Effects of Hyperglycemia and Hyperinsulinemia on Circulating Tissue Factor Procoagulant Activity and Platelet CD40 Ligand

A total of 29 healthy volunteers participated in four different studies. Each volunteer was studied once. The subjects’ ages, weights, heights, and BMIs are shown in Table 1. None of the subjects had a family history of diabetes or any other endocrine disorder, and none were taking any medication. Informed written consent was obtained from each participant after explanation of the nature, purpose, and potential risks of these studies. The study protocol was approved by the institutional review board of Temple University Hospital. Subjects were admitted to Temple University Hospital’s General Clinical Research Center on the evening before the studies.

The studies began at ∼8:00 a.m. after an overnight fast with the subjects reclining in bed. A short polyethylene catheter was inserted into an antecubital vein for infusion of test substances. Another catheter was placed into the contralateral forearm vein for blood sampling. This arm was wrapped with a heating blanket (∼70°C) to arterialize venous blood. The following four studies were performed.

A 20% glucose solution was infused intravenously at variable rates that were adjusted to maintain plasma glucose at ∼200 mg/dl (∼11 mmol/l). Small blood samples (0.25 ml) were collected every 30–60 min initially and every 1–2 h later for measurement of blood glucose concentrations. Subjects were fasting but were allowed to drink water ad libitum and remained at bed rest for the duration of the study. Plasma electrolytes were monitored every 6 h; body weight every 12 h; and fluid balances every 6 h. Potassium (20 mg) and magnesium (400 mg) were given orally every 12 h.

Regular human insulin (Humulin; Eli Lilly, Indianapolis, IN) was infused intravenously at a rate of 12 pmol · kg⁻¹ · min⁻¹ (2 mU · kg⁻¹ · min⁻¹). Glucose was maintained at ∼5.5 mmol/l (100 mg/dl) by feedback controlled glucose infusions. Glucose concentrations were determined every 15–30 min at the beginning and at 1–2 h intervals later with a glucose analyzer, and the glucose infusion rates were adjusted as needed. Plasma electrolyte and fluid balances were monitored as described above.

A 10% glucose infusion was infused at variable rates to maintain blood glucose levels at ∼11 mmol/l. Stimulation of endogenous insulin secretion was prevented by continuous infusion of somatostatin (305 nmol/h). Basal plasma insulin and glucagon levels were maintained by continuous intravenous infusion of insulin (0.33 pmol · kg⁻¹ · min⁻¹) and glucagon (0.25 ng · kg⁻¹ · min⁻¹).

A 5% glucose solution was infused intravenously at variable rates to maintain blood glucose concentrations at ∼100 mg/dl (∼5.5 mmol/l) for 24 h. The rest of the study was as described above.

Blood samples were collected from antecubital veins without tourniquet-induced venostasis at 0, 6, 12, 18, and 24 h. Plasma glucose was measured with a glucose analyzer using the glucose oxidase method and serum insulin by radioimmunoassay using an antiserum with minimal (<0.2%) cross-reactivity with proinsulin (Linco Research, St. Charles, MO). Serum triglycerides were measured enzymatically. Electrolytes were measured at the Temple University Hospital Chemistry Laboratory.

All of the following were measured in plasma harvested from blood collected into 0.1 volume 3.8% sodium citrate. FVIIa activity was measured by a clotting assay using recombinant soluble tissue factor (Diagnostica Stago, Parsipanny, NJ). FVII antigen (FVIIAg) was measured by an enzyme-linked immunosorbent assay (Diagnostica Stago). FVII activity (FVIIc) was measured by a one-stage clotting assay using a mechanical timer Fibrometer (BBL Microbiology System, Cocksville, MD), Simplastin Excel (Biomerieux, Durham, NC), and FVII-deficient plasma (George King Biomedical, Overland Park, KS). Native coagulation factor VIII (FVIII) coagulant activity was measured with an activated partial thromboplastin time-based one-stage clotting assay using the activated partial thromboplastin time reagent from Biomerieux. FVIII-deficient plasma and pooled normal plasma were purchased from George King Biomedical. Tissue factor pathway inhibitor (TFPI) antigen levels were measured by the Imubind total TFPI ELISA from American Diagnostica (Greenwich, CT). Thrombin generation was assessed by determination of prothrombin fragment 1.2 and thrombin-antithrombin (TAT) complexes in plasma using ELISAs (Enzygnost; Dade Behring, Marburg, Germany).

Tissue factor PCA was measured in whole-blood cell lysates with a two-stage clotting assay using recombinant FVIIa (American Diagnostica), Factor X (Enzyme Research Laboratories, South Bend, IN), and normal human plasma containing phospholipid vesicles, according to the method described by Key et al. (17). This assay measures cell-bound and microparticle-associated tissue factor in lysed whole blood. Blood samples (1 ml) were drawn into 0.1 volume 3.8% sodium citrate as anticoagulant were frozen at −70°C until assayed. Blood lysates and cellular components were collected from aliquots of whole blood subjected to three cycles of freezing and thawing followed by centrifugation and washing, and they were finally suspended in HBSA buffer (10 mmol/l HEPES, 137 mmol/l NaCl, 5.38 mmol/l KCl, 5.55 mmol/l glucose, and 0.1% bovine serum albumin, pH 7.5) as described previously (17). The final membrane pellet resuspended in HBSA buffer was used as a source of tissue factor. Tissue factor PCA was measured using a two-stage clotting assay. In the first stage, 20 μl sample, 5 nmol/l FVIIa, and 250 nmol/l human factor X were incubated for 5 min at 37°C. CaCl₂ (8.3 mmol/l) was added and incubated for 3 min. In the second stage, 100 μl normal human plasma containing phospholipid vesicles (75% phosphatidylcholine:25% phosphatidylserine prepared by weight; Avanti Polar Lipids, Alabaster, AL) and 100 μl 25 mmol/l CaCl₂ were added, and the clotting time was recorded in a mechanical fibrometer (BBL Microbiology System). Recombinant human tissue factor Ortho Recombiplastin (Ortho Diagnostic Systems, Raritan. NJ) was used as a standard. The tissue factor concentration in the standard was determined by using IMUBIND Tissue Factor antigen ELISA (American Diagnostica). Repeated measurements of 1 arbitrary unit/ml Ortho Recombiplastin gave 1.34 pg/ml. A log plot of tissue factor versus clotting time was linear in the 1–1,000 units/ml range (r = 0.99). The interassay coefficient of variation (CV) was 8.6%, the intra-assay CV was 7.0%.

Tissue factor expression on monocyte surface was measured by flow cytometry. Blood samples collected into 0.1 volume 3.8% sodium citrate were diluted 1:3 with modified HEPES-Tyrodes buffer (10 mmol/l HEPES, 137 mmol/l NaCl₂, 2.8 mmol/l KCl, 1 mmol/l MgCl₂, 12 mmol/l NaHCO₃, 0.4 mmol/l Na₂HPO₄, 0.35% BSA, and 5.5 mmol/l glucose, pH 7.4) and incubated (20 min, at room temperature) with fluorescein isothiocynate (FITC)-labeled anti-human CD14 (monocytes) and phycoerythrin-labeled anti-human CD142 (tissue factor) antibodies. As controls, corresponding matched isotype antibodies FITC anti-mouse IgG_2a and phycoerythrin mouse IgG_1K were used. All monoclonal antibodies were purchased from BD Biosciences, Pharmingen (San Diego, CA). The samples were fixed with 1.1% formaldehyde (Sigma, St. Louis, MO) for 10 min at room temperature, diluted fourfold with distilled water, and analyzed using a BD FACScan flow cytometer (Becton Dickinson Immunocytometry Systems, San Jose, CA) within 2 h of sample preparation. The monocyte population was identified by gating CD14-positive events and tissue factor expression was determined as percent CD142-positive cells. Four thousand monocytes were analyzed. The positive delineator for tissue factor–expressing monocyte population was determined by gating ∼1% background staining on both FITC- and phycoerythrin-labeled isotype controls.

Platelet-rich plasma was obtained from citrated whole blood by centrifugation at 200g for 15 min. Platelet-rich plasma was diluted threefold with modified HEPES-Tyrodes buffer and incubated with phycoerythrin-labeled anti-human CD154 (CD40 ligand [CD40L]) or corresponding matched isotype control phycoerythrin mouse IgG_1K antibody for 20 min. Cells were fixed in 1% formaldehyde for 10 min at room temperature. The samples were diluted 10-fold with HEPES-Tyrodes buffer and analyzed (10,000 platelets) within 2 h in BD FACScan flow cytometer. The CD40L-positive platelets were determined by gating on 1% of isotype control–positive cells.

Mononuclear cells were isolated from 20 ml freshly heparinized whole blood by the Ficol-Paque TM Plus gradient method according to the manufacturer’s instructions (Amersham, Piscataway, NJ). Monocytes were positively selected using 20 μl CD14 microbeads per 10⁷ mononuclear cell according the manufacturer’s protocol (Miltenyi Biotech, Auburn, CA).

Total RNA isolation and one-step SYBR Green real-time RT-PCR were performed as described previously (28). The following primer sequences were used. For tissue factor (gene ID NM 001993): forward, 5′-CAGTGATTCCCTCCCGAACA-3′ and reverse 5′-TGCCTTTCTACACTT GTGTAGAG-3′ (amplicon size 173 bp). For 18sRNA: Competitimer Primer Set (catalog no. 5103G; Ambion, Austin, TX).

A standard curve of cycle threshold versus concentration was obtained using serial dilutions of control total RNA isolated from tissue factor–expressing human breast cancer cells (MDA-MB-231) (29). Relative concentrations of the tissue factor amplicon were determined from the standard curve, and the after-to-before ratio was normalized with the after-to-before ratio of the internal standard (18srRNA).

A one-way ANOVA was used to test for significant differences between studies with Student-Newman-Keuls post hoc analysis. If data were not normally distributed, the Kruskal-Wallis one-way ANOVA with Dunn’s post hoc analysis was used. To test for differences across time, a one-way repeated-measures ANOVA with Student-Newman-Keuls post hoc analysis was used. If data were not normally distributed, the Friedman repeated-measures ANOVA on ranks with Student-Newman-Keuls post hoc analysis was used. Statistical differences between day 1 (baseline) and day 2 (24 h) were determined with a paired t test and, if not normally distributed, with the Wilcoxon Signed Rank test. Statistical analyses were performed using SigmaStat for Windows (version 2.0; SPSS, Chicago, IL). Statistical significance was defined as P < 0.05. All results are presented as means ± SE.

Plasma glucose concentrations remained basal (∼5.5 mmol/l or ∼100 mg/dl) for 24 h in the selective hyperinsulinemia (normal glucose/high insulin) and the control (normal glucose/normal insulin) clamps and were raised by infusion of glucose to ∼11 mmol/l (∼200 mg/dl) for 24 h in the selective hyperinsulinemia (high glucose/normal insulin) and the high glucose/high insulin clamps.

Serum insulin concentrations remained basal (∼50 pmol/l or ∼8 μU/ml) for 24 h in the selective hyperglycemia and control clamps and were raised by infusion of insulin to ∼1,200 pmol/l (∼200 μU/ml) in the selective hyperinsulinemia and high glucose/high insulin clamps.

During the control study (normal glucose/normal insulin), basal tissue factor PCA remained unchanged at ∼20 units/ml for 24 h. During selective hyperglycemia, tissue factor PCA increased approximately twofold (from 14.2 ± 1.5 to 33.3 ± 5.7 units/ml, P < 0.05); during selective hyperinsulinemia, tissue factor PCA rose approximately sixfold (from 13.6 ± 1.3 to 83.9 ± 5.7, P < 0.001); and during combined hyperglycemia and hyperinsulinemia, it rose approximately ninefold (from 19.7 ± 1.1 to 172.4 ± 5.9, P < 0.001). In all groups, except control subjects, significant increases were observed starting at 6 h.

Because much of circulating tissue factor PCA is associated with monocytes, tissue factor expression on monocytes was assessed by flow cytometry at baseline and at 24 h. During high glucose/high insulin, tissue factor–positive monocytes increased from 19.8 ± 5.5 to 31.9 ± 5.0% (P < 0.002). There was no increase in the mean fluorescence intensity of tissue factor expression on monocytes. During selective hyperinsulinemia, tissue factor–positive monocytes increased from 12.5 ± 3.9 to 17.5 ± 3.3% (P < 0.02). In contrast, there was no change in the control study (normal glucose/normal insulin). Tissue factor–positive monocytes were not determined during selective hyperglycemia. Tissue factor mRNA in monocytes obtained from three volunteers increased 1.9 ± 0.3-fold between 0 and 24 h of high glucose/high insulin (P < 0.04) (Fig. 3).

FVIIa, the activated form of FVII, remained unchanged for 24 h during the control study. Selective hyperinsulinemia and high glucose/high insulin were both associated with large decreases in FVIIa, whereas FVIIa decreased less during selective hyperglycemia (Fig. 4A and B). Plasma FVIIc decreased most during high glucose/high insulin and to a lesser degree during selective hyperinsulinemia and selective hyperglycemia and did not change during the control studies (Fig. 4C and D). Similarly, FVIIAg levels declined during the high glucose/high insulin studies, selective hyperinsulinemia, and selective hyperglycemia but remained unchanged during the control studies (Fig. 4E and F). Factor VIII increased continuously during the high glucose/high insulin studies but did not change significantly during the other three studies (Fig. 4G and H).

There were statistically significant inverse relationships between tissue factor PCA and FVIIa in the high glucose/high insulin (r = −0.52, P < 0.001), the selective hyperinsulinemia (r = −0.51, P = 0.002), and the control (r = −0.52, P = 0.007) studies. FVIII levels correlated positively with tissue factor PCA (r = 0.57, P < 0.0001) during the high glucose/high insulin studies.

TAT, a sensitive indicator of thrombin formation, did not change during the control study. There were, however, large and significant increases in TAT during the selective hyperinsulinemia and during the high glucose/high insulin studies (Fig. 5A and B). The TAT levels during these three studies correlated positively with tissue factor PCA levels (r = 0.49, P = 0.006; r = 0.35, P = 0.04; and r = 0.57, P < 0.0001, respectively). Fragment 1.2, another indicator of thrombin formation, also remained unchanged during the control studies but rose during selective hyperinsulinemia and during high glucose/high insulin (Fig. 5C and D).

Plasma TFPI antigen levels tended to decline in all four studies, but these changes reached statistical significance only in the selective hyperinsulinemia group, where the levels declined from 86.0 ± 9.6 to 69.6 ± 7.4 ng/ml at 18 h (P < 0.001) and to 65.7 ± 3.0 ng/ml at 24 h (P < 0.05).

No change occurred during the control study (28.1 ± 2.1 vs. 29.0 ± 2.1%, NS). In contrast, during high glucose/high insulin, CD40L-positive platelets increased from 32.9 ± 3.0 to 46.1 ± 3.7%, (P < 0.001). CD40L-positive platelets also increased during selective hyperinsulinemia (from 31.2 ± 2.0 to 37.5 ± 3.1%, P = 0.01) and during selective hyperglycemia (from 33.2 ± 6.5 to 37.2 ± 7.4%, P < 0.05), indicating that platelets had been activated. There was no significant increase in the mean fluorescence intensity of platelet CD40L expression in these groups.

Tissue factor is the physiological initiator of blood coagulation (13). The earlier concept that blood coagulation is initiated after vascular damage when blood is exposed to “extrinsic” tissue factor has been broadened by the observations that tissue factor is also present in circulating blood (13,14,16,17) and that hematopoietic cell–associated tissue factor is important in thrombogenesis (18). For instance, circulating tissue factor has been associated with increased blood thrombogenicity in patients with cardiovascular disease (30), sickle cell disease (17), antiphospholipid antibody syndrome (31), hyperlipidemia (20), and disseminated intravascular coagulation (32). In addition, we (19) and others (20) have reported that patients with type 2 diabetes have increased blood levels of tissue factor PCA. This is of considerable clinical interest because these patients are known to be at greatly increased risk for ASVD and acute arterial events. In the current study, we have examined in healthy subjects the hypothesis that the hyperglycemia and/or hyperinsulinemia, such as is seen in many patients with type 2 diabetes, may increase circulating tissue factor PCA. Exposure for 24 h to selective hyperglycemia resulted in an approximate twofold rise, exposure to selective hyperinsulinemia in an approximate sixfold rise, and exposure to combined high glucose/high insulin in an approximate ninefold rise in circulating tissue factor PCA (Fig. 2). (Direct comparison of the magnitude of changes observed in the selective hyperglycemia group with the other three groups needs to be made with some caution because of the additional infusion of somatostatin in the selective hyperglycemia protocol.) These results suggested that the effects of selective high glucose and high insulin levels were additive and that the circulating tissue factor PCA may have been even greater had the studies lasted >24 h. They are consistent with our previously reported findings of elevated tissue factor PCA in patients with type 2 diabetes compared with nondiabetic control subjects (72.5 ± 12.6 vs. 23.2 ± 0.2 units/ml, P < 0.001) (19). The levels of plasma TAT were also elevated in these patients.

Monocytes are the only hematopoietic cells with established ability to synthesize tissue factor (13,14). During high glucose/high insulin and during selective hyperinsulinemia, tissue factor–positive monocytes increased significantly (Fig. 3). In addition, monocyte tissue factor mRNA increased approximately twofold during high glucose/high insulin, indicating increased tissue factor transcription.

The modest increases in monocyte surface expression of tissue factor and mRNA may not be sufficient to explain the marked increase in circulating tissue factor PCA. A possible explanation for this discrepancy is that tissue factor–containing microparticles may have been released from activated monocytes on exposure to high glucose and/or high insulin and may have accumulated in the blood during the 24-h experiments. They are measured in the tissue factor PCA assay together with tissue factor on lysed cell membranes but are not detected by flow cytometry, which measured only monocyte surface tissue factor.

Several lines of evidence point to an important role of blood-borne tissue factor in thrombus formation. Circulating tissue factor is biologically active in converting factor X to Xa (16). Tissue factor–bearing microparticles are highly procoagulant and have been shown to transfer tissue factor to platelets and to propagate further thrombus growth (13,16,18,33). Even where thrombus initiation was mediated by vessel wall tissue factor, the propagation of the initial thrombus depended on the recruitment of blood-borne tissue factor (13,14,16,18,34). Also, enhanced thrombus formation occurred when blood from patients with type 2 diabetes and elevated circulating tissue factor levels was perfused over a collagen-coated surface (20). Moreover, in patients undergoing angioplasty or stent implantation, elevated whole-blood tissue factor PCA present before the procedure predicted restenosis (30). We therefore evaluated effects of increases in circulating tissue factor PCA on other coagulation markers. We found that TAT and fragment 1.2, two sensitive indicators of thrombin generation, rose continuously during selective hyperinsulinemia and more so during the combined high glucose/high insulin studies (Fig. 5). Plasma factor VIII also rose in the latter group (Fig. 4). It needs to be noted that at least some of the monocyte membrane-bound tissue factor is encrypted, i.e., it can bind FVIIa without leading to activation of coagulation (35). This and the inhibitory effect of TFPI may have limited thrombin generation in our studies.

Plasma levels of FVIIa, FVIIc, and FVIIAg paradoxically declined during the high glucose/high insulin–stimulated rise of tissue factor PCA (Fig. 4), reflecting a decrease in total plasma FVII. Similar decreases in FVIIa have been reported after bolus infusions of endotoxin (36) and in sepsis (37). Because tissue factor is the principal ligand for FVIIa, we believe that the decrease in plasma FVII levels reflect accelerated clearance as a result of binding to an increased number of available tissue factor sites.

Thus, our studies provided evidence of increased circulating tissue factor PCA and FVIII associated with increased thrombin generation during hyperglycemia/hyperinsulinemia. These studies were performed in healthy subjects, but comparable findings have been reported in diabetic patients (19). The implications are that hyperglycemia/hyperinsulinemia induced a hypercoagulable state with the potential for enhanced thrombus formation, particularly in the context of plaque rupture. This is likely to play a role in the development of acute vascular events in patients with diabetes who are known to have a higher incidence of thrombus formation than nondiabetic patients (5,38).

CD40L is a 39-kDa glycoprotein belonging to the tumor necrosis factor superfamily (23–25). Most (>95%) of CD40L in circulation is stored in platelets, is expressed rapidly on platelet surfaces after activation, and is then released into the circulation (25). CD40L expressed on platelets interacts with cells that constitutively display its receptor CD40, including endothelial cells, monocytes, and macrophages, and incites a cascade of inflammatory responses, including expression of adhesion factors (including e-selectin and vascular cell adhesion molecule-1), expression of tissue factor, and release of chemokines and cytokines (including monocyte chemoattractant protein-1, interleukin-6, and interleukin-8) (23–25).

In our studies, CD40L-bearing platelets increased during the combined high glucose/high insulin studies with a somewhat lesser increase during selective hyperglycemia and selective hyperinsulinemia (Fig. 6). These findings provide strong evidence of in vivo platelet activation. They are relevant for patients with type 2 diabetes because CD40L triggers inflammatory reactions and is a major contributor to atherogenesis (24,39,40). Serum CD40L levels are elevated in diabetic patients (41). The upregulation of CD40L in platelets in our studies also suggested a mechanism for the increased monocyte tissue factor expression. Tissue factor is by itself proinflammatory (42) and in turn stimulates thrombosis and platelet release of CD40L, thereby sustaining a vicious cycle of inflammation and thrombosis (43). The sequence of events is summarized in Fig. 7.

In summary, combined elevation of plasma insulin and glucose levels for 24 h in healthy volunteers produced 1) a dramatic (ninefold) increase in circulating tissue factor PCA associated with an increase in monocyte tissue factor surface expression and mRNA and 2) changes in other components of blood coagulation, suggesting that the coagulation system had been activated, including increases in plasma TAT complexes, prothrombin fragment 1.2, and FVIII and decreases in FVIIa and FVIIc activities. In addition, there was an increase in platelet CD40L, indicating platelet activation. Effects of selective hyperinsulinemia and selective hyperglycemia were less dramatic but appeared to be additive. We conclude that hyperinsulinemia and hyperglycemia, but particularly the combination of both, create a prothrombotic state and may in addition be proinflammatory and proatherogenic by virtue of the recognized actions of CD40L and tissue factor.

G.B. has received National Institutes of Health Grants R01-DK-58895, R01-AG-15353, R01-HL-733267, and R01-DK-066003 and a Mentor-Based Training grant from the American Diabetes Association. A.K.R. has received National Institutes of Health Grant R01-DK-58895. The Molecular Core Facility of the Center for Substance Abuse at Temple University has received grant P30-DA-13429.

We thank Karen Kresge for outstanding technical assistance, Mayumi Katoaka for assistance with the flow cytometry studies, and Constance Harris Crews for typing the manuscript. We also thank the Molecular Core Facility of the Center for Substance Abuse at Temple University for letting us use their Roche Light-Cycler for the real-time RT-PCR measurements.