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Am J Physiol Renal Physiol 294: F1381-F1387, 2008. First published April 2, 2008; doi:10.1152/ajprenal.00003.2008
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Increased inorganic phosphate induces human endothelial cell apoptosis in vitro

G. S. Di Marco,1 M. Hausberg,1 U. Hillebrand,1 P. Rustemeyer,2 W. Wittkowski,2 D. Lang,1,* and H. Pavenstädt1,*

1Department of Internal Medicine and 2Institute of Anatomy, University Clinics Muenster, Muenster, Germany

Submitted 3 January 2008 ; accepted in final form 28 March 2008


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Chronic kidney disease with hyperphosphatemia is associated with accelerated atherosclerosis and endothelial dysfunction. However, the contribution of high serum phosphate levels to endothelial injury is incompletely understood. The aim of this work was to evaluate the responses of endothelial cells to elevated levels of extracellular phosphate in vitro. High phosphate in concentrations similar to those observed in uremia-associated hyperphosphatemia (>2.5 mM) induced apoptosis in two endothelial cell lines (EAhy926 cells and GM-7373 cells). This effect was enhanced when cells were incubated for 24 h in the presence of 2.8 mM calcium instead of 1.8 mM. By treating cells with 0.5 or 1.0 mM phosphonoformic acid, an inhibitor of the phosphate transporter, death was completely prevented. The process of phosphate-induced apoptosis was further characterized by increased oxidative stress, as detected by increased ROS generation and disruption of the mitochondrial membrane potential at ~2 h after treatment, followed by caspase activation. These findings show that hyperphosphatemia causes endothelial cell apoptosis, a process that impairs endothelial integrity. Endothelial cell injury induced by high phosphate concentrations may be an initial event leading to vascular complications in patients with chronic kidney disease.

atherosclerosis; ROS; mitochondrial dysfunction; endothelial dysfunction

VASCULAR CHANGES IN CHRONIC kidney disease and end-stage renal failure are associated with increased atherosclerosis, ischemic heart disease, and vascular stiffening. Prospective studies and registry data suggest a strong role of hyperphosphatemia and an increased calcium-phosphate product as significant predictors of cardiovascular mortality, potentially acting as progression factors of unwanted atherosclerosis and calcifications in uremia (26). In addition, it has been very recently shown that an increased phosphate concentration is an independent risk factor for higher mortality during the predialysis phase, suggesting that phosphate levels within the normal range are likely of vital importance in predialysis patients (32).

Moreover, it is now widely accepted that oxidative stress and the ensuing endothelial dysfunction play a key role in the pathogenesis of atherosclerosis and cardiovascular disease. Specifically, endothelial dysfunction caused by an excess of reactive oxygen species (ROS) precedes and promotes atherosclerosis (18, 25). The high turnover of endothelial cells in atherosclerosis also suggests that apoptosis may contribute to the pathology (4). In this context, common risk factors for atherosclerosis are associated with increased generation of ROS and mitochondrial dysfunction, which can lead to activation of the mitochondrial apoptotic pathway (4, 18, 25). Thus mitochondria have been proposed as an important link among risk factors, oxidative damage, endothelial dysfunction, and apoptosis, and the initiation and development of atherosclerotic lesions.

If a number of observations confirm the importance of endothelial cell apoptosis in the pathogenesis of atherosclerosis (8, 14, 25, 29), there are only sparse data on the effects of phosphate, in concentrations as observed in uremia-related hyperphosphatemia, on endothelial cell function and death. Thus, in the present study, we aimed to ascertain whether increased phosphate concentrations alone or in combination with increased calcium concentrations are able to modulate endothelial cell apoptosis and to explore the mechanisms involved in this process.


   MATERIALS AND METHODS
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 MATERIALS AND METHODS
RESULTS
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All experiments were performed under institutional ethical approval of the protocol.

Endothelial cell culture. Human endothelial cells [EAhy926; a permanent human cell line derived from human umbilical vein endothelial cells that express highly differentiated functions characteristic of human vascular endothelium] (1, 5, 6, 24) and bovine aortic endothelial GM-7373 cells (DSMZ, Braunschweig, Germany) (24) were used.

EAhy926 cells were grown in DMEM (Biochrom, Berlin, Germany) containing 5% FCS (PAA Laboratories, Pasching, Austria), 2 mM L-glutamine, and 50 U/ml each of penicillin/streptomycin at 37°C in an atmosphere of 5% CO2 in air. GM-7373 cells were cultured in MEM supplemented with 20% FCS, nonessential amino acids, MEM vitamins, and penicillin/streptomycin as recommended by the manufacturer. Cell cultures between passages 10 and 20 and 8 and 13 were used, respectively. For most of the experiments, cells were seeded in 24-well plates. Twenty-four hours before apoptosis induction, the growth medium was replaced; and just before each specific treatment, cells were rinsed with fresh medium and treated for the indicated times.

Treatment protocols. Medium supplemented with 5% FCS was set as the control (phosphate and calcium concentrations in the culture medium are 1.0 and 1.8 mM, respectively). In treated groups, control medium was supplemented with 133 µl of a stock solution of sodium phosphate (1.0 M Na+; 0.6 M POFormula) and 217 µl of a stock solution of calcium gluconate/calcium saccharate (0.23 M Ca2+) to raise the phosphate concentration to 2.5 mM and calcium concentration to 2.8 mM. Phosphonoformic acid (PFA; 0.1–1.0 mM, Sigma, St Louis, MO), a specific inhibitor of the sodium-phosphate transporter, was used to block phosphate transport into the cells. Cells were incubated in the presence of 10 mM N,N'-dimethyl-thioharnstoff 99%, 1.3-dimethyl-2-thiourea, 99% (DMTU; Sigma-Aldrich), a superoxide scavenger, to block ROS generation. Z-VAD-FMK (50–100 µM, R&D Systems, Minneapolis, MN), a cell-permeable and irreversible general caspase inhibitor, was employed to investigate the involvement of caspases in phosphate- and calcium-phosphate-induced apoptosis; before the addition of apoptogens, cells were incubated for 45 min with Z-VAD-FMK following the manufacture's instructions. When necessary, the bacterial alkaloid staurosporine (STS; Calbiochem, La Jolla, CA), a potent well-known apoptogen, was used at 1.0 µM for 2 h.

RT-PCR. Total RNA was extracted from cells using a QIA shredder followed by an RNeasy mini kit (Qiagen, Valencia, CA). First-strand cDNA was synthesized on 1 µg total RNA in a 10-µl volume containing 1x Moloney murine leukemia virus (M-MuLV) Reverse Transcriptase buffer (Stratagene, Cedar Creek, TX), 5 µM random hexamers, 1 mM dNTP mix, and 25 U M-MuLV Reverse Transcriptase (Stratagene). The reaction was performed for 1 min at 23°C and 60 min at 37°C and stopped by 5-min incubation at 99°C. PCR was performed using one-tenth of the RT reaction in 1x PCR Rxn Buffer (Invitrogen, Paisley, UK), 0.5 µM primers, 2 mM MgCl, 0.4 mM dNTP mix, and 0.5 U Taq DNA Polymerase (Invitrogen). Human Pit-1 cDNA was amplified using the following primer sequences: forward 5'-TACCATCCTCATCTCGGTGG-3' and reverse 5'-TGACGG-CTTGACTGAACTGG-3' (10). PCR products were run on 1.5% agarose gels.

Immunofluorescence microscopy. To investigate possible morphological alterations, confocal fluorescence microscopy was employed. Cells were grown directly on 12-mm coverslips, washed three times with PBS, and fixed with 4% paraformaldehyde in PBS for 10 min at 4°C. Coverslips were washed again with PBS and incubated sequentially with 0.1% Triton in PBS for 10 min and 0.2% gelatin in PBS for 20 min. Cells were then reacted with Alexa Fluor 591-conjugated phalloidin (1:1,000) (Sigma-Aldrich) and a monoclonal antibody to integrin-β1 (1:35) in 0.2% gelatin in PBS (ab3167; Abcam, Cambridge, UK) for 90 min at room temperature. Alexa Fluor 488 secondary antibody was used at 1:500 dilution. 4',6-Diamidino-2'-phenylindol-dihydrochloride (DAPI) was used to analyze the appearance of apoptotic nuclei. Images were generated with a laser-scanning fluorescence microscope (Fluoview, Olympus) or with a fluorescence microscope (Carl Zeiss).

Apoptosis assay: phosphatidylserine externalization. Apoptotic cells were measured by staining with annexin V-FITC and subsequent flow cytometry (12). Briefly, cells were treated for 24 h as described above. Cells were harvested using cold PBS on ice, scraped, and collected by centrifugation. After being washed with staining buffer (PBS with calcium and magnesium supplemented with 0.5% FCS and 0.5% NaN3), cells were incubated with 5 µl annexin V-FITC (BD Biosciences Pharmingen, Erembodegem, Belgium) and 5 µg/ml propidium iodide in 100 µl staining buffer, for 30 min, at 4°C, in the dark. Cells were washed again, resuspended in 250 µl staining buffer, and analyzed by flow cytometry on a FACScan flow cytometer (Becton-Dickinson, Mountain View, CA). Ten thousand events were triggered by the forward-scatter and side-scatter light, and samples were analyzed by using CellQuest software (Becton-Dickinson). Cells stained positive for annexin V-FITC and negative for propidium iodide were considered as apoptotic.

Transmission electron microscopy. To evaluate the morphological features of the treated cells, endothelial cells were treated with 2.5 mM phosphate and 2.8 mM calcium for 24 h as indicated above. The culture dishes were washed twice with PBS, and cells were scraped in the presence of PBS, centrifuged, fixed in 1.25% glutaraldehyde/0.1 M phosphate buffer, and postfixed in 4% OsO4/0.1 M phosphate buffer. Samples were dehydrated in an ascending ethanol series, treated with propylene oxide, and embedded in epoxy resin (Epon 812). Alternating semithin and ultrathin sections were cut. Semithin sections were stained with toluidine blue for gross orientation. Ultrathin sections were contrasted with uranyl acetate and lead citrate, mounted on 150-mesh Formvar-coated copper grids, and examined with a Philips electron microscope (CM 10) (2).

Mitochondrial membrane potential determination. To analyze a disruption of the mitochondrial potential, cells were harvested by scraping cells in cold PBS on ice or by trypsinization and collected by centrifugation. Cells were then incubated with 20 nM tetramethylrhodamine ethyl ester (TMRE: Fluka, Buchs, Switzerland) for 15 min, at 37°C, in PBS containing 130 mM KCl to abolish plasma membrane potential. After incubation, cells were washed once in PBS and analyzed for TMRE red fluorescence by flow cytometry. Five thousand events were analyzed. Live cells rapidly and reversibly take up TMRE, and accumulation of the dye in mitochondria has been shown to be potential driven (23).

ROS generation measurement. The measurement of ROS generation was performed as described by Mone et al. (21) after minor modifications. After treatment with 2.5 mM phosphate in the presence or not of increased calcium concentration, cells were incubated with 2.5 µM dihydroethidine (DHE; Invitrogen) during the last 30 min in culture medium at 37°C. The cells were washed with PBS, harvested by scraping, and collected by centrifugation. After resuspension in PBS, the cells were immediately analyzed by flow cytometry using an FL-3 filter. Gates for intact cells were determined based on light-scattering properties of untreated cells and left unchanged for the analysis of the treated cells.

For in situ detection of ROS, cells were grown directly on 12-mm coverslips. They were then incubated with control medium, 2.5 mM phosphate, and 2.5 mM phosphate in the presence of DMTU. After 75 min, cells were washed two times with PBS and fixed with 4% paraformaldehyde in PBS for 10 min at 4°C. Cells were examined with a fluorescence microscope as described above.

Statistical analysis. Values are means ± SE. Multiple comparisons among groups were carried out by one-way ANOVA with Tukey's significant difference as the post hoc test. A level of P < 0.05 was accepted as statistically significant.


   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
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Phosphate induces apoptosis. We first exposed endothelial cells to phosphate at 2.5 mM, a concentration that correlates with serum levels found in uremia-related hyperphosphatemia. Morphologically, by confocal microscopy, cells exposed for 24 h to high phosphate in the presence of normal (1.8 mM) or increased (2.8 mM) calcium concentration exhibited a shrinkage in size, membrane blebbing, and loss of their attachment to the substratum compared with untreated control cells (1.0 mM phosphate and 1.8 mM calcium) (Fig. 1A). In control cells, integrin-β1 (green fluorescence) is mainly expressed in the filopodia, playing an important role in the attachment of cells to other cells and substratum. In treated cells, we can observe retrieval of the filopodia and loss of cell contact; however, integrins are still expressed in the cell membrane, only presenting a distinct distribution compared with control situation.


Figure 1
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  		Fig. 1. High phosphate concentration induces endothelial cell apoptosis. A: morphological features of cells exposed to 1.0 mM phosphate and 1.8 mM calcium (Control), 2.5 mM phosphate and 1.8 mM calcium (Pi), and 2.5 mM phosphate and 2.8 mM calcium (CaPi). The photographs were taken 24 h after exposure and evidenced changes such as cell shrinkage, rounding, and loss of cell contact with and attachment to the substratum. Images are the result of overlaying F-actin (red fluorescence) and integrin-β1 (green fluorescence) stainings. Magnification: x63. B: effect of phosphate and calcium on phosphatidylserine externalization. Cells were treated for 24 h with increasing phosphate concentration (1.0–7.0 mM) and calcium (1.8 or 2.8 mM). Apoptosis was assessed by annexin V staining followed by flow cytometric analyses. Values are means ± SE (n = 4–6) expressed as control-fold increase. Control values (1 mM phosphate, 1.8 mM calcium) were used to normalize all experimental data. *P < 0.05 compared with control. §P < 0.05 compared with the correspondent 1.8 mM calcium-treated cells. C: 4',6-diamidino-2'-phenylindol-dihydrochloride (DAPI) staining and transmission electron microscopy. Death through apoptosis was further confirmed by the presence of changes in nuclear morphology, such as the presence of pyknotic nuclei, condensed chromatin at the periphery of the nucleus and in the nucleoplasm, apoptotic bodies, and intense vacuolization after 24-h incubation with 2.5 mM phosphate independently of calcium concentration. Magnification: x63 for DAPI staining and x4,600 for electron microscopy.

 
For quantitative assessment of phosphate-induced apoptosis, cells were incubated for 24 h with increasing phosphate concentrations (1–7 mM) and analyzed by means of phosphatidylserine externalization. We observed that physiological levels of phosphate (<1.4 mM) present no effects on endothelial cell death. The phosphate threshold level for inducing apoptosis was 2.5 mM (Fig. 1B); with an increase of almost 40% in the number of apoptotic cells compared with control levels. An increase of >100% could be observed when cells were incubated at concentrations of phosphate >5 mM. Concerning the influence of extracellular calcium, we noted that at a phosphate concentration of 1 or 1.5 mM in the culture medium, an increase in the calcium level from 1.8 to 2.8 mM did not affect the extent of endothelial cell death. However, the elevation of phosphate concentration to levels >2.5 mM and a simultaneous elevation of the calcium concentration from 1.8 to 2.8 mM resulted in a significant increase in apoptosis compared with elevated phosphate concentration only (Fig. 1B).

At an ultramicroscopic level, transmission electron microscopic images of the treated cells provide further evidence that high phosphate concentration (2.5 mM) alone and in combination with elevated calcium (2.8 mM) mediated endothelial cell apoptosis. Following treatment, endothelial cells displayed condensation of the chromatin, intense vacuolization, and numerous secretory vacuoles and ribosomes, as well as the presence of apoptotic bodies (Fig. 1C, bottom). Changes in nuclear morphology can also be observed by DAPI staining (Fig. 1C, top).

Expression of sodium-phosphate transporter on endothelial cells and effect of phosphate transport blockage on phosphate- and calcium-phosphate-induced apoptosis. Figure 2A shows that human endothelial cells (EAhy926) express at the mRNA level at least one kind of phosphate transporter, Pit-1 (sodium-phosphate transporter type III).


Figure 2
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  		Fig. 2. Effect of phosphonoformic acid (PFA) on phosphate-calcium- and phosphate-induced endothelial cell apoptosis. A: Pit-1 expression in endothelial cells. RNA was extracted from control cells, and RT-PCR was performed. Human kidney and human embryonic kidney (HEK) cells were used as the positive control. Lane 1, molecular weight; lanes 2 and 3, human kidney and HEK cells, respectively; lanes 4 and 5, endothelial cells. B: cells were treated with increasing concentrations of PFA in the presence of 5 mM phosphate and 1.8 or 2.8 mM calcium for 24 h. Apoptosis was assessed by annexin V staining followed by flow cytometric analyses. Values are means ± SE (n = 6) expressed as control fold-increase. *P < 0.05 compared with control. C and D: representative density plots showing the percentage of cells presenting positive staining for annexin V and negative staining for propidium iodide Shown in C are plots made after 24-h incubation with control medium (Control), phosphate (Pi), and phosphate in the presence of 1 mM PFA (Pi+PFA), and shown in D are Control, staurosporine (STS; 1 µM), and STS in the presence of 1 mM PFA (STS+PFA). Note that incubation with PFA presented no effect on STS-induced apoptosis, showing that the inhibitory effect of PFA on phosphate-induced apoptosis is related to phosphate transport blockage.

 
To determine then whether apoptosis is phosphate dependent, we incubated cells with PFA, an inhibitor of phosphate transport. Specifically for inhibition studies, cells were treated with high phosphate concentration (5 mM) or with calcium-phosphate (5 mM phosphate, 2.8 mM calcium ), and apoptosis was analyzed in the presence of increasing concentrations (0.1–1 mM) of PFA. Figure 2, B and C, shows that PFA caused a dose-dependent decrease in the percentage of apoptotic cells in phosphate- and calcium-phosphate-treated cells. At 0.5 or 1.0 mM, PFA was able to abolish apoptosis in both treated cell groups (n = 6) (Fig. 2B). Incubation with PFA alone had no effect on cell apoptosis (data not shown).

To ascertain that apoptosis inhibition by PFA was specifically related to phosphate transport, we analyzed the effect of PFA on staurosporine-induced apoptosis. Figure 2D shows that treatment with 1 mM PFA was not able to prevent endothelial cell apoptosis induced by 1 µM staurosporine (n = 6).

Disruption of mitochondrial membrane potential and ROS generation. To explore the mechanism of apoptosis, control and treated cells were probed with TMRE, a mitochondrial membrane voltage dye, and DHE, a ROS reporter. Figure 3A shows that cells treated with phosphate (2.5 mM) or calcium-phosphate (2.5 mM phosphate, 2.8 mM calcium) presented a significant reduction in TMRE uptake (n = 6/group) compared with control (n = 8), evidencing changes in mitochondrial membrane potential 2 h after incubation. As expected, the disruption of mitochondrial membrane potential was preceded by induction of ROS generation. Figure 3B shows that, in the presence of both treatments, the relative levels of ROS started elevating at 60 min after treatment, presenting a great increase at 75 min. The levels of ROS in untreated control cells were unchanged over all of the incubation time periods (n = 7–14). Figure 3C contains typical fluorescence histograms showing that the exposure to 2.5 mM phosphate (in the presence or not of increased calcium concentration) induces a rightward shift of the DHE fluorescence curve, which is avoided in the presence of 1 mM PFA. Figure 3D shows in situ staining of DHE in control and cells treated with 2.5 mM phosphate in the presence or not of DMTU, a superoxide scavenger. Weak superoxide signals were detected in control and DMTU-treated cells, while intense production was observed when cells were treated with a high phosphate concentration for 75 min.


Figure 3
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  		Fig. 3. Effect of phosphate and calcium-phosphate on mitochondrial membrane potential (MMP) and reactive oxygen species (ROS) generation. Endothelial cells were treated with 1.0 mM phosphate and 1.8 mM calcium (Control), 2.5 mM phosphate and 1.8 mM calcium (Pi), and 2.5 mM phosphate and 2.8 mM calcium (CaPi) for different periods of time. A: dissipation of MMP was assessed by tetramethylrhodamine ethyl ester (TMRE) staining followed by flow cytometric analyses. Representative histograms show an increased number of cells with reduced MMP after 2-h incubation with phosphate or calcium-phosphate. Values are means ± SE; n = 8 for control and n = 6 for phosphate- and calcium-phosphate-treated groups. B: time course studies of phosphate- and calcium-phosphate-induced elevation of ROS generation as determined by dihydroethidine (DHE) staining. Relative levels of ROS started elevating after 60-min incubation and presented a great increase at 75 min. Values are means ± SE (n = 7–14) expressed as control fold-increase. *P < 0.05 compared with control. C: representative histograms of ROS after 75-min incubation with phosphate and calcium-phosphate in the presence or not of 1 mM PFA. Note that PFA avoided the induction of ROS generation, presenting the same fluorescence curve as found for control. D: in situ detection of superoxide. Fluorescence microscopic images of endothelial cells labeled with the oxidative dye DHE (red fluorescence when oxidized to ethidium bromide by superoxide) are shown. Weak signals were detected in control and cells treated with 2.5 mM phosphate in the presence of DMTU, a superoxide scavenger, while intense production was observed when cells were treated with 2.5 mM phosphate for 75 min.

 
Effect of caspase inhibition on phosphate- and calcium-phosphate-induced apoptosis and mitochondrial membrane potential. As a subsequent step in the sequence of events related to the mitochondrial apoptotic pathway is the activation of caspases, we preincubated endothelial cells with the pan-caspase inhibitor Z-VAD-FMK for 45 min before adding phosphate and calcium-phosphate to the culture medium. We thus observed that the inhibitor was able to avoid endothelial cell apoptosis as measured by means of phosphatidylserine externalization (n = 6 for all groups) (Fig. 4A).


Figure 4
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  		Fig. 4. Effect of caspase inhibitor on phosphate- and calcium-phosphate-induced endothelial cell apoptosis and MMP. A: endothelial cells were treated with increasing concentrations of Z-VAD-FMK for 45 min before addition of 5 mM phosphate (Pi) in the presence of 1.8 or 2.8 mM calcium for 24-h. Apoptosis was assessed by annexin V staining followed by flow cytometric analyses. Note that blocking caspase activation prevented phosphatidylserine exposure, indicating that phosphate and calcium-phosphate trigger caspase-dependent pathways in endothelial cells. B: dissipation of MMP was assessed by TMRE staining followed by flow cytometric analyses. Endothelial cells were treated with 5 mM phosphate alone or in the presence of 100 µM Z-VAD-FMK or 1 mM PFA. Values are means ± SE expressed as control-fold increase. For apoptosis studies, n = 6 in all groups; for MMP evaluation, n = 8 for control and phosphate-treated groups and n = 5 for Z-VAD-FMK- and PFA-treated groups. *P < 0.05 compared with control.

 
In contrast, under the same experimental condition, we found that Z-VAD-FMK did not avoid the disruption of mitochondrial membrane potential induced by high phosphate concentrations (Fig. 4B, n = 5 for Z-VAD-FMK-treated group and n = 8 for control and phosphate-treated groups). The mitochondrial membrane depolarization was only avoided when cells were incubated in the presence of 1 mM PFA (Fig. 4B, n = 5). These two lines of data suggest that caspase activation, which is responsible for phosphatidylserine externalization, is a downstream event resulting from ROS and disruption of mitochondrial function.


   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Endothelial cell dysfunction and apoptosis are crucial events in the pathogenesis of atherosclerosis. Apoptosis of endothelial cells in atherosclerosis may be caused by the generation of oxidative stress and/or inflammatory mediators, for example (18, 25, 27). In uremia, hyperphosphatemia is highly correlated with the extent of vascular calcification and vascular disease (15–17). Moreover, the effect of hyperphosphatemia may not be limited to renal disease patients because, also in patients with normal renal function and coronary heart disease, the serum phosphate concentration has been suggested to be a strong predictor of the severity of vascular constriction, suggesting a more general role of serum phosphate in the development of coronary plaques (22).

Because endothelial cells first sense phosphate variations occurring in the plasma and may respond initially by modulation of their functions, we examined the role of high phosphate concentrations in endothelial cell injury. We demonstrated here that phosphate induces endothelial cell apoptosis, an effect that is substantially enhanced by increased calcium concentrations, as verified from morphological observations and the assessment of phosphatidylserine externalization by FACS analysis. The induction of apoptosis by high phosphate is not only the effect of binding and precipitation of calcium, since acute effects of hyperphosphatemia on cell shrinkage and stiffening can be discriminated from apoptosis (Hillebrand U, Lang D, unpublished observations). In addition, our experiments showed disruption of mitochondrial function, via the observed elevation of ROS production, and subsequent caspase activation. The time course studies showed that ROS generation and disruption of the mitochondrial membrane potential occurred as early as 75–120 min post-exposure to high phosphate or calcium-phosphate concentrations, indicating that these events were earlier than that of the apoptotic execution phase, such as phosphatidylserine externalization. In addition, we found that all of these events were inhibited in the presence of PFA, a well-known phosphate transport blocker, suggesting that an increase in the intracellular phosphate concentration is a primary and essential death signal.

In this study, we chose to use a permanent human cell line (EAhy926) derived from human umbilical vein endothelial cells because 1) it is extensively described in the literature that these cells express highly differentiated functional characteristic of human vascular endothelium (1, 5, 6); and 2) they can be used as a homogeneous experimental cell line, which permits more consistent responses to specific variables and greater reproducibility of data. Moreover, using bovine aortic endothelial cells (GM-7373), we were able to confirm that the responses to high phosphate (or calcium-phosphate) observed here probably reflect endothelial behavior, at least at in vitro level.

Since the intracellular concentrations of phosphate and calcium are regulated by the mitochondria, and mitochondrial dysfunction is known to trigger apoptosis (7, 33), we examined mitochondrial function in phosphate- and calcium-phosphate-treated endothelial cells. Indeed, ROS, which are the mitochondrial by-products of normal cellular oxidative processes, have been suggested as regulating the process involved in the initiation of mitochondrial apoptotic signaling (30).

In this study, we demonstrate that phosphate and calcium-phosphate exposure results in an elevation of ROS generation in a time dependent manner. However, it is not exactly known how these treatments could directly induce ROS generation. As it occurs before there is any evidence of apoptosis, including any noticeable disruption in the mitochondrial membrane potential, it is unlikely to be produced by mitochondrial dysfunction. However, as ROS generation was blocked in the presence of PFA, it is evident here that an increase in ROS is related to the accumulation of phosphate in the cells and that ROS accumulation will lead to oxidative stress.

However, these phenomena, increased ROS generation and mitochondrial dysfunction, seem to be a normal response of mitochondria to increased intracellular phosphate. A number of studies show that isolated mitochondria treated with elevated phosphate concentrations present irreversible changes in mitochondrial oxidative-phosphorylating activity, inducting common steps in the apoptosis pathway (3, 9, 11).

Thus, to prove that caspase activation was also involved in phosphate- and calcium-phosphate-mediated apoptosis, we used the cell-permeable caspase inhibitor Z-VAD-FMK. This inhibitor avoided apoptosis in endothelial cells, indicating that phosphate and calcium-phosphate trigger caspase-dependent pathways in these cells. However, caspase inhibition was not able to prevent the loss of mitochondrial membrane potential, confirming that caspase activation is downstream of ROS generation and mitochondrial dysfunction in phosphate-induced apoptosis in endothelial cells (13). Our data support the concept that mitochondrial damage represents a key step in the course of apoptosis induced by phosphate and calcium-phosphate.

Finally, the use of PFA provided interesting insights into the mechanism of phosphate action. At 0.5 or 1.0 mM PFA, endothelial cells were protected against phosphate-induced apoptosis, suggesting that phosphate, as well as phosphate in the presence of increased calcium concentrations, mediates apoptosis through a plasma membrane transport mechanism. These results are in line with previous studies showing that phosphate and the ion pair calcium-phosphate are a potent apoptogen for osteoblasts, chondrocytes, and vascular smooth muscle cells and that PFA was able to block phosphate-induced apoptosis in all these different cell types (19, 20, 28). Herein, as we have not aimed to determine the specific requirement for Pit-1, but rather the effect of decreased phosphate uptake on endothelial cell apoptosis, we opted to use PFA instead of the RNA interference approach. As recently published by Villa-Bellosta et al. (31), RNA interference for Pit-1 in vascular smooth muscle cells correlates with only a 23% inhibition of total phosphate uptake, suggesting that this small amount of transport inhibition observed is enough to bring the phosphate transport beneath a "threshold" such that the osteogenic signaling pathways are not activated. This would explain then the prevention of smooth muscle cell calcification with PFA, despite the fact that PFA is described as a weak inhibitor of type III phosphate transporters. The same line of thought could be applied to our findings, which support the idea that exposure to high extracellular phosphate concentration causes a rise in the intracellular phosphate level, leading to endothelial cell apoptosis.

In summary, human endothelial cells challenged with high phosphate and calcium-phosphate concentrations present increased ROS generation, mitochondrial membrane depolarization, and, consequently, caspase activation. Since endothelial cell apoptosis could be blocked by PFA, it is likely that phosphate mediates cell death by raising the intracellular phosphate concentration and activating downstream effectors of the apoptotic process.

Thus it is conceivable that hyperphosphatemia, by inducing endothelial cell apoptosis, may provide an early contribution to atherogenesis. Indeed, additional insights into the molecular basis of endothelial cell apoptosis could facilitate specific therapeutic interventions slowing the progression of uremia-associated vascular disease.


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This study was supported by a grant from the Else-Kroener Fresenius Foundation (P37/2004) and by Novartis.


   ACKNOWLEDGMENTS
 
The authors thank Petra Haussmann and Katrin Beul for general technical assistance.


   FOOTNOTES
 

Address for reprint requests and other correspondence: G. S. Di Marco, Experimentelle Nephrologie, Universitätsklinikum Münster, Domagkstraβe 3a, D48149 Münster, Germany (e-mail: giodimarco{at}gmail.com)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

* D. Lang and H. Pavenstädt contributed equally to this work. Back


   REFERENCES
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 DISCUSSION
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  1. Ben-Yosef Y, Lahat N, Shapiro S, Bitterman H, Miller A. Regulation of endothelial matrix metalloproteinase-2 by hypoxia/reoxygenation. Circ Res 90: 784–791, 2002.[Abstract/Free Full Text]
  2. Bock N, Bockers TM, Bockmann J, Nowak P, Buse E, Wittkowski W. The Pars tuberalis of the monkey (Macaca fascicularis) hypophysis: cell types and hormone expression. Cells Tissues Organs 169: 55–63, 2001.[CrossRef][Web of Science][Medline]
  3. Bravo C, Chavez E, Rodriguez JS, Moreno-Sanchez R. The mitochondrial membrane permeability transition induced by inorganic phosphate or inorganic arsenate. A comparative study. Comp Biochem Physiol B Biochem Mol Biol 117: 93–99, 1997.[CrossRef][Medline]
  4. Davidson SM, Duchen MR. Endothelial mitochondria: contributing to vascular function and disease. Circ Res 100: 1128–1141, 2007.[Abstract/Free Full Text]
  5. Edgell CJ, Haizlip JE, Bagnell CR, Packenham JP, Harrison P, Wilbourn B, Madden VJ. Endothelium specific Weibel-Palade bodies in a continuous human cell line, EAhy926. In Vitro Cell Dev Biol 26: 1167–1172, 1990.[CrossRef]
  6. Edgell CJ, McDonald CC, Graham JB. Permanent cell line expressing human factor VIII-related antigen established by hybridization. Proc Natl Acad Sci USA 80: 3734–3737, 1983.[Abstract/Free Full Text]
  7. Green DR, Reed JC. Mitochondria and apoptosis. Science 281: 1309–1312, 1998.[Abstract/Free Full Text]
  8. Inoue T, Node K. Vascular failure: a new clinical entity for vascular disease. J Hypertens 24: 2121–2130, 2006.[Web of Science][Medline]
  9. Jassem W, Heaton ND. The role of mitochondria in ischemia/reperfusion injury in organ transplantation. Kidney Int 66: 514–517, 2004.[CrossRef][Web of Science][Medline]
  10. Jono S, McKee MD, Murry CE, Shioi A, Nishizawa Y, Mori K, Morii H, Giachelli CM. Phosphate regulation of vascular smooth muscle cell calcification. Circ Res 87: E10–E17, 2000.[Web of Science][Medline]
  11. Kowaltowski AJ, Castilho RF, Grijalba MT, Bechara EJ, Vercesi AE. Effect of inorganic phosphate concentration on the nature of inner mitochondrial membrane alterations mediated by Ca2+ ions. A proposed model for phosphate-stimulated lipid peroxidation. J Biol Chem 271: 2929–2934, 1996.[Abstract/Free Full Text]
  12. Lang D, Dohle F, Terstesse M, Bangen P, August C, Pauels HG, Heidenreich S. Down-regulation of monocyte apoptosis by phagocytosis of platelets: involvement of a caspase-9, caspase-3, and heat shock protein 70-dependent pathway. J Immunol 168: 6152–6158, 2002.[Abstract/Free Full Text]
  13. Ling YH, Liebes L, Zou Y, Perez-Soler R. Reactive oxygen species generation and mitochondrial dysfunction in the apoptotic response to Bortezomib, a novel proteasome inhibitor, in human H460 non-small cell lung cancer cells. J Biol Chem 278: 33714–33723, 2003.[Abstract/Free Full Text]
  14. Littlewood TD, Bennett MR. Apoptotic cell death in atherosclerosis. Curr Opin Lipidol 14: 469–475, 2003.[CrossRef][Web of Science][Medline]
  15. London GM. Cardiovascular calcifications in uremic patients: clinical impact on cardiovascular function. J Am Soc Nephrol 14: S305–S309, 2003.[Abstract/Free Full Text]
  16. London GM. Vascular disease and atherosclerosis in uremia. Nefrologia 25, Suppl 2: 91–95, 2005.[Web of Science][Medline]
  17. London GM, Marchais SJ, Guerin AP, Metivier F. Arteriosclerosis, vascular calcifications and cardiovascular disease in uremia. Curr Opin Nephrol Hypertens 14: 525–531, 2005.[Web of Science][Medline]
  18. Madamanchi NR, Runge MS. Mitochondrial dysfunction in atherosclerosis. Circ Res 100: 460–473, 2007.[Abstract/Free Full Text]
  19. Mansfield K, Teixeira CC, Adams CS, Shapiro IM. Phosphate ions mediate chondrocyte apoptosis through a plasma membrane transporter mechanism. Bone 28: 1–8, 2001.[Medline]
  20. Meleti Z, Shapiro IM, Adams CS. Inorganic phosphate induces apoptosis of osteoblast-like cells in culture. Bone 27: 359–366, 2000.[Medline]
  21. Mone AP, Huang P, Pelicano H, Cheney CM, Green JM, Tso JY, Johnson AJ, Jefferson S, Lin TS, Byrd JC. Hu1D10 induces apoptosis concurrent with activation of the AKT survival pathway in human chronic lymphocytic leukemia cells. Blood 103: 1846–1854, 2004.[Abstract/Free Full Text]
  22. Narang R, Ridout D, Nonis C, Kooner JS. Serum calcium, phosphorus and albumin levels in relation to the angiographic severity of coronary artery disease. Int J Cardiol 60: 73–79, 1997.[CrossRef][Web of Science][Medline]
  23. Narvaez CJ, Welsh J. Role of mitochondria and caspases in vitamin D-mediated apoptosis of MCF-7 breast cancer cells. J Biol Chem 276: 9101–9107, 2001.[Abstract/Free Full Text]
  24. Oberleithner H, Riethmuller C, Schillers H, MacGregor GA, de Wardener HE, Hausberg M. Plasma sodium stiffens vascular endothelium and reduces nitric oxide release. Proc Natl Acad Sci USA 104: 16281–16286, 2007.[Abstract/Free Full Text]
  25. Puddu P, Puddu GM, Galletti L, Cravero E, Muscari A. Mitochondrial dysfunction as an initiating event in atherogenesis: a plausible hypothesis. Cardiology 103: 137–141, 2005.[CrossRef][Web of Science][Medline]
  26. Reynolds JL, Joannides AJ, Skepper JN, McNair R, Schurgers LJ, Proudfoot D, Jahnen-Dechent W, Weissberg PL, Shanahan CM. Human vascular smooth muscle cells undergo vesicle-mediated calcification in response to changes in extracellular calcium and phosphate concentrations: a potential mechanism for accelerated vascular calcification in ESRD. J Am Soc Nephrol 15: 2857–2867, 2004.[Abstract/Free Full Text]
  27. Simionescu M. Implications of early structural-functional changes in the endothelium for vascular disease. Arterioscler Thromb Vasc Biol 27: 266–274, 2007.[Abstract/Free Full Text]
  28. Son BK, Kozaki K, Iijima K, Eto M, Nakano T, Akishita M, Ouchi Y. Gas6/Axl-PI3K/Akt pathway plays a central role in the effect of statins on inorganic phosphate-induced calcification of vascular smooth muscle cells. Eur J Pharmacol 556: 1–8, 2007.[CrossRef][Web of Science][Medline]
  29. Stoneman VE, Bennett MR. Role of apoptosis in atherosclerosis and its therapeutic implications. Clin Sci (Lond) 107: 343–354, 2004.[Medline]
  30. Tan S, Sagara Y, Liu Y, Maher P, Schubert D. The regulation of reactive oxygen species production during programmed cell death. J Cell Biol 141: 1423–1432, 1998.[Abstract/Free Full Text]
  31. Villa-Bellosta R, Bogaert YE, Levi M, Sorribas V. Characterization of phosphate transport in rat vascular smooth muscle cells: implications for vascular calcification. Arterioscler Thromb Vasc Biol 27: 1030–1036, 2007.[Abstract/Free Full Text]
  32. Voormolen N, Noordzij M, Grootendorst DC, Beetz I, Sijpkens YW, Manen JG, Boeschoten EW, Huisman RM, Krediet RT, Dekker FW. High plasma phosphate as a risk factor for decline in renal function and mortality in pre-dialysis patients. Nephrol Dial Transplant 22: 2909–2916, 2007.[Abstract/Free Full Text]
  33. Zamzami N, Marchetti P, Castedo M, Zanin C, Vayssiere JL, Petit PX, Kroemer G. Reduction in mitochondrial potential constitutes an early irreversible step of programmed lymphocyte death in vivo. J Exp Med 181: 1661–1672, 1995.[Abstract/Free Full Text]



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