessel biology.

Time for primary review 27 days.


   Acknowledgements
 
This work was supported by grants DFG Sp-502/2 of the Deutsche Forschungsgesellschaft and fortüne Sp-504/98 of the University of Tübingen. The assistance of Heike Runge for isolation of human endothelial cells and Anke Pfister for maintaining cell cultures is greatly appreciated.


   References
Top
 Abstract
 1 Introduction
 2 Methods
 3 Results
 4 Discussion
 References
 

  1. Vermes I., Haanen C., Steffens-Nakken H., Reutelingsperger C. A novel assay for apoptosis: flow cytometric detection of phosphatidylserine expression on early apoptotic cells using fluorescein labelled annexin V. J Immunol Methods (1995) 184:39–51.[CrossRef][ISI][Medline]
  2. Sherr C.J. G1 phase progression: cycling on cue. Cell (1994) 79:551–555.[CrossRef][ISI][Medline]
  3. Sherr C.J. Cancer cell cycles. Science (1996) 274:1672–1677.[Abstract/Free Full Text]
  4. Long X., Boluyt M.O., de Lourdes Hipolito M., Lundberg M.S., Zheng J.S., O'Neill L., et al. p53 and the hypoxia-induced apoptosis of cultured neonatal rat cardiomyocytes. J Clin Invest (1997) 11:2635–2643.
  5. Kenyon B.M., Voest E.E., Chen C.C., Flynn E., Folkman J., D'Amato R.J. A model of angiogenesis in the mouse cornea. Invest Ophthalmol Vis Sci (1996) 37:1625–1632.[Abstract/Free Full Text]
  6. Brooks P.C., Clark R.A.F., Cheresh D.A. Requirement of vascular integrin {alpha}vβ3 for angiogenesis. Science (1994) 264:569–571.[Abstract/Free Full Text]
  7. Friedlander M., Brooks P.C., Shaffer R.W., Kincaid C.M., Varner J.A., Cheresh D.A. Definition of two angiogenic pathways by distinct {alpha}v integrins. Science (1995) 270:1500–1502.[Abstract/Free Full Text]
  8. Strömblad S., Becker J.C., Yebra M., Brooks P.C., Cheresh D.A. Suppression of p53 activity and p21WAF1/CIP1 expression by vascular cell integrin {alpha}vβ3 during angiogenesis. J Clin Invest (1996) 98:426–433.[ISI][Medline]
  9. Santana P., Pena L.A., Haimovitz-Friedman A., Martin S., Green D., McLoughlin M., et al. Acid sphingomyelinase-deficient human lymphoblasts and mice are defective in radiation-induced apoptosis. Cell (1996) 86:189–199.[CrossRef][ISI][Medline]
  10. Auge N., Andrieu N., Negre-Salvayre A., Thiers J.C., Levade T., Salvayre R. The sphingomyelin-ceramide signaling pathway is involved in oxidized low density lipoprotein-induced cell proliferation. J Biol Chem (1996) 32:19251–19255.
  11. Harada-Shiba M., Kinoshita M., Kamido H., Shimokado K. Oxidized low density lipoprotein induces apoptosis in cultured human umbilical vein endothelial cells by common and unique mechanisms. J Biol Chem (1998) 273:9681–9687.[Abstract/Free Full Text]
  12. Marathe S., Schissel S.L., Yellin M.J., Beatini N., Mintzer R., Williams K.J., et al. Human vascular endothelial cells are a rich and regulatable source of secretory sphingomyelinase. J Biol Chem (1998) 273:4081–4088.[Abstract/Free Full Text]
  13. Spyridopoulos I., Principe N., Krasinski K., Kearney M., Isner J.M., Losordo D.W. Restoration of E2F expression rescues vascular endothelial cells from TNF-induced apoptosis. Circulation (1998) 98:2883–2890.[Abstract/Free Full Text]
  14. Slowik M.R., De Luca L.G., Min W., Pober J.S. Ceramide is not a signal for tumor necrosis factor-induced gene expression but does cause programmed cell death in human vascular endothelial cells. Circ Res (1996) 79:736–747.[Abstract/Free Full Text]
  15. Obeid L.M., Linardic C.M., Karolak L.A., Hannun Y.A. Programmed cell death induced by ceramide. Science (1993) 259:1769–1771.[Abstract/Free Full Text]
  16. Axel D.I., Brehm B.R., Wolburg-Buchholz K., Betz E.L., Köveker G., Karsch K.R. Induction of cell-rich and lipid-rich plaques in a transfilter coculture system with human vascular cells. J Vasc Res (1996) 33:327–339.[ISI][Medline]
  17. Buttke T.M., McCubrey J.A., Owen T.C. Use of an aqueous soluble tetrazolium/formazan assay to measure viability and proliferation of lymphokine-dependent cell lines. J Immunol Methods (1993) 157:233–240.[CrossRef][ISI][Medline]
  18. Spyridopoulos I., Sullivan A.B., Kearney M., Isner J.M., Losordo D.W. Estrogen-receptor-mediated inhibition of human endothelial cell apoptosis. Estradiol as a survival factor. Circulation (1997) 95:1505–1514.[Abstract/Free Full Text]
  19. Sasaguri T., Ishida A., Kosaka C., Nojima H., Ogata J. Phorbol ester inhibits the phosphorylation of the retinoblastoma protein without suppressing cyclin D-associated kinase in vascular smooth muscle cells. J Biol Chem (1996) 271:8345–8351.[Abstract/Free Full Text]
  20. Zhou W., Takuwa M., Kumada M., Takuwa Y. Protein kinase C-mediated bidirectional regulation of DNA synthesis, Rb protein phosphorylation, and cyclin-dependent kinases in human vascular endothelial cells. J Biol Chem (1993) 268:23041–23048.[Abstract/Free Full Text]
  21. Rudolph B., Saffrich R., Zwicker J., Henglein B., Müller R., Ansorge W., et al. Activation of cyclin-dependent kinases by Myc mediates induction of cyclin A, but not apoptosis. EMBO J (1996) 15:3065–3076.[ISI][Medline]
  22. O'Dwyer P.J., Moyer J.D., Suffness M., Harrison S.D.J., Crysyk R., Hamilton T.C., et al. Antitumor activity and biochemical effects of aphidicolin glycinate (NSC 303812) alone and in combination with cisplatin in vivo. Cancer Res (1994) 54:724–729.[Abstract/Free Full Text]
  23. Verheij M., Bose R., Lin X.H., Yao B., Jarvis W.D., Grant S., et al. Requirement for ceramide-initiated SAPK/JNK signalling in stress-induced apoptosis. Nature (1996) 380:75–79.[CrossRef][Medline]
  24. Hannun Y.A. Functions of ceramide in coordinating cellular responses to stress. Science (1996) 274:1855–1859.[Abstract/Free Full Text]
  25. Jayadev S., Linardic C.M., Hannun Y.A. Identification of arachidonic acid as a mediator of sphyngomyelin hydrolysis in response to tumor necrosis factor {alpha}. J Biol Chem (1994) 269:5757–5763.[Abstract/Free Full Text]
  26. Jayadev S., Liu B., Bielawska A.E., Lee J.Y., Nazaire F., Pushkareva M.Y., et al. Role of ceramide in cell cycle arrest. J Biol Chem (1995) 270:2047–2052.[Abstract/Free Full Text]
  27. Wolff R.A., Dobrowsky R.T., Bielawska A., Obeid L.M., Hannun Y.A. Role of ceramide-activated protein phosphatase in ceramide-mediated signal transduction. J Biol Chem (1994) 269:19605–19609.[Abstract/Free Full Text]
  28. Nickels J.T., Broach J.R. A ceramide-activated protein phosphatase mediates ceramide-induced G1 arrest of Saccharomyces cerevisiae. Genes Dev (1996) 10:382–394.[Abstract/Free Full Text]
  29. Spyridopoulos I., Sullivan A., Kearney M., Losordo D. Loss of anchorage in TNF treated endothelial cells correlates to downregulated cyclin A promoter activity. Circulation (1997) 95(Suppl_I):I–7.
  30. Gervais J.L.M., Seth P., Zhang H. Cleavage of CDK inhibitor p21Cip1/Waf1 by caspases is an early event during DNA damage-induced apoptosis. J Biol Chem (1998) 273:19207–19212.[Abstract/Free Full Text]
  31. Berndt N. Protein dephosphorylation and the intracellular control of the cell number. Front Biosci (1999) 4:d22–42.[CrossRef][Medline]
  32. Favre B., Turowski P., Hemmings B.A. Differential inhibition and posttranslational modification of protein phosphatase 1 and 2A in MCF7 cells treated with calyculin-A, okadaic acid and tautomycin. J Biol Chem (1997) 272:13856–13863.[Abstract/Free Full Text]
  33. Li H., Zhao L.L., Funder J.W., Liu J.P. Protein phosphatase 2A inhibits nuclear telomerase activity in human breast cancer cells. J Biol Chem (1997) 272:16729–16732.[Abstract/Free Full Text]
  34. Weinberg R.A. E2F and cell proliferation: a world turned upside down. Cell (1996) 85:457–459.[CrossRef][ISI][Medline]
  35. Field S.J., Tsai F.Y., Kuo F., Zubiaga A.M., Kaelin W.G. Jr., Livingston D.M., et al. E2F-1 functions in mice to promote apoptosis and suppress proliferation. Cell (1996) 85:549–561.[CrossRef][ISI][Medline]
  36. Schulze A., Zerfass-Thome K., Spitkovsky D., Middendorp S., Berges J., Helin K., et al. Cell cycle regulation of the cyclin A gene promoter is mediated by variant E2F site. Proc Natl Acad Sci USA (1995) 92:11264–11268.[Abstract/Free Full Text]
  37. Henglein B., Chenivesse X., Wang J., Eick D., Brechot C. Structure and cell cycle-regulated transcription of the human cyclin A gene. Proc Natl Acad Sci USA (1994) 91:5490–5494.[Abstract/Free Full Text]
  38. Desdouets C., Matesic G., Molina C.A., Foulkes N.S., Sassone-Corsi P., Brechot C., et al. Cell cycle regulation of cyclin A gene expression by the cyclic AMP-responsive transcription factors CREB and CREM. Mol Cell Biol (1995) 15:3301–3309.[Abstract]
  39. Guadagno T.M., Ohtsubo M., Roberts J.M., Assoian R.K. A link between cyclin A expression and adhesion-dependent cell cycle progression. Science (1993) 262:1572–1575.[Abstract/Free Full Text]

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