Am J Physiol Cell Physiol AJP: Gastrointestinal and Liver Physiology
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Am J Physiol Cell Physiol 289: C312-C322, 2005. First published March 16, 2005; doi:10.1152/ajpcell.00107.2004
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RECEPTORS AND SIGNAL TRANSDUCTION

Pathologically elevated cyclic hydrostatic pressure induces CD95-mediated apoptotic cell death in vascular endothelial cells

Cornelia Hasel, Susanne Dürr, Anke Bauer, Rene Heydrich, Silke Brüderlein, Tabe Tambi, Umesh Bhanot, and Peter Möller

Institute of Pathology, University of Ulm, Ulm, Germany

Submitted 25 February 2004 ; accepted in final form 11 March 2005


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
We describe cyclic hydrostatic pressure of 200/100 mmHg with a frequency of 85/min as a hemodynamically relevant pathological condition enforcing apoptosis in endothelial cells (EC) after 24 h of treatment. This went along with an increase of CD95 and CD95L surface expression, shedding of CD95L into the supernatant, cleavage of caspase-3 and caspase-8, and elevated JNK-2, c-Jun, and CD95L mRNA expression. Furthermore, increased DNA-binding activity of the AP-1 transcription factor family members FRA-1 and c-Jun was observed. This activation was reduced by inhibition of JNK, which subsequently prevented elevated CD95L mRNA expression. Caspase inhibitors and a CD95L-neutralizing antibody also reduced EC apoptosis. Most of the pressure-induced events were most prominent at 24 and 48 h. However, after 48 h, the CD95/CD95L expression pattern switched back to CD95–/CD95L+ and the specific death rate decreased. Cyclic pathological hydrostatic pressure is a novel type of stress to EC that renders them susceptible to CD95/CD95L-mediated autoapoptosis and/or paracrine apoptosis accompanied by upregulation of intracellular molecules known to trigger both apoptosis and survival.

hypertension; CD95/CD95L

MECHANICAL FORCES HAVE BEEN SHOWN to contribute to the development and aggravation of atherosclerosis by triggering proliferation or apoptosis in intimal and medial compartments of the arterial wall (11). In addition, apoptotic endothelial cells (EC) accumulate in atherosclerotic lesions with enhanced expression of proapoptotic factors (4), affirming a critical role for EC in vascular disease. Most studies to date have concentrated on fluid shear stress and cyclic strain as the main mechanical forces affecting EC, mainly via modulation of cytokine release, activation of intracellular kinases, and altered gene expression (1, 3, 12). It was shown that high shear stress occurring in laminar flow led to protection from apoptosis, while reduced laminar flow and turbulent flow promoted EC apoptosis and proliferation (24). Cyclic strain arising from a periodic change in vessel diameter due to pulsatile blood flow also has been shown to influence EC turnover, depending on the amount of strain applied (19, 28). While physiological amounts of cyclic strain inhibit EC apoptosis, e.g., by activation of the phosphatidylinositol 3-kinase pathway, pathological cyclic strains lead to induction of apoptosis (19). Very little is known how hydrostatic pressure affects EC turnover. Sustained hydrostatic pressure has been shown to inhibit or stimulate the proliferation of cells, largely depending on cell type, culture conditions, and pressure regimen. Vouyouka et al. (30) first presented data regarding pulsatile hydrostatic pressure leading to a decrease in cell numbers without affecting the cell viability under 160/100 mmHg. In cultured cardiomyocytes, pressure overload led to activation of the mitogen-activated protein kinase family member JNK, surprisingly mediated by CD95, a member of the TNF receptor family known to induce apoptosis upon binding CD95L and subsequent trimerization (2). In contrast to the well-studied role of CD95/CD95L in triggering apoptosis, there is growing evidence that CD95 signaling is also involved in proliferation and inflammatory processes via cross talk with other intracellular signaling pathways such as proteasome degradation and JNK activation (32). To study the effects of alterations in hydrostatic pressure on EC in vitro, we developed a pressurized chamber based on the Flexcell Strain Unit as described previously (8). With this device, the exertion of pulsatile hydrostatic pressure similar to in vivo situations with respect to amplitude, frequency, and duration is possible. We undertook this study to examine the effects of pathological hydrostatic pressure (200/100 mmHg) with a frequency of 85/min, corresponding to a clinically relevant hypertensive situation. We report herein that pulsatile pathological hydrostatic pressure itself is able to induce apoptosis in EC.


   MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell lines and culture conditions. Human umbilical vein endothelial cells (HUVECs) as well as human aortic endothelial cells (HAOECs) were obtained from PromoCell (Heidelberg, Germany), grown in endothelial cell basal medium (PromoCell) supplemented with 10% fetal calf serum (PAA, Linz, Austria), ECGS-heparin (PromoCell), 100 U/ml penicillin, 100 µg/ml streptomycin (BioWhittaker, Heidelberg, Germany), 50 ng/ml gentamicin, and 50 µg/ml amphotericin B (Biochrom, Berlin, Germany) and used for experiments within passages 6–8. All cells were maintained at 37°C in 5% CO2 atmosphere. Cells (106/well) were seeded onto Flexcell plates coated with collagen I (Dunn Laboratories, Ansbach, Germany) and allowed to attach for 24 h. All EC cultures showed a consistent spontaneous apoptotic death rate of ~20% independent of the detachment procedure, consistent with data reported in the literature (10, 17).

Cytospin preparations. An aliquot of complete cell suspension was poured into cytospin chambers and pelleted at 650 rpm for 3 min before cells were placed onto slides. The slides were air dried, fixed in methanol, and then stained with May-Grünwald (Merck, Darmstadt, Germany) for 5 min and 1.5% Giemsa (Merck) in H2O for 15 min. The slides were rinsed in H2O and air dried before examination.

Physiological and pathological hydrostatic pressure conditions. In short, the pressurized chamber based on the Flexcell Strain Unit enables the exertion of cyclic hydrostatic pressure on cells in vitro with a dynamic airflow and a defined membrane extension regulated by spacers. During operation up to 220 mmHg, O2 partial pressure and pH in the cell culture medium do not change compared with control cultures kept at normal atmosphere (8). Cells were placed in this pressurized chamber and subjected to the following pressure profiles: 0 mmHg, 120/80 mmHg, 160/80 mmHg, and 200/100 mmHg, all with a frequency of 85/min. The prechosen triangular pressure profile led to a pressure rise in the first 25% of each cycle duration, followed by a decrease for 75% of each cycle duration. The resulting pressure curve nearly equaled the arterial blood pressure curve in humans (Fig. 1). These parameters do not change in different pressure profiles up to 240 mmHg unless the frequency is altered. In all experiments described in this report, membrane extension was fixed at 0 using a spacer inserted underneath the flexible membrane to restrict the effect to pure hydrostatic pressure. In one set of experiments, cells were exposed to the above-mentioned pressure profiles and either 50 µM Z-Val-Ala-D,L-fluoromethylketone (ZVAD-fmk), a broad spectrum caspase inhibitor, 100 µM Ac-Asp-Glu-Val-aspartic acid aldehyde (Ac-DEVD-CHO), a caspase-3 inhibitor (Bachem, Heidelberg, Germany), or 1 µM D-JNKI1, a protease-resistant JNK inhibitor (Alexis Biochemicals, Grünberg, Germany). In blocking experiments, the CD95L-neutralizing monoclonal antibody (MAb) NOK-1 (Pharmingen, San Diego, CA) was added before as well as every 24 h afterward at a concentration of 1 µg/ml. To detect early responses, additional cell samples were obtained 2, 4, 6, 8, 10, and 12 h after the onset of the experimental condition. Cells were washed once with PBS without Ca2+ or Mg2+ (Life Technologies) and then harvested with 500 mg/l trypsin diluted 1:250 with 200 mg/l EDTA (BioWhittaker). Cells were resuspended in the supernatant, centrifuged at 2,000 g for 5 min at room temperature, and further prepared for flow cytometric analysis of DNA content.



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  		Fig. 1. Triangular pressure profile leads to a pressure rise in the first 25% of each cycle duration, followed by a decrease during 75% of each cycle duration. The resulting pressure curve mimics the arterial blood pressure curve in humans.

 
Flow cytometric analysis of annexin V binding. To detect annexin V binding as a marker of early apoptosis, cells exposed to pathological hydrostatic pressure (200/100 mmHg) for 24 h and were harvested as described above. Cells kept under normal atmospheric pressure served as controls. After being washed twice in PBS, cells were incubated with 5 µl of a fluorescein isothiocyanate (FITC)-labeled mouse anti-human annexin V MAb (IgG1 isotype; Pharmingen) and 2 µl of propidium iodide (50 mg/ml) for 15 min in the dark. After being gated for propidium iodide negativity to exclude nonapoptotic dead cells, 105 events were examined for each determination. Flow cytometry was performed on a FACSCalibur device with CellQuest software (Becton Dickinson, Mountain View, CA).

Flow cytometric analysis of DNA fragmentation. To quantify cells with advanced DNA degradation, we used a procedure described by Nicoletti et al. (22). In short, ~106 cells per sample were gently resuspended in 500 µl of hypotonic fluorochrome solution containing 0.1% Triton X-100, 0.1% sodium citrate, and 50 µg/ml propidium iodide (Sigma). The cell suspensions were kept at 4°C in the dark overnight before flow cytometric analysis was performed. For each determination, 104 events were examined. Percentage of specific death was defined as the percentage of DNA fragmentation in the presence of pathophysiological hydrostatic pressure compared with the percentage of DNA fragmentation of the untreated control and cells treated with physiological hydrostatic pressure. Flow cytometry was performed on a FACSCalibur device equipped with CellQuest software.

Flow cytometry of surface and cytoplasmic CD95/CD95L expression. For analysis of cytoplasmic flow cytometric measurements, cells were fixed in 2% paraformaldehyde and incubated for 15 min in permeabilization solution containing 0.2% Tween 20. The following primary mouse anti-human MAbs used were CD3 (Leu4, IgG1 isotype; Dako, Copenhagen, Denmark) as an isotype-matched negative control, CD95L antibody G247-4 (IgG1 isotype; Pharmingen), and CD95 (anti-Apo-1, IgG1 isotype; Dako). Immunofluorescent staining was performed in polystyrene round-bottomed tubes (Falcon, San Jose, CA). For dilution and washing, HBSS containing 2% bovine serum albumin (BSA) and 0.1% sodium acid, referred to as FACS medium, was used. After exposure of cells to cyclic pathological hydrostatic pressure (200/100 mmHg) for 24 and 48 h, 106 cells per sample were resuspended in FACS medium and incubated on ice with the appropriate volume of each MAb. After 1 h, cells were washed twice in FACS medium and 2 µg of FITC-labeled F(ab')2 goat anti-mouse immunoglobulins (Dako) were added for another 30 min. Cells were washed twice and resuspended in 300 µl of FACS medium containing 1 µg/ml propidium iodide (Sigma) to allow selective gating of nonapoptotic dead cells (i.e., propidium iodide-positive cells).

Human soluble CD95L enzyme-linked immunosorbent assay. To determine protein concentrations of CD95L in the supernatant of HUVECs treated with 200/100 mmHg for 2–24 h, supernatants were sampled at different time points. We applied a commercial enzyme-linked immunosorbent assay (ELISA) kit using a standard of known CD95L concentration (EuroClone, Devon, UK). Each protein lysate (100 µl), together with a dilution series of the standard, was dispensed in triplicate in 96-well microtiter plates precoated with a MAb against human CD95L for antigen capture. Simultaneously, a biotinylated MAb against human CD95L was added for detection. Plates were sealed and incubated for 3 h at room temperature. After repeated washing of the plates, we added streptavidin-conjugated horseradish peroxidase (HRP), resealed and incubated the plates for 30 min at room temperature, and then washed the plates again. A ready-to-use 3,3',5,5'-tetramethylbenzidine solution was used as the substrate. This reaction leads to the formation of a colored product absorbing light at 450 nm. The color reagent was dispensed in each well. The light absorption was measured and calibrated against the standard using an MRX microplate reader (Dynatech Laboratories, Chantilly, VA). All wells were measured in triplicate. Results were analyzed using Revelation software (Dynatech), and data are means ± SD expressed in picograms per milliliter.

Preparation of RNA and cDNA synthesis. Total RNA was extracted using TRIzol reagent (Life Technologies) according to the manufacturer's instructions, precipitated with 1 volume of 2-propanol, and rinsed with 70% ethanol. Total RNA was digested with RNAse-free DNAse I (Boehringer Mannheim, Mannheim, Germany) for 30 min at 37°C and precipitated with 3 volumes of ethanol at –20°C for 1 h. The RNA pellet was air dried and dissolved in diethyl pyrocarbonate (DEPC) water. Optical density (OD) was measured at 260 and 280 nm. RNA was quantified as OD260 = 1 = 40 µg/ml. The OD260/280 ratio showed values between 1.6 and 1.8 as required for pure RNA content. To ensure the purity of RNA, PCR was performed as described below using an SP1 primer and an RNA template, which yielded no amplification product (data not shown). Total RNA (5 µg) was incubated with 1 µl of poly(dT)15 (500 µg/ml) and denatured at 80°C for 5 min to ensure linear cords, followed by brief centrifugation and quick chilling on ice. First-strand buffer (5x), 0.1 M DTT, 10 mM dNTP mix, 1 U of SuperScript reverse transcriptase (Life Technologies), 40 U of RNAsin (Promega, Madison, WI), and DEPC water were added. cDNA synthesis was performed with a DNA Thermal Cycler (PerkinElmer, Norwalk, CT).

Real-time PCR. Real-time semiquantitative analysis of CD95L mRNA was performed using the iCycler IQ detection system and software (Bio-Rad Laboratories, Munich, Germany). Commercially available primers and probe for CD95L and cyclophilin were purchased from Applied Biosystems (Branchburg, NJ). First, the absence of nonspecific amplification was confirmed by analyzing the PCR amplification products using agarose gel electrophoresis. Amplicons generated from cDNA were also tested against no template control and RNA. The curves were checked for low cycle threshold (CT) and fast rising, and they were analyzed using agarose gel electrophoresis for confirmation. Real-time PCR was performed using 4 µl of cDNA (12.5 ng/µl), 4 µl of CD95L, c-Jun NH2-terminal kinase (JNK2), and c-Jun primer probe mix (Applied Biosystems, Foster City, CA), 12 µl of sterile distilled water, and 20 µl of PCR Universal Master Mix (Applied Biosystems) per reaction. The following cycling conditions were set: denaturation at 95°C for 2 min, followed by 45 cycles at 95°C for 15 s and 60°C for 1 min. Gene expression of CD95L, JNK2, and c-Jun in HUVEC subjected to cyclic pathological pressure (200/100 mmHg) for 6, 12, 24, and 48 h was measured relative to untreated cells as the calibrator sample. All quantitations were also normalized to cyclophilin as an endogenous control to account for variability in the initial concentration of total RNA. Analysis of quantitation was performed by calculating 1) the mean CT value of two replicates/sample, 2) the difference between mean CT values of samples for each target and those of the endogenous controls (CT), 3) the difference between mean CT values of the samples for each target and the mean CT value of the corresponding calibrator (CT). The quantitation is expressed as 2–{Delta}{Delta}CT to allow graphic presentation and shown as x-fold expression of the target gene in controls compared with stimulated cells. All experiments were performed in triplicate, and data are expressed as means ± SE.

Immunoblotting. Cytosolic proteins were extracted according to Dignam's protocol (5). Protein concentration was determined using Bradford reagent (Bio-Rad). Every total protein (10 µg) was separated on 10–20% Tricine precast gel (Novex, San Diego, CA) and transferred onto a polyvinylidene difluoride membrane. Membranes were incubated with the mouse anti-human MAbs anti-caspase-3/CPP 32 (IgG1 isotype; Transduction Laboratories) and anti-caspase-8/C15 (IgG2a isotype; generous gift from G. Moldenhauer, DKFZ, Heidelberg, Germany). Primary antibodies were then incubated overnight at 4°C after being blocked in PBS containing 0.05% Tween 20, followed by incubation with mouse anti-human biotinylated immunoglobulins (1:5,000 dilution; Dako) and streptavidin (1:5,000 dilution; Dako). For detection of phosphorylated JNK2, a rabbit anti-human polyclonal HRP-linked antibody was used (Cell Signaling, Beverly, MA). After another three washes, the blots were developed by performing enhanced chemiluminescence using the ECL system (Amersham).

AP-1 family transcription factor assay. DNA binding activity of different members of the activator protein (AP)-1 transcription factor family in HUVECs treated with 220/100 mmHg for 2–48 h was determined using an ELISA-based assay kit (TransAM kit) obtained from Active Motif (Rixensart, Belgium). In brief, the nuclear extracts were added to microwells coated with a cold oligonucleotide containing the consensus binding site for AP-1. After 1-h incubation at room temperature, the microwells were washed three times with washing solution. Antibodies directed against phosphorylated c-Jun (JunB, JunD, Fra-1, Fra-2, c-fos, and FosB) were used to label the AP-1 dimers bound to the oligonucleotide, followed by a secondary antibody conjugated to HRP. Finally, the results were quantified using a chromogenic reaction. The results were analyzed using Revelation software (Dynatech), and data are expressed in picograms per milliliter as means ± SE.

Electrophoretic mobility shift assay. Nuclear protein extract prepared according to the Dignam protocol (5) was used in the protein binding reactions. Complementary oligonucleotide for AP-1 [5'-(AGG) CGG TTG CTC ACT AAT TG-3'; 5'-(AGG) CT ATT AGT GAG CAA CCG-3'] and CD95L –120 [5'-(AGG) TCA GCT GCA AAG TGA GTG GGT GTT TCT TTG AG-3'; 5'-(AGG) CT CAA AGA AAC ACC CAC TCA CTT TGC AGC TGA-3'] probes were annealed and end labeled with [{alpha}-32P]dCTP using Klenow enzyme (Amersham). The specific activity of the probes used in the assays was adjusted to 50,000 cpm/0.1 pmol DNA. Oligonucleotides were incubated with 5 µg of nuclear protein extract in a buffer consisting of 10 mM HEPES, pH 7.9, 250 mM KCl, 5 mM EDTA, 20% Ficoll, 5 mM DTT, 0.2 µg/µl poly(dI-dC), and 1 µg/µl BSA in a total volume of 20 µl. Samples were loaded onto 4.5% polyacrylamide gels and run at 10 V/cm for 2 h in 0.5x Tris-borate-EDTA buffer. Gels were dried and exposed to X-ray film. Densitometric analysis of the DNA protein complexes was performed using the captured images with ImageMaster VDS software (Amersham Biosciences).

Statistical analysis. The Wilcoxon rank-sum test was applied for statistical comparison of untreated controls and cells exposed to cyclic hydrostatic pressure. The results of this test were labeled significant at P ≤ 0.05.


   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
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DISCUSSION
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Cyclic pathological hydrostatic pressure induces apoptotic cell death. HUVECs subjected for 24 h to cyclic pathological hydrostatic pressure of 200/100 mmHg with a frequency of 85/min, regarded as corresponding to a clinically relevant hypertensive state (also referred to herein as pathological hydrostatic pressure), showed cytomorphology of apoptotic cell death with typical membrane blebbing, nuclear condensation, and fragmentation (Fig. 2A). Prolonged exposure to this pressure regimen led to a recovery of the cell culture. We thus set out to study this seemingly transient effect in more detail and used annexin V binding and DNA fragmentation as assays for early and late apoptosis, respectively. After 24 h, 48.2 ± 5.2 of the cells treated with a cyclic pressure of 200/100 mmHg showed annexin V binding, which was more than twofold compared with untreated controls (19.2 ± 6.2) (Fig. 2B). Apoptosis induction was a rather late effect because cells treated with 200/100 mmHg for 6 and 12 h featured apoptotic rates in the range of 25.5 ± 5.1% and 15.7 ± 7.2%, respectively. Correspondingly, the percentage of DNA fragmentation increased significantly (P < 0.03) from 20.6 ± 6.5% in untreated control cells to 71.0 ± 7.8% in cells treated with high pressure (200/100 mmHg). Under these conditions, the apoptosis rate further increased to 82.8 ± 2.3% after 48 h and subsequently declined to 32.3 ± 4.6% after 72 h and to 42 ± 5.1% after 96 h. At the later time points, cell cultures were not depleted but tended morphologically to proliferate and reattain confluence (Fig. 2A). This pressure-induced apoptosis proved to be pulse dependent because static pressure of 200 mmHg up to 96 h did not affect the viability of HUVECs (Fig. 3). Furthermore, apoptosis induction was observed only under pathological hydrostatic pressure; no changes were noted at 120/80 mmHg (24 h: 25.0 ± 0.5%; 48 h: 24.3 ± 2.4%) and 160/80 mmHg (24 h: 23.2 ± 3.1%; 48 h: 24.7 ± 2.4%). To evaluate whether HUVECs are comparable to human aortic ECs (HAOECs), we also subjected HAOECs to 200/100 mmHg at the rate of 85/min for 24 h. These conditions led to an apoptotic death rate comparable to that in HUVECs as shown using annexin V binding (72.8 ± 5.6% vs. 25.5 ± 4.1% in untreated controls) (Fig. 2C). In summary, pathological hydrostatic pressure induced apoptosis in ECs starting after 12 -h exposure and peaking at 48 h.



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  		Fig. 2. A: exposure to pathological hydrostatic pressure led to an increase in endothelial cell (EC) size and induced morphological features of apoptosis after 24 h. Treatment for 72 h revealed a decreasing amount of apoptotic cells accompanied by an increase in mitotic activity (May-Grünwald/Giemsa staining). B and C: effects of pathological hydrostatic pressure on annexin V binding of human umbilical vein endothelial cells (HUVECs) (B) and human aortic endothelial cells (HAOECs) (C) after 24 h revealed using flow cytometry.

 


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  		Fig. 3. A: physiological cyclic pressure (120/80 mmHg). B: elevated systolic pressure (160/80 mmHg). C: pathological cyclic pressure (200/100 mmHg) (*P < 0.03). D: static pressure of 200 mmHg. Percentage of DNA fragmentation (means ± SE) after 24, 48, 72, and 96 h of pathological hydrostatic pressure compared with physiological cyclic pressure and untreated cells. The degree of DNA fragmentation was determined by using flow cytometry to measure hypodiploid events after propidium iodide staining as an indirect measure of the apoptotic death rate.

 
Caspases are critically involved in apoptosis by pathological hydrostatic pressure. To further substantiate our observation, we performed caspase inhibition assays. HUVECs were subjected to cyclic pressure (200/100 mmHg) with and without caspase inhibitors ZVAD-fmk and Ac-DEVD-CHO, respectively. After 24 h, cells were harvested and the sub-G1 peak was determined using flow cytometry. Each of these inhibitors extensively, although not completely, blocked cell death induced by pathological hydrostatic pressure. The broadly reacting caspase inhibitor ZVAD-fmk reduced specific death to 28.7 ± 5.1% (P ≤ 0.01) compared with control cells with ZVAD-fmk (24.7 ± 8.1%) and untreated controls (20.6 ± 6.5%). Caspase-3 inhibitor Ac-DEVD-CHO was slightly less effective and reduced specific death to 36.2 ± 6.3% (controls with Ac-DEVD-CHO; 29.3 ± 8.1%; P ≤ 0.01) (Fig. 4A). Application of elevated pressure also led to the cleavage of death receptor-associated caspase-8, which was detectable after 24 and 48 h, with no detectable levels at 72 and 96 h (Fig. 4B). Caspase-8 cleavage was accompanied by a loss of the noncleaved form of effector caspase-3 at 24 h, reverting to normal conditions after 48 h. This strongly suggests a death receptor-dependent signaling pathway that is operative in the induction of apoptosis by cyclic, pathologically elevated hydrostatic pressure.



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  		Fig. 4. A: caspase inhibition reduces the apoptotic death rate. HUVECs were subjected to pathological hydrostatic pressure with or without different caspase inhibitors, Z-Val-Ala-D,L-fluoromethylketone (ZVAD-fmk) and Ac-Asp-Glu-Val-aspartic acid aldehyde (Ac-DEVD-CHO), for 24 h. The degree of DNA fragmentation (means ± SE) was determined by using flow cytometry to measure hypodiploid events after propidium iodide staining. B: simultaneously, cleavage of caspase-8 (arrow) and loss of noncleaved caspase-3 were detected using immunoblotting at 24 and 48 h.

 
CD95/CD95L system is involved in apoptosis triggered by pathological hydrostatic pressure. Cytoplasmic and surface expression of CD95 and CD95L were evidenced by FACS analysis (Fig. 5A). Constitutively, HUVECs showed moderate CD95 expression in the cytoplasm and low levels of cell surface expression. Treatment with cyclic pressure of 200/100 mmHg did not alter this expression pattern after 6 and 12 h. However, after 24 h, there was a switch with increased surface and decreased cytoplasmic CD95 levels. After 48 h of treatment, surface and cytoplasmic CD95 expression returned to almost initial levels (Fig. 5A). Next, we examined the expression of cytoplasmic and surface CD95L expression. Untreated HUVECs showed constitutive CD95L expression in the cytoplasm, with a slight decrease after 24 h under cyclic high pressure accompanied by neoexpression of CD95L on the cell surface. After 48 h, this expression pattern returned, by and large, to initial levels. Again, these changes were not detectable after 6 and 12 h of treatment. This argues for a strong stress response within 24 h of exposure to pathological hydrostatic pressure, with the surviving cells trying to adapt to this condition, resulting in an only slightly increased apoptotic death rate after 48 h. Correspondingly, CD95L mRNA was significantly increased after 24 h of pathological hydrostatic pressure, with only slightly elevated levels at 48 h compared with untreated cells and cells treated for 6 and 12 h (Fig. 5B). In contrast, cells exposed to physiological hydrostatic pressure showed a decrease of CD95L mRNA to almost undetectable levels after 48 h (data not shown).



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  		Fig. 5. A: CD95L is expressed on the surface of ECs after 24 h of pathological hydrostatic pressure, reverting to initial levels after 48 h. Cytoplasmic CD95L expression levels were not changed. CD95 is expressed on the EC surface after 24 h of pathological hydrostatic pressure, reverting to initial levels after 48 h. This was paralleled by significantly decreased CD95 expression in the cytoplasm, followed by a return to close to initial levels. Analysis of both surface and cytoplasmic CD95/CD95L after 6 and 12 h showed no changes in the expression pattern compared with untreated controls. After 6, 12, 24, and 48 h of exposure, living cells were stained by fluorescein isothiocyanate (FITC)-labeled CD95 monoclonal antibody (MAb). The percentage of FITC-positive cells was determined by performing flow cytometry and compared with untreated cells. B: correspondingly, CD95L mRNA was significantly increased after 24 h but not after 6 or 12 h of pathological hydrostatic pressure, with only slightly elevated levels observed at 48 h as revealed using real-time PCR.

 
CD95L is detectable in the supernatant after 12 h of pathological hydrostatic pressure. To determine whether CD95L is shed into the supernatant during treatment with pathological cyclic pressure, we analyzed CD95L in supernatants at 0, 2, 4, 6, 8, 10, 12, 24, and 48 h (Fig. 6). CD95L protein was first detected after 12 h of stimulation with 200/100 mmHg as revealed using ELISA. Specifically, CD95L protein content was 0 pg/ml (SE ± 0.00) from 0 to 10 h with increasing quantities at 12 h (4.32 ± 0.01 pg/ml), 24 h (6.86 ± 0.01 pg/ml), and 48 h (10.67 ± 0.01 pg/ml) (Fig. 6).



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  		Fig. 6. CD95L protein was first detected in the supernatant after 12 h, with a further increase observed after 24 and 48 h of stimulation with 220/100 mmHg as revealed using enzyme-linked immunosorbent assay (ELISA).

 
Next, we applied the NOK-1 antibody, which efficiently neutralizes CD95L (14). As shown in Fig. 7, NOK-1 antibody, when added to the medium, substantially reduced the apoptotic death rate under cyclic high pressure after 24 h (25.4 ± 6.7%) and 48 h (44.5 ± 7.8%). This indicates that the peak of CD95 and CD95L expression on the cell surface of EC parallels the maximum sensitivity toward CD95-mediated apoptosis.



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  		Fig. 7. CD95L-neutralizing antibody significantly reduces apoptotic cell death triggered by pathological hydrostatic pressure. ECs were exposed to 200/100 mmHg pressure for 24 and 48 h in the presence vs. absence of the CD95L-inhibitory MAb NOK-1. The degree of DNA fragmentation (means ± SE) was determined using flow cytometry to measure hypodiploid events after propidium iodide staining.

 
Involvement of stress-associated proteins. Because members of the stress-associated protein kinase pathway are known to play a role in both apoptotic signaling and the regulation of death receptor/ligand expression, we determined RNA expression of JNK2 and its target c-Jun, which, upon phosphorylation, can act as a transcription factor involved in CD95L gene expression (9). JNK2 transcripts increased after 12-h treatment, peaked at 24 h, and returned to 12-h levels after 48 h (Fig. 8A). JNK phosphorylation was present at very low levels in controls with comparable levels at 2, 6, 8, and 12 h. After 24 h, there was a significant increase in JNK phosphorylation, which was sustained up to 48 h (Fig. 8B). Simultaneously, a significant increase in c-Jun RNA expression was observed at 24 and 48 h (Fig. 8C).



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  		Fig. 8. Pathological hydrostatic pressure induces stress-associated proteins JNK and c-Jun. A: at the mRNA level, a significant increase in JNK expression was first detected after 12 h and a peak was observed at 24 h, reverting to 12-h levels at 48 h. B: JNK phosphorylation was detectable at very low levels up to 12 h, with a distinct increase observed after 24 h and sustained levels noted at 48 h. UV light-treated Jurkat cells were used as a positive control. C: correspondingly, c-Jun mRNA expression was increased at 24 h, with similar expression levels observed at 48 h. JNK and c-Jun mRNA was detected after real-time PCR amplification. As an intrinsic standard, cyclophilin was coamplified.

 
Pathological hydrostatic pressure modulates DNA binding activity of members of the AP-1 transcription factor family. To determine whether this increase in JNK2/c-Jun expression also leads to altered DNA binding activity of different members of the AP-1 family (c-Jun, JunB, JunD, Fra-1, Fra-2, c-fos, and FosB), we performed an ELISA-based transcription factor assay. In contrast to c-fos, fos-B, Fra-2, Jun B, and JunD, which remained largely unaffected after 2, 4, 6, 8, 10, 12, 24, and 48 h of treatment with pathological cyclic pressure (200/100 mmHg), c-Jun and Fra-1 DNA binding activity increased after 24 h (Fig. 9A). To further substantiate these findings, we added the JNK-specific inhibitor JNKI1, which almost completely abolished activation of Fra-1, JunD, and c-Jun during treatment with 200/100 mmHg (Fig. 9B).



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  		Fig. 9. Pathological hydrostatic pressure induces induction of the activator protein (AP)-1 family members Fra-1, JunD, and c-Jun. A: in contrast to c-fos, fos-B, Fra-2, Jun B, and JunD, which largely remained unaffected by treatment with pathological cyclic pressure (200/100 mmHg), c-Jun and Fra-1 showed the most prominent increase in DNA binding activity after 24 h, with a decrease observed at 48 h. B: addition of the JNK-specific inhibitor D-JNKI1 almost completely abolished activation of Fra-1, JunD, and c-Jun.

 
Because c-Jun is also known to play a role in the regulation of CD95L expression, we further tested binding of nuclear extracts to a regulatory element in its promoter, using an oligonucleotide sequence described by Li-Weber et al. (20). After 24 h under cyclic high pressure, there was a strong induction of DNA binding activity compared with undetectable levels in control cells (Fig. 10A). The specificity of DNA binding to the CD95L promoter sequence was confirmed by the observation that a 100-fold excess of unlabeled probe inhibited the binding of the labeled probe to the nuclear proteins tested (data not shown). Moreover, the increase in CD95L mRNA expression after 24 h of treatment with a pressure of 200/100 mmHg was significantly reduced by addition of the JNK inhibitor. This strongly suggests a role of this kinase and the subsequent activation of c-Jun in restoring the initial levels of CD95L (Fig. 10B).



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  		Fig. 10. A: pathological hydrostatic pressure affects CD95L-regulatory element DNA-binding activity. Electrophoretic mobility shift assay (EMSA) was performed using nuclear extracts prepared from stimulated vs. untreated HUVECs after 24 and 48 h. Nuclear extracts were incubated with either 32P-labeled AP-1 or CD95L –120 (–129 to –98) binding nucleotide and subjected to electrophoresis on 4.5% polyacrylamide gel. B: increase in CD95L mRNA expression after 24 h of treatment with 200/100 mmHg could be reduced significantly by addition of the JNK-specific inhibitor D-JNKI1 as revealed by performing real-time PCR.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Chronically sustained arterial hypertension is one of the major risk factors in the development and progression of atherosclerotic lesions because it inflicts a smoldering injury on the intimal cell layer, leading to endothelial dysfunction and denudation of the intimal barrier (21). Emerging evidence suggests that EC apoptosis may be the causative event in the development of atherosclerosis (4, 7, 27). Various proatherogenic factors such as oxidized low-density lipoproteins, hypoxia, and inflammatory cytokines have been shown to be potent inducers of EC apoptosis (4). In contrast, mechanical forces such as fluid shear stress, cyclic strain, and hydrostatic pressure seem to act in a modulating way in inducing or preventing EC apoptosis, with the net effect being highly dependent on the amount of force applied (6, 15, 19, 28). Whether the initial injury of ECs is due to mechanical forces, metabolic factors, or a combination thereof requires further examination. Herein, we have shown that pathological hydrostatic pressure without additional strain or fluid shear stress is able to induce apoptosis in ECs within 24 h after onset. Caspase inhibitors and the addition of CD95L neutralizing antibody, with the latter being even more effective at 48 h of high pressure, significantly reduced the apoptotic death rate. Consistent with our findings, untreated ECs have been reported to show moderate CD95L and almost no CD95 expression on the cell surface (25). Untreated ECs further seem to be resistant to CD95L-mediated apoptosis, possibly because of the lack of receptor expression or the previously reported constitutive expression of c-FLIP, an inhibitor of caspase-8. The sensitivity toward CD95-mediated apoptosis also was not enhanced by increased CD95 surface expression after stimulation with IFN-{gamma} (25), indicating that, also in this cell type, CD95 surface expression does not necessarily parallel increased sensitivity toward CD95-mediated apoptosis (29). In our experimental setting, high-pressure-stressed ECs committed CD95-mediated suicide by release of blockable, soluble CD95L. Transferred to the natural context, this would imply the release of soluble CD95L into the arterial bloodstream in response to hypertension. Janin et al. (13) showed disseminated EC apoptosis in mice after application of agonistic CD95-specific antibody or soluble multimeric CD95L prevented by caspase inhibitors. Furthermore, a role of the CD95/CD95L system in atherogenesis has recently been emphasized by the finding that intimal medial thickening is associated with elevated soluble CD95L blood levels (23). We have shown that treatment of ECs with a pathologically elevated cyclic hydrostatic pressure profile corresponding to a hypertensive state leads to a transient switch in the expression of CD95 and CD95L from cytoplasm to surface after 24 h, with a return to approximately basal levels after 48 h. The change in the CD95/CD95L expression pattern was accompanied by an increase in JNK-2 and c-Jun mRNA expression after 12 h (peaking at 24 h), followed by a significant increase in JNK phosphorylation. This in turn led to an increase in DNA-binding activity among the AP-1 family members c-Jun and Fra-1. AP-1 is composed of heterodimeric protein complexes derived from the Fos and Jun families, and from those Jun proteins they form transcriptionally active dimers (26). Enhanced c-Jun binding also has been shown to activate transcriptional upregulation of CD95L expression after ligation of CD95, resulting in a so-called autostimulatory loop (9, 16). Furthermore, increased DNA binding to a regulatory element of the CD95L promoter identified by Li-Weber et al. (20) was induced by pathological hydrostatic pressure. On the other hand, activation of JNK with or without involving c-Jun has been reported to play a role in cell death as well as in cell survival (18). Until recently, the CD95/CD95L system was basically studied in terms of its capability to induce apoptosis in a variety of cell types and diseases. Yet, there is growing evidence that CD95 activation can also elicit nonapoptotic responses such as cytokine release, proliferation, and activation of transcription factors, e.g., NF-{kappa}B and AP-1. The signaling pathways involved in these CD95-mediated nonapoptotic responses have only been poorly analyzed. CD95-triggered proliferation has been shown to play a role in activated T cells via activation of caspases. In diploid fibroblasts, CD95 stimulation led to either apoptosis or proliferation, depending on the conditions used. CD95-mediated activation of JNK by, for example, cleavage and activation of MEKK-1 through active caspases has been reported to contribute to CD95-mediated apoptosis by phosphorylation of apoptosis-related proteins or activation of AP-1 (31, 32). Thus the prolonged activation of caspase-8 and the restoration of caspase-3 may point to an additional function of caspase-8, e.g., activation of JNK. The relatively early activation of c-Jun and JNK-2 in our experiments strongly suggests an initial role for c-Jun in increased CD95L synthesis, also referred to as autoamplification. These findings imply different possible mechanisms triggered by cyclic pathological pressure: First, upregulation of CD95 and increased sensitivity of ECs to CD95-mediated apoptosis are paralleled by increased CD95L surface expression and its possible release into the supernatant, leading to autocrine or paracrine apoptotic cell death. Second, the slightly higher apoptotic death rate after 48 h is paralleled by a decrease of CD95 surface expression as well as autoamplification of CD95L possibly triggered by c-Jun, which may contribute either to ongoing apoptosis or to the restoration of basic cell surface levels, thereby ending the vulnerable phase. Within this time frame, ECs may also lose their immunoprivileged status, potentially resulting in vulnerability to leukocyte attack (33). Pro- and antiapoptotic effects of increased JNK expression (18) have not yet been identified in this context, but the peak mRNA expression after 24 h in relation to the just slightly increased apoptotic death rate at this time point suggests a role of this kinase in the cell's adaptation to pathological hydrostatic pressure, imparting relative resistance to this mode of stress, similar to the proliferative function of JNK in pressure-stimulated cardiomyocytes (2, 34).

In conclusion, cyclic, pathological hydrostatic pressure is a novel type of stress to ECs that renders them susceptible to CD95/CD95L-mediated apoptosis accompanied by upregulation of intracellular molecules known to trigger both apoptosis and survival. Blocking the apoptotic machinery strikingly augments the resistance to this stress and leads to survival. In the context of atherosclerosis, these findings may further contribute to a better understanding of the role of endothelial injury in the development of atherosclerosis and may help to form a rational basis for therapeutic intervention.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by a grant from the Deutsche Forschungsgemeinschaft SFB 518/A13 (to P. Möller) and by Landesforschungsschwerpunkt Project Baden-Württemberg Fehlregulation von Apoptose (to C. Hasel and P. Möller).


   FOOTNOTES
 

Address for reprint requests and other correspondence: P. Möller, Dept. of Pathology, Univ. of Ulm, Albert-Einstein-Allee 11, D-89081 Ulm, Germany (e-mail: peter.moeller{at}medizin.uni-ulm.de)

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.


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