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We previously demonstrated that coagulation factor VIII (FVIII) accelerates proteolytic cleavage of von Willebrand factor (VWF) by A disintegrin and metalloprotease with thrombospondin type 1 repeats (ADAMTS13) under fluid shear stress. In this study, the structural elements of FVIII required for the rate-enhancing effect and the biological relevance of this cofactor activity are determined using a murine model. An isolated light chain of human FVIII (hFVIII-LC) increases proteolytic cleavage of VWF by ADAMTS13 under shear in a concentration-dependent manner. The maximal rate-enhancing effect of hFVIII-LC is ~8-fold, which is comparable with human full-length FVIII and B-domain deleted FVIII (hFVIII-BDD). The heavy chain (hFVIII-HC) and the light chain lacking the acidic (a3) region (hFVIII-LCΔa3) have no effect in accelerating VWF proteolysis by ADAMTS13 under the same conditions. Although recombinant hFVIII-HC and hFVIII-LCΔa3 do not detectably bind immobilized VWF, recombinant hFVIII-LC binds VWF with high affinity (KD, ~15 nm). Moreover, ultra-large VWF multimers accumulate in the plasma of fVIII−/− mice after hydrodynamic challenge but not in those reconstituted with either hFVIII-BDD or hFVIII-LC. These results suggest that the light chain of FVIII, which is not biologically active for clot formation, is sufficient for accelerating proteolytic cleavage of VWF by ADAMTS13 under fluid shear stress and (patho) physiological conditions. Our findings provide novel insight into the molecular mechanism of how FVIII regulates VWF homeostasis.
Proteolytic cleavage of ultra-large von Willebrand factor (VWF)3 on endothelial cells (1, 2) and in flowing blood (3, 4) by a plasma metalloprotease, ADAMTS13, is crucial for normal hemostasis. ADAMTS13 cleaves VWF at the specific Tyr1605–Met1606 bond in the central A2 domain (5, 6). This cleavage is dramatically accelerated by fluid shear stress (4, 6, 7) or mild denaturization with urea (5, 8) or guanidine (6, 9), which alters VWF conformation and exposes binding and cleavage sites. Inability to cleave VWF as a result of a severe deficiency of plasma ADAMTS13 leads to an accumulation of ultra-large VWF on endothelial cells or in blood (10). This triggers spontaneous platelet aggregation and disseminated microvascular thrombosis, characteristic of thrombotic thrombocytopenic purpura (11, 12). Moreover, mild to moderate deficiency of plasma ADAMTS13 has been shown to be a risk factor for cardiovascular events such as myocardial infarction (13–16) and ischemic cerebral stroke (17, 18).
In addition to fluid shear stress, we and others have shown that cleavage of soluble multimeric VWF by ADAMTS13 under fluid shear stress is dramatically accelerated by coagulation factor VIII (FVIII) (19), platelets (20, 21), and glycoprotein 1bα (21, 22). FVIII and platelets appear to synergistically accelerate cleavage of VWF by ADAMTS13 under these conditions (21). We have previously shown that the FVIII B-domain is not required, but the a3 in the context of two chain B-domainless FVIII, which binds VWF with high affinity, is required for the rate-enhancing effect on proteolytic cleavage of VWF by ADAMTS13 under shear stress (19).
Here, we show that an isolated light chain of FVIII, which is biologically inactive for clot formation, is sufficient for accelerating proteolytic cleavage of VWF by ADAMTS13 in vitro using a fluid shear-based assay and in vivo using fVIII−/− mice expressing FVIII variants via a hydrodynamic approach. This rate-enhancing effect by FVIII light chain also depends on its high affinity binding with VWF; a light chain of FVIII lacking the a3 region and a heavy chain of FVIII, which do not bind VWF detectably, exhibit no effect on ADAMTS13-mediated VWF proteolysis under the same conditions. Our findings may shed more light on the structure-function relationship of FVIII in regulation of the VWF-ADAMTS13 axis, which helps in understanding the clinical heterogeneity of patients with severe hemophilia A.
The cDNA fragments encoding a heavy chain (hFVIII-HC), a light chain (hFVIII-LC), and a light chain lacking a3 (hFVIII-LCΔa3) of human FVIII were amplified by PCR using hFVIII-BDD as a template (19). The primer pairs were as follows: hFVIII-HC (5′-GAGTACTCCCTCTCAAAAGCGGGCATG-3′ and 5′-GGAGAAGCTTCTTGGTTCAAT-3′); hFVIII-LC (5′-GAAATAACTCGTACTACTCTT-3′ and 5′-GTAGAGGTCCTGTGCCTCGCA-3′); and hFVIII-LCΔa3 (5′-AGCTTTCAAAAGAAAACACGA-3′ and 5′-GTAGAGTCCTGTGCCTCGCA-3′). The amplified hFVIII-LC and hFVIII-LCΔa3 fragments were cloned into pSecTag/FRT/V5-His TOPO vector (Invitrogen). An hFVIII-HC fragment was cloned into pcDNA3.1 V5-His TOPO vector (Invitrogen) according to the manufacturer's recommendation. All three constructs were tagged at their C termini with a V5-His epitope (Fig. 1A). Plasmids were sequenced at the Nucleic Acid Core Facility at the Children's Hospital of Philadelphia to confirm the accuracy. Plasmids encoding a B-domain deleted human factor VIII (hFVIII-BDD) (19) or canine FVIII (cFVIII-BDD) (23) were described in other studies.
Baby hamster kidney cells were used to express human recombinant hFVIII-BDD and cFVIII-BDD, as described previously (19, 23, 26). Human embryonic kidney (HEK293) cells were used to express recombinant hFVIII-LC, hFVIII-LCΔa3, and hFVIII-HC. Cells were grown in Dulbecco's modified Eagle's medium (DMEM) (Invitrogen) containing 10% of FetalPlex (Gemini BioProducts, West Sacramento, CA). The cells were transfected with LipofectAMINE2000 and plasmid DNA in serum-reduced Opti-MEM. For those vectors without the G418-resistent gene, co-transfection with pcDNA3.1 (10:1 ratio) containing the neomycin resistance gene (Invitrogen) was carried out. Stable clones were selected after culturing cells for 10 days in the presence of 0.5–1.0 mg/ml of G418 (Invitrogen). The positive clones were identified by Western blotting and immunofluorescent staining with anti-FVIII IgG (ESH-8) that recognizes light chain or anti-V5 IgG (Invitrogen) for V5-His-tagged proteins as described previously (25).
Serum-free condition medium of stable cell lines expressing recombinant FVIII or variants was collected daily for a total of 2.5–10 liters. Recombinant hFVIII-BDD and cFVIII-BDD were purified and quantified as described previously (19, 23, 26). Recombinant hFVIII-HC without the C-terminal epitope was kindly provided by Dr. Philip Fay at the University of Rochester, New York. Recombinant hFVIII-LC, hFVIII-LCΔa3, and hFVIII-HC were purified by SP-Sepharose ion exchange chromatography, followed by a nickel-nitrilotriacetic acid affinity column as described previously (19).
The purity and integrity of the purified recombinant FVIII variants were determined by 10% SDS-polyacrylamide gel with Coomassie Blue staining. All purified recombinant FVIII variants except for hFVIII-HC were quantified by absorbance at 280 nm corrected with light scattering at 320 nm. Partially purified hFVIII-HC (V5-His tagged) was quantitated by Western blotting with monoclonal anti-V5 IgG using a positopeTM as a reference (Invitrogen) (25).
Human VWF was purified from plasma by cryoprecipitation followed by gel filtration on a Sephacryl-300 column (2.5 × 100 cm) (GE Healthcare) as described previously (21). Recombinant human ADAMTS13 (V5-His tagged) was expressed from stably transfected HEK293 cells and purified by Q-fast flow ion exchange followed by a nickel-affinity chromatography as described previously (7, 21).
Purified plasma VWF (37.5 μg/ml) was incubated in a PCR tube (Fisher Scientific, Newark, DE) for 10 min with 50 nm recombinant human ADAMTS13 in the absence or in the presence of FVIII variants at various concentrations in 20 mm HEPES, 0.15 m NaCl, 5 mm CaCl2, and 1.0 mg/ml BSA, pH 7.5. The reaction mixture (20 μl) was subjected to constant vortexing at 2,500 rpm for 10 min at room temperature as described previously (7, 19, 21). The reaction was quenched by boiling the sample with an equal volume of sample buffer (125 mm Tris, 10% glycerol, 2% SDS, and 0.01% bromphenol blue, pH 6.8) for 5 min. A fraction of sample was fractionated on a 5% Tris-glycine SDS-polyacrylamide gel. Alternatively, samples were denatured at 60 °C for 20 min with sample buffer (60 μl) (70 mm Tris, pH 6.8, 2.4% SDS, 0.67 m urea, and 4 mm EDTA, 10% glycerol, 0.01% bromphenol blue). Denatured sample (10 μl, ~90 ng of VWF) was fractionated on a mini-gel containing 1% agarose (Lonza, Rockland, ME). After being transferred onto a nitrocellulose membrane (Bio-Rad), VWF cleavage product or multimers were determined by Western blotting. Membrane was blocked with 1% casein in TBST for 30 min and incubated with rabbit anti-VWF IgG (Dako, Carpinteria, CA) (1:5,000) in 1% casein/TBST, followed by IRDye 800CW-labeled goat anti-rabbit IgG (LI-COR Bioscience, Lincoln, NE) (1:20,000) in the same buffer. When resolved by SDS-PAGE, the cleavage product at 350 K was quantified by densitometry using ImageJ software as described previously (19, 21). When resolved by agarose gel electrophoresis, the ratio of the low molecular weight bottom band to the high molecular weight bands was quantified in a similar manner.
A microtiter plate was coated with 100 μl of human VWF (2 μg/ml) and blocked with 1% casein in PBS, pH 7.4. Recombinant FVIII variants at various concentrations diluted with PBS containing 0.2% casein were added and incubated at 25 °C for 1 h. After being washed with PBS, bound FVIII variants were detected by peroxidase-conjugated mouse anti-human monoclonal anti-FVIII IgG (ESH-8HR) (1:3,000) that recognizes the C2 domain in the light chain of FVIII (American Diagnostica Inc., Stamford, CT) or peroxidase-conjugated monoclonal anti-V5 (1:1,000) (Invitrogen). A pre-mixed chromogenic substrate, 3,3′,5,5′-tetramethylbenzidine (TMB) (Thermo Fisher Scientific, Rockford, IL), was added for color reaction. The absorbance was determined at 450 nm in a SpectroMax microtiter plate reader (Molecular Device, Sunnyvale, CA).
The Institutional Animal Care and Use Committee (IACUC) at the University of Pennsylvania and the Children's Hospital of Philadelphia approved the protocol for the mouse study. FVIII-deficient mice (fVIII−/−) in a C57BL6/129 strain with exon-16 deletion were described previously (27). Mice at the age of 6–8 weeks were injected with 2 ml of saline alone or saline containing 100 μg of plasmid DNA (endotoxin-free) via a tail vein within 5 s as described previously (28). 48 h after injection, whole blood (200 μl) was collected after tail clip and anti-coagulated with sodium citrate (3.8%). Platelet-poor plasma was obtained after centrifugation at 10,000 rpm for 10 min and stored in small aliquots at −80 °C until use.
Plasma antigen levels of FVIII variants expressed after hydrodynamic injection were determined by a home grown ELISA. A microtiter plate was coated with polyclonal goat anti-FVIII IgG (1:2,000) at 25 °C for 1 h. The plate was blocked with 1% (w/v) casein in PBST for 30 min. Purified recombinant hFVIII-BDD or cFVIII-BDD (0, 5, 10, 20, 40, 80, 160, and 320 ng/ml) or mouse plasma (1:20 or 1:40) in PBS was incubated in antibody-coated wells for 2 h. After being washed with PBS, bound FVIII variants (hFVIII-BDD, cFVIII-BDD, hFVIII-LC, and hFVIII-LCΔa3) were detected by peroxidase-conjugated monoclonal anti-FVIII IgG (ESH-8HR) (1:1,000) (American Diagnostica, Stamford, CT) for 1 h. For quantification of plasma hFVIII-HC antigen, mouse plasma was incubated in a nickel-coated microtiter plate for 2 h. The bound hFVIII-HC was detected by monoclonal anti-V5 IgG (Invitrogen), followed by a horseradish peroxidase-conjugated rabbit anti-mouse IgG (DAKO, Carpinteria, CA). A pre-mixed TMB solution was added for color development, and absorbance at 450 nm was determined by a SpectroMax microtiter plate reader as described above.
Plasma VWF antigen was quantified by an ELISA as described previously (29). A microtiter plate was coated with rabbit anti-vWF IgG (Dako, Carpinteria, CA) (1: 2,000) and blocked with 1% casein in PBS containing 0.05% Tween 20 (TBST). Mouse plasma diluted (1:20 and 1:40) with 0.2% casein in TBST was added and incubated for 1 h. After being washed with PBS, bound VWF was detected by peroxidase-conjugated rabbit anti-VWF IgG (1:2,000) (Dako, Carpinteria, CA). Pooled murine plasma from C57BL6 was used as a reference. A pre-mixed TMB solution was used for color development. The absorbance at 450 nm was determined with a SpectroMax microtiter plate reader (Molecular Device, Sunnyvale, CA).
Mouse plasma (1 μl) was denatured at 60 °C for 20 min with 10 μl of sample buffer (70 mm Tris, pH 6.8, 2.4% SDS, 0.67 m urea, and 4 mm EDTA, 10% glycerol, 0.01% bromphenol blue). The proteins were fractionated on a 1% agarose gel (7 × 8 cm) on ice. After being transferred to a nitrocellular membrane, VWF multimers were detected by incubation with rabbit anti-VWF IgG (1:5,000) (Dako, Carpinteria, CA), followed by IRDye 800CW-labeled goat anti-rabbit IgG (1:20,000) in 1% casein/TBST. Odyssey imaging analysis (LI-COR Bioscience, Lincoln, NE) was used to scan the membrane (19, 30). Densitometry analysis using ImageJ software determined the formation of cleavage product or the ratio of high to low molecular weight VWF multimers.
The difference in means between the control and experimental groups was determined by one-way analysis of variance using Minitab16 software. p values less than 0.05 and 0.01 are considered to be statistically significant and highly significant, respectively.
We previously found that full-length FVIII accelerates proteolytic cleavage of VWF by ADAMTS13 under fluid shear stress (19). However, the domain components of FVIII required for the cofactor activity to enhance VWF proteolysis are not fully understood. We therefore prepared various recombinant FVIII variants, including hFVIII-HC, hFVIII-LC, and hFVIII-LCΔa3, in addition to hFVIII-BDD and cFVIII-BDD (Fig. 1A). All variants except for hFVIII-BDD and cFVIII-BDD contained a V5-His epitope at their C-terminal end to facilitate purification and detection (Fig. 1A). All variants except for hFVIII-HC were purified to homogeneity as demonstrated by SDS-PAGE and Coomassie Blue staining (Fig. 1B). The hFVIII-HC was only partially purified due to low secretion of this variant from stably transfected cells (data not shown). Therefore, a purified preparation of hFVIII-HC (without V5-His) was obtained from Dr. Phillip J. Fay, University of Rochester, School of Medicine and Dentistry, Rochester, NY.
As shown, cFVIII-BDD was purified as a single chain protein (Mr ~160,000) (Fig. 1B, 1st lane), whereas hFVIII-BDD was purified as a two-chain protein (Mr ~90,000 and ~80,000) (Fig. 1B, 2nd lane). The hFVIII-HC and hFVIII-LC were secreted and purified as a single chain protein with Mr of ~95,000 and ~84,000, respectively (Fig. 1, B, 3rd and 4th lanes, and C, 6th and 7th lanes). Interestingly, purified hFVIII-LCΔa3 exhibited two different sizes (~80,000 and ~75,000) (Fig. 1B, 5th lane), both of which were recognized by monoclonal anti-V5 IgG on a Western blot (Fig. 1C, 8th lane), suggesting potentially aberrant N-terminal processing of hFVIII-LCΔa3 during biosynthesis.
Moreover, using a one-stage clotting assay, we had previously shown that hFVIII-BDD exhibited specific clotting activity, comparable with full-length FVIII (19), whereas cFVIII-BDD exhibited an ~3-fold increase in specific activity compared with hFVIII-BDD (23). No clotting activity was detected with hFVIII-LC, hFVIII-HC, and hFVIII-LCΔa3 (data not shown).
To determine whether the isolated light chain of FVIII, a biologically inactive FVIII variant for clot formation, accelerates VWF proteolysis by ADAMTS13, we incubated a fixed concentration of VWF (150 nm) with ADAMTS13 (50 nm) in the presence of various concentrations of recombinant hFVIII-LC (0, 0.5, 1.0, 2.5, 5.0, and 10 nm) for 10 min under constant vortex at the rotation rate of 2,500 rpm. The proteolytic cleavage of VWF was determined by SDS-PAGE and Western blotting as described under “Experimental Procedures.” Recombinant hFVIII-LC increased the formation of the proteolytic cleavage product (~350,000, a dimer of C-terminal fragments) in a concentration-dependent manner (Fig. 2, A and B). The maximal rate-enhancing effect was ~8-fold (Fig. 2B). Similar fold of rate-enhancing effect by hFVIII-LC without a V5-His epitope on the cleavage of VWF by ADAMTS13 under the same conditions was observed (data not shown). The concentration of hFVIII-LC achieving 50% of the maximal enhancing effect (C50) was ~1.0 nm (Fig. 2B), quite similar to that of full-length FVIII and hFVIII-BDD that we previously reported (19). Addition of EDTA (15 mm) into the reaction completely inhibited proteolytic cleavage of VWF by ADAMTS13 even in the presence of 10 nm hFVIII-LC (Fig. 2, A, 2nd lane, and B). These results demonstrate for the first time that the isolated light chain of FVIII is sufficient for accelerating proteolytic cleavage of VWF by ADAMTS13 under fluid shear stress.
To determine whether the light chain of FVIII or the heavy chain, both lacking the high affinity binding site for VWF, was able to enhance VWF proteolysis by ADAMTS13 under the same conditions, a fixed concentration of human VWF (150 nm) and human ADAMTS13 (50 nm) was incubated with various concentrations of recombinant hFVIII-LCΔa3 or hFVIII-HC (0, 0.5, 1.0, 2.5, 5.0, and 10 nm) for 10 min under constant vortexing at a rotation rate of 2,500 rpm. The proteolytic cleavage of VWF was determined by SDS-polyacrylamide gel and Western blotting. As shown, recombinant hFVIII-LCΔa3 (Fig. 2, C and D) or hFVIII-HC (Fig. 2, E and F) at any given concentration (up to 10 nm) did not exhibit a rate-enhancing effect on cleavage of VWF by ADAMTS13 under the same conditions. Recombinant hFVIII-BDD (20 nm) used as a positive control dramatically increased the formation of the proteolytic cleavage product (Fig. 2, 1st lane). These results suggest that the acidic a3 region in the light chain of FVIII is required for accelerating VWF proteolysis by ADAMTS13 under shear stress.
To assess the binding affinity between FVIII variants and VWF, increasing concentrations of recombinant cFVIII-BDD, hFVIII-BDD, hFVIII-LC, hFVIII-HC, and hFVIII-LCΔa3 (0, 1.56, 3.1, 6.25, 12.5, 25, 50, and 75 nm) were incubated with human VWF immobilized on a microtiter plate (1.0 μg/well). The bound FVIII variants were determined by anti-FVIII IgG or anti-V5 IgG (if V5-His tagged) as described under “Experimental Procedures.” We showed that cFVIII-BDD (Fig. 3A), hFVIII-BDD (Fig. 3B), and hFVIII-LC (Fig. 3C) bound human VWF in a concentration-dependent manner. The dissociation constants (KD) for cFVIII-BDD, hFVIII-BDD, and hFVIII-LC binding to immobilized human VWF were ~1.1, ~1.3, and ~15.6 nm, respectively (Table 1). These results indicate that the heavy chain may contribute to the overall binding affinity of FVIII to immobilized VWF. Moreover, the KD values for cFVIII-BDD, hFVIII-BDD, and hFVIII-LC to bind immobilized murine VWF was 1.9, 1.7, and 26 nm, respectively. These results suggest that there is little species difference between VWF and FVIII binding. As predicted, no detectable binding was observed between hFVIII-LCΔa3 (or hFVIII-HC) and immobilized human VWF and murine VWF under the same conditions (data not shown). These results indicate that the light chain, particularly the a3 region in the light chain of FVIII, contains the major binding site for VWF.
Although full-length FVIII, B-domainless FVIII variant, and the isolated light chain of FVIII are able to accelerate proteolytic cleavage of VWF by ADAMTS13 under mechanically induced shear stress in vitro, the physiological relevance of such an enhancing effect has never not been determined in vivo. Using a hydrodynamic approach, we were able to reconstitute plasma FVIII in fVIII−/− mice with various FVIII variants, including cFVIII-BDD, hFVIII-BDD, hFVIII-LC, and hFVIII-LCΔa3 at levels between 0.7 and 2.8 μg/ml measured 48 h after injection (Table 2). Plasma VWF antigen and multimer distribution in fVIII−/− mice were determined after 48 h of post-reconstitution. We showed that although plasma VWF antigen did not change dramatically in different groups of mice, the ratios of high to low molecular weight VWF multimers were significantly higher in fVIII−/− mice receiving saline alone (1.7 ± 0.6, n = 10) than those receiving plasmids encoding cFVIII-BDD (0.4 ± 0.4, n = 16) (p < 0.01), hFVIII-BDD (0.9 ± 0.2, n = 9) (p < 0.01), hFVIII-LC+HL (0.4 ± 0.2, n = 5) (p < 0.01), and hFVIII-LC (1.0 ± 0.3, n = 9) (p < 0.01) (Fig. 4 and Table 2). In contrast, the ratios of high to low molecular weight) VWF multimers in the fVIII−/− mice receiving plasmids encoding hFVIII-LCΔa3 (1.4 ± 0.4, n = 11) and hFVIII-HC (not shown) were not significantly different from those receiving saline alone (Fig. 4 and Table 2). These results suggest that the isolated light chain of FVIII is required and sufficient for accelerating VWF proteolysis by ADAMTS13 under (patho) physiological conditions, thus altering the distribution of VWF multimers in circulation. However, the isolated heavy chain or light chain of FVIII lacking the a3 region has no enhancing effect in VWF proteolysis under the same conditions.
In this study, we demonstrate both in vitro and in vivo that an isolated light chain of FVIII, which is biologically inactive for enhancing clot formation, appears to be sufficient for accelerating proteolytic cleavage of VWF by ADAMTS13. The maximal rate enhancing effect with 5 nm of hFVIII-LC was ~8-fold, comparable with that of hFVIII-BDD (Fig. 2, A and B). The C50 is estimated to be ~1.0 nm (Fig. 2, A and B), which is within the physiological ranges of FVIII in human plasma. In contrast, a light chain of FVIII lacking the acidic a3 region (hFVIII-LCΔa3) and hFVIII-HC, both of which do not bind VWF, exhibit no enhancing effect under the same conditions (Fig. 2, C–F). These results indicate that high affinity binding of FVIII to VWF through the acidic a3 region may be critical for accelerating VWF proteolysis.
However, the rate-enhancing effect of FVIII on VWF proteolysis does not appear to be in a linear relationship with its VWF binding affinity. For instance, although hFVIII-LC binds human VWF (KD = 15 nm) and murine VWF (KD = 26 nm) ~10 times less than binding of hFVIII-BDD and full-length FVIII to human VWF (KD = 1.3 nm) and murine VWF (KD = 1.9 nm) (Table 1), the rate-enhancing activity is quite similar to that of hFVIII-BDD (Fig. 2) and full-length FVIII (19).
The biological relevance of FVIII and its variants on regulating VWF proteolysis by ADAMTS13 was further assessed using a murine model. At the steady state, plasma levels of VWF antigen in fVIII−/− mice are increased by ~2-fold when compared with wild-type mice in the same genetic background C57BL6/129 (data not shown), whereas the ratio of high to low molecular weight VWF multimers was not statistically different between the two groups of mice. Similar increases in plasma levels of VWF antigen and ristocetin-cofactor activity were observed in patients with severe hemophilia A compared with healthy controls (32). These findings suggest that FVIII may be involved in regulating VWF homeostasis under physiological conditions. However, the interpretation of these data may be complicated by the fact that plasma VWF antigen levels differ significantly among various strains of mice or human individuals.
Although hydrodynamic challenge is considered to be a nonphysiological transfection method, it activates endothelial cells and triggers the release of ultra-large VWF from endothelial cells, resulting in an accumulation of ultra-large VWF multimers in plasma of the fVIII−/− mice receiving saline alone. However, these ultra-large VWF multimers were not observed in the same mice receiving plasmids encoding cFVIII-BDD, hFVIII-BDD, and hFVIII-LC or hFVIII-LC+HC (Fig. 4 and Table 2). These results demonstrate for the first time that reconstitution of functional and nonfunctional FVIII variants restores the distribution of plasma VWF multimers in severe hemophilia A mice.
Consistent with the in vitro data, reconstitution in the fVIII−/− mice with a plasmid encoding hFVIII-LCΔa3 (Fig. 4 and Table 2) has no effect on plasma VWF multimer distribution. However, the in vivo effect of hFVIII-HC alone remains to be determined because plasma levels of the expressed hFVIII-HC were low, only 1/10 of the plasma levels of other constructs after the hydrodynamic injection (data not shown). The low expression of the heavy chain is due to inefficient secretion of this chain when it is expressed alone (24, 26). Interestingly, co-expression of hFVIII-LC dramatically improves the secretion of the heavy chain, thereby synergistically enhancing proteolytic cleavage of VWF by ADAMTS13 in vivo. The efficacy of the co-expressed hFVIII-LC and hFVIII-HC appears to be similar to that of hFVIII-BDD but better than that of hFVIII-LC alone (Table 2), suggesting that the heavy chain is able to stabilize the light chain to improve the light chain function.
The implication of these findings is not clear. Patients with severe hemophilia A (with FVIII activity <1%) are heterogeneous in their clinical presentations (31). Further investigation of plasma VWF multimer distribution in correlation with the genetic basis that results in severe FVIII deficiency may shed new light on how FVIII-dependent proteolysis of VWF may play a role in the modification of the clinical phenotype in patients with severe hemophilia A.
In conclusion, our findings provide novel insight into the structure-function relationship of FVIII in the regulation of ADAMTS13-mediated proteolysis in vitro under fluid shear stress and in vivo under (patho) physiological conditions. This may help explain the heterogeneity of plasma VWF multimer distribution and clinical phenotype in patients with hemophilia A.
We thank Philip J. Fay (Professor of Biochemistry, University of Rochester, School of Medicine and Dentistry, Rochester, NY) for providing us with purified recombinant hFVIII-HC (no tag) (data not shown). We also thank Sriram Krishnaswamy (Professor, Division of Hematology, The Children's Hospital of Philadelphia) for key reagents and helpful comments and discussion.
*This work was supported, in whole or in part, by National Institutes of Health Grants HL079027 and HL074124. This work was also supported by American Heart Association Postdoctoral Fellowship 09POST2250397 (to W. C.), Bayer Hemophilia awards, and American Heart Association Grant 0940100N (to X. L. Z.).
3The abbreviations used are: