Published online before print January 18, 2008
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* Department of Immunology and Cell Biology, Research Center Borstel, Borstel, Germany; and
District Hospital Segeberger Kliniken, Bad Segeberg, Germany
1 Correspondence: Department of Immunology and Cell Biology, Research Center Borstel, Parkallee 22, D-23845 Borstel, Germany. E-mail: fpetersen{at}fz-borstel.de
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Key Words: macrophages • chemokines • cytotoxicity
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Besides inducing short-term responses, PF4 contributes to long-term regulatory processes of cell activation and differentiation, such as the support of the survival of hematopoietic stem cells and progenitor cells [16 ]. Furthermore, PF4 acts as a potent inhibitor of T cell activation [17 , 18 ] and EC proliferation [19 ]. Although the effect on the former cells is mediated indirectly by interference with the autocrine IL-2 loop, the regulation of EC appears to be more complicated. The latter involves competition of PF4 with growth factors for binding to coreceptors, direct interaction with growth factors, as well as direct binding to CXCR3B, a recently described splice variant of CXCR3 [20 21 22 23 ]. Furthermore, it has been shown that PF4 reversibly abolishes the entry of these cells into S-phase and thereby inhibits DNA synthesis [24 ].
Although in most cell types, PF4 exclusively mediates long-lasting or short-term biological responses, a complex spectrum of different consecutive functions is observed in monocytes. During an initial phase of up to 60 min, PF4 induces the generation of oxygen radicals and phagocytosis in these cells [25
]. This period is followed by an intermediary phase of 2–8 h, which is characterized by functions such as tight adhesion of monocytes and the release of TNF-
. Thereafter, PF4 initiates a cellular program, which prevents monocytes from undergoing spontaneous apoptosis and induces differentiation of these cells into a specific subtype of macrophage. Different from GM-CSF- or M-CSF-derived macrophages, PF4-treated cells lack surface expression of HLA-DR but show an up-regulation of the costimulatory molecule B7-2 [26
]. Moreover, the latter cell type displays enhanced innate immune functions such as phagocytosis as well as the generation of reactive oxygen species (ROS) [27
].
The interaction of monocytes with EC of the vessel wall is a crucial process in tissue homeostasis as well as under pathological conditions of inflammation or tissue injury. Depending on the respective situation, monocytes may migrate through this barrier, remove damaged cells, and initiate wound healing and angiogenesis. On the other hand, activated monocytes are involved in vascular diseases such as reperfusion injury and atherosclerosis (reviewed in ref. [28 ]). In the present study, we have investigated the molecular basis as well as the functional consequences of the interaction of EC with PF4-activated monocytes. Here, we can show that human blood monocytes triggered with PF4 have a considerable cytotoxic effect on resting EC. This cytotoxic effect is mediated by the formation of oxygen radicals leading to the induction of programmed cell death of EC.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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).
Cell preparation and culture
Human EC were isolated from umbilical cord veins by collagenase treatment and cultured in dishes precoated with 0.04% gelatine, according to Jaffe et al. [33
], with minor modifications described by Schönbeck et al. [34
]. Briefly, the cells were maintained in the EC culture medium EBM in 25 cm2 cell culture flasks. For experiments, cells were subcultured after trypsinization in 96-well microtiter plates in 200 µl at a density of 5000 cells/cm2. Cells were cultured until confluence and used throughout passages 1 and 2.
Monocytes were obtained from peripheral blood of healthy volunteer donors by Ficoll-Paque gradient centrifugation and subsequent counterflow centrifugation as described earlier [35 ]. The purity of the monocytes in all experiments was more than 95%. Approval for these studies was obtained from the Institutional Review Board at the University of Lübeck (Germany), and informed consent was provided according to the declaration of Helsinki.
The human lung epithelial cell line A549 (American Type Culture Collection (Manassas, VA, USA) was cultured in DMEM supplemented with 10% FCS, 1% L-Glu, and 1% PEST.
For coculture with EC, 100 µl aliquots of monocytes (1.25x105/ml) were added to microtiter plates onto confluent EC layers in coculture medium (M199/0.1% BSA/1% L-Glu/1% PEST). In some experiments, concentrations of monocytes varied from 0.625 x 105/ml up to 20 x 105/ml. Unless otherwise indicated, cocultures were performed in the presence or absence of PF4 (4 µM) or GM-CSF (500 U/ml) for 18 h. Alternatively, cocultures were performed with cells of the epithelial cell line A549 instead of EC. In some experiments, the cell cultures were treated additionally with DPI (4 µM), apocynin (1 mM), or antibodies directed against CD18, CD54, or CD102 at concentrations of 10 µg/ml for each antibody.
To prevent direct cell contact between monocytes and EC, some cocultures were performed in transwell units (pore size 0.2 µM, Nunc, Wiesbaden, Germany). Therefore, EC were grown on the bottom of the plates, and monocytes were placed in the transwell units.
In a separate set of experiments, EC were cultured with monocyte-conditioned medium. Therefore, increasing concentrations of monocytes (0.625x105/ml–2.5x105/ml) were stimulated for 18 h with 4 µM PF4 or remained unstimulated. Cell-free supernatants were collected and subsequently added to EC.
Cytotoxicity assay
The cytotoxic effect of monocytes against EC and epithelial cell line A549 was determined by using the UptiBlue reagent. The indicator is added to the cells in its oxidized form and subsequently becomes reduced by mitochondrial enzyme activity of the viable cells. The chemical reduction exhibits a change in color that can be quantified colorimetrically [36
]. According to the recommendations of the manufacturer, 10 µl of the reagent was added to each well containing monocytes and EC and was incubated for an additional 3 h. Thereafter, absorbance was recorded by an automatic photometer at wavelengths of 570 and 600 nm, and the relative cytotoxicity was calculated according to the following formula: Percent cytotoxicity = [1–((At–Am)/Ab)] x 100; where is A = difference between the absorbance measured at 570 and 600 nm; t = A in the coculture of monocytes and EC/A549; m = A in the culture of monocytes alone; and b = A in the culture of EC/A549. Ab was determined in parallel in cultures that received medium alone or a corresponding amount of PF4. No differences in cell viability were observed between both cultures. Routinely, cocultures were additionally analyzed for cell integrity by phase-contrast microscopy, and photographs of representative sections were taken.
Oxygen radical formation
Generation of ROS by monocytes was determined by measurement of chemiluminescence in the presence of luminol as described elsewhere [37
]. Briefly, 200 µl aliquots of monocytes (1x106/ml) were distributed into an intransparent, 96-well microtiter plate (Nunc) on the plastic surface or on confluent EC, and chemiluminescence was recorded for 2 h. Results were expressed as relative light units (RLU). Some experiments were performed in the presence of the NADPH oxidase inhibitors DPI (0.5–8 µM) and apocynin (0.25–4 mM), which were added 15 min before PF4 treatment.
Evaluation of cell viability and detection of apoptosis
Determination of apoptotic cells was done by double-staining with Annexin-V-FITC and propidium iodide (PI) using an Annexin-V kit. Briefly, after the detachment of EC and monocytes by exposure to cell dissociation solution, cells were incubated with Annexin-V-FITC in binding buffer (provided by the manufacturer) for 20 min on ice, washed, and resuspended in the same buffer as described by the manufacturer. Monocytes were identified by staining with APC-labeled anti-CD14 and excluded from further analysis. PI (1 µg/ml finally) was added immediately before flow cytometrical analysis, performed with a FACSCalibur (Becton Dickinson, Heidelberg, Germany).
Statistics
All experiments were performed at least three times with cells from different individuals, and the data are expressed, unless indicated otherwise, as mean of the replicates ± SD. Statistical significance was analyzed by Wilcoxon-matched pairs test or by Kruskal-Wallis test.
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Figure 1. Effect of monocytes on the integrity of EC. Monocytes (5x105/well) were cultured in the absence [unstimulated (unst.); A] or in the presence of 500 µ/ml GM-CSF (B) or 4 µM PF4 (C) on confluent layers of EC for 18 h in 24-well plates. In control cultures, EC received 4 µM PF4 in the absence of monocytes [without monocytes (w/o Monoc.; D]. Photographs of representative sections were taken by phase-contrast microscopy (20-fold magnification). The data from one representative experiment out of five are given.
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Figure 2. Dependency of PF4-induced monocyte cytotoxicity on stimulus concentration, effector cell density, culture time, and target cell type. (A) Monocytes (0.125x105 cells/well) were exposed for 18 h to confluent layers of EC in 96-well plates in the absence or presence of increasing concentrations of PF4. (B) Monocytes (0.125x105/well) were left unstimulated (open bars) or stimulated with 4 µM PF4 (black bars) or 500 µ/ml GM-CSF (gray bars) and cultured for different time periods on EC layers. Alternatively, increasing numbers of monocytes were stimulated with 4 µM PF4 (solid bars) or left untreated (open bars), followed by 18 h exposure to EC (C) or to A549 cells (D). Cytotoxicity was calculated on the basis of residual living cells determined by using the UptiBlue system, where EC, cultured with or without PF4 in the absence of monocytes, was set as 100% living cells. Data represent mean ± SD of four (B) and three (A, C, and D) independent experiments. Statistically significant differences between cytotoxicity of PF4-treated or untreated approaches are indicated (*, P 0.01).
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Figure 3. Role of cell contact and adhesion molecules in PF4-mediated cytotoxicity. Monocytes (0.125x105/well) were stimulated with 4 µM PF4 (black bars), 500 µ/ml GM-CSF (gray bars), or left untreated (open bars). Subsequently, EC were cocultured in the presence of monocytes or exposed to cell-free supernatants derived from stimulated or unstimulated monocytes cultured for 18 h (A). Alternatively, stimulated or unstimulated monocytes were cocultured with EC under conditions that allowed a direct cell contact or where cell contact was prevented by a transwell membrane (0.2 µM pore size; B). Furthermore, PF4-treated as well as untreated monocytes were cocultured on EC layers in the presence of antibodies directed against the common chain of β2 integrin (10 µg/ml; C) or ICAM-1 as well as ICAM-2 (10 µg/ml, respectively; D) or corresponding isotype controls. After 18 h, cytotoxicity was measured by UptiBlue reagent. Data represent mean ± SD of three (B and D) or five (A and C) independent experiments.
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Taken together, we conclude that PF4-mediated cytotoxicity of monocytes against EC depends on a direct cell contact, which is mediated by ICAM-1 on EC and β2 integrins on monocytes.
Monocytes kill EC by the formation of radical oxygen species
Next, we investigated the mechanisms by which monocytes mediate their cytotoxic effects on EC. In preliminary experiments, we could show that 5% FCS was able to completely abrogate the cytotoxic effect of the chemokine (data not shown). As serum is known to contain several protease inhibitors, we first investigated the role of proteases in this regard. However, neither inhibitors of serine proteases or cysteine proteases (aprotinin and leupeptin, 0.25–1 µg/ml, respectively) nor a general inhibitor of endoproteases (2-macroglobulin, 1 mg/ml) showed any effect on the PF4-mediated cytotoxicity (data not shown). As this set of inhibitors covers all prominent monocyte proteases, we exclude a relevant role for these enzymes. As it is known that serum, besides inhibitors of proteases, also contains scavengers for ROS, we examined next whether these short-lived products of monocytes are involved in the cytotoxic process. In a first approach, we tested whether PF4-stimulated monocytes, when attached to EC, are able to produce ROS. As a control, monocytes were allowed to settle on a plastic surface prior to stimulation. Under both conditions, approximately within 2–5 min after contact with PF4, monocytes started to release ROS, which reached a maximum after 20 min of stimulation. However, although ROS production in monocytes attached to plastic surfaces returned to background levels after 80 min, in the presence of EC, release of oxygen metabolites remained nearly unchanged at high levels for over 2 h of data recording (Fig. 4A
). This indicates that EC cocultured with PF4-stimulated monocytes may be exposed to oxidative stress for rather long time periods. Statistical analysis of the integrated data based on four experiments revealed that during the first 60 min, no significant difference between cells attached to plastic and EC occurred (P=0.1157). However, ROS production of monocytes adherent to EC was significantly higher than that by cells layered on plastic when the calculation was based on an interval of 120 min of stimulation (P<0.048). Interestingly, under the same conditions, no difference between monocytes attached to plastic or to EC was seen when fMLP (1 µM) was used for monocyte activation (data not shown), and GM-CSF (250–1000 U/ml) was incapable of inducing any ROS production (not shown). Most strikingly, comparable with their effect on PF4-induced cytotoxicity, antibodies directed against ICAM-1 or CD18 also inhibited PF4-induced ROS production. Although in the presence of anti-ICAM-1, burst rates were reduced by more than 60%, formation of ROS was completely abrogated in samples that received antibodies to CD18 (Fig. 4A)
. Corresponding antibodies of the same isotype were without effect on PF4-induced release of oxygen radicals (data not shown). To prove that ROS are responsible for the cytotoxic effect of monocytes on EC, two inhibitors of NADPH oxidase, apocynin and DPI, were used. Both inhibitors were able to block oxygen radical formation induced by PF4 in a dose-dependent manner, with complete inhibition observed at 1 mM apocynin or 8 µM DPI (Fig. 4B)
. Consequently, both inhibitors were used in cocultures of monocytes and EC in the presence of PF4. However, as these substances interfere with the detection of viable cells by redox indicators such as UptiBlue and formazan [45
], analyses were performed by phase-contrast microscopy. In the absence of inhibitors, PF4-activated monocytes completely destroyed the EC layer (Fig. 4C
, left panel). Conversely, treatment of cultures with apocynin or DPI (Fig. 4C
, middle and right panels, respectively) drastically inhibited the PF4 effect. Approximately 80–90% of the EC layer remained intact compared with a parallel culture without PF4. Both inhibitors were without effect on EC control cultures, receiving unstimulated monocytes or PF4 in the absence of these cells (data not shown). These data provide direct evidence that ROS released from PF4-activated monocytes are responsible for the cytotoxic effect on EC.
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Figure 4. The role of monocyte-derived ROS in PF4-mediated cytotoxicity. (A) Monocytes attached to a plastic surface or to an EC layer were stimulated with 4 µM PF4 (solid lines), and formation of oxygen radicals was visualized by chemiluminescence. Additionally, PF4-stimulated monocytes adherent to EC received antibodies (10 µg/ml) directed against ICAM-1 (dotted line) or CD18 (dashed line). (B) Monocytes were left untreated or pretreated with increasing concentrations of apocynin (gray bars) or DPI (black bars), followed by stimulation with 4 µM PF4. The formation of oxygen radicals was determined by chemiluminescence, and the data were recorded for 60 min. The data were integrated and shown as mean ± SD of three independent experiments. (C) Monocytes (3x105/well) were stimulated with 4 µM PF4 and cultured in the absence or presence of 1 mM apocynin or 8 µM DPI on confluent layers of EC. After 12 h of incubation, photographs of representative sections were taken by phase-contrast microscopy (20-fold original magnification). The data from one representative experiment out of three are given.
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Figure 5. PF4-stimulated monocytes induce apoptosis in EC. Monocytes (3x105/well) were added to EC layers in 24-well plates in the absence or presence of 4 µM PF4. After 12 h of incubation, EC were nonenzymatically detached, and numbers of apoptotic cells were determined by Annexin-V/PI double-staining by flow cytometry. Monocytes were identified in the same samples by using a CD14-APC antibody and excluded from further analysis. Data from one representative experiment out of nine are shown (A). In a parallel set of experiments, untreated (gray bars) or PF4-stimulated (black bars) monocytes (Mo) were cultured in the presence of 1 mM apocynin or 8 µM DPI on confluent EC, and EC cultured without monocytes (open bars) served as a control. Analysis of apoptotic EC was performed as described above. Data represent mean ± SD derived from nine independent experiments (B). Statistically significant differences between PF4-treated cocultures alone and those that received additional inhibitors are indicated (*, P<0.021).
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The generation of ROS, which occurs at low levels in nearly all cell types, is one of the fundamental defense mechanisms of phagocytic leukocytes against microbial invaders. However, uncontrolled production of ROS is involved in severe tissue damage, as observed in emphysema, reperfusion injury, or rheumatoid arthritis [47 ]. The mechanism underlying ROS-mediated damage of cells and tissues is poorly understood. Recent data provide evidence that oxidative stress may act as an inducer of apoptosis. Pajusto and coworkers [48 ] could demonstrate a proapoptotic effect of ROS on T cells, and Hansson et al. [49 ] demonstrate that monocytes are able to initiate apoptosis in autologous NK cells through a release of oxygen radicals. Furthermore, Warren et al. [50 ] observed that exposure of bovine EC to chemically generated ROS leads to programmed cell death in these cells. In a more detailed analysis, Madesh and colleagues [51 ] described that artificially generated ROS mediate apoptosis in rat EC via elevation of intracellular calcium concentration, leading to membrane depolarization of mitochondria and caspase activation. However, a clear cellular source of ROS as well as physiological stimuli, which are able to induce an appropriate ROS production in these cells leading to EC apoptosis, remained to be defined until now. Here, we could show for the first time that human monocytes activated by the platelet-derived chemokine PF4 induce a ROS-dependent programmed cell death in human EC. Interestingly, fMLP, as a potent inducer of rather short-lived ROS production in monocytes, failed to provoke EC cytotoxicity. This indicates that the duration of the stimulus-induced ROS release may play a critical role in the induction of the latter process. After activation with different soluble stimuli, monocytes produce oxygen radicals for a defined time period. Although the fMLP- and IL-8-induced responses are restricted to the first few minutes after stimulation, PF4-mediated oxidative burst occurs later in its onset and is remarkably longer in duration (lasting up to 60 min) [52 , 53 ]. In addition to this, we show here that as compared with monocytes attached to plastic surfaces, contact with EC results in a significantly sustained monocyte oxidative burst response in PF4-treated monocytes (Fig. 4) . Interestingly, irrespective of whether the monocytes were in contact to EC or plastic, ROS release induced by fMLP was not affected (data not shown). This suggests that only long-lasting, oxidative stress, as observed with PF4-activated monocytes attached to EC, is able to mediate killing of the latter cells.
Besides its capacity to induce oxygen radicals, PF4 also mediates the differentiation of monocytes into macrophages. However, this process appears not to be linked to monocyte cytotoxicity, as the development of macrophages induced by PF4 requires at least 48 h of stimulation ([54 ] and unpublished results), and a first cytotoxic effect became visible already after 12 h of culture (Fig. 2B) . Furthermore, GM-CSF, which is a potent elicitor of monocyte differentiation but did not induce any production of ROS in these cells (data not shown), failed to promote a cytotoxic effect against EC (Figs. 1 2 3) .
According to our data, direct cell–cell contact between monocytes and EC is an indispensable prerequisite for PF4-mediated cytotoxicity against EC (Fig. 3)
. This is in line with findings by others, showing that leukocyte oxidative burst requires adhesion of these cells [55
]. However, we could show that simply physical contact between EC and monocytes is not sufficient to induce the monocytic response. Moreover, our studies using inhibitory antibodies against adhesion molecules revealed that killing of EC by the monocytes strictly depends on the binding of monocyte β2-integrins and EC-expressed ICAM-1 (Fig. 3)
. In all probability, β2-integrin binding to its counter-receptor provides a costimulatory signal in monocytes, leading to prolonged production of ROS. Indeed, we found that the presence of antibodies against CD18 and ICAM-1 drastically reduces PF4-induced ROS generation of monocytes attached to EC (Fig. 4A)
. In an earlier study, we reported that PF4-mediated adherence of neutrophils to EC also involves β2-integrins and that these integrins provide a costimulatory signal leading to neutrophil exocytosis [56
]. Furthermore, Elner et al. [57
] described that the induction of apoptosis in human retinal pigment epithelial cells mediated by IFN-
-stimulated monocytes also depends on the interaction of these molecules, and in an early report, Jonjic et al. [58
] showed the essential role of β2-integrins in the IFN-/LPS-induced cytotoxic effect of monocytes on EC. It should be noted that the experimental settings in the latter study were different from those applied by us. Although Jonjic and coworkers [58
] activated and differentiated the monocytes for 20 h with IFN- and LPS before coculture with EC, we used fresh and undifferentiated blood monocytes. Moreover, although we observed a cytotoxic effect already after 15 h of coculture (Fig. 2)
, EC killing was described by Jonjic and coworkers [58
] to occur between 24 h and 48 h of culture, indicating that different cellular mechanisms of cytotoxicity may be involved in the two experimental systems.
Beside oxidative stress, other potential cytotoxic mechanisms of PF4-activated monocytes were explored. As described earlier, the time-course of PF4-induced cytotoxicity parallels that of TNF release [59 ]. However, as neither supernatants of PF4-activated monocytes (Fig. 3) nor stimulated monocytes/EC cocultures in a transwell system led to EC killing, a potential, direct cytotoxic effect by TNF or other soluble factors induced by PF4 in monocytes can be excluded. Furthermore, these findings also eliminate the potential participation of soluble monocyte proteases in PF4-mediated cytotoxicity of EC. This is in line with our observation that protease inhibitors of broad and different specificities were without effect on the killing process.
Cytotoxicity of stimulated monocytes can be targeted against different cell types, for example, tumor cells or epithelial cells [57 , 60 ]. In this work, we have shown that PF4-activated monocytes develop cytotoxicity against EC but not against cells of the lung epithelial cell line A549. Surprisingly, A549 cells express constitutively high amounts of ICAM-1 ([61 ] and our own observations), indicating that a lack of interaction with the corresponding β2-integrins cannot be the reason for their insensitivity to ROS attack. It is possible that survival of lung epithelial cells in their physiological environment requires a higher resistance to oxidative stress than that of blood vessel-derived EC.
So far, the in vivo physiological situation, where PF4-mediated cytotoxicity may play a role, is still speculative. Our observation, that the presence of serum drastically reduces EC killing, argues for strict, local restriction of this potentially harmful, biological effect to regions where serum components are absent. Such serum-free microenvironments may arise through platelet activation, leading to the formation of thrombi, which are normally firmly attached to the vessel wall. Thrombus formation would not only exclude serum-derived inhibitors but as a source for PF4, would also allow the formation of high, local concentrations of the chemokine required to induce monocyte cytotoxicity. As normal serum concentrations of PF4 may reach 0.9–1.9 µM [3
], the amounts released in platelet aggregates can be assumed to be much higher. With regard to this, the relatively high threshold of 
0.125 µM PF4 for the induction of a cytotoxic effect could serve to prevent unintended activation of monocytes by low concentration of the chemokine released from decaying platelets. Together, these mechanisms would strictly localize cytotoxicity and prevent extensive damage of the endothelium. On the other hand, such selective activation of cytotoxic monocytes in close proximity to thrombi could serve to effectively eliminate damaged EC, and removal of intact EC would facilitate the passage of inflammatory leukocytes and mediators, accelerating the formation of an inflammatory focus.
Oxidative stress has been identified as a central, pathogenic factor of many diseases, such as arteriosclerosis, heart attacks, ischemia/reperfusion injury, or chronic inflammatory diseases (reviewed in refs. [62 , 63 ]). Strikingly, practically all of these disorders are accompanied by an acute or chronic activation of thrombocytes. Furthermore, it was shown that in cystathioninsynthase-deficient mice, which display enhanced thrombus formation, injuries of the endothelium are accompanied by enhanced oxidative stress [64 ].
In summary, our data provide evidence that the long-lasting release of ROS by PF4-activated monocytes induces apoptosis selectively in EC. This cytotoxic effect requires direct cell contact between effector and target cells, which is mediated by β2-integrins as well as their corresponding counter ligand. A future goal will be to understand if and how ROS formation induced by mediators of activated platelets is involved in pathological processes. In further studies, we will investigate the role of platelet-induced ROS production in vivo in animal models of atherosclerosis and antibody-induced autoimmune diseases.
Received September 3, 2007; revised December 14, 2007; accepted December 17, 2007.
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