Abstract
Abstract. The pathogenesis of the epidemic form of hemolytic uremic syndrome is characterized by endothelial cell damage. In this study, the role of apoptosis in verocytotoxin (VT)-mediated endothelial cell death in human glomerular microvascular endothelial cells (GMVEC), human umbilical vein endothelial cells, and foreskin microvascular endothelial cells (FMVEC) was investigated. VT induced apoptosis in GMVEC and human umbilical vein endothelial cells when the cells were prestimulated with the inflammatory mediator tumor necrosis factor-α (TNF-α). FMVEC displayed strong binding of VT and high susceptibility to VT under basal conditions, which made them suitable for the study of VT-induced apoptosis without TNF-α interference. On the basis of functional (flow cytometry and immunofluorescence microscopy using FITC-conjugated annexin V and propidium iodide), morphologic (transmission electron microscopy), and molecular (agarose gel electrophoresis of cellular DNA fragments) criteria, it was documented that VT induced programmed cell death in microvascular endothelial cells in a dose- and time-dependent manner. Furthermore, whereas partial inhibition of protein synthesis by VT was associated with a considerable number of apoptotic cells, comparable inhibition of protein synthesis by cycloheximide was not. This suggests that additional pathways, independent of protein synthesis inhibition, may be involved in VT-mediated apoptosis in microvascular endothelial cells. Specific inhibition of caspases by Ac-Asp-Glu-Val-Asp-CHO, but not by Ac-Tyr-Val-Ala-Asp-CHO, was accompanied by inhibition of VT-induced apoptosis in FMVEC and TNF-α-treated GMVEC. These data indicate that VT can induce apoptosis in human microvascular endothelial cells.
The hemolytic uremic syndrome (HUS) is characterized by hemolytic anemia, thrombocytopenia, and acute renal failure (1). The pathogenesis of HUS is characterized by endothelial cell damage of glomeruli and/or arterioles of the kidney (2). In severe cases of HUS, endothelial cell damage is not limited to the kidney but extends to other organs, such as the brain and pancreas (3). The epidemic form of HUS is strongly associated with verocytotoxin (VT)-producing Escherichia coli infection (4). VT are exotoxins that consist of an enzymatic A-subunit and five receptor-binding B-subunits. The latter bind specifically to globotriaosylceramide (Gb3), which contains the carbohydrate sequence Galα1—4Gal in a terminal position (5). The biologic activity of the toxins is thought to involve inhibition of overall protein synthesis through enzymatic inactivation of the 60S ribosomal units (6). Induction of apoptosis has also been reported for a variety of cell types (7,8,9), including renal tubule-derived epithelial cells (10). The presence of apoptotic endothelial cells in the glomeruli of kidney biopsy specimens from three patients with the epidemic form of HUS was recently reported (11). However, whether VT can directly induce apoptosis in endothelial cells has not been studied. Apoptosis (programmed cell death) is a highly regulated process characterized by morphologic, functional, and molecular features (12, 13). Apoptosis occurs not only under physiologic conditions, such as embryogenesis, morphogenesis, and other processes of tissue formation and cell renewal, but also under pathologic conditions. A number of physical and chemical agents, including a variety of toxins, have been reported to induce apoptosis (7,8,9,10, 14). Apoptosis can be induced by several activation pathways, which cause the activation of procaspases into active caspases (13, 15). These activities converge in the activation of the so-called death caspases (caspases 3, 6, and 7), which initiate an irreversible process that leads to DNA fragmentation, cell detachment, and death (15, 16). The presence of caspase 1-like activity and caspase 3 has been demonstrated in human umbilical vein endothelial cells (HUVEC) (17).
In vitro studies using HUVEC and glomerular microvascular endothelial cells (GMVEC) have indicated that VT susceptibility requires additional stimulation by inflammatory mediators for induction of a sufficiently large number of specific VT receptors on these cells (5, 18). Because these mediators may themselves induce both stimulators and inhibitors of apoptosis in endothelial cells (19,20,21), it would be extremely difficult to discriminate between direct and indirect effects of VT on apoptosis in tumor necrosis factor-α (TNF-α)-stimulated GMVEC. In our studies, we observed that unstimulated human foreskin microvascular endothelial cells (FMVEC) are very sensitive to VT cytotoxicity in vitro. These cells provided a means to evaluate whether VT could induce apoptosis in human microvascular endothelial cells without prior stimulation with TNF-α. Using morphologic, functional, and molecular criteria, we demonstrate that VT induces apoptosis in these cells and also induces apoptosis in TNF-α-stimulated GMVEC. Using specific inhibitors, we investigated whether caspase 3 (CPP32, apopain) and caspase 1-like [interleukin-1β (IL-1β)-converting enzyme (ICE)-like] proteases play a role in VT-induced apoptosis.
Materials and Methods
Materials
Purified VT-1, VT-2, and VT-2c (1.0 to 1.2 mg protein/ml) were kindly provided by Dr. M. A. Karmali (Department of Microbiology, The Hospital for Sick Children, Toronto, Canada). The endotoxin content of the VT preparations was <0.05 EU/ml, as assessed using a Limulus amebocyte lysate assay with a detection level of 0.05 to 0.10 EU/ml. All experiments were performed with VT-2c (unless otherwise mentioned), which is structurally and functionally related to VT-1 and VT-2 and is associated with human disease (22) (Table 1).
Effects of various VT on apoptosis in human FMVECa
Endothelial Cell Cultures
FMVEC were isolated and purified according to methods previously described by Davison et al. (23) and Voyta et al. (24). FMVEC were seeded on gelatin (1%; Fluka Biochemicals, Buchs, Switzerland)-coated, six-well plates (Costar, Cambridge, MA) and cultured in M199 (BioWhittaker, Walkersville, MD) supplemented with 10% (vol/vol) human serum (local blood bank), 10% (vol/vol) newborn calf serum (NBCS) (Life Technologies, Grand Island, NY), 2 mM L-glutamine (ICN Biomedicals, Zoetermeer, The Netherlands), 5 U/ml heparin (Leo Pharmaceutical, Weesp, The Netherlands), 100 IU/ml penicillin/0.1 mg/ml streptomycin (Yamanouchi Pharma B.V., Leiderdorp, The Netherlands), and 150 mg/liter crude endothelial cell growth factor [extracted from bovine brains as described by Maciag et al. (25)]. Cells were subcultured with mild trypsinization (3 to 5 min), using trypsin/ethylenediaminetetraacetate (EDTA) (0.5 g/liter trypsin, 1:250, and 0.2 g/liter EDTA; Life Technologies), after which the cells were replated with a split ratio of 1:3. For experiments, FMVEC were used after nine to 11 passages.
GMVEC were isolated and purified as previously described by van Setten et al. (18). HUVEC were isolated from umbilical cords according to the method described by Jaffe et al. (26). GMVEC and HUVEC were used after eight to 10 and two to four passages, respectively.
All endothelial cells used displayed the presence of von Willebrand factor, platelet endothelial cell adhesion molecule-1, and V- and E-cadherin at all passages used. No immunoreactivity to the anticytokeratin 20 antibody or the anti-α-smooth muscle actin antibody was observed, excluding the possibility of contaminating epithelial and mesangial cells, respectively (18).
Cytotoxicity Assays
Microvascular endothelial cells were cultured in complete medium on gelatin-coated, 24-well plates and were grown until confluence. Five days after reaching confluence, cells were preincubated with or without TNF-α (10 ng/ml) for 24 h. The next day, the medium was aspirated and fresh medium with different concentrations of VT was added. VT was diluted in medium with 20% fetal calf serum instead of 10% NBCS and 10% human serum, because previous studies indicated that NBCS and human serum may have neutralizing activity against VT. After 24 h, the cells were washed with phosphate-buffered saline (PBS) and released with trypsin/EDTA. Viable, trypan blue-excluding cells were then counted with a hemocytometer.
Protein Synthesis
Protein synthesis was assessed by assaying the incorporation of 35S-labeled methionine (0.25 μCi/ml complete medium) into 35S-proteins during 24-h incubations with different concentrations of VT. After incubation, the cells were washed and 35S-labeled cellular proteins and 35S-labeled proteins present in the medium were precipitated with the addition of TCA (10 and 20%, respectively). Precipitated proteins were dissolved in 0.3 ml of 0.3 M NaOH, 60 μl of 1.5 M HCl was added, and precipitated radioactivity was counted in a liquid scintillation counter (18,27).
Glycolipid Extraction and Thin Layer Chromatography
FMVEC were cultured in complete medium on gelatin-coated, six-well plates (Costar) and were grown to confluence. Highly confluent cells were exposed to complete medium alone or complete medium supplemented with TNF-α (10 ng/ml). After 24 h, the cells were trypsinized and washed three times with ice-cold PBS. Subsequently, glycolipids were extracted, separated, and assayed for 125I-VT binding, as described previously (18,27). Thin layer chromatograms were analyzed with a Fuji BAS 1000 PhosphorImager (Fuji Photo Film Co., Tokyo, Japan).
Apoptosis Assays
Fluorescence-Activated Cell Sorting Analyses. FMVEC, HUVEC, and GMVEC were grown on gelatin-coated, 12-well plates (Costar) until they reached confluence. Highly confluent HUVEC and GMVEC were prestimulated with TNF-α (10 ng/ml) for 24 h, to upregulate VT receptors and induce VT susceptibility (18,27). Because FMVEC display VT susceptibility under basal conditions, these cells were not pretreated with TNF-α. Subsequently, unstimulated FMVEC and TNF-α-stimulated HUVEC and GMVEC were incubated for 4 or 24 h with complete medium, with complete medium supplemented with various concentrations of VT (0.1 pM to 10 nM), B-subunit (5 to 130 nM), or cycloheximide (CHX) (0.01 to 1 μg/ml), or with M199 alone. After the incubation period, detached cells were collected and pooled with trypsinized adherent cells. Cells were centrifuged at 200 × g for 5 min at 4°C, and the supernatant was removed. The cells were then washed with ice-cold binding buffer (10 mM Hepes, [pH 7.4], 150 mM NaCl, 5 mM KCl, 1 mM MgCl2, 1.8 mM CaCl2), as previously described by Koopman et al. (28), supplemented with 0.1% bovine serum albumin (BSA) (ICN Biomedicals, Zoetermeer, The Netherlands). Cells were resuspended in 500 μl of binding buffer with 0.1% BSA, of which 445 μl was transferred into a tube suitable for fluorescence-activated cell sorting (FACS) analysis. Five microliters of FITC-conjugated annexin V (2 μg/ml, diluted in binding buffer with 0.1% BSA; Bender Medsystems, Vienna, Austria) and 50 μl of propidium iodide [100 μg/ml, diluted in RPMI DM (Flow Inc.) supplemented with 5% fetal calf serum and 2 mM CaCl2; Sigma] were added, after which the cell suspension was gently mixed and incubated in the dark on ice for 10 min (28,29). Samples were assayed for viable, apoptotic, and necrotic cells by FACS analysis (Coulter 7 Epics 7 XL-MCL, Beckman Coulter Inc., Mijdrecht, The Netherlands). Necrotic cells were defined as cells demonstrating positive staining for both FITC-conjugated annexin V and propidium iodide. Viable cells were not positive for either FITC-conjugated annexin V or propidium iodide. Apoptotic cells were defined as cells exhibiting positive staining for FITC-conjugated annexin V and negative staining for propidium iodide. Fluorescence was measured on a double-parameter histogram, using logarithmic scales. For each tube, 5000 events were analyzed.
To assess the inhibition of the apoptosis-inducing effect by reversible inhibitors, FMVEC, cultured under identical conditions, were preincubated for 1 or 24 h with various concentrations of ICE inhibitor I [Ac-Tyr-Val-Ala-Asp-CHO (YVAD-CHO); Calbiochem, San Diego, CA] or CPP32/apopain inhibitor [Ac-Asp-Glu-Val-Asp-CHO (DEVD-CHO); Calbiochem] (30,31). After preincubation with these inhibitors, cells were rinsed once with M199, after which the cells were exposed for 16 h to complete medium or complete medium supplemented with VT (1 nM), in the presence of the same reversible inhibitor. Apoptosis was evaluated by FACS analysis as described previously. To detect the presence of caspase 3, caspase 6, or caspase 7 in FMVEC and GMVEC, Western blot analysis was performed according to established procedures (31), using primary antibodies against human CPP32/p20 (caspase 3) (N-19), Mhc2/p20 (caspase 6) (K-20), or ICE-LAP3 (caspase 7) (C-18) (Santa Cruz Biotechnology, Santa Cruz, CA) and a horseradish peroxidase-conjugated anti-goat IgG antibody (diluted 1:5000 in PBS with 0.5% BSA and 0.05% Tween-20; Nordic Immunology, Tilburg, The Netherlands) as a secondary antibody.
Fluorescence Microscopy Using Triple Staining of Cell Monolayers. Cells were seeded onto gelatin-coated glass coverslips, as described previously (18), and were grown to confluence. Highly confluent FMVEC were exposed to complete medium alone or complete medium supplemented with various concentrations of VT. After an incubation period of 16 h, adherent cells were stained with 500 μl of Hoechst 33342 (1 μg/ml in complete medium; Sigma) for 15 min at 37°C. Subsequently, cells were rinsed once with M 199 and incubated, on ice, with 500 μl of complete medium containing FITC-conjugated annexin V (1 μg/ml) and propidium iodide (1 μg/ml). Cells were analyzed by fluorescence microscopy within 15 min, using filters of 365, 490, and 560 nm for Hoechst 33342, FITC-conjugated annexin V, and propidium iodide staining, respectively. Definitions of viable, apoptotic, and necrotic cells were similar to those described for the FACS analyses.
DNA Fragmentation Analyses. FMVEC were grown in complete medium on gelatin-coated, six-well plates until they reached confluence. Highly confluent cells were exposed to complete medium or complete medium supplemented with various concentrations of VT or B-subunit. After 16 h of incubation, detached cells were collected on ice and pooled with trypsinized adherent cells. Cells were subjected to centrifugation at 200 × g for 5 min at 4°C, and the pellets were washed twice with ice-cold PBS. Subsequently, cells were lysed with 10 mM Tris (Boehringer Mannheim, Mannheim, Germany), pH 8.0, 10 mM EDTA (Life Technologies), 0.5% Triton X-100 (Boehringer Mannheim), as previously described by Bissonnette et al. (32). To separate fragmented DNA from intact chromatin, cells were centrifuged at 15,000 × g for 20 min at 4°C. Soluble fragmented DNA was transferred into a new tube and treated with RNase A (100 μg/ml; Sigma) for 1 h at 37°C, followed by treatment with proteinase K (200 μg/ml; Sigma) in 1% sodium dodecyl sulfate (Merck, Darmstadt, Germany) for 2 h at 50°C. DNA was precipitated with 0.1 volume of 3 M sodium acetate and 2 volumes of 96% ethanol for 16 h, after which the samples were centrifuged at 15,000 × g for 10 min at 4°C. Subsequently, DNA pellets were air-dried and dissolved in Tris-EDTA solution, pH 8.0 (10 mM Tris-HCl and 0.1 mM EDTA, both at pH 8.0). Loading buffer was added to each sample, and electrophoresis was performed at 80 V for 1.5 to 2 h on a 1.5% agarose gel containing 0.4 μg/ml ethidium bromide (Merck), with 1 × Tris/borate/EDTA running buffer (22.5 mM Tris-borate, 0.5 mM EDTA, pH 8.0). DNA was observed under ultraviolet light (350 nm) and photographed. DNA size calibration was performed using a 100-bp marker (Pharmacia, Roosendaal, The Netherlands).
Transmission Electron Microscopy. To evaluate the apoptosis-inducing effect of VT on adherent endothelial cells by transmission electron microscopy, FMVEC were cultured in complete medium on gelatin-coated, 12-well plates. After they reached confluence, the cells were exposed to control medium or control medium supplemented with various concentrations (0.1 pM to 10 nM) of VT. After 16 h of incubation, detached cells were pooled with trypsinized adherent cells. Cells were pelleted and fixed for 2 h at room temperature with 2.5% glutaraldehyde in cacodylate buffer. The cells were then carefully rinsed with 0.1 M cacodylate buffer for 30 min, after which the cells were immersed and pelleted in 15% gelatin. After immersion, alternate fixation with 1% OsO4 in cacodylate buffer was performed for 30 min. Cells were then selectively dehydrated in graded ethanol and embedded in Epon 812. Thin sections were cut with an ultramicrotome and stained with uranyl acetate and lead citrate. Sections were examined with a Jeol 1200 EX electron microscope (Jeol Europe bv., Schilphol-Rijk, The Netherlands).
Additional Analytic Procedures
To determine whether the cultured cells exhibited signs of endogenous activation, the expression of monocyte chemoattractant protein-1 (MCP-1) (33), urokinase-type plasminogen activator (u-PA) (34), and vascular cell adhesion molecule-1 (VCAM-1) (18) in culture supernatants (u-PA and MCP-1) or on the cell surface (VCAM-1) was assessed according to established procedures.
Statistical Analyses
The statistical significance of differences between groups was analyzed by ANOVA followed by Bonferroni's modified t test. Differences were considered significant at P < 0.05 (two-sided).
Results
Effects of VT on Unstimulated FMVEC
Under basal conditions, human FMVEC were very sensitive to VT, in contrast to GMVEC and HUVEC, which required preexposure to TNF-α or IL-1 to become sensitive to VT. Exposure of unstimulated FMVEC to various concentrations of VT (0.01 fM to 1 nM) for 24 h produced concentration-dependent cell toxicity and inhibition of overall protein synthesis (Figure 1). This high sensitivity to VT was related to high levels of specific VT binding to these cells, as assayed in binding experiments with 125I-VT. Thin layer chromatographic analyses of cellular neutral glycolipids extracted from unstimulated FMVEC revealed strong binding of 125I-VT to glycolipid species in the Gb3 region (Figure 2), with an additional increase after stimulation with TNF-α.
Cytotoxic effects of verocytotoxin-2c (VT-2c) on foreskin microvascular endothelial cells (FMVEC) and glomerular microvascular endothelial cells (GMVEC). Human FMVEC and GMVEC were grown to confluence and maintained confluent for 5 d. Subsequently, FMVEC (○), control GMVEC (□), and tumor necrosis factor-α (TNF-α) (10 ng/ml)-treated GMVEC (▪) were incubated with different concentrations of VT (0.01 fM to 10 nM). After 24 h, the number of viable cells (A) and the incorporation of [35S]methionine into proteins (B) were determined as described in Materials and Methods. Data are expressed as the mean ± SD of two independent experiments, with two different donors [percentage of control (% of C)].
125I-VT binding to glycolipids extracted from human GMVEC and FMVEC. Glycolipids were extracted from highly confluent endothelial cells, separated by thin layer chromatography, assayed for 125I-VT binding, and observed by PhosphorImaging. Lanes A and B, glycolipid extracts of 20 cm2 of human GMVEC from one representative donor; lane A, control GMVEC; lane B, TNF-α (10 ng/ml)-treated GMVEC. Lanes E and F, glycolipid extracts of 20 cm2 of human FMVEC from one representative donor; lane E, control FMVEC; lane F, TNF-α (10 ng/ml)-treated FMVEC. Lanes C and G, standard neutral glycolipids (2 μg of each glycolipid). Lanes D and H, orcinol staining of standard neutral glycolipids (Gb1, galactosylceramide; Gb2, lactosylceramide; Gb3, globotriaosylceramide; Gb4, globotetraosylceramide; Gb5, Forssman pentasaccharide).
To investigate whether FMVEC had spontaneously acquired the activation phenotype that is induced by TNF-α or IL-1, we measured the levels of several compounds that are induced in endothelial cells by TNF-α and IL-1. Values for VCAM-1 (not detectable), MCP-1 (<0.3 ng/ml), and u-PA (0.2 ng/ml) in unstimulated FMVEC were essentially identical to those found in unstimulated GMVEC and HUVEC, and values increased during 24-h exposure to TNF-α (VCAM-1 clearly expressed; MCP-1, >2000 ng/ml; u-PA, 1.7 ng/ml; average of three independent determinations).
VT Induction of Apoptosis in FMVEC
To evaluate whether apoptosis is involved in VT-mediated FMVEC cell death, we initially performed FACS analyses using a dual-staining method with FITC-conjugated annexin V and propidium iodide. This method discriminates among viable, apoptotic, and necrotic cells (28,29) (see the Materials and Methods section for details). Figure 3 presents representative cytograms for control (Figure 3A) and VT-treated (Figure 3B) FMVEC. Whereas in control cells the numbers of apoptotic and necrotic cells were rather small (3 and 5%, respectively), VT-treated (0.1 nM for 24 h) cells demonstrated an impressive increase in apoptotic cells (40.7 ± 4.5%, n = 3) and a slight increase in necrotic cells (12.3 ± 2.9%, n = 3).
Flow cytometric analysis of apoptosis in control and VT-treated FMVEC. FMVEC were grown to confluence and maintained confluent for 5 d. Subsequently, cells were exposed to control medium alone (A) or control medium supplemented with 1 nM VT (B). After 16 h of incubation, detached cells were pooled with trypsinized adherent cells and assayed for the occurrence of apoptosis by fluorescence-activated cell sorting (FACS) analysis, as described in Materials and Methods. The lower left quadrant of each panel represents viable cells [annexin V (ANNEX)-negative/propidium iodide (PI)-negative]. The upper right quadrant represents nonviable necrotic cells (annexin V-positive/propidium iodide-positive). The lower right quadrant represents apoptotic cells (annexin V-positive/propidium iodide-negative). Data for a representative donor are presented; similar results were obtained in three other experiments with cells from two different donors.
The apoptosis-inducing effect of VT in FMVEC was time-and concentration-dependent (Figure 4). The apoptosis-inducing effect of VT was observed as early as 4 to 6 h after the beginning of the VT incubation, with an increase in the number of apoptotic cells after 24 h of VT exposure. The threshold dose for VT to induce apoptosis was 0.1 pM. Identical results were obtained when VT-1, VT-2, and VT-2c were compared (Table 1). The B-subunit of VT induced apoptosis only at very high concentrations. Exposure of the cells to 50 and 130 nM B-subunit for 24 h resulted in 29 ± 8% and 40 ± 8% apoptotic cells (mean ± SD of two independent experiments with two different donors), respectively. The percentages of necrotic cells were 9 ± 0.2% and 11 ± 2%, respectively.
Dose-response curves for VT-mediated apoptosis in human FMVEC. Human FMVEC were grown to confluence and maintained confluent for 5 d. Subsequently, cells were exposed to various concentrations of VT (0 to 10 nM) during a 5-h (○) or 24-h () incubation period. Detached and adherent cells were obtained and assayed for the occurrence of apoptosis by FACS analysis, as described in Materials and Methods. Data are expressed as the mean ± SD of three independent experiments with cells from two different donors.
VT-induced apoptosis in FMVEC was confirmed by direct observation, using a triple-staining method. As shown in Figure 5, highly confluent control and VT-exposed cells were stained with Hoechst 33342 (Figure 5, A and B) or FITC-conjugated annexin V and propidium iodide (Figure 5, C and D). Staining of control cells with Hoechst 33342 resulted in homogeneous diffuse nuclear staining (Figure 5A). After incubation of FMVEC with VT (10 nM) for 16 h, some of the adherent cells displayed condensed and fragmented nuclear chromatin, a characteristic feature of apoptosis (Figure 5B, arrows). Whereas almost all (>99%) control cells were negatively stained with annexin V and propidium iodide (Figure 5C), indicating that they were viable, a major fraction of VT-treated FMVEC demonstrated positive staining with annexin V and negative staining with propidium iodide, indicating apoptosis (Figure 5D, arrows). A minority of VT-exposed cells exhibited intracellular staining with annexin V and propidium iodide, which is characteristic of necrotic cells (Figure 5D, star).
Analysis of apoptosis in control- and VT-treated FMVEC by fluorescence microscopy, using triple-staining. FMVEC were grown to confluence and maintained confluent for 5 d. Subsequently, the cells were incubated for 16 h with control medium (A and C) or control medium supplemented with VT (10 nM) (B and D). After the incubation period, cells were stained with Hoechst 33342 (A and B) or FITC-conjugated annexin V and propidium iodide (C and D), as described in Materials and Methods. Magnification, ×200 in A to D. FMVEC indicated by arrows and stars represent apoptotic (annexin V-positive/propidium iodide-negative, approximately 24%) and necrotic (annexin V-positive/propidium iodide-positive) cells, respectively.
DNA Fragmentation Analysis and Ultrastructural Morphologic Features of VT-Exposed FMVEC
Observations made using FACS analysis and fluorescence microscopy were extended by DNA fragmentation analyses. Endonuclease-induced cleavage of nuclear DNA into mono-and oligonucleosomal fragments with approximate molecular sizes of multiples of 180 nucleotides is a characteristic feature of programmed cell death (35). Figure 6 demonstrates the electrophoretic patterns of DNA extracted from control and VT holotoxin-treated FMVEC. Whereas DNA from control cells remained intact throughout the 16-h incubation period (Figure 6, lane C), DNA from VT holotoxin (0.1 fM to 1 nM)-exposed cells clearly exhibited the DNA “ladder” pattern of multiples of 180-bp fragments. Exposure of the cells to 130 nM VT B-subunit induced a similar DNA ladder pattern (data not shown).
DNA fragmentation in VT-treated FMVEC, as determined by agarose gel electrophoresis. FMVEC were grown to confluence and maintained confluent for 5 d. Subsequently, FMVEC were incubated for 16 h with control medium alone or control medium supplemented with VT (0.1 fM to 1 nM), after which DNA was extracted, centrifuged, and analyzed for mono-/oligonucleosomal fragments by agarose gel electrophoresis, as described in Materials and Methods. Note that DNA fragmentation with the characteristic ladder pattern was observed only in VT-treated cells, whereas DNA from control cells remained intact. Shown is one representative experiment of two. M, marker; C, control untreated FMVEC.
In addition, the apoptosis-inducing effect of VT on FMVEC was studied by transmission electron microscopy. At the ultrastructural level, apoptotic cells exhibit distinctive alterations, primarily of the nucleus, which can be discriminated from events of necrosis (35). Representative micrographs of control and VT-treated FMVEC are presented in Figure 7. The morphologic features of control FMVEC in culture were maintained; cells exhibited nuclei with normal heterogeneous chromatin and a nuclear envelope, normal intact organelles, and normal plasma membranes (Figure 7A). In contrast, FMVEC exposed to VT (1 nM) for 16 h exhibited varying degrees of nuclear condensation (Figure 7, B and C), and even cellular fragmentation was noted (Figure 7D). Furthermore, we observed abundant cytoplasmic vacuolization and the presence of apoptotic bodies; however, cellular organelles of VT-treated FMVEC appeared normal. Although the integrity of the plasma membrane remained intact, blebbing of the plasma membrane occurred in VT-exposed FMVEC. These morphologic changes are indicative of apoptosis and are clearly different from those exhibited by cells undergoing necrosis.
Ultrastructural morphologic features of control and VT-treated FMVEC, as assessed by transmission electron microscopy (TEM). FMVEC were grown to confluence and maintained confluent for 5 d. Subsequently, FMVEC were exposed for 16 h to control medium alone or control medium supplemented with VT (1 nM), after which the cells were prepared for TEM and subsequently examined as described in Materials and Methods. (A) Ultrastructural morphologic features of a representative normal control cell. (B, C, and D) Ultrastructural morphologic features of VT-treated cells, including features characteristic of apoptosis. Magnification, ×5800 in A; ×7500 in B through D.
Comparison of Apoptosis Induced by VT and CHX
To evaluate whether the induction of apoptosis by VT merely reflects inhibition of protein synthesis, we compared the effect of VT with that of the protein synthesis inhibitor CHX. For proper comparisons, we determined which CHX concentration inhibited protein synthesis to a similar extent, compared with VT (0.1 pM), in wells cultured in parallel. Because 0.1 μg/ml CHX and 0.1 pM VT both inhibited protein synthesis by 30%, their effects on apoptosis induction were compared. Only VT induced apoptosis (14 ± 3% for VT versus 2 ± 1% for CHX; mean of two independent experiments). These data indicate that VT is a more potent inducer of apoptosis than is CHX at comparable potencies for protein synthesis inhibition.
Apoptosis of TNF-α-Pretreated GMVEC and HUVEC after Exposure to VT
The effect of VT on the induction of apoptosis among GMVEC and HUVEC was subsequently investigated by FACS analysis, using a dual-staining method with FITC-conjugated annexin V and propidium iodide. VT did not induce significant cell death among untreated GMVEC or HUVEC (data not shown). However, when GMVEC or HUVEC were preincubated with TNF-α for 24 h, 4- to 6-h exposure of the cells to VT (0.1 to 10 nM) induced a dose-dependent increase in the number of apoptotic cells (Figure 8). Although no further increase in the percentage of apoptotic cells was observed after 24 h of VT exposure, the percentage of necrotic cells increased significantly.
Apoptosis of TNF-α-pretreated human umbilical vein endothelial cells (HUVEC) and GMVEC. Cells were grown to confluence. After 5 d of confluence, HUVEC and GMVEC were prestimulated with TNF-α (10 ng/ml) for 24 h. After pretreatment with TNF-α, the endothelial cells were incubated with VT (□, 0 nM; ▪, 0.1 nM; , 1 nM;
, 10 nM) for 5 h. Subsequently, detached cells and trypsinized adherent cells were pooled and assayed for the presence of apoptotic cells by FACS analysis, as described in Materials and Methods. (A and C) Data are expressed as the mean ± SD of four (HUVEC) (A) or three (GMVEC) (C) independent experiments with cell cultures from different donors. (B and D) Representative flow cytometric analyses of control and VT (10 nM)-treated, TNF-α-stimulated HUVEC (B) and GMVEC (D) are presented.
Effects of Caspase Inhibitors on VT-Induced Apoptosis of FMVEC and GMVEC
Caspases such as caspase 1 (ICE) and caspase 3 (CPP32) have been implicated in the complex cascade of events that results in apoptosis (15). To determine whether these cysteine proteases play a role in VT-mediated apoptosis, DEVD-CHO and YVAD-CHO, specific inhibitors that discriminate between different caspases (inhibiting caspase 3 and caspase 1-like proteases, respectively), were used. FMVEC incubated with DEVD-CHO (3 to 300 μM) demonstrated dose-dependent inhibition of VT-induced apoptosis. The threshold dose to decrease the percentage of apoptotic cells was 30 μM; a maximal response of 60 to 70% inhibition of VT-mediated apoptosis was obtained with 300 μM DEVD-CHO (Tables 2 and 3). Similarly, DEVD-CHO partly inhibited VT-induced apoptosis in TNF-α-exposed GMVEC (Table 2). Administration of YVAD-CHO (3 to 300 μM) did not result in significant inhibition of VT-induced apoptosis in FMVEC (Table 2). Western blotting of cell extracts demonstrated that both FMVEC and unstimulated and TNF-α-stimulated GMVEC contained caspase 3, whereas caspases 6 (Mhc-2) and 7 (ICE-LAP, Mhc-3) could not be detected (data not shown). These data are consistent with a possible involvement of caspase 3 in VT-induced apoptosis in human microvascular endothelial cells.
Effects of the caspase inhibitors YVAD-CHO and DEVD-CHO on VT-mediated apoptosis in human microvascular endothelial cellsa
Effects of different concentrations of DEVD-CHO on the apoptosis-inducing effect of VTa
Discussion
VT-producing E. coli infections are strongly implicated in the pathogenesis of the epidemic form of HUS, which is characterized by endothelial cell damage. Using morphologic, functional, and molecular criteria, we demonstrated in this study that apoptosis plays a role in VT-mediated endothelial cell death. This involvement was demonstrated in HUVEC and GMVEC that had been preexposed to the inflammatory mediator TNF-α and in unstimulated FMVEC and probably involves the activation of caspase 3.
Endothelial cell damage of predominantly glomerular capillaries is a characteristic feature in the pathogenesis of the epidemic form of HUS (2). From a pathogenetic point of view, it is generally assumed that VT is potentially cytopathic for endothelial cells. Several in vitro observations have indicated that priming of the endothelial cells by inflammatory mediators is required for VT cytotoxicity. These mediators cause an increase in VT susceptibility via enhancement of the number of VT-binding receptors (18,27). The mechanisms reported for VT-induced cell death in a variety of cell types, including endothelial cells, involve A-subunit-dependent inhibition of overall protein synthesis through the enzymatic inactivation of 60S ribosomal units (6). Observations in Vero cells (7,9), Burkitt's lymphoma cells (8), and renal tubule-derived epithelial cells (10) have indicated that VT also induces programmed cell death. VT-mediated apoptosis in cultured endothelial cells has not yet been reported.
Here we demonstrate that VT induces apoptosis in TNF-α-treated GMVEC and HUVEC. Because these cells require prestimulation with TNF-α to acquire increased numbers of VT receptors and high sensitivity to VT (18,27), a direct effect of VT on apoptosis is difficult to establish. On one hand, TNF-α itself has been reported to induce apoptosis in lymphocytes (36) and bovine endothelial cells (19,20). On the other hand, TNF-α has been reported to induce the expression of A1, a Bcl-2 homolog that is known to block programmed cell death in human endothelial cells (21). Subsequent exposure to protein synthesis inhibitors, including VT, may cause a decrease in the expression of this apoptosis-blocking homolog, resulting in the induction of apoptosis. It would thus be extremely difficult to establish whether VT directly or indirectly induces apoptosis. In our studies, we observed that unstimulated FMVEC demonstrated high levels of VT binding and high VT sensitivity, with only minor increases after stimulation with the inflammatory mediator TNF-α. The reason why unstimulated FMVEC express many VT receptors (Gb3 molecules) is unknown. It is not a reflection of general TNF-α-like activation of the cells but may be attributable to reduced catabolism of Gb3 or markedly reduced conversion of Gb3 to Gb4. It is also not known whether the large number of VT receptors in these cells represents an in vitro feature of these cells or also occurs in vivo. Nevertheless, in various assays with these cells, we obtained evidence that VT can directly induce apoptosis in endothelial cells, even without prior exposure to TNF-α. These observations make it likely that VT can also directly induce apoptosis in TNF-α-treated GMVEC.
Previous observations indicated that certain cell populations undergo apoptosis when exposed to inhibitors of protein synthesis (37,38,39). These data may indicate that VT-mediated apoptosis in FMVEC results from the inhibitory effect of VT on protein synthesis. Interestingly, whereas partial inhibition of protein synthesis by VT was associated with a considerable number of apoptotic cells, comparable inhibition of protein synthesis by CHX, which is known to block the peptidyl transferase reaction on ribosomes, was not. The finding that high concentrations of the B-subunit of VT alone induced similar features of apoptosis in FMVEC, compared with the holotoxin, further suggests that additional pathways, independent of protein synthesis inhibition, may contribute to VT-mediated apoptosis. The latter finding is closely related to observations made by Mangeney et al. (8), who reported that the B-subunit of VT induced apoptosis in Burkitt's lymphoma cells. However, in Vero cells, which, like FMVEC, are highly susceptible to VT, only VT holotoxin and not the B-subunit induced programmed cell death (40). This leaves open the possibility that the effect of high concentrations of the B-subunit occurs in addition to the effect of VT itself.
In our studies, we used primarily VT-2c, which is structurally and functionally related to VT-1 and VT-2 (22). VT-1, VT-2, and VT-2c are associated with the epidemic form of HUS in childhood, but the former two are more commonly associated with human disease than is VT-2c. As pointed out in our comparative studies, the apoptosis-inducing effect of VT-2c is similar to that of VT-1 and VT-2. Identical effects of different forms of VT were previously demonstrated for inhibition of protein synthesis (18) and cell proliferation (33).
Cell death by apoptosis involves the activation of caspases, followed by caspase-mediated proteolysis and nucleosomal DNA fragmentation. Similar to the coagulation system, a cascade of proteolytic activations occurs, beginning with the activation of procaspases with a long prodomain and converging in the activation of procaspases with a short prodomain (15,41). These latter caspases, i.e., caspases 3, 6, and 7, are also referred to as death caspases, because they initiate processes that irreversibly cause cell death. Our observation that a peptide aldehyde inhibitor of caspase 3 and related caspases, such as caspases 7 and 8, and not an inhibitor of caspase 1 inhibited VT-mediated apoptosis in FMVEC and TNF-α-stimulated GMVEC indicates that VT-mediated apoptosis is linked to activation of the former caspases and not that of caspase 1. Caspase 3 was present in relatively large quantities in both FMVEC and GMVEC, in agreement with previous observations in HUVEC (17,31,42). Whereas caspase 3 was demonstrated in cultured endothelial cells, it could not be demonstrated in quiescent endothelial cells in vivo (43). However, the induction of caspase 3 in pathologic conditions in vivo has not yet been evaluated. Future studies may provide further insight into the complex molecular mechanisms underlying the apoptosis-inducing effect of VT in endothelial cells. In particular, it may be of interest to study the roles of glycolipid breakdown products, such as ceramide and sphingosine, and their phosphorylation products. These products have been reported to be involved in programmed cell death in other cell types (44).
Apoptosis is thought to represent an important defense mechanism in the event of cell damage. In contrast to necrosis, this type of cell death is usually followed by the removal of unwanted cells without the induction of an inflammatory response. Therefore, apoptosis of endothelial cells may allow uncompromised restoration of the endothelial cell layer. However, inflammation seems to precede VT-induced cell death in GMVEC. Therefore, we hypothesize that apoptosis is part of a complex cascade that takes place inside the glomerular capillaries and finally leads to kidney failure. Because apoptotic cells have been reported to be procoagulant (45), they may contribute to thrombotic events in HUS and thus are likely to amplify glomerular endothelial cell injury.
Mitra et al. (46) reported that plasma from four patients with thrombotic thrombocytopenic purpura (which is related to HUS) could induce apoptosis in restricted lineages of microvascular endothelial cells. This could not be demonstrated with plasma from one patient with HUS associated with diarrhea. However, whether VT is present at sufficient levels in the plasma of patients with HUS remains uncertain. Recently, Karpman et al. (11) reported for the first time the presence of apoptotic endothelial cells in the glomeruli of kidney biopsy specimens from three patients with the epidemic form of HUS. The occurrence of apoptotic features in the epidemic form of HUS may be underestimated. Descriptions of the pathologic features of HUS included primarily autopsy or biopsy findings for kidney specimens obtained at an advanced stage of the disease. In such samples, necrosis secondary to apoptosis, as well as phagocytosis of small numbers of apoptotic cells by macrophages and adjacent cells, might have occurred. In addition, most pathologic studies used light microscopy, with which it is difficult to detect and distinguish apoptotic and necrotic cells. Therefore, advanced techniques applied early in the course of the disease are needed to specifically detect apoptotic cells in tissue samples.
We conclude from this study that VT can induce apoptosis in human microvascular endothelial cells by a mechanism that possibly involves caspase 3. These observations may provide further insight into the pathogenesis and pathophysiologic features of VT-mediated microvascular endothelial cell damage in HUS.
Acknowledgments
We thank Mario Vermeer (Gaubius Laboratory TNO Prevention and Health, Leiden, The Netherlands) for excellent technical assistance. This study was supported by a grant from the Dutch Kidney Foundation (Grant C94.1344). Drs. Pijpers and van Setten contributed equally to this study.
- © 2001 American Society of Nephrology