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. 2002 Mar;127(3):445–454. doi: 10.1046/j.1365-2249.2002.01769.x

Intravenous immunoglobulin (IVIG) preparations induce apoptosis in TNF-α-stimulated endothelial cells via a mitochondria-dependent pathway

K NAKATANI 1, S TAKESHITA 1, H TSUJIMOTO 1, I SEKINE 1
PMCID: PMC1906310  PMID: 11966760

Abstract

Endothelial cells (ECs) are a target in inflammation, and the death of EC is regulated by various factors. Although intravenous immunoglobulin (IVIG) preparations are known to be beneficial therapeutic agents for the treatment of autoimmune diseases and systemic inflammatory disorders, their mechanism of action have not yet been completely elucidated. The aim of the present study is to investigate the possible role of IVIG in EC apoptosis. We demonstrate herein that IVIG induced the apoptosis of human umbilical vein ECs (HUVECs) prestimulated by TNF-αin vitro, but not in unstimulated HUVECs, in a dose- and time-dependent manner, using a proportion of cells with hypodiploid DNA, DNA ladder formation and morphological changes. Anti-Fas MoAbs had no effect on the IVIG-induced apoptosis in the TNF-α-stimulated HUVECs. IVIG decreased the intracellular expression of anti-apoptotic proteins of the Bcl-2 family (A1 and Bcl-XL) while IVIG increased the intracellular expression of pro-apoptotic proteins (Bax and Bcl-XS) in the TNF-α-stimulated HUVECs. Furthermore, IVIG increased the intracellular production of reactive oxygen species and decreased the mitochondrial membrane potential (Δψm). Caspase-inhibitors inhibited the IVIG-induced apoptosis of the TNF-α-stimulated HUVECs. The present results show a novel action in which IVIG can induce the apoptosis of TNF-α-stimulated HUVECs through a mitochondrial apoptotic signalling pathway. These observations suggest that the clinical use of IVIG preparations may thereby regulate the cell death of activated ECs in inflammation.

Keywords: apoptosis, Bcl-2, endothelial cells, intravenous immumoglobulin

INTRODUCTION

Endothelial cells (ECs) play a pivotal role in inflammation, haemostasis and angiogenesis [1,2]. EC activation is an integral component of the inflammatory response to tissue injury by releasing such mediators as cytokines and adhesion molecules [3]. The homeostasis of ECs is dependent upon the balance of cell proliferation and death. To maintain the integrity of the vascular barrier, ECs are usually resistant to cell death. Apoptosis, programmed cell death, is now believed to be involved in the homeostasis of tissues as well as in the aetiology and pathology of several diseases [4,5]. Apoptosis in ECs is thought to play a significant role in the pathogenesis of such diseases as atherosclerosis, allograft vasculopathy, hypertension, sepsis and associated syndromes [3].

EC apoptosis is induced by LPS [6], TNF-α[7], interferon-γ[8], H2O2[9] and transforming growth factor β1 [10], while EC apoptosis is inhibited by fibroblast growth factor (FGF) [11], vascular endothelial growth factor (VEGF) [12] and endothelin [13]. EC apoptosis may also contribute to the regulation of inflammatory responses including vascular restructuring [3]. Recently, EC apoptosis-inducing drugs, such as endostatin [14], angiostatin [15] and chloroquine [16] have been reported to demonstrate an anti-inflammatory effect by repressing angiogenesis. EC apoptosis is mediated via such death domains as Fas and TNF receptor-1 and/or via the mitochondria-dependent pathway [17]. The mitochondrial apoptotic pathway is regulated mainly by the relative expression of specific genes such as Bcl-2 family [18]. Bcl-2, Bcl-XL and A1 are the anti-apoptotic proteins which promote the EC survival, while Bcl-XS and Bax are pro-apoptotic proteins which are known to accelerate EC apoptosis [18]. EC apoptosis is also regulated by reactive oxygen species (ROS) [19].

The therapeutic administration of intravenous immunoglobulin (IVIG), prepared from pools of plasma of several thousand healthy donors, has been reported to be an effective treatment for severe bacterial and viral infections [20], and also for immune-mediated inflammatory disorders including autoimmune diseases [21,22] and systemic vasculitis [23,24]. Although several mechanisms of action for IVIG have been proposed by many investigators, they have still not been elucidated fully. IVIG is believed to exhibit a broad spectrum of immunomodulatory activities both in vitro and in vivo[25]. Recently, IVIG is reported to induce apoptosis in human lymphocytes and monocytes via the Fas-mediated pathway [26] and in a human T cell line via the mitochondria-dependent pathway [27]. It has also been shown to inhibit the EC proliferation and mRNA expression of cytokines, adhesion molecules and chemokines in ECs [28]. However, so far no studies have yet been made to clarify the effect of IVIG on EC apoptosis. The aims of the present study are to investigate whether IVIG induces the apoptosis of human ECs in vitro and, if so, to elucidate the mechanism of the IVIG action.

METHODS

Reagents

The following reagents were prepared: recombinant human TNF-α from R&D Systems, Minneapolis, MN, USA; recombinant human bFGF from Kokusai Shiyaku Co, Tokyo, Japan; fetal bovine serum (FBS), trypsin, EDTA and HBSS from Gibco BRL Life Technologies Inc., Rockville, MD, USA; protease inhibitors cocktail, 2′,7′- dichlorofluorescin diacetate (DCF-DA), N-acetyl l-cysteine (NAC), catalase, 3,3′-dihexyloxacarbocyanine iodide (DiOC6(3)) and carbonyl cyanide m-chlorophenylhydrazone (CCCP) from Sigma Chemical Co., St Louis, MO, USA; RNase A and d-sorbitol from WAKO Chemical Co., Tokyo, Japan; propidium iodide (PI) from Molecular Probes Inc., Eugene, OR, USA; phycoerythrin (PE)-conjugated anti-Fas MoAb (clone UB2) and anti-Fas blocking MoAb (clone ZB4) from Immunotech Co., Fullerton, CA, USA; anti-FasL MoAb (clone 4H9), anti-Fas agonistic MoAb (clone CH-11), anti-Bax MoAb (clone 4F11), caspase inhibitors (Z-VAD-fmk) (non-specific inhibitor), Z-DEVD-fmk (caspase-3 inhibitor), Z-IETD-fmk (caspase-8 inhibitor) and Z-IEHD-fmk (caspase-9 inhibitor) from BML, Nagoya, Japan; anti-Bcl-2 MoAb (clone 124) from Dako, Glostrup, Denmark; antihuman EC MoAb (clone P1H12) and anti-Bcl-XL MoAb (clone 7B2·5) from Chemicon International Inc., Temecula, CA, USA; anti-Bcl-XL/S polyclonal antibody (pAb), anti-A1 pAb and HRP-conjugated antigoat donkey IgG from Santa Cruz Biotechnology Inc, Santa Cruz, CA, USA; ECLTM Western blotting detection system, HRP-conjugated antirabbit donkey IgG and HRP-conjugated antimouse sheep IgG from Amersham Life Science, Brussels, Belgium; IVIG (Venoglobulin-IH, lot no. E001VHT, 90 mg/ml, Welfide Corporation, Osaka, Japan, used in the form sent by the manufacturer), F(ab′)2 fragment and Fc fragment from Welfide Corporation, Osaka, Japan. We evaluated the levels of TNF-α, IFN-γ and IL-6 by ELISA (Quantikine, R&D Systems, Minneapolis, MN, USA) in the IVIG preparations, used in the present study, and the levels of these cytokines were < 5·0 pg/ml. F(ab′)2 and Fc fragments were purified using column chromatography following the digestion of the IVIG preparations with pepsin and papain, respectively. The purity of each fragment was confirmed by a single band in an immunoblot analysis.

Isolation and culture of human endothelial cells

The HUVECs were isolated according to the methods of Jaffe et al.[29]. The cells were suspended in M199 (Gibco BRL, Life Technologies, Inc., Rockville, MD, USA) containing 10% FBS, 10 ng/ml of bFGF and 100 μ/ml of penicillin and streptomycin and were cultured in a collagen-type 1-coated plastic dish (BIOCOAT®, Becton Dickinson, San Jose, CA, USA) at 37°C in a 5% CO2 atmosphere. The culture medium was changed the following day and thereafter twice weekly. When they became 70–80% confluent, the HUVECs were treated with 0·05% trypsin/0·53 mm EDTA, resuspended in the culture medium, and plated at 1105/ml. Only the second and third passage cells were used in the present experiments. When the HUVECs became 90% confluent, the medium was exchanged with a bFGF-free medium containing TNF-α (1, 10, 50, 100 and 500 ng/ml). After culturing for 6 h, IVIG was added at the concentration of 5, 10, 20, 30 and 40 mg/ml, and then the cells were cultured for 12, 24, 36, 48 and 72 h.

To investigate effect of anti-Fas MoAbs, antioxidants and caspase inhibitors on TNF-α-stimulated HUVECs, an agonistic anti-Fas MoAb (CH-11, 500 ng/ml), a blocking anti-Fas MoAb (ZB4, 200 ng/ml), 30 mm NAC, 250 U/ml of catalase or 10 μm caspase inhibitors (Z-VAD-fmk, Z-DEVD-fmk, Z-IETD-fmk and Z-IEHD-fmk) was added to the culture medium and it was then cultured for 1 h, followed by incubation with IVIG for 24–36 h.

Analysis of apoptotic cells by a flow cytometer

After the HUVECs were cultured at several conditions, both detached and adherent cells were collected and centrifuged at 200 g for 5 min The pellet was treated with a hypotonic solution (0·1% sodium citrate, 0·1% TritonX-100 and 20 μg/ml of RNaseA), and the cells were then stained with propidium iodide (PI). Apoptotic cells were quantified by flow cytometric determination of proportion of cells with hypodiploid DNA, as described previously by others [30]. We used a FACSCalibur flow cytometer (Becton Dickinson, San Jose, CA, USA), and the data were analysed using the CellQuest software package (Becton Dickinson).

DNA fragmentation assay

DNA from HUVECs were extracted by a Quick Apoptosis DNA Ladder Detection Kit (MBL, Nagoya, Japan). Samples were then loaded onto 1·5% agarose gel. After electrophoresis at 10 V for 2 h, the gel was subsequently stained with ethidium bromide and visualized under UV illumination.

SDS-PAGE and immunoblotting

HUVECs were washed with PBS and lysed with a lysis buffer (20 mm Tris-HCl, 1% Triton-X, 0·5% deoxycholic acid, 150 mm NaCl) containing protease inhibitor cocktail. SDS-PAGE was performed using the Bio-Rad Mini Protein II apparatus. After transfer, the membranes (polyvinylidene fluoride) were blocked with PBS-0·1% Tween 20 (PBS-T) with 5% skim milk. After being washed with PBS-T, the membranes were incubated with the first antibody (anti-Bcl-2, Bcl-XL, A1, Bax, Bcl-XS MoAbs). Next, blots were developed with ECLTM Western blotting detection system using an HRP-conjugated second antibody.

Confocal laser microscopy

Intracellular ROS was measured as described previously by others [31]. Briefly, HUVECs were exposed to DCF-DA (5 μg/ml) for 5 min The fluorescence of DCF was observed under a confocal laser microscopy (LSM410, Carl Zeiss Inc., Oberkokken, Germany). Furthermore, the fluorescence intensity was quantified on an arbitrary scale (0–256), and the mean fluorescence intensity in 100 random cells were measured and averaged in each condition.

Analysis of the mitochondrial membrane potential (Δψm) by flow cytometer

Δψm was measured as described previously by others [32]. Briefly, HUVECs were exposed to 10 mm DiOC6(3) for 15 min at 37°C. The cells were harvested by trypsinization and were then resuspended in PBS, followed by the determination of Δψm using a flow cytometer. A mitochondrial uncoupler, 50 μm CCCP, was used as a positive control for the detection of decreased Δψm.

Statistical analysis

All data are expressed as the mean ± SD, and the differences analysed by the Mann–Whitney test. A P-value < 0·05 was considered significant.

RESULTS

EC apoptosis in after stimulation of TNF-α and treatment of IVIG

HUVECs prestimulated with or without TNF-α (100 ng/ml) were cultured in the presence and absence of IVIG (20 mg/ml). In the following experiments, the HUVECs were divided into four groups: (1) unstimulated, TNF-α(−) and IVIG(−); (2) unstimulated + IVIG, TNF-α(−) and IVIG(+); (3) TNF-α only, TNF-α(+) and IVIG(−); (4) TNF-α + IVIG, TNF-α(+) and IVIG(+). The DNA content was analysed by flow cytometer, and the proportion of apoptotic HUVECs was presented as the percentage of cells with hypodiploid DNA (Fig. 1). No significant differences in the proportion of cells with hypodiploid DNA were seen among the treatments with unstimulated, unstimulated + IVIG and TNF-α only. In contrast, a substantial loss (~46% apoptosis) of the DNA content was observed after the TNF-α + IVIG treatment. In addition, agarose gel electrophoresis demonstrated characteristic DNA-ladder formation in the HUVECs treated with TNF-α + IVIG (Fig. 2). As a result, IVIG induces apoptosis in TNF-α-stimulated HUVECs, but not in unstimulated HUVECs.

Fig. 1.

Fig. 1

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Flow cytometric analysis of propidium iodide (PI)-stained HUVECs. After the HUVECs were stimulated with or without TNF-α (100 ng/ml) for 6 h, the cells were incubated in the presence and absence of IVIG (20 mg/ml) for 36 h. Representative histograms of the DNA content in HUVECs from a single donor are demonstrated. The percentage (%) of the cells with hypodiploid DNA was regarded as the proportion of apoptotic cells.

Fig. 2.

Fig. 2

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DNA ladder formation of HUVECs.DNAs from floating and remaining adherent HUVECs were extracted, and each sample was electrophorized onto a 1·5% agarose. A molecular-weight marker of 100-bp DNA ladder was run as a control.

Time- and dose-dependent induction of EC apoptosis by IVIG

After the HUVECs were stimulated with TNF-α for 6 h, the cells were cultured in the presence of IVIG for 72 h. The time course for the percentage (%) of cells with hypodiploid DNA was plotted in Fig. 3a. IVIG had little effect on the percentage of cells with hypodiploid DNA in the unstimulated HUVECs. On the other hand, IVIG induced a significant increase in the percentage of cells with hypodiploid DNA in the TNF-α-stimulated HUVECs at the incubation time of 24, 36 and 48 h. Since the percentage of apoptotic cells observed at 72 h was very high, we could not rule out the possibility that necrotic cells and late apoptotic cells were contaminated. As a result, IVIG enhanced the apoptosis of the TNF-α-stimulated HUVECs time-dependently by 48 h. Furthermore, the HUVECs were stimulated with TNF-α and then treated with IVIG at diverse concentrations (Fig. 3b). When the HUVECs were stimulated by TNF-α at concentrations of 50, 100 and 500 ng/ml, IVIG showed an increase in the percentage of cells with hypodiploid DNA in a dose-dependent fashion.

Fig. 3.

Fig. 3

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Time- and dose-dependent effects of IVIG on HUVEC apoptosis. (a) TNF-α (100 ng/ml)-stimulated or unstimulated HUVECs were incubated in the presence and absence of IVIG (20 mg/ml) for 72 h. The percentage of cells with hypodiploid was plotted in 0, 6, 12, 24, 36, 48 and 72 h after the addition of IVIG. •, Unstimulated; ○, unstimulated + IVIG; ■, TNF-α only; □, TNF-α + IVIG. *P < 0·05 versus unstimulated, unstimulated + IVIG and TNF-α only. (b) After the HUVECs were stimulated with TNF-α (0, 1, 10, 50, 100 and 500 ng/ml) for 6 h, the cells were incubated with IVIG (5, 10, 20, 30 and 40 mg/ml) for 24 h. The control indicates no treatment with IVIG. *P < 0·05 versus control in the same group. The data were expressed as the mean of samples ± SD from five experiments. □, Control; Inline graphic, IVIG: 5 mg/ml; Inline graphic, IVIG: 10 mg/ml; Inline graphic, IVIG: 20 mg/ml; ■, IVIG: 30 mg/ml; Inline graphic, IVIG: 40 mg/ml.

Effect of F(ab′)2 and Fc fragments on EC apoptosis

HUVECs stimulated with TNF-α were cultured with equimolar amounts of intact IVIG, F(ab′)2 fragments, Fc fragments and albumin (Fig. 4). The cells were also treated with d-sorbitol, which is contaminated in the therapeutic preparations of IVIG and has an equal osmolarity with physiological saline (0·9% NaCl). IVIG, F(ab′)2 fragments and Fc fragments induced a significant increase in the percentage of cells with hypodiploid DNA in the TNF-α-stimulated HUVECs but albumin and d-sorbitol did not, in comparison to the HUVECs without these treatments. However, the percentage of cells with hypodiploid DNA tended to be low in the HUVECs treated with F(ab′)2, Fc fragments and F(ab′)2 + Fc fragments, compared with intact IVIG. This result indicates that intact Ig has a stronger capacity to induce the IVIG-induced apoptosis of HUVECs than F(ab′)2 fragments and Fc fragments.

Fig. 4.

Fig. 4

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Effect of F(ab′)2, Fc fragments, albumin and d-sorbitol on HUVEC apoptosis.After HUVECs were stimulated with or without TNF-α (100 ng/ml) for 6 h, the cells were incubated with equimolar (0·12 mm) of IVIG, F(ab′)2 fragments, Fc fragments and albumin and 20 mg/ml of d-sorbitol for 36 h. The percentage of cells with hypodiploid DNA was expressed as the mean of samples ± SD from five experiments. The control indicates no treatment with IVIG, F(ab′)2 and Fc fragments, albumin and d-sorbitol. *P < 0·05 versus control. □, Control; ■, IVIG; Inline graphic, F(ab)2; Inline graphic, Fc; Inline graphic, F(ab′)2 + Fc; Inline graphic, albumin; Inline graphic, d-sorbitol.

Involvement of Fas-mediated pathway in IVIG-induced EC apoptosis

First, we analysed the cell surface expression of Fas and FasL on the surface of the HUVECs, but IVIG induced no changes of the surface expression of Fas and FasL on the HUVECs in unstimulated and TNF-α-stimulated HUVECs. Secondly, to investigate any possible involvement of Fas-dependent pathway in the IVIG-induced apoptosis, either agonistic anti-Fas MoAb (CH-11) or blocking anti-Fas MoAb(ZB4) was added to the TNF-α-stimulated HUVECs 1h before the IVIG was administered (Fig. 5). CH-11 induced a significant increase in the percentage of cells with hypodiploid DNA in the HUVECs treated with TNF-α only, but not in the HUVECs treated with unstimulated, unstimulated + IVIG and TNF-α + IVIG. Furthermore, ZB4 did not significantly decrease the percentage of cells with hypodiploid DNA in the HUVECs treated with TNF-α + IVIG. Therefore, these results indicate that the Fas-mediated pathway may play only a small role in the apoptosis of TNF-α-stimulated HUVECs.

Fig. 5.

Fig. 5

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Effect of anti-Fas MoAbs (CH-11 and ZB4) on HUVEC apoptosis. After the HUVECs were stimulated with or without TNF-α (100 ng/ml) for 5 h, the cells were cultured in the presence and absence of anti-Fas MoAbs (500 ng/ml of CH-11 and 200 ng/ml of ZB4) for 1 h, followed by the incubation with IVIG (20 mg/ml) for 24 h. The percentage of cells with hypodiploid DNA was expressed as the means of the samples ± SD from five experiments. Control indicates no treatment of anti-Fas MoAbs. *P < 0·05 versus control in the same group. □ Control; Inline graphic CH-11; ■ ZB4.

Effect of IVIG on the expression of Bcl-2 family in EC

To investigate whether IVIG treatment could influence the expression of the anti- and pro-apoptotic Bcl-2 homologues, an immunoblotting analysis was carried out (Fig. 6). All proteins (Bcl-2, A1, Bcl-XL, Bax and Bcl-Xs) were expressed constitutively. In an anti-apoptotic subfamily, the expression of A1 and Bcl-XL increased in the HUVECs treated with TNF-α only, while such expression decreased in the HUVECs treated with TNF-α + IVIG. There was no change in the expression of Bcl-2. In a pro-apoptotic subfamily, the expression of Bax and Bcl-XS showed no dramatic change in the unstimulated HUVECs, HUVECs treated with unstimulated + IVIG and HUVECs treated with TNF-α only, but the expressions of both Bax and Bcl-XS increased markedly in the HUVECs treated with TNF-α + IVIG. As a result, IVIG induced either a decrease in the expression of antiapoptotic proteins (A1 and Bcl-XL) or an increase in the expression of pro-apoptotic proteins (Bax and Bcl-XS) in the TNF-α-stimulated HUVECs.

Fig. 6.

Fig. 6

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Intracellular expression of Bcl-2 family proteins in HUVECs. After HUVECs were stimulated with or without TNF-α (100 ng/ml) for 6 h, the cells were incubated in the presence and absence of IVIG (20 mg/ml) for 12 h. The cell lysates were analysed by SDS-PAGE and immunoblotting. Pre indicates the HUVECs before TNF-α stimulation.

Effect of IVIG on intracellular ROS in EC

To investigate the possible role for ROS in the IVIG-induced apoptosis of TNF-α-stimulated HUVECs, intracellular ROS was measured by confocal laser microscopy using DCF fluorescence (Fig. 7a). The levels of ROS were low in the unstimulated HUVECs and HUVECs treated with unstimulated + IVIG. An increase in ROS production was seen in the HUVECs stimulated with TNF-α only, and TNF-α + IVIG treatment produced a significantly higher level of ROS than TNF-α stimulation only.

Fig. 7.

Fig. 7

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Levels of intracellular ROS and effect of antioxidants on IVIG-induced HUVEC apoptosis. (a) After the HUVECs were stimulated with or without TNF-α (100 ng/ml) for 6 h, the cells were incubated in the presence and absence of IVIG (20 mg/ml) for 6 h. Thereafter, the HUVECs were exposed to DCF-DA and were observed under confocal laser microscopy. (b) After HUVECs were stimulated with or without TNF-α (100 ng/ml) for 6 h, the cells were treated with NAC (30 mm) and catalase (250 U/ml) for 1 h, followed by the incubation with IVIG (20 mg/ml) for 36 h. The control indicates no treatment with NAC or catalase. *P < 0·05 versus control. □, Control; Inline graphic, NAC; ■, catalase; Inline graphic, NAC + catalase.

We next investigated whether IVIG-induced production of ROS might be involved in HUVEC apoptosis. After the antioxidants (NAC and catalase) were added to the cells before IVIG treatment, the percentage of cells with hypodiploid DNA was determined (Fig. 7b). Although NAC or catalase alone did not show a significant decrease in the percentage of cells with hypodiploid DNA, NAC + catalase significantly reduced the percentage of cells with hypodiploid DNA. As a result, ROS may thus play some role in the IVIG-induced apoptosis of TNF-α-stimulated HUVECs.

Kinetics of IVIG-induced mitochondrial permeability transition in EC

The induction of mitochondrial permeability transition has been suggested in the apoptosis process. To characterize apoptosis signalling in TNF-α-stimulated HUVECs by IVIG, we measured the mitochondrial membrane potential (Δψm) based on the uptake of the mitochondrial-specific dye, DiOC6 (Fig. 8). IVIG induced an increase in the percentage of cells with a decreased Δψm in the TNF-α-stimulated HUVECs, but not in the unstimulated HUVECs.

Fig. 8.

Fig. 8

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Cytofluorometric analysis of the mitochondrial transmembrane potential (δψm).After the HUVECs were stimulated with or without TNF-α (100 ng/ml) for 6 h, the cells were incubated in the presence (bold lines) and absence (shaded histograms) of IVIG (20 mg/ml) for 24 and 36 h. After the HUVECs were exposed to DiOC6 and analysed by a flow cytometer. The proportion (%) of cells in M1 indicates the loss of δψm, represented by a decrease in the fluorescence (FL1-H). As a positive control, CCCP (50 μm) was able to cause a complete loss of δψm at 0 h (dotted lines). All data are representative of five independent experiments giving similar results. Inline graphic, IVIG (−); —, IVIG (+); …, positive control.

Inhibitory effect of caspase inhibitors on IVIG-induced EC apoptosis

To investigate the involvement of caspases on IVIG-induced EC apoptosis, we tested whether the IVIG-induced apoptosis of HUVECs could be inhibited by specific caspase inhibitors (Fig. 9). IVIG-induced apoptosis was significantly attenuated after the administration of broad spectrum caspase inhibitor (C.I.) and caspase-3, 8 and 9 inhibitors. The IVIG-induced apoptosis of HUVECs stimulated with TNF-α was attenuated significantly by treatments with all caspase inhibitors. As a result, the activation of caspase may mediate the IVIG-induced apoptosis of TNF-α-stimulated HUVECs.

Fig. 9.

Fig. 9

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Caspase activation and the effect of caspase inhibitors on the IVIG-induced HUVEC apoptosis.HUVECs were stimulated with or without TNF-α (100 ng/ml) for 6 h, the cells were treated with caspase inhibitors (10 μm) for 1 h, followed by incubation with IVIG (20 mg/ml) for 36 h. The percentage of cells with hypodiploid DNA was expressed as the mean of the samples ± SD from five experiments. C.I., non-specific caspase inhibitor; C.I.3, caspase-3 inhibitor; C.I.8., caspase-8 inhibitor; C.I.9, caspase-9 inhibitor. The control indicates no treatment with caspase inhibitors. *P < 0·05 versus control. □, Control; ■, C.I.; Inline graphic, C.I.3; Inline graphic, C.I.8; Inline graphic, C.I.9; Inline graphic, C.I.8 + C.I.9.

DISCUSSION

In the present study, IVIG induced apoptosis in the TNF-α-stimulated HUVECs, but not in the unstimulated HUVECs, in a dose- and time-dependent fashion. Anti-Fas MoAbs (CH-11 and ZB4) neither enhanced nor inhibited the apoptosis of TNF-α-stimulated HUVECs. IVIG increased the expression of such pro-apoptotic proteins as Bcl-XS and Bax, while IVIG decreased the expression of such anti-apoptotic proteins as Bcl-XL and A1. Furthermore, IVIG increased the intracellular production of ROS and decreased the Δψm in the TNF-α-stimulated HUVECs. The IVIG-induced apoptosis was inhibited by caspase inhibitors.

Recent data suggest that apoptosis takes place through a death receptor-dependent pathway and/or through a mitochondria-dependent pathway [17]. Since the expression of Fas receptor is reported to be present at low levels on the surfaces of ECs, ECs are thought to be resistant to Fas-mediated cell death under normal conditions [32,33]. Sata et al. reported that TNF-α decreased the expression level of FasL on ECs while ECs are sensitive to Fas-mediated apoptosis after TNF-α stimulation [33]. Prasad et al. revealed that anti-Fas antibodies contained in IVIG preparations can induce lymphocyte apoptosis via the Fas-mediated pathway [26]. In the present study, however, blocking anti-Fas MoAb (ZB4) did not inhibit IVIG-induced HUVEC apoptosis (Fig. 6b). Therefore, the IVIG-induced apoptosis in the TNF-α-stimulated ECs may not be dependent on the Fas-mediated pathway. As an alternative pathway, many Bcl-2 family proteins are predominantly located in the outer mitochondrial membrane, and the relatively abundant expression of pro- and anti-apoptotic proteins determines the susceptibility to cell death [18]. Bcl-XL and A1 have been reported to inhibit TNF-α-induced EC apoptosis [34,35]. Bcl-XL and Bax are known to be involved in the LPS-induced EC apoptosis [36]. Vitamin C and E inhibit the LPS-induced apoptosis in ECs by modulating the expression of Bcl-2 and Bax [37]. The present study demonstrated that IVIG induced the apoptosis of the TNF-α-stimulated HUVECs by altering the balance of expression between anti-apoptotic proteins (A1 and Bcl-XL) and pro-apoptotic proteins (Bax and Bcl-XS), and that IVIG induced the disruption of the mitochondrial membrane. These findings thus indicate that IVIG-induced EC apoptosis is dependent on the mitochondrial signalling pathway.

Although TNF-α is a pro-apoptotic factor of ECs [3,7], most normal ECs in culture are resistant to TNF-α-induced apoptosis [35,38]. However, TNF-α can cause EC apoptosis when ECs were co-cultured with the protein synthesis inhibitor (cycloheximide) [39,40]. TNF-α has been reported to induce EC apoptosis by increasing the caspase activity via TNF-α receptor signalling pathway [41], while ECs produce de novo anti-apoptotic proteins [40], such as Bcl-XL[42] and A1 [43] in Bcl-2 family protein, after TNF-α stimulation. Since TNF-α is thus able to elicit both cell survival and death signals at the same time, ECs hardly undergo apoptosis in response to TNF-α alone. The present study also demonstrated that treatment with TNF-α did not induce HUVEC apoptosis (Figs 1–3). Furthermore, treatment with TNF-α increased only the expression of Bcl-XL and A1 without the increased expression of Bax and Bcl-Xs (Fig. 6). As a result, HUVECs may be prevented from undergoing apoptosis by increasing the expression of antiapoptotic proteins, when HUVECs are stimulated with TNF-α only.

Several mechanisms of IVIG action have been proposed to explain the immunomodulatory properties [25]. As a plausible mechanism, therapeutic concentrations of IgG block Fc receptors on phagocytes and thus inhibit antibody-dependent cytotoxity by cellular effectors [44]. IVIG has also been demonstrated to down-regulate the proliferation and functions of activated B and T cells [45,46] and to reduce the cytokine productions from immuno-effector cells [47,48]. The present study demonstrated a new action that IVIG can induce the apoptosis of TNF-α-stimulated HUVECs via a mitochondria-dependent pathway. Since the IVIG concentrations (5–40 mg/ml) used in the present study in vitro are equivalent to the physiological concentrations after high dose IVIG therapy in vivo, IVIG may thus be potentially useful as an EC apoptosis-inducing drug. Although hyperosmotic shock, such as a high concentration of NaCl, has been reported to induce apoptosis [49], the d-sorbitol used as a solvent in the present IVIG preparations did not induce HUVEC apoptosis (Fig. 4), thus indicating that IgG induces apoptosis in HUVECs. Furthermore, F(ab′)2 and Fc fragments of IgG induced the apoptosis of TNF-α-stimulated HUVECs in the present study, although their effect was lesser than the effect of intact IgG. The underlying mechanism of IVIG action for EC apoptosis remains unknown in the present study, but both F(ab′)2 and Fc fragments of IgG are suggested to be involved in this function. It is possible that Fc receptors on ECs may be involved in apoptosis and/or that antibody activity may merely be in part lost by the enzyme digestion. Xu et al. also reported the inhibitory effect of IVIG on EC proliferation to be dependent on both F(ab′)2 and Fc fragments [28].

Intracellular ROS is involved in EC apoptosis through a mitochondrial pathway, by decreasing the Δψm[18]. Inorganic iron and heat shock [50] and high concentration of glucose [51] induce EC apoptosis via an ROS-dependent mechanism. The stimulation of TNF-α only has also been reported to increase the ROS production in ECs [52]. In the present study, treatment with TNF-α + IVIG produced a larger amount of intracellular ROS in HUVECs than treatment with TNF-α only. Furthermore, the addition of antioxidants (NAC and catalase) partly inhibited HUVEC apoptosis. These findings suggest that an increase of intracellular ROS is in part involved in IVIG-induced EC apoptosis. Furthermore, the treatment with caspase inhibitors suppressed the proportion of apoptotic cells in the HUVECs stimulated with TNF-α + IVIG, thus indicating that the activation of caspase cascades is necessary for IVIG-induced EC apoptosis to occur.

The pathophysiological significance of IVIG-induced EC apoptosis in vivo may be interpreted in different ways: the clinical use of IVIG may have either an advantageous or disadvantageous effect on the human body. EC is a focus in inflammation, and activated ECs secrete cytokines [53], chemokines [54], histamine [55] and matrix metalloproteinase [56], and these inflammatory mediators may contribute to the tissue injury [3]. Activated ECs also increase the expression of adhesion molecules, accelerating the neutrophil-mediated EC injury [57]. Cytokine-activation may shift anticoagulant to the pro-coagulant surface on ECs, thus inducing intracellular thrombus formation [58]. As a result, the activated ECs may contribute to the deterioration of inflammation. Therefore, if IVIG could selectively induce the apoptosis of the cytokine-activated ECs in vivo, this phenomenon may thus be a suitable mechanism to limit inflammatory process. Furthermore, EC apoptosis inhibits angiogenesis and vasculogenesis and may thereby prevent the tumour blood supply thus leading to tumour regression [59]. Since IVIG is suggested to repress angiogenesis by inhibiting EC proliferation [27], IVIG may also provide a safeguard against the deregulated cell proliferation. In contrast, IVIG may have an unfavourable effect on EC integrity. EC apoptosis is reported to play an important role in the early phase of atherosclerosis [3,17], and apoptotic ECs have also been shown to become procoagulant [60]. In sepsis and the associated syndrome including systemic inflammatory response syndrome (SIRS), ARDS and multiple organ failure (MOF), the EC apoptosis induced by pro-apoptotic factors such as LPS and TNF-α is suggested to be involved in the pathology of tissue injury [3,6]. As a result, IVIG-induced EC apoptosis may promote the destruction of the vascular barrier functions in ECs. Malek et al. suggest that clinical use of the diuretic mannitol have a deleterious effect on ECs by inducing EC apoptosis [61]. Therefore, to clarify the significance of the IVIG-induced EC apoptosis in vivo, further studies are called for in the future.

In summary, IVIG induced the apoptosis of TNF-α-stimulated ECs in a dose- and time-dependent fashion. These mechanisms are caused by altering the expression levels between anti- and pro-apoptotic proteins, by increasing the intracellular production of ROS, and by the resultant collapse of Δψm. Therefore, IVIG induces apoptosis in TNF-α-stimulated ECs through a mitochondria-dependent pathway. These observations may provide new insight into the mechanism of IVIG action, although the therapeutic significance in IVIG-induced EC apoptosis should still be further investigated.

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