Hyperglycemia Potentiates Collagen-Induced Platelet Activation Through Mitochondrial Superoxide Overproduction

Collagen, thenoyltrifluoroacetone (TTFA), carbonyl cyanide m-chlorophenylhydrazone (CCCP), bovine serum albumin, NP-40, β-mercaptoethanol, leupeptin, phenylmethylsulfonyl fluoride (PMSF), deoxycholate, Na₃VO₄, aprotinin, and pepstatin were purchased from Sigma Chemical (St. Louis, MO). Mn(III)tetrakis (4-benzoic acid) porphyrin (MnTBAP) was purchased from Calbiochem (La Jolla, CA).

Whole blood was drawn from normal individuals and collected in test tubes containing 3.8% sodium citrate (Vacutainer; Becton Dickinson, Frankin Lakes, NJ). Platelet-rich plasma (PRP) was harvested from this anticoagulated whole blood after centrifugation at 2,500 rpm for 1 min at room temperature. Platelet-poor plasma was obtained by spinning the rest of the blood at 5,000 rpm for 10 min at room temperature. The PRP sample was adjusted using platelet-poor plasma to a platelet concentration of 2 × 10⁸ cells/ml before use.

PRP was preincubated with 5 or 30 mmol/l glucose in the presence or absence of 10 μmol/l TTFA or 0.1 μmol/l CCCP for 60 min. Platelets were sedimented by centrifugation at 5,000 rpm for 5 min and then washed with HEPES buffer (137 mmol/l NaCl, 2.7 mmol/l KCl, 1 mmol/l MgCl₂, and 3.8 mmol/l HEPES, pH 6.5) containing 0.35% bovine serum albumin and 0.4 mmol/l EGTA. The washed platelets were incubated at 37°C with 10 μmol/l of the fluorescent probe CM-H₂DCFDA (Molecular Probes, Eugene, OR) with or without 200 μg/ml collagen for 30 min, and then the intracellular formation of ROS was measured in an HTS 7000 Bio Assay Fluorescent Plate Reader using the HTSoft program (Perkin Elmer, Branchburg, NJ).

Platelet aggregation was monitored using Born’s turbidimetric method (15) with a dual-channel Lumi-aggregometer (Payton Associates, Buffalo, NY). Cuvettes containing 480 μl PRP were stirred at 1,000 rpm for 1 min at 37°C before addition of 20 μl of 5 mg/ml collagen. The change in light transmission was recorded for 5 min. The maximum aggregation (%) was determined as the peak light transmission after the addition of collagen.

PRP was incubated with 5 or 30 mmol/l glucose in the presence or absence of 10 μmol/l TTFA, 0.1 μmol/l CCCP, or 100 μmol/l MnTBAP, a stable, cell-permeable superoxide dismutase mimetic, for 60 min and for the last 10 min with 200 μg/ml collagen. After centrifugation at 5,000 rpm for 10 min, PDGF-AB content in the supernatant was analyzed using an enzyme-linked immunosorbent assay (ELISA) kit derived from R&D Systems (Minneapolis, MN), according to the manufacturer’s instruction.

PRP was incubated with 5 or 30 mmol/l glucose in the presence or absence of 10 μmol/l TTFA, 0.1 μmol/l CCCP, or 100 μmol/l MnTBAP for 60 min. Then, platelets were sedimented and resuspended in 500 μl Tyrode’s buffer, pH 7.4, containing 137 mmol/l NaCl, 2 mmol/l KCl, 1 mmol/l MgCl₂, 0.3 mmol/l NaH₂PO₄, 12 mmol/l NaHCO₃, and 5 mmol/l HEPES and were lysed with an equal volume of lysis buffer, pH 7.4 (2% NP-40, 10 mmol/l HEPES, 2 mg/ml bovine serum albumin, 0.2% β-mercaptoethanol, 200 mmol/l leupeptin, and 4 mmol/l PMSF). PTP activity in the lysate was analyzed using an ELISA kit from Boehringer Mannheim (Mannheim, Germany), according to the manufacturer’s instruction.

PRP was incubated with 5 or 30 mmol/l glucose in the presence or absence of 0.1 μmol/l CCCP or 100 μmol/l MnTBAP for 60 min. Then, platelets were washed and resuspended in 250 μl HEPES buffer, pH 7.4, containing 138 mmol/l NaCl, 2.9 mmol/l KCl, 2 mmol/l MgCl₂, 12 mmol/l NaHCO₃, and 50 mmol/l HEPES. The platelets were incubated with 200 μg/ml collagen at 37°C for 10 min, and then were lysed with an equal volume of RIPA buffer (50 mmol/l Tris-HCl, pH 7.4, 150 mmol/l NaCl, 1% NP-40, 0.25% deoxycholate, 1 mmol/l EDTA, 1 mmol/l PMSF, 1 mmol/l Na₃VO₄, 1 mmol/l NaF, 1 μg/ml aprotinin, 1 μg/ml leupeptin, 1 μg/ml pepstatin). After centrifugation, the supernatants were collected and precleared by the addition of protein A-Agarose (Boehringer Mannheim). After 2 h, the cleared lysates were incubated overnight with polyclonal anti-Syk antibody (Santa Cruz Biotechnology, Santa Cruz, CA) and protein A-Agarose at 4°C. Immunoprecipitated proteins were eluted with 1 × Laemmli buffer and subjected to SDS-PAGE and transferred to nitrocellulose membranes. Membranes were incubated with either polyclonal anti-phosphotyrosine antibody (Upstate Biotechnology, Lake Placid, NY) or anti-Syk antibody, and blots were visualized using the enhanced chemiluminescence detection system (Amersham, Buckinghamshire, U.K.) (6). Signal intensities of phosphotyrosine-Syk bands were normalized by those of Syk protein bands.

PRP was incubated with 5 or 30 mmol/l glucose in the presence or absence of 10 μmol/l TTFA or 0.1 μmol/l CCCP. After 60 min, platelets were sedimented and incubated with 0.5 μmol/l JC-1 (Molecular Probes) and then fluorescence at 590 and 525 nm (510 nm excitation) was monitored.

Data were analyzed using the one-factor analysis of variance procedure to compare the means of all the groups. The Tukey-Kramer multiple comparisons procedure was used to determine which pairs of means were different. Data are shown as mean ± SE.

As shown in Fig. 1, hyperglycemia (30 mmol/l glucose) increased ROS generation in human platelets, and this effect was additive with that of collagen. Furthermore, both TTFA, an inhibitor of mitochondria electron transport chain complex II, and CCCP, an uncoupler of oxidative phosphorylation, completely prevented the effects of hyperglycemia on ROS production in platelets, suggesting that ROS generation arises from the mitochondrial electron transport chain (Fig. 2).

As shown in Fig. 3, 200 μg/ml collagen alone did not significantly induce platelet aggregation. However, marked irreversible aggregation was induced by collagen when PRP was preincubated with high glucose. The hyperglycemia-induced potentiation of platelet aggregation in response to collagen was completely prevented by either TTFA or CCCP.

Hyperglycemia increased collagen-induced PDGF-AB release from platelets approximately threefold. TTFA, CCCP, or MnTBAP completely inhibited the hyperglycemia-induced PDGF-AB release from platelets (Fig. 4).

Hyperglycemia decreased PTP activity in platelets to ∼50% of that of control. TTFA, CCCP, or MnTBAP completely reversed the hyperglycemic effects, suggesting that hyperglycemia-induced mitochondria superoxide inactivates PTP in human platelets (Fig. 5).

Tyrosine phosphorylation of Syk is an early event in collagen-induced platelet activation and aggregation (6). Therefore, we investigated whether hyperglycemia increased Syk phosphorylation induced by collagen. As shown in Fig. 6, hyperglycemia increased Syk phosphorylation in human platelets by fourfold. CCCP or MnTBAP completely inhibited the effects of hyperglycemia.

JC-1 is a fluorescent probe that is sensitive to metabolic inhibitors of mitochondrial oxidative phosphorylation. Increased mitochondrial membrane potential derived from increased oxidative phosphorylation promotes the formation of J-aggregates, which exhibited fluorescence at 590 nm (16). As shown in Fig. 7, TTFA or CCCP was found to completely inhibit the hyperglycemia-induced increase of mitochondrial membrane potential derived from increased oxidative phosphorylation in platelets.

In this study, we found for the first time that hyperglycemia increased mitochondrial ROS generation in human platelets and that this effect was additive with that of collagen. Furthermore, this hyperglycemia-induced ROS generation was found to potentiate both platelet aggregation and the subsequent release of PDGF-AB by platelets exposed to a nonaggregating subthreshold concentration of collagen.

We have recently found that hyperglycemia increases mitochondrial superoxide production inside cultured endothelial cells, providing a causal link between elevated glucose and each of the main pathways responsible for hyperglycemic damage: glucose-induced activation of protein kinase C, formation of advanced glycation end products, sorbitol accumulation, and NF-κB activation (9). The data presented in this paper demonstrate that hyperglycemia-induced mitochondrial ROS overproduction also underlies diabetic platelet hyperaggregation.

Numerous studies have observed enhanced activity of the arachidonate pathway and increased thromboxane A₂ formation by platelets from diabetic patients, which could be responsible for the hypersensitivity of platelets to several agonists, including collagen, in diabetes (17,18,19). Syk activation and the subsequent phosphorylation of PLCγ2 are essential for arachidonic acid release, 5-hydroxytryptamine secretion, and ultimately platelet aggregation in response to collagen (6,20). Therefore, inhibition of redox-sensitive PTP activity by hyperglycemia-induced mitochondrial ROS and the subsequent increased tyrosine phosphorylation and activation of Syk may be an initial event leading to platelet dysfunction in diabetes. ROS have also been implicated in the inhibition of PTP activity and tyrosine phosphorylation of Syk in lymphocytes and macrophages (13,14,21). Hyperglycemia-induced mitochondrial superoxide overproduction may thus potentially play an important role in activation of these inflammatory cells as well as platelets, which would also participate in the pathogenesis of accelerated atherosclerosis in diabetes (22).

PDGF, a potent mitogen and chemotactic factor to smooth muscle cells, is also a key factor mediating restenosis after percutaneous transluminal coronary angioplasty (23,24). Recently, abciximab, a platelet glycoprotein IIb/IIIa inhibitor, was found to improve outcomes in diabetic patients after percutaneous transluminal coronary angioplasty (25). These observations suggest that hyperglycemia-induced platelet hyperaggregation and the subsequent release of PDGF may be one of the potential causes of the high frequency rates of restenosis in patients with diabetes (26,27). Inhibition of hyperglycemia-induced mitochondrial ROS overproduction may provide a new therapeutic strategy for the treatment of diabetic patients with acute coronary syndromes.

This study was supported by grants from the National Institutes of Health.

We thank David Samson for his assistance.