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(1991), as the adverse health effects of lead poisoning become widely recognized (1). Lead poisoning can directly cause peripheral neuropathy and...
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Chem. Res. Toxicol. 2007, 20, 38-43

Lead-Induced Procoagulant Activation of Erythrocytes through Phosphatidylserine Exposure May Lead To Thrombotic Diseases Jung-Hun Shin, Kyung-Min Lim, Ji-Yoon Noh, Ok-Nam Bae, Seung-Min Chung, Moo-Yeol Lee, and Jin-Ho Chung* College of Pharmacy, Seoul National UniVersity, Seoul 151-742, Korea ReceiVed May 26, 2006

Lead (Pb) is a ubiquitous heavy metal pollutant in various environmental media, especially in food and drinking water. In human blood, about 95% of lead is associated with erythrocytes, suggesting that erythrocytes could be an important target of lead toxicity in the cardiovascular system. Recent studies suggested that erythrocytes could contribute to blood coagulation via phosphatidylserine (PS) exposure and resultant procoagulant activation. We investigated the effects of lead on the procoagulant activity of erythrocytes using in Vitro human erythrocyte and in ViVo rat models. In a flow cytometric analysis, lead (Pb2+) enhanced PS exposure on human erythrocytes in a concentration- and time-dependent manner. The concentration of lead (1-5 µM) used in the current investigation is well within the ranges observed in blood from lead-exposed populations. PS exposure by lead appeared to be mediated by increased intracellular calcium levels as shown by 19F-NMR and intracellular ATP depletion. Consistent with these findings, the activity of scramblase, which is important in the induction of PS exposure, was enhanced, whereas the activity of flippase, which translocates exposed PS to inner membrane, was inhibited by lead treatment. Furthermore, lead-exposed erythrocytes increased thrombin generation as determined by a prothrombinase assay and accelerated the coagulation process initiated by tissue factor in plasma. These procoagulant activations by lead were also confirmed in ViVo. Administration of lead significantly enhanced PS exposure on erythrocytes and, more importantly, elevated thrombus formation in a rat venous thrombosis model. These results suggest that lead exposure can provoke procoagulant activity in erythrocytes by PS exposure, contributing to enhanced clot formation. These data will provide new insights into the mechanism of lead-induced cardiovascular diseases. Introduction Lead is one of the most abundant heavy metals on earth. It has been widely used throughout human history, posing a serious health problem to susceptible populations, such as children or occupationally exposed people. The Center for Disease Control (CDC) defines lead poisoning as the blood lead level (BLL1) exceeding 10 µg/dL, whereas the normal BLL is less than 5 µg/dL. This advisory BLL has been continually declining over the past few decades, from 60 µg/dL (1960-1970), to 30 µg/ dL (1970-1985), 25 µg/dL (1985-1991), and to 10 µg/dL (1991), as the adverse health effects of lead poisoning become widely recognized (1). Lead poisoning can directly cause peripheral neuropathy and chronic renal disease. The cardiovascular system, gastrointestinal tract, reproductive tissue and skeletal tissues can also be affected by lead exposure (2, 3). Individuals with BLLs of 20 to 29 µg/ dL in 1976 to 1980 (15% of the US population at that time) experienced significantly increased circulatory, cardiovascular, and total mortality from 1976 through 1992 (4). The most wellknown cardiovascular complication of lead poisoning is hyper* Corresponding author. Tel: +82-2-880-7856. Fax: +82-2-885-4157. E-mail: [email protected]. 1 Abbreviations: ACD, acid citrate dextrose; BLL, blood lead level; C6-NBD-PC, 1-oleoyl-2-[6-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]hexanoyl]-sn-glycero-3-phosphocholine; C6-NBD-PS, 1-palmitoyl-2-[6-[(7nitro-2-1,3-benzoxadiazol-4-yl)amino]hexanoyl]-sn-glycero-3-phospho-Lserine; 5F-BAPTA AM, 5,5′-difluoro BAPTA acetoxymethyl ester; MV, microvesicles; PBS, phosphate buffered saline; PS, phosphatidylserine; TBS, Tris buffered saline.

tension (5, 6). The link between hypertension and lead poisoning is confirmed by an in ViVo animal model (7). Lead is also known to induce a procoagulant state through suppression of tissue plasminogen activator release from endothelial cell and resultant retardation of fibrinolysis (8). In addition, some epidemiological studies reported the connection between BLL and an increased prevalence of peripheral arterial diseases (9). The diversity of cardiovascular complications associated with chronic lead exposure raises the possibility that lead may induce the procoagulant activity of blood cells directly leading to thrombosis. In human blood, about 95% of lead is accumulated in erythrocytes (3) suggesting that erythrocytes could be an important target of lead toxicity in the cardiovascular system (10). In erythrocytes, lead could degenerate the lipid and protein component (11), suppress hemoglobin synthesis (12, 13), induce lipid peroxidation (14, 15), inhibit superoxide dismutase, and lower the intracellular glutathione level (16, 17). Lead is also known to induce shape change in erythrocytes from biconcave normocyte to echinocyte (18). Erythrocytes constitute the majority of cellular blood components and act as oxygen carriers. Although they were commonly believed in the past to be inert in hemostasis and thrombosis, recent studies showed that erythrocytes can contribute to thrombus formation (19) and have effects on hemostasis through interaction with endothelial cells, platelets, and macrophages (20). It has been reported that alterations in the erythrocyte membrane, such as phosphatidylserine (PS) externalization, and PS-bearing microvesicle (MV) generation enable

10.1021/tx060114+ CCC: $37.00 © 2007 American Chemical Society Published on Web 01/16/2007

Procoagulant ActiVation of Erythrocytes by Lead

erythrocytes to participate in thrombus formation (21). PS exposed on erythrocytes provides a site for the assembly of the prothrombinase and tenase complex, leading to thrombin generation and clotting (21, 22). Furthermore, erythrocytes that express PS on the outer surface become more adhesive to endothelial cells, contributing to vaso-occlusion (23). In addition, PS-bearing microvesicles shed from the erythrocyte membrane are also shown to have procoagulant activity (24, 25). Very recently, it was reported that PS exposure could be induced in erythrocytes by lead ions (18). However, the biological significance of PS exposure was not pursued. We investigated whether PS exposure or MV generation could be induced by lead in human erythrocytes and if lead-exposed erythrocytes could enhance thrombin generation in Vitro. Finally, the clinical relevancy of lead exposure to procoagulant activation was examined using an in ViVo rat model.

Experimental Procedures Materials. Lead(II) acetate, the ATP bioluminescent assay kit, and purified human thrombin were purchased from Sigma Chemical Co. (St. Louis, MO). 5,5′-Difluoro BAPTA acetoxymethyl ester (5F-BAPTA AM) was from Molecular Probes (Eugene, OR), D2O was from Merck (Darmstadt, Germany), fluorescein-isothiocyanate (FITC)-labeled annexin V (annexin V-FITC) was obtained from Pharmingen (San Diego, CA), and phycoerythrin-labeled monoclonal antibody against human glycophorin A (anti-glycophorinA-RPE) was from Dako Cytomation (Glostrub, Denmark). 1-Palmitoyl-2-[6-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]hexanoyl]-snglycero-3-phospho-L-serine (C6-NBD-PS) and 1-oleoyl-2-[6-[(7nitro-2-1,3-benzoxadiazol-4-yl)amino]hexanoyl]-sn-glycero-3phosphocholine (C6-NBD-PC) were from Avanti Polar Lipids (Alabaster, AL). Purified human prothrombin, factor Xa and factor Va were from Hematologic Technologies, Inc. (Essex Junction, VT), and S2238 was from Chromogenix (Milano, Italy). Human recombinant tissue factor (Recombiplastin) was obtained from Instrumentation Laboratory (Lexington, MA) and thromboplastin (Simplastin Excel) was from Biomerieux (Durham, NC). Preparation of Erythrocytes. With the approval from the Ethics Committee of Health Service Center at Seoul National University, human blood was obtained from healthy male donors (18-25 years old) using a vacutainer with acid citrate dextrose (ACD) and a 21 gauge needle (Becton Dickinson, U.S.A.) on the day of each experiment. Platelet rich plasma and buffy coat were removed by aspiration after centrifugation at 200g for 15 min. Packed erythrocytes were washed 3 times with phosphate buffered saline (PBS: 1.06 mM KH2PO4, 154 mM NaCl, 2.96 mM and Na2HPO4 at pH 7.4) and once more with Tris buffered saline (TBS: 15 mM Tris-HCl, 150 mM NaCl, 5 mM KCl, and 2 mM MgCl2 at pH 7.4). Washed erythrocytes were resuspended in TBS buffer to a cell concentration of 5 × 107 cells/mL, and the final CaCl2 concentration was adjusted to 1 mM prior to use. Flow Cytometric Analysis of Phosphatidylserine Exposure and Microvesicle Generation. Annexin V-FITC was used as a marker for phosphatidylserine (PS) detection, whereas anti-glycophorin A-RPE was used as an identifier of erythrocytes. Negative controls for annexin V binding were stained with annexin V-FITC in the presence of 2.5 mM EDTA instead of 2.5 mM CaCl2. Samples were analyzed on the flow cytometer FACScalibur (Becton Dickinson, U.S.A.) equipped with an argon-ion laser emitting at 488 nm. The light scatter and fluorescence channels were set on a log scale. Data from 20,000 events were collected and analyzed using CellQuest Pro software. Microvesicles (MV) were identified on the basis of forward scatter characteristics after calibration by 1 µm standard beads. Measurement of Cytosolic Calcium in Erythrocytes. Isolated erythrocytes from heparinized blood were washed three times with Na+ wash buffer (145 mM NaCl, 5 mM HEPES, and Tris-base at pH 7.4) and once more with a loading buffer (145 mM NaCl, 5

Chem. Res. Toxicol., Vol. 20, No. 1, 2007 39 mM sodium pyruvate, 10 mM HEPES, 95% O2/5% CO2 bubbling, and Tris-base at pH 7.4), as described previously (26). Cell counts were adjusted to 5 × 107 cells/mL with loading buffer, and CaCl2 was added to a final concentration of 1.25 mM. After the Pb2+ or deionized water addition, erythrocyte suspension was incubated for 1 h and spun at 500g for 5 min. The resultant erythrocyte pellet was washed three times with CaCl2 added to the loading buffer. The final erythrocyte pellet was resuspended again with loading buffer to be 10-fold concentrated and loaded with 50 µM 5FBAPTA AM by incubation at 37 °C for 20 min. Subsequently, the cells were washed once with loading buffer and incubated at 37 °C for 1 h. The cells were washed and concentrated stepwise to 4-fold concentrated and treated with D2O saline. Intracellular calcium was measured using 19F-NMR (Bruker DRX 500 1H/19F Dual Probehead spectrometer, Germany). One pulse sweep width was (100,000 Hz, and acquisition time was 655.4 ms. Delay time was 1 s, and each spectrum acquisition was 32,768. All experiments were performed at 25 °C under spinning conditions. D2O saline was added for field/frequency locking. Phospholipid Translocation Measurement. Erythrocytes (1 × 108 cells/mL) were incubated with Pb2+ for 30 min at 37 °C and then loaded with 0.5 µM C6-NBD-PS or C6-NBD-PC. Aliquots from the cell suspension were removed at the indicated time intervals and placed on ice for 20 min in the presence or absence of 1% bovine serum albumin (BSA). Pellets obtained after 1 min of centrifugation at 12,000g were lyzed in 1% Triton X-100, and the fluorescence intensities were measured (λex 485 nm, λem 535 nm). The amount of internalized probe was determined by comparing the fluorescence intensity associated with the cells before and after back-exchange. Intracellular ATP Level Measurement. After incubation with Pb2+ for 4 h, the erythrocyte suspension was centrifuged at 3,000g for 1 min. The resultant erythrocyte pellet was washed and resuspended in TBS containing 1 mM CaCl2. The aliquot was mixed vigorously with 10% TCA in Tris-acetate buffer (3:2) and cooled in ice for 20 min. The sample was centrifuged at 13,000g for 2 min, and the aliquot of the resultant supernatant was mixed with cold Tris-acetate buffer (100 mM Tris-acetate and 2 mM EDTA at pH 7.8). ATP was quantified by a chemiluminescence method using an ATP bioluminescent assay kit according to the instruction manual. Chemiluminescence was measured with AutoLumat LB953 (Berthold, Germany). The ATP level was expressed as % of control erythrocytes. Prothrombinase Assay. After incubation with Pb2+ for 4 h, the erythrocyte suspension was centrifuged, and the resultant erythrocyte pellet was washed twice with TBS. After the pellet was resuspended to a cell concentration of 5 × 107 cells/mL, an aliquot was incubated with 5 nM factor Xa and 10 nM factor Va in Tyrode buffer (134 mM NaCl, 10 mM HEPES, 5 mM glucose, 2.9 mM KCl, 1 mM MgCl2, 12 mM NaHCO3, 0.34 mM Na2HPO4, 0.3% BSA, and 2 mM CaCl2 at pH 7.4) for 3 min at 37 °C. Thrombin formation was initiated by addition of 2 µM prothrombin. Exactly 3 min after the addition of prothrombin, an aliquot of the suspension was transferred to a tube containing EDTA stop buffer (50 mM Tris-HCl, 120 mM NaCl, and 2 mM EDTA at pH 7.9). Thrombin activity was determined using the chromogenic substrate S2238. The rate of thrombin formation was calculated from the change in absorbance at 405 nm, using a calibration curve generated with active site-titrated thrombin. Measurement of Thrombin Generation in Plasma. Thrombin generation in plasma was measured according to the methods previously described (27). TBS or erythrocytes were added to plasma, and the mixture was pre-warmed for 2 min at 37 °C. Thrombin formation was initiated under gentle magnetic stirring by adding Recombiplastin diluted (1:3200) in TBS containing 100 mM CaCl2. The aliquots were collected at the indicated time and transferred to a tube containing TBS with 20 mM EDTA. Thrombin activity was determined using the chromogenic substrate S2238, as previously described in the prothrombinase assay. Detection of PS Exposure Following Lead Administration in Rats. Male Sprague-Dawley rats were obtained from Dae Han

40 Chem. Res. Toxicol., Vol. 20, No. 1, 2007 BioLink Co. (Chungbuk, Korea) and were housed in groups of four or five, for at least 1 week prior to the experiment. The animals were fed a standard laboratory diet from Purina Korea and had access to food and water ad libitum. At the time of experiment, rats (250-300 g) were randomly grouped for control (saline), 25, 50, and 100 mg/kg lead(II) acetate doses. Four hours after lead(II) acetate or saline was administered intraperitoneally, blood was collected from the abdominal aorta using 3.8% trisodium citrate as anticoagulant. An aliquot of the blood sample was diluted 200fold with the following buffer (10 mM HEPES-Na, 136 mM NaCl, 2.7 mM KCl, 2.0 mM MgCl2, 1.0 mM NaH2PO4, 5.0 mM dextrose, 5 mg/mL BSA, and 2.5 mM CaCl2 at pH 7.4) and was stained with annexin V-FITC for 15 min in the dark. PS exposure was measured as described above. Venous Thrombosis Animal Model. Thrombus formation was induced by stasis combined with hypercoagulability. Male Sprague-Dawley rats (300-400 g) were anesthetized with urethane (1.25 g/kg, i.p.). The abdomen was surgically opened, and the vena cava was exposed after careful dissection. Two loose cotton threads were prepared 16 mm apart around the vena cava. All side branches were ligated tightly with cotton threads. One hour after the intravenous injection of saline or lead(II) acetate (10 or 25 mg/kg) into a left femoral vein, 500-fold diluted thromboplastin was infused for 1 min to induce thrombus formation. Stasis was initiated by tightening the two threads, first the proximal and the distal thereafter. The abdominal cavity was provisionally closed, and blood stasis was maintained for 15 min. After the abdomen was reopened, the ligated venous segment was excised and opened longitudinally to remove the thrombus. The isolated thrombus was blotted of excess blood and immediately weighed. Statistical Analysis. Data are expressed as mean ( SEM. The difference between the two groups was evaluated using the Student’s t-test. When more than three groups were compared, the ANOVA test was conducted, followed by Duncan’s multiple range test. Significance was acknowledged when the p value was less than 0.05.

Results Phosphatidylserine (PS) exposure and microvesicle (MV) generation were measured by flow cytometry analysis after erythrocytes were incubated with Pb2+ (lead acetate) or deionized water for 4 h at 37 °C. PS exposure increased in a concentration-, and time-dependent manner over the concentration range 1-5 µM Pb2+; the maximum increase being 26.8 ( 3.15% (Figure 1A). This finding agrees with a previous report using lead(II) nitrate (18). No significant generation of microvesicles was observed following a 4 h exposure to Pb2+ at concentrations ranging up to 5 µM (Figure 1B). There was no hemolysis at the concentration of Pb2+ up to 5 µM (data not shown). Because a calcium-dependent pathway is considered to be a major pathway for PS exposure in erythrocytes, we investigated whether intracellular calcium can be increased in erythrocytes by exposure to Pb2+. To avoid the interference by Pb2+, 19FNMR was adopted with 5F-BAPTA AM for analysis of Ca2+ in this study (28). After erythrocytes were treated with 5 µM Pb2+ for 1 h at 37 °C, the level of intracellular Ca2+ was measured. The greatest concentration of the free form of 5FBAPTA was observed at 0 ppm, whereas peak concentrations of Ca2+- and Pb2+-bound 5F-BAPTA appeared at 4.8 and 4.0 ppm, respectively. Intracellular calcium was increased by treatment of Pb2+ (6.88 ( 1.13 µM for Pb2+ vs 0.24 ( 0.21 µM for control) (Figure 2). An increase in intracellular calcium levels can activate scramblase, which can induce PS exposure and inhibit flippase, resulting in maintenance of the exposed PS on the outer membrane. A 30 min pretreatment interval was selected for this

Shin et al.

Figure 1. Effects of Pb2+ on phosphatidylserine exposure and microvesicle generation in human erythrocytes. After erythrocytes were treated with DW (vehicle) or 1, 2, and 5 µM Pb2+ (lead(II) acetate) for 4 h at 37 °C, the extent of phosphatidylserine (PS) exposure and microvesicle (MV) generation were measured by flow cytometry. (A) Percentage of cells exposing PS following Pb2+ treatment. (B) (Left) Erythrocytes and MV identified by forward scatter characteristics in a representative dot plot; (Right) number of MV released from erythrocytes. Values are the mean ( SEM of three to four independent experiments. (*) Significantly different from the control (p < 0.05).

Figure 2. Increase of intracellular Ca2+ level by Pb2+ in human erythrocytes. After erythrocytes were treated with DW (vehicle) or 5 µM Pb2+ for 1 h at 37 °C, the level of intracellular Ca2+ was measured as described in Experimental Procedures. The representative 19F-NMR charts and the level of intracellular Ca2+ are shown. Values are the mean ( SEM of two independent experiments.

study because pretreatment intervals of 1 to 4 h as shown in Figure 1 did not permit enough time to internalize C6-NBD PS over the 60 min time period in untreated erythrocytes. In fact, the extent of PS exposure by 30 min pretreatment with 20 µM Pb2+ was similar to that by 4 h pretreatment with 5 µM Pb2+ (data not shown). Consistent with the observed increase in intracellular calcium, the activity of scramblase was enhanced as measured by C6-NBD PC translocation, whereas the activity of flippase was inhibited by Pb2+ as measured by C6-NBD PS translocation (Figure 3). Although the inward movement of PS was inhibited at up to 10 µM Pb2+, significant inward movement

Procoagulant ActiVation of Erythrocytes by Lead

Figure 3. Effects of Pb2+ on phospholipid translocation in human erythrocytes. After erythrocytes were treated with DW (vehicle) or 5, 10, and 20 µM Pb2+ for 30 min at 37 °C, the extent of phospholipid translocation was measured as described in Experimental Procedures. (A) C6-NBD PS translocation by flippase measured as described in Experimental Procedures. (B) C6-NBD PC translocation by scramblase determined in Pb2+-treated erythrocytes. Values are the mean ( SEM of three independent experiments. (*) Significantly different from the control (p < 0.05).

Chem. Res. Toxicol., Vol. 20, No. 1, 2007 41

Figure 5. Enhancement of thrombin generation in prothrombinase assay and in plasma by Pb2+-treated erythrocytes. (A) After erythrocytes were treated with DW (vehicle) or 1, 2, and 5 µM Pb2+ for 4 h at 37 °C, aliquots were subjected to the prothrombinase assay as described in Experimental Procedures. (B) Pb2+-treated RBCs were added to human plasma and thrombin generation was initiated by recombiplastin. Values are the mean ( SEM of four independent experiments. (*) Significantly different from the control (p < 0.05).

increased thrombus formation with the thrombus weights increasing from 7.40 ( 2.64 mg in untreated rats to 12.53 ( 2.03 and 27.72 ( 5.23 mg in animals given 10 or 25 mg lead (II) acetate/kg, respectively (Figure 7).

Discussion

Figure 4. Effects of Pb2+ on ATP levels in human erythrocytes. After erythrocytes were treated with DW (vehicle) or 1, 2, and 5 µM Pb2+ for 4 h at 37 °C, the levels of intracellular ATP were measured by a luciferin-luciferase assay. Values are the mean ( SEM of four independent experiments. (*) Significantly different from the control (p < 0.05).

occurred at 20 µM Pb2+ (Figure 3A). This result might be due to the activation of nonspecific lipid scramblase as shown in Figure 3B. In addition to the calcium increase, the activity of flippase is inhibited by the depletion of cellular ATP levels (29). Treatment of Pb2+ for 4 h resulted in ATP depletion in a concentration-dependent manner (Figure 4). The significance of Pb2+-induced PS exposure in procoagulant activation was investigated. Erythrocytes were incubated with Pb2+ for 4 h, and potentiated thrombin generation was observed by the prothrombinase assay in a concentration-dependent manner (Figure 5A). Furthermore, Pb2+-treated erythrocytes accelerated the coagulation process initiated by recombinant human tissue factor in plasma (Figure 5B). These findings were in good accordance with the results shown in Figure 1. To confirm if these in Vitro effects can be observed in ViVo, SD male rats were administered intraperitoneally with saline or 25, 50, and 100 mg/kg lead acetate and the extent of PS exposure in erythrocytes was determined. At first, rat erythrocytes showed a concentration-dependent response to in Vitro lead exposure similar to that observed in human erythrocytes (Figure 6A). Consistent with in Vitro results, PS exposure in erythrocytes was significantly (g50 mg/kg) enhanced by lead administration in a dose-dependent manner (Figure 6B). To assess the clinical relevancy of these observations, the effects of lead on thrombus formation were examined using a rat venous thrombosis model. Intravenous lead administration to rats

In the present study, we demonstrated that lead (Pb2+) potently enhanced PS exposure in human erythrocytes mediated through calcium increase and ATP depletion. These in Vitro observations were confirmed in ViVo using a rat venous thrombosis model. Because the blood lead level (BLL) of lead poisoning is 10 µg/dL (approximately 0.5 µM) or greater, concentrations of lead (1-5 µM) used in the current investigation are well within the achievable range in blood from leadexposed (poisoned) humans. There was no hemolysis or microvesicle generation (a marker for shape change in erythrocytes), suggesting that these observations were relatively specific from the adverse action of lead on erythrocytes and not due to the nonspecific cytotoxicity or hemolytic effect. In erythrocytes, other substances, such as arachidonic acid and U46619 (a thromboxane A2 mimic), could also induce PS exposure on erythrocyte surfaces (30). However, the extent of PS exposure induced by those substances was minimal (less than 5% of total erythrocyte population). The results from this study showed that Pb2+ could increase PS exposing cells up to 25%, positioning Pb2+ as one of the most potent substances that induce PS exposure in erythrocytes. Considering the fact that the extent of PS exposure in erythrocytes in sickle cell disease or chronic renal failure patients is around 2 to 3% (31, 32) and that there is a potential thrombogenic effect of focused localization of abnormal erythrocytes (32), these PS exposure effects of lead could accelerate the coagulation process in ViVo. PS exposure by Pb2+ appeared to be mediated by increased intracellular Ca2+ level as shown by 19F-NMR using 5F-BAPTA AM. A previous study measured the intracellular Ca2+ increase in the presence of 1 µM Pb2+ using fluo-3 loaded erythrocytes in flow cytometry (18). Our preliminary study, however, revealed that treatment with 5 µM Pb2+ could interfere with the accurate measurements of Ca2+ because of the binding of Pb2+ to the fluo-3 dye (28). Accordingly, we adopted 19F-NMR with 5F-BAPTA AM for the measurements of Ca2+ increase in erythrocytes, which could bypass the interference by Pb2+ (33, 34). This method could also allow the quantitative measurement of intracellular Ca2+ levels, showing that Pb2+

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Figure 6. Effects of Pb2+ on PS exposure in rat red blood cells. (A) (Left) After erythrocytes isolated from rats and humans were treated with 5 µM of Pb2+ for 4 h at 37 °C, the representative fluorescence histograms by flow cytometry were shown to present the binding of annexin V-FITC to erythrocytes; (Right) Effects in Vitro of Pb2+ on PS exposure were compared in isolated rat and human erythrocytes. Values are the mean ( SEM of three to four independent experiments. (B) Extent of PS exposure on erythrocytes was measured by whole blood flow cytometry 4 h after the i.p. administration of saline (vehicle), 25, 50, and 100 mg/kg of lead(II) acetate. Values are the mean ( SEM of four independent experiments. (*) Significantly different from the control (p < 0.05).

Figure 7. Effects in vivo of Pb2+ on thrombus formation in a rat animal model. After the i.v. administration of saline (vehicle) and lead(II) acetate (10 and 25 mg/kg), thrombus formation was induced by the infusion of thromboplastin in a rat venous thrombosis model. Values are the mean ( SEM of four to five independent experiments. (*) Significantly different from the control (p < 0.05).

treatment elevated calcium levels up to 6.88 ( 1.13 µM (vs 0.24 ( 0.21 µM for control) in erythrocytes. This finding agrees well with the action of Pb2+ studied in other cells, such as osteoblastic bone cells (28, 35) or platelets (36), where intracellular Ca2+ was increased as shown by 19F-NMR analysis. A previous study reported that lead-induced erythrocyte shrinkage mediated by the activation of the Ca2+-sensitive K+

channel enhanced PS exposure (18). In the current study, however, the activities of scramblase and flippase, the major regulating enzymes for PS exposure, were directly determined. Consistent with the intracellular calcium increase and ATP depletion, the activity of scramblase was enhanced, whereas the activity of flippase was inhibited by Pb2+, which was associated with the strong perturbation of membrane asymmetry and resultant potent exposure of PS. The most significant findings of the current investigation are that Pb2+-exposed erythrocytes become procoagulant and could actively contribute to thrombus formation. Pb2+-exposed erythrocytes potentiated the thrombin generation as determined by the prothrombinase assay and accelerated the coagulation process initiated by recombinant human tissue factor in plasma. These procoagulant activities of lead on erythrocytes were also confirmed in in ViVo rat models. Administration of lead significantly enhanced PS exposure and, more importantly, elevated thrombus formation in a dose-dependent manner. In contrast to erythrocytes, platelets, important blood cells in thrombus formation, did not show PS exposure following lead treatment (data not shown), suggesting that erythrocytes are a primary target of lead for thrombus formation. In conclusion, our study suggests that lead exposure can provoke procoagulant activity by PS exposure in erythrocytes,

Procoagulant ActiVation of Erythrocytes by Lead

leading to thrombus formation. Along with the hypertensive effect and anti-fibrinolytic effect of lead, procoagulant activation of erythrocytes could substantially contribute to the overall cardiovascular adverse effect and circulatory mortality rise by lead poisoning. These results will provide new insights into the mechanism of lead-induced cardiovascular disease. Acknowledgment. This work was supported by Korea Research Foundation Grant funded by the Korean Government (MOEHRD) (KRF-2006-E00162). Supporting Information Available: This material is available free of charge via the Internet at http://pubs.acs.org.

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