Monoclonal antibodies to αVβ3 (7E3 and LM609) inhibit sickle red blood cell–endothelium interactions induced by platelet-activating factor: Presented in part at the INSERM Symposium, Paris, France, September 2, 1995 (ref ²⁴), and at the 40th Annual Meeting of the American Society of Hematology, Miami, FL, December 7, 1998.

F(ab′)₂ fragments of MoAbs 7E3 (anti-αIIbβ3 + αVβ3),44 10E5 (anti-αIIbβ3),45 and the control irrelevant antibody OC125 were generously provided by Centocor, Inc (Malvern, PA). 7E3 F(ab′)₂ reacts with rat αIIbβ3 and αVβ3 (K_D 7 and 9 μg/mL).46 MoAb LM609 (anti-αVβ3) was kindly provided by Dr David Cheresh (Scripps Institute, La Jolla, CA). It is not known whether 7E3 and LM609 bind to the ligand-bound forms of the αVβ3 receptors.

Heparinized blood was obtained with informed consent from normal (AA) adults (n = 3) and from sickle cell anemia patients (n = 11) who were not in crisis and had not received a blood transfusion in the preceding 4 months.

After removal of the buffy coat, blood was washed three times in normal saline, once in bicarbonate Ringer-albumin solution (118 mmol/L NaCl, 5 mmol/L KCl, 2.5 mmol/L CaCl₂, 0.64 mmol/L MgCl₂, 27 mmol/L NaHCO₃, 0.5% bovine albumin, equilibrated with 95% O₂ and 5% CO₂; pH 7.4; osmolarity, 295 mosm/kg), and resuspended in Ringer-albumin solution. In each case, hematocrit (Hct) was adjusted to 30% for perfusion studies.

Perfusion studies were performed in the isolated, acutely denervated, and artificially perfused rat mesocecum vasculature (n = 46) according to the method of Baez et al47 as modified by Kaul et al6,48 for the infusion of erythrocytes. Details of the procedure have been described elsewhere.48 Arterial perfusion pressure in the mesocecum was maintained at 60 mm Hg, and venous outflow pressure was kept at 3.8 mm Hg. During perfusion with Ringer-albumin solution containing 3% bovine albumin, a 0.2-mL bolus of a given red blood cell sample (Hct 30%) was injected over ∼5 seconds. Peripheral resistance units (PRU) were determined as described49 and expressed in mm Hg/mL/min/g. PRU = ΔP/Q, where ΔP is the arteriovenous pressure difference and Q is the rate of venous outflow (mL/min) per gram of tissue weight. Pressure flow recovery time (Tpf), defined as the time (seconds) required for the arterial pressure and the venous outflow to return to their baseline levels, was determined after the red blood cell injection.

Direct intravital microscopic observations and simultaneous video-recording of the microcirculatory events were performed with an Olympus microscope (model BH-2; Olympus Corp, Lake Success, NY) equipped with a television camera (Cohu, 5000 Series; Cohu, San Diego, CA) and a Sony U-matic video recorder (model VO5800; Sony, Teaneck, NJ). The number of adherent SS cells per 100 μm² was calculated from the counts of individual adherent cells and the surface area (μm²) of the inner wall of the vessel segment as described previously.6 Adhesion data for each experimental group were pooled for statistical comparisons.

PAF supplied in chloroform solution (Sigma, St Louis, MO) was first diluted in dimethyl sulfoxide (DMSO; Sigma), followed by serial dilutions in Ringer-albumin. Rat mesocecum was isolated and perfused with 40 mL of Ringer-albumin containing PAF (200 pg/mL) for 10 minutes. After a 5-minute incubation period, the preparation was perfused as above, and a bolus of AA or SS RBC was injected.

In experiments designed to evaluate the effects of 7E3 F(ab′)₂ or 10E5 F(ab′)₂, the preparation was first infused with the respective antibody (50 μg/mL in 5 mL Ringer-albumin). Control experiments were performed using a control MoAb (OC125) F(ab′)₂ of the same subtype (IgG₁) as 7E3. After a 30-minute incubation at room temperature, the preparation was perfused with PAF solution (200 pg/mL in 40 mL) containing the MoAb (50 μg/mL). Thereafter, a bolus of RBC was injected. In experiments using the MoAb LM609, the preparation was first infused with the MoAb (50 μg/mL in 3 mL) and incubated for 30 minutes as above. This was followed by infusion with PAF (200 pg/mL in 40 mL). A bolus of RBCs was then injected. The control for these experiments was an intact mouse IgG MoAb at the same concentration.

Cryostat sections (6 μm) of fresh-frozen tissue were postfixed in acetone. Sections were treated with 3% hydrogen peroxide, and then immunoperoxidase staining was performed using primary rabbit antihuman vWf polyclonal antibody (Dakopatts, Glostrup, Denmark), diluted 1:400 in phosphate-buffered saline (PBS) containing 5% bovine serum albumin (BSA; Sigma). Control sections were incubated with normal rabbit Ig (Dakopatts) in BSA-PBS. After a 30-minute incubation, the sections were treated with goat antirabbit IgG conjugated with horseradish peroxidase (Dakopatts) at 1:250 (30 minutes). After three washes in PBS, sections were treated for 2 to 3 minutes with freshly prepared 3′3′-diaminobenzidine (DAB [Sigma]). The slides were counterstained with 0.05% toluidine blue. Sections were mounted with Cytoseal mounting medium (Stephens Scientific, Riverdale, NJ).

Paired t-test or unpaired Student's t-test was applied to analyze PRU data as indicated in the tables and Results. Linear regression line analysis of the number of adhered cells/100 μm² (Y) versus the venular diameter (X) was performed. Comparisons of the regression lines between experimental groups were performed using multiple linear regression analysis50 as indicated in Results. The various statistical tests or tests for hypotheses were performed using a Type I error of .05 and were two-tailed. The statistical analysis was performed using the Statgraphic Plus (version 3.1) program for Windows (Manugistics, Inc, Rockville, MD).

In untreated preparations, immunoperoxidase staining for vWf in postfixed frozen sections showed a positive granular pattern in the cytoplasm and often a linear pattern on the luminal surface of endothelial cells (Figure 1A). There was little or no reactivity at interendothelial cell junctions. In contrast, in PAF-treated vasculature, distinct endothelial contraction and separation of endothelial cell junctions were observed. Heavy deposition of vWf could be seen on the luminal aspect of many endothelial cells as well as in the interendothelial cell gaps (Figure1B,arrows); this was accompanied by almost complete depletion of vWf inside some cells (Figure 1B, arrowheads).

Table 1 depicts hemodynamic parameters in the absence or presence of PAF in the ex vivo mesocecum vasculature. In infusion groups 1 and 3, which comprised the untreated preparations, the PRU values were 4.0 and 4.2 mm Hg/mL/min/g, respectively, when Ringer-albumin solution was perfused through the vasculature. In infusion groups 2 and 4, which comprised the preparations treated with PAF, the baseline PRU values were 5.1 and 6.4 mm Hg/mL/min/g, respectively, 40% higher on average than the PRU values in the untreated preparations. Because the dose of PAF employed does not alter arteriolar diameter (as determined in separate preparations), the increase in PRU with PAF treatment reflects the effect on venous outflow of tissue edema secondary to PAF-induced increases in venular permeability.

When AA RBCs were infused into untreated preparations, the PRU values increased by 15.0% ± 2.3% compared with the baseline (Table 1, infusion group 1; Figure 2), and when AA RBCs were infused into the PAF-treated preparations (infusion group 2), the PRU values increased by 17.6% ± 1.7% (Figure 2). In contrast, when SS RBCs were infused into the untreated preparations (infusion group 3), the PRU values increased 25.2% ± 3.0%, which is a 68% greater increase than the increase with AA RBCs (P< .002). Similarly, when SS RBCs were infused into the PAF-treated preparations (infusion group 4), the PRU values increased by 40.5% ± 6.7%, which is a 130% greater increase than when AA RBCs were infused into PAF-treated preparations (infusion group 2;P < .002), and a 61% greater increase than when SS RBCs were infused into untreated preparations (infusion group 3; P = .024).

The Tpf values correlated with the PRU responses. Thus, with AA RBCs, PAF treatment only modestly increased the Tpf (from 31.5 ± 1.9 seconds to 37.0 ± 6.3 seconds, a nonsignificant difference; Table1, infusion groups 1 and 2). The Tpf of untreated preparations infused with SS RBCs was 49.9 ± 2.3 seconds, which was significantly greater than the Tpf of untreated preparations infused with AA RBCs (infusion group 1; P < .0001). The longest Tpf was observed with the infusion of SS RBCs into PAF-treated preparations (90.8 ± 13.9 seconds), which was significantly longer than the values with either AA RBC infusion into PAF-treated preparations (infusion group 2;P < .001) or SS RBC infusion into untreated preparations (infusion group 3; P = .013).

Direct microscopic observations and analysis of videotapes revealed that AA RBCs showed little or no adhesion to postcapillary venules in untreated or PAF-treated preparations. In sharp contrast, infusion of SS RBCs into untreated preparations resulted in prominent adhesion of these cells, which occurred exclusively to the venular endothelium. In the affected venules, adhesion was inversely related to the vessel diameter, which is in accord with our previous observations.6,15 In preparations pretreated with PAF, the increase in adhesion of SS RBC was most prominent and frequently led to obstruction of small-diameter venules (Figure 3). As we previously showed,6,7 the postcapillary blockage may involve contribution from SS RBCs adherent to the endothelium and subsequent trapping of dense SS RBCs. The number of adherent SS RBC/100 μm² in individual venules was plotted as a function of venular diameter. As shown in Figure 4, the number of adherent SS RBCs/100 μm² increased as the venular diameter decreased. Linear regression analysis of the data, using the equation Y = a + bX, confirmed a strong inverse correlation between the number of adherent RBC/100 μm²and the vessel diameter in both control and PAF-treated preparations (control/SS, r = −.80, P < .00001; PAF/SS,r = −.86, P < .00001). Furthermore, a comparison of the two regression lines confirmed significantly increased adhesion of SS RBC in the PAF-treated group as revealed by differences in Y intercepts (P < .0001; Figure 4).

Control antibody OC125 F(ab′)₂ and 7E3 F(ab′)₂ (antiαIIbβ3 + αVβ3) were tested in five different experiments in combination with PAF; in each experiment, the blood from a single SS patient was tested in both a preparation pretreated with control F(ab′)₂ and a preparation pretreated with 7E3 F(ab′)₂ (total, 5 patients and 10 preparations). In four additional experiments, 10E5 F(ab′)₂ (anti-αIIbβ3) was tested with the SS RBCs from two SS patients. Briefly, the preparation was first treated with a given antibody, and then perfused for 10 minutes with a combination of the antibody and PAF (see “Materials and Methods”).

The baseline PRU values in these experiments (Table 2A) (7.3 ± 1.6, 6.2 ± 1.6, and 8.8 ± 4.3 mm Hg/mL/min/g for preparations pretreated with control antibody, 7E3, or 10E5, respectively) were somewhat higher than those for the untreated preparations in the previous experiments (Table 1), reflecting interpreparation variations. When SS RBCs were infused into the control antibody–pretreated and 10E5 F(ab′)₂–pretreated preparations, the PRU increased by 37.8% ± 12.7% and 34.7% ± 6.8% (Figure 5), values similar to the 40.5% ± 6.7% increase in PRU in experiments using PAF and SS RBCs without antibody reported in Table 1 and Figure 2. In contrast, the PRU values increased by only 23.5% ± 7.2% in the preparations pretreated with 7E3 F(ab′)₂, which is significantly less than the increases with either the control antibody (P < .0013) or 10E5 F(ab′)₂ (P < .05). The Tpf results were in accord with the PRU data (Table 2) because the Tpf values of both the control antibody–pretreated preparation (97.0 ± 9.7 seconds) and the 10E5-pretreated preparation (105.0 ± 31.0 seconds) were significantly greater than the Tpf values of the 7E3 F(ab′)₂–pretreated preparation (62.0 ± 5.7 seconds; P < .003 and P < .02, respectively).

In preparations pretreated with the control antibody and PAF, there was pronounced adhesion of SS RBC to the venular endothelium (Figure 3A and C). In marked contrast, in preparations pretreated with 7E3 F(ab′)₂, there was minimal SS RBC adhesion in four experiments and moderate SS RBC adhesion in one experiment (Figure 3D and F). Quantitative analysis of the number of adherent SS RBCs per 100 μm² as a function of venular diameter revealed marked inhibition of SS RBC adhesion by 7E3 F(ab′)₂ even in the smaller-diameter venules, the sites of maximal adhesion. As a result, the Y-intercept of the data plotted in Figure 6 was markedly different for the preparations treated with the control F(ab′)₂ (0.55 SS RBC/100 μm²) and the 7E3 F(ab′)₂(0.02 SS RBC/100 μm²; P < .0001). The slopes were also significantly different (P < .00001) because the 7E3 F(ab′)₂–treated preparations did not demonstrate an inverse relationship between SS RBC adhesion and venular diameter. Furthermore, no obstruction of small-diameter postcapillary venules was evident in the presence of 7E3.

Microscopic analysis of the preparations treated with 10E5 F(ab′)₂ revealed results similar to those seen with the control antibody, with pronounced adhesion of SS RBC in venules and frequent postcapillary blockage.

To further test the role of αVβ3 in SS RBC adhesion, we pretreated four preparations with control IgG and four preparations with LM609 IgG (each 50 μg/mL); all of the preparations were then treated with PAF (200 pg/mL). With the control antibody, the infusion of a bolus of SS RBCs resulted in a 50.3% ± 8.7% increase in PRU, whereas with LM609, the increase was only 25.1% ± 8.0% (P < .006; Table 2B). Similarly, the Tpf was signficantly greater with the control antibody (117.5 ± 9.6 seconds) than with LM609 IgG (82.0 ± 9.3 seconds, P < .02). Microscopic examination revealed that LM609, like 7E3 F(ab′)₂, dramatically inhibited SS RBC adhesion (Figure 7; P < .0001 for the Y-intercept values and slopes).

Our data show that: (1) in the ex vivo mesocecum vasculature, PAF promotes SS RBC–endothelium interactions, and this is associated with increased endothelial surface expression of vWf; and (2) antibodies directed against αVβ3 inhibit adhesion of SS RBC to the PAF-treated vasculature.

We chose PAF to stimulate endothelial cells in our studies because it has been shown to be elevated in the plasma of patients with sickle cell anemia43 and because it has been shown to cause release of endothelial vWf,41 an agent that we and others have reported increases SS RBC adhesion to endothelium.21,22 The concentration of PAF used in this ex vivo, plasma-free study is ∼50% of normal plasma PAF levels (393 ± 65 pg/mL), and ∼25% of the levels found in SS patients (797 ± 62 pg/mL).43 We previously showed that desmopressin, another agent that causes release of vWf from endothelial cells, also results in enhanced SS RBC–endothelial interaction.22 

Because αVβ3 can bind vWf and because endothelial cells have αVβ3 on their luminal surface, we tested whether antibodies to αVβ3 would inhibit adhesion to the postcapillary venules. Importantly, both anti-αVβ3 antibodies we tested (7E3 and LM609) dramatically inhibited SS RBC adhesion under shear flow conditions. Because 7E3 also reacts with αIIbβ3,44 we also tested an antibody (10E5) that inhibits αIIbβ3,45 but not αVβ3. This antibody did not inhibit SS RBC adhesion, supporting the hypothesis that it is 7E3's anti-αVβ3 reactivity that results in the decrease in SS RBC adhesion. Likewise, TSP-enhanced adhesion of SS RBCs to cultured endothelium is abolished by anti-αVβ3 antibodies.34 Taken together, these results indicate that interfering with TSP and vWF binding to the endothelium can effectively inhibit SS RBC adhesion.

Consistent with our previous findings,6,9 the small-diameter venules, probably because of the diameter constraint, low red blood cell velocities, and perhaps endothelial differences compared with arterioles, were the sites of maximal adhesion and frequent blockage in PAF-treated preparations. The inhibition of SS RBC adhesion by anti-αVβ3 antibodies almost completely abolished postcapillary blockage in the present studies. The inhibition of adhesion also resulted in improved hemodynamic behavior of SS RBCs as shown by significantly smaller increases in the peripheral resistance and shortened pressure-flow recovery times. Thus, effective inhibition of SS RBC adhesion can alleviate adhesion-induced vasooclusion.

Although we used washed SS RBC and perfused the vasculature with plasma-free buffer, we cannot rigorously exclude a contribution from a small number of platelets that may have remained or the products they may have released, including TSP, vWf, and fibrinogen. Because TSP, vWf, and fibrinogen can bind to αVβ3, it is possible that some of the effects of the anti-αVβ3 antibodies were caused by inhibition of binding of these adhesive glycoproteins. Similarly, we cannot rigorously exclude the possibility that very small amounts of thrombin may have been generated in our near plasma-free system. Studies by Manodori et al51 found that thrombin can enhance SS RBC adhesion to endothelium with a peak effect at ∼0.1 U/mL, via a mechanism that involves interendothelial cell gap formation.

In conclusion, we have shown that PAF causes a significant increase in SS RBC adhesion, accompanied by an increase in vWf expression on the endothelial surface. This indicates that PAF could be a potential in vivo participant in the pathophysiology of sickle cell vasoocclusion. Pretreatment of the ex vivo vasculature with either of two different monoclonal antibodies to αVβ3 results in a significant inhibition of adhesion and a smaller increase in peripheral resistance. Thus, blockade of αVβ3 receptors may constitute a potential therapeutic approach to prevent SS RBC–endothelium interactions and related vasoocclusion in sickle cell anemia.

Note added in proof. After this manuscript was submitted for publication, Solovey et al52 reported that circulating endothelial cells from patients with sickle cell anemia exhibit increased expression of αVβ3, lending support to a role for αVβ3 in human SS red blood cell adhesion to the vasculature.