Endothelial cells and megakaryocytes
Since VWF is being synthesized exclusively by either endothelial cells or megakaryocytes, these primary cells would be the most suitable and effective in vitro system to study VWD. However, due to elements such as location and culture issues, the use of these cells appears to be difficult.
Human umbilical vein endothelial cells (HUVECs) are frequently used to study VWF synthesis and secretion in the healthy situation. HUVECs from healthy individuals are readily available, as umbilical veins are resected after childbirth, and require simple isolation techniques.2 Besides the use of these normal HUVECs for VWF focused research,9–11 there have been studies performed on HUVECs of newborns with inherited VWD. This has been successful for several types of VWD, looking at different aspects, such as cell morphology, VWF production, storage, secretion and activity levels, multimerization patterns, factor VIII production, and platelet binding.12–16
Even though HUVECs with a VWD phenotype have contributed to our knowledge of this bleeding disorder, instances where these disease-specific cells are available are rare, especially when one is interested in studying a specific VWF defect. Therefore, patient-derived (adult) endothelial cells would be a valuable source to study all VWD (sub)types, however, due to their anatomical location, there is the need for invasive procedures to access these cells.17 Vascular cell isolation can be done by limb vein stripping, but only when patients undergo surgeries such as coronary artery bypass grafting, thus limiting the availability of tissues with a VWD phenotype.18 Furthermore, in in vitro cultures primary endothelial cells readily dedifferentiate and go into senescence, which is a major drawback of these cells.19
Megakaryocytes and platelets
Like primary endothelial cells, invasive procedures (bone marrow puncture) are needed to collect megakaryocytes, the precursor cells that form and release platelets. Megakaryocytes use similar VWF processing steps to endothelial cells, whilst storing VWF in the α-granules.3 These vesicles appear to be heterogeneous in cargo and, in contrast to WPBs, not all α-granules contain VWF.20
There have been few studies using megakaryocytes, either isolated from bone marrow21 or differentiated from circulating hematopoietic stem (CD34+) cells in peripheral blood, from VWD patients. A study in 2010 compared the production of platelets from differentiated megakaryocytes and isolated megakaryocytes from a patient with VWD type 2. They showed the regulatory role of VWF-GPIbα interactions in megakaryocytopoiesis and that this was aberrant leading to abnormal platelet structure and production, typical of type 2B VWD.22
However, in-depth knowledge about the production, storage and secretion of VWF in megakaryocytes and platelets in regards to VWD is restricted. Only a few papers have been published recently and most of these reports are on platelets, reviewed in 2010, with the majority of references from before 1990.23 The limited research concerning VWF in megakaryocytes and platelets is most likely linked to the difficulties the collection and isolation bring. Megakaryocytes represent a population of less than 0.05% of nucleated cells in the bone marrow, and therefore hampered by location as well as scarcity.24 This can be overcome by generating megakaryocytes from CD34+ blood progenitor cells from either peripheral or cord blood. However, these hematopoietic stem cells circulate in low numbers, which means these stem cells need to be expanded in vitro first before being efficiently differentiated into megakaryocytes, and subsequently platelets.
Furthermore, when able to isolate or generate megakaryocytes and/or platelets, large scale production might be problematic. This approach requires continuous supply of donor material due to limited expansion potential. Because there is no optimal expansion protocol for in vitro megakaryocyte and platelet production, this is currently not a suitable source to generate a reproducible, high throughput cell model for VWD.
Heterologous cell systems
In view of the limited accessibility and challenges of primary cells expressing VWF, in vitro transfection experiments in heterologous cell systems have become an attractive alternative to study the pathophysiology of VWD. Because not all cell types transfected with VWF act similarly regarding the biosynthetic pathway of VWF, this can be applied to study specific mechanisms of VWF mutations.25 These systems are also an effective approach for co-transfections of alleles, making it possible to look at the effect of heterozygous variants in the VWF gene known to be causative of VWD.
Cell types used in these experiments can be divided into different groups, based on the processing and storage pathways of VWF (Table 2). Certain cells, such as COS (monkey kidney tissue) and CHO (Chinese hamster ovary epithelium) cell lines, do produce VWF upon transfection, but lack a regulated secretory pathway and are therefore hindered to store VWF. These lines have shown to be suitable for experiments specifically looking at VWF synthesis, multimerization and constitutive basal secretion.26 Particularly, COS-1 and COS-7 lines have been instrumental in elucidation of mechanisms of VWF multimerization; when transfected transiently with VWF, it was revealed that in addition to the propeptide, both the D’ and D3 domains are required for multimer assembly.27,28
More specific, COS cells transfected with VWF harboring cysteine mutations in the D3 domain (p.C1157F or p.C1234W), displayed (and released) only the LMWMs and showed intracellular retention of these mutated forms of VWF in pre-Golgi compartments.29 Co-transfections with wild-type and mutant VWF alleles in COS cells showed that both cysteine substitutions reduced the release of wild-type VWF in a dose dependent manner and these cells failed to form HMWMs. Therefore, even though these cells do not store VWF in vesicles, they are still a valuable tool to study the conformation of the VWF molecules required for a normal transport pathway, maturation and constitutive secretion.
Interestingly, in some of these cell lines, heterologous VWF expression can induce the formation of de novo WPB-like storage vehicles. This group of cells can give insights in the storage and regulated secretion of VWF and this has been shown for cell lines such as HEK293 (human embryonic kidney cells) (Fig. 2) and CV-1 (monkey kidney cells).25 Especially HEK293 cells have been used extensively in VWF research and are still a frequent and preferred cell line of choice. For example, this approach has led to the discovery that the non-covalent interaction between the propeptide (D1-D2 domains) and the D’-D3-A1 domains of VWF is essential for tubulation and storage of VWF into WPBs.30,31 These cells have also been used to look at specific known VWD causing mutations, by transfecting HEK293 cells with modulated VWF constructs.32–34 Our group has recently applied transfected HEK293 cells to investigate a potential small interfering RNA (siRNA) based therapeutic approach for VWD.35 Here we applied siRNAs targeting common heterozygous single nucleotide variants (SNVs) to distinguish alleles harboring heterozygous VWF mutations to inhibit the production of mutated alleles. This allele-specific knockdown of the mutant allele resulted in an improved multimerization pattern, which is aberrant in VWD type 2A.
Finally, there are the cell lines that comprise a regulated secretory pathway, containing endogenous secretory granules, where the exogenous VWF is stored. AtT20 (mouse pituitary tumor) and RIN-5F (rat pancreas tumor) lines belong to this group of cells. RIN-5F cells store the transfected VWF in their vesicles containing other (endogenous) proteins, whereas AtT20 cells produce granules containing merely VWF.36,37 A study with AtT20 cells showed that the VWF propolypeptide is necessary for the formation of the VWF-containing granules in the AtT20 cells, contributing to the knowledge about the generation of WPBs.38
Staining with VWF antibodies in these cell systems show that, even though containing VWF, most of these storage vesicles, both endogenous and exogenous, are roundly shaped. However, in combination with electron microscopy, there are rod-shaped structures detected with tubulated VWF. These resemble endothelial WPBs and are known as pseudo-WPBs.25,38
Altogether, these heterologous cell systems have provided an enormous amount of knowledge on the synthesis, storage and secretion of VWF and thus the pathophysiology of VWD. Regardless of the value, these cell models have obvious limitations since only few non-endothelial cell lines can target recombinant VWF to WPB-like organelles, restricting the applications.39 This needs to be taken into consideration, as WPBs are an important aspect in the processing of VWF. Moreover, WPBs are actually dependent on the tubular assembly of VWF, which drives the formation of these storage vesicles. Because these transfection experiments show that not all cells are able to form WPBs after VWF expression, it is suggested that these cell types may lack the necessary chaperone or adaptor proteins.40 Another obvious drawback is the possible aberrant effects overexpression of recombinant VWF with viral promoters will have upon transfection in (non-endothelial) cells.
Furthermore, VWD patients are mostly heterozygous for VWF mutations, which can be modeled in cell systems by co-transfections. It is not a guarantee that co-transfections with wild-type and mutant alleles will lead to an even ratio in expression between the alleles. However, acknowledging the shortcomings, these heterologous cell systems, especially HEK293 cells, have and continue to contribute tremendously towards VWF and VWD research.
Endothelial colony forming cells (ECFCs)
VWF transfection in HEK293 cells has been one of the most recognized model to study VWD for many years. However, in 2000 Hebbels group identified endothelial cells that can be isolated and cultured from peripheral blood, with acceptable expansion capacity in vitro.41 These cells, by consensus recently named endothelial colony forming cells (ECFCs), but also previously referred to as blood outgrowth endothelial cells (BOECs) or late outgrowth endothelial cells,42 represent a population of endothelial cells with a progenitor status, harboring clonal proliferation. They contain the distinct properties and features of endothelial cells such as a cobblestone morphology, expression of endothelial cell surface antigens, and the presence of endogenous WPBs. Likewise, these organelles in ECFCs store VWF, which is released after stimulation, indistinguishable from other endothelial cells43 (Fig. 3). These endothelial-like characteristics, in combination with the simple venepuncture to collect, make ECFCs ideal to study VWF and VWD (and other bleeding/vascular disorders) together with possible therapeutic assessment. We and others have described the use of ECFCs as a feasible cell model to study the effect of VWF mutations on the synthesis, storage, secretion, and string formation of VWF, but also on abnormal cell proliferation, migration, and increased Ang-2 secretion.44–51
VWF and angiogenesis
VWF has been described as a negative regulator of angiogenesis, and ECFCs have been applied to investigate the pathogenesis of VWD and angiodysplasia.51 Starke et al found in HUVECs that knock-down of VWF expression leads to an increase in angiogenesis, which was also shown with ECFCs isolated from patients with VWD, showing enhanced angiogenesis.51 Our group applied ECFCs to investigate the potential pathogenic effect of specific VWF mutations on angiogenesis.46 ECFCs, isolated from a type 3 and type 2B patient, displayed increased migratory velocity, and in the majority of VWD ECFCs (8 out of 10) directional migration was impaired. This shows that ECFCs directly isolated from VWD patients displayed the disease phenotype and serve as a good source to study VWF and VWD.45,49,51
Recently, Selvam et al used ECFCs derived from healthy donors to investigate and determine the normal range of angiogenesis. Subsequently, this was compared to angiogenesis levels in ECFCs from VWD patients (all types and subtypes).49 ECFCs were assessed for VWF and Ang-2 gene expression, secretion and storage and characterized for cellular proliferation, matrix protein adhesion, migration, and tube formation. Overall, the results indicated that there is great variability in the angiogenic properties of both control and VWD ECFCs.
Challenges and opportunities of ECFCs
Even though VWD ECFCs reflect the pathogenic effects of VWF mutations, significant heterogeneity was observed among individual VWD phenotypes. For ECFCs to be a more robust cell model for use in disease studies, the extent of variability of cellular phenotype needs to be understood in more detail. We recently studied a cohort of separate ECFC clones derived from six healthy donors and observed large variations between ECFCs. Not only from different donors, but also amongst clones from individual donors. This variability both in morphology and proliferative potential is being investigated and the origin and age of the ECFC might be involved. However, this should be taken into account when using ECFCs as a cell model.52
There is also the need for standardization of protocols to be able to compare findings across laboratories.53 ECFC isolation and culture methods vary between research groups, and this might affect the cells proliferative capacity and possibly phenotype.50 An additional challenge is the rather low success rate of ECFC isolations from peripheral blood (between 40% and 60%). An explanation for these low scores could be the general scarcity of ECFCs in peripheral blood. Kolbe et al showed that ECFCs represent a small fraction of mononuclear cells and their numbers vary considerably between donors54 which might also be influenced by pathological conditions, affecting the number of ECFCs circulating in the blood.55
In conclusion, there are some challenges and unknowns involved in the isolation and usage of ECFCs such as success rate, low numbers of circulating ECFCs, and whether this is person specific or disease related. Nevertheless, by the use of these cells in in vitro studies and disease modeling, ECFCs show potential in applications such as drug screening, bioengineering approaches, as well as cell and gene therapies.56 ECFCs have made it possible to profile the synthesis and storage of VWF in endothelial cells from individual VWD patients44,45 and these results could be of use in the choice of patient-specific therapeutic approaches. This is not only applicable to VWD but also relevant for other diseases that have been studied using ECFCs, such as sickle cell anemia, myeloproliferative neoplasms, hereditary hemorrhagic telangiectasia, and venous thromboembolic disease (reviewed in57). In addition to these forward developments, ECFCs have also shown to be a source to reprogram into iPSCs.58
Induced pluripotent stem cells (iPSCs)
Reprogramming and differentiation of somatic cells can overcome the lack of disease-specific cells and tissues in disorders like VWD. The generation of iPSCs from human fibroblasts was first achieved in 2007,59 and several different cell types, such as peripheral blood mononuclear cells (PBMCs) and keratinocytes, have since been reprogrammed.60,61 Nowadays, other cell types, with simple non-invasive collection procedures, have also been reprogrammed, such as cells from hair follicles, urine, and dental pulp from milk teeth.62,63 Generated iPSCs are capable of self-renewal and have the potential to differentiate into almost any cell type, and can thereby create a disease in a dish model. This is especially relevant for cells of the internal organs for which biopsies are not routinely available, such as megakaryocytes and endothelial cells. Acquiring these cells through patient-specific iPSC differentiation can enable better insights into VWD and other bleeding disorders, in combination with additional aims such as (high-throughput) drug screening, development and cell therapy.
ECFCs as reprogramming source
ECFCs have been used as a reprogramming source, with various reprogramming efficiency and results depending on the method to deliver the reprogramming factors58,62,64 compared to fibroblast or PBMCs. Regardless, these ECFC-derived iPSC colonies (ECFC-iPSCs) express embryonic stem cell markers and show differentiation potential in all three germ layers in vitro, similar to other somatic cell sources. Furthermore, it has been shown that reprogrammed ECFCs show slightly lower rates of acquired chromosomal abnormalities (compared to the parental cells) in contrast to the other somatic cells tested.58 However, to obtain the required cell number, ECFCs need a longer culture period than most other somatic cells such as fibroblast, which could induce genetic changes.
Unfortunately, limited studies have been published where reprogrammed ECFCs have been used in differentiation assays. Orlova et al differentiated ECFC-iPSCs into endothelial cells and showed that these cells expressed similar levels of endothelial-specific surface antigens and perform similar in functional assays compared to endothelial cells from either fibroblast-iPSCs and or differentiated from human embryonic stem cells.65
Furthermore, there are reports that endothelial cell-derived iPSCs have a higher tendency to differentiate to endothelial cells, compared to fibroblast-iPSCs.66 This suggests that ECFCs as an iPSC source might be beneficial when differentiation into the endothelial lineage is required, with regards to epigenetic memory and has been reported for a variety of cell types, which predispose iPSCs to favor differentiation towards their cell of origin (reviewed in 67). This is an important trait when considering the use of iPSCs in disease modeling, drug screening and future (autologous) cell transplantation therapies.
The fact that iPSCs are self-proliferative, in contrast to ECFCs which will go into senescence and/or loose the endothelial morphology after multiple passages, ECFCs as a somatic source could therefore have an advantage and preference when differentiating into endothelial cells for further applications and assays.
Differentiation of iPSCs
There are several protocols and approaches to differentiate human iPSCs from different somatic cell sources into endothelial cells, such as three-dimensional embryoid bodies or co-cultures with stromal cells to induce endothelial cell lineage differentiation.68 However, most groups use feeder-free monolayer differentiation on matrix coated culture plates, such as Matrigel or fibronectin, using specific culture mediums with sequentially added recombinant growth factors.69,70 The majority of this protocol can be divided into a mesoderm and endothelial differentiation phase. Two populations are generated after the first differentiation round, based on CD31 positive and negative populations. The CD31+ cells are further differentiated into endothelial cells, while the CD31- group can be differentiated into pericytes for co-culture purposes. This will generate substantial numbers of both cell types that can be derived in only 2 to 3 weeks. These cells show the typical endothelial cell-like morphology and express endothelial markers such as vascular endothelial (VE-) cadherin and VWF, and have been used in functional studies (Fig. 3).71
Regardless, there are some concerns about the efficiency and maturity of iPSC-derived endothelial cells using these protocols. Most endothelial differentiation protocols that have been developed to date generate low endothelial cell yields. Like primary endothelial cells in vitro, these iPSC-derived endothelial cells have restricted proliferative potential and either undergo senescence or endothelium-to-mesenchymal transition after multiple passages. Because these are differentiated in vitro, they are not exposed to impacts from the (tissue) specific environment, such as blood flow and pressure that play roles in endothelial cell differentiation in vivo. This might explain the intermediate (heterogenous) phenotype seen in the population of differentiated endothelial cells; neither committed fully to an arterial or venous fate.72 Evidence has also emerged that endothelial cells reprogrammed from iPSCs possess slightly different gene expression and epigenetic patterns compared to primary endothelial cells.73 However, research into the maturation of these differentiated endothelial cells into more specific types of endothelium (arterial, venous, lymphatic) through the manipulation of culture media is ongoing. To mimic these in vivo environments more closely, micro-fluidic 3D systems, like organ-on-a-chip, have been developed.
To generate platelets, pluripotent cells need to be differentiated into megakaryocytes first and this can be done through CD34+ progenitor cells derived from sources such as peripheral or umbilical blood, bone marrow, or iPSCs.74–76 Most of these sources have limitations for large scale regeneration of megakaryocytes and platelets and require a continuous supply of donors. However, iPSCs show capacity to serve as a renewable and unlimited cell source that can be expanded in culture and differentiated into megakaryocytes and platelets. The attractiveness of this system is that the same patient-specific iPSCs batch can be used to differentiate into a variety of cell types of interest. With regards to VWD, besides endothelial cell differentiation, these iPSCs can also be used to generate megakaryocytes and subsequently platelets.
Recently, megakaryocytes have been differentiated from CD34+ cells from patients with Roifman syndrome, which is a rare congenital disorder characterized by growth retardation, cognitive delay and in some patients thrombocytopenia.77 The patient-specific CD34+ cells showed defects in megakaryocytes differentiation, with inadequate generation of proplatelets which is a characteristic of this syndrome. This indicates that a differentiation approach from iPSCs into megakaryocytes can be used for disease modeling.
By reprogramming the same somatic cell source and using identical protocols, the differentiation of cells into endothelial cells, might reduce the variation seen in ECFCs and therefore possibly lead to a model for vascular disease.
Potential drawbacks of iPSCs
Even though human iPSCs have emerged as a promising candidate for vascular regeneration medicine,70,78 there are still some issues concerning the use of pluripotent cells as a source for cell therapies. Things like low reprogramming efficiency, genetic instability, in vivo functionality and the risk of teratoma formation from undifferentiated iPSCs are serious matters that are currently under investigation and need to be resolved before further application into the clinic. Research is developing rapidly, to improve the differentiation efficiency of endothelial cells and megakaryocytes from iPSCs by new signaling pathways and novel culture conditions. A deeper understanding of the development of the endothelial cell lineage is required for differentiated cells to become a robust model for vascular diseases and the potential to the safe use of these cells as a patient-specific cell therapy in future.
For some research questions cell models are not applicable and therefore animal models are required. Naturally, occurring VWD has been described amongst several animal species, especially mammals, such as dogs and pigs.79–82 These animal models have contributed heavily to the understanding of various aspects of VWF and VWD that cannot be addressed through in vitro approaches.
VWD in an animal was first reported in 1941, when a bleeding disorder was described in swine by Hogan et al,80 making this the oldest known animal model of a human bleeding disorder. Later on, it was revealed that the disease was transmitted as an autosomal recessive trait and identified as a VWD type 3 model.83 The pig is a good model of hemorrhagic disorders since its clotting and platelet characteristics resemble those of humans.84 However, there are some important differences when comparing this to human VWD. Homozygous pigs are not totally deficient in VWF, and low but significant amounts of VWF antigen can be detected both in platelets and in endothelial cells from the pulmonary artery and inferior vena cava.85 Another restriction of this model is its size and housing cost of the animals and nowadays, research has been restricted with this animal model due to new and better alternatives.
High costs also apply to the canine model of VWD. The first dog reported with this bleeding disorder was in 1970 in a German Shepherd family86 and over the years many other dog breeds have been identified with all three subtypes of VWD. However, canine VWD is a very heterogeneous group of diseases with different subtypes and modes of inheritance87 and limited research has been done using dogs as a VWD model in contrast to the porcine model. In 2006, De Meyer et al transduced ECFCs, isolated from type 3 VWD dogs, with VWF, showing that gene therapy of type 3 VWD is feasible.88
For murine VWD, both naturally and genetically engineered VWF knockout mouse models are available.89–91 The naturally occurring RIIIS/J mouse, is characterized by typical VWD features such as a prolonged bleeding time and low levels of plasma factor VIII and VWF, however, presenting with a normal VWF multimer distribution.91 Genetic linkage did not show a correlation between VWF antigen level and genotype, suggesting that VWD in this mouse strain is caused by a defect at a novel genetic locus, distinct from the murine VWF gene.92
By breeding the genetically engineered VWF knockout mouse, a model has been designed that mimics VWF deficiency (type 1 and 3). The generated VWF knockout mouse has been used to generate type 2B VWD models. This has been a great advantage in VWD research as naturally occurring VWD type 2 in animals is very rare. VWD type 2B in humans is characterized by gain-of-function mutations leading to increased affinity for its platelet-receptor, GPIbα. This phenotype has first been replicated in mice by hydrodynamic injection of mutant VWF constructs into the Vwf-/- mice.93,94 Even though mimicking VWD type 2B, showing enhanced platelet binding but normal multimerization, the mutant VWF proteins with this approach are (exclusively) expressed by the hepatocytes and therefore only present in the plasma, and not in endothelial cells or platelets. More recently, 2 groups have successfully engineered a VWD type 2B knock-in model in mice.95,96 Adam et al introduced the VWD causing mutation p.V1316 M into the murine Vwf locus and these mice display human VWD-type 2B-like characteristics, such as macrothrombocytopenia, deficiency of HMWM VWF, reduction of active VWF levels, circulating platelet-aggregates and a severe bleeding tendency.95
Mouse models have advantages such as characterized genetic backgrounds, optimized genome editing approaches, and low cost and maintenance of housing, making this an attractive model to study disease. However, many disorders lack a suitable animal model or are not feasible to use for these experiments. Additionally, there are differences between various species and care needs to be taken when extrapolating results obtained in these studies to humans. Nevertheless, animal models will most likely remain an important model and a requirement in many aspects of disease and drug modeling for the coming years.
Altogether, the collection of these disease models, both in vitro and in vivo, has led to a major part of the knowledge on VWF and the mechanisms underlying VWD (Fig. 1). The generation and differentiation of iPSCs, along with the availability of the human genome and genome editing tools has transformed disease research immensely, leading to the development of new strategies to treat or study vascular diseases.97 This approach has already been applied in combination with human iPSCs to correct defective genotypes in vitro for several diseases and conditions, such as hemophilia A.98 Studies like these are good examples of combining patient-specific vascular regenerative medicine (in vitro) approaches and animal models. These developments, in combination with three dimensional models, such as organ-on-a-chip and drug screens, could be at the basis to generate novel (cell) therapies for VWD.
1. Leebeek FW, Eikenboom JC. Von Willebrand's disease. N Engl J Med.
2. Jaffe EA, Nachman RL, Becker CG, et al. Culture of human endothelial cells derived from umbilical veins. Identification by morphologic and immunologic criteria. J Clin Invest.
3. Sporn LA, Chavin SI, Marder VJ, et al. Biosynthesis of von Willebrand protein by human megakaryocytes. J Clin Invest.
4. Wagner DD. Cell biology of von Willebrand factor. Annu Rev Cell Biol.
5. Springer TA. von Willebrand factor, Jedi knight of the bloodstream. Blood.
6. Cramer EM, Breton-Gorius J, Beesley JE, et al. Ultrastructural demonstration of tubular inclusions coinciding with von Willebrand factor in pig megakaryocytes. Blood.
7. Gralnick HR, Williams SB, McKeown LP, et al. Platelet von Willebrand factor: comparison with plasma von Willebrand factor. Thromb Res.
8. Sadler JE, Budde U, Eikenboom JC, et al. Update on the pathophysiology and classification of von Willebrand disease: a report of the Subcommittee on von Willebrand Factor. J Thromb Haemost.
9. Mourik MJ, Faas FG, Zimmermann H, et al. Content delivery to newly forming Weibel-Palade bodies is facilitated by multiple connections with the Golgi apparatus. Blood.
10. Valentijn KM, van Driel LF, Mourik MJ, et al. Multigranular exocytosis of Weibel-Palade bodies in vascular endothelial cells. Blood.
11. Wang JW, Valentijn JA, Valentijn KM, et al. Formation of platelet-binding von Willebrand factor strings on non-endothelial cells. J Thromb Haemost.
12. Booyse FM, Quarfoot AJ, Chediak J, et al. Characterization and properties of cultured human von Willebrand umbilical vein endothelial cells. Blood.
13. de Groot PG, Federici AB, de Boer HC, et al. von Willebrand factor synthesized by endothelial cells from a patient with type IIB von Willebrand disease supports platelet adhesion normally but has an increased affinity for platelets. Proc Natl Acad Sci U S A.
14. Ewenstein BM, Inbal A, Pober JS, et al. Molecular studies of von Willebrand disease: reduced von Willebrand factor biosynthesis, storage, and release in endothelial cells derived from patients with type I von Willebrand disease. Blood.
15. Federici AB, de Groot PG, Moia M, et al. Type I von Willebrand disease, subtype ‘platelet low’: decreased platelet adhesion can be explained by low synthesis of von Willebrand factor in endothelial cells. Br J Haematol.
16. Levene RB, Booyse FM, Chediak J, et al. Expression of abnormal von Willebrand factor by endothelial cells from a patient with type IIA von Willebrand disease. Proc Natl Acad Sci U S A.
17. Waldo SW, Brenner DA, McCabe JM, et al. A novel minimally-invasive method to sample human endothelial cells for molecular profiling. Plos One.
18. Perry L, Flugelman MY, Levenberg S. Elderly patient-derived endothelial cells for vascularization of engineered muscle. Mol Ther.
19. Nguyen MTX, Okina E, Chai X, et al. Differentiation of human embryonic stem cells to endothelial progenitor cells on laminins in defined and xeno-free systems. Stem Cell Rep.
20. Italiano JE Jr, Richardson JL, Patel-Hett S, et al. Angiogenesis is regulated by a novel mechanism: pro- and antiangiogenic proteins are organized into separate platelet alpha granules and differentially released. Blood.
21. Bury L, Malara A, Momi S, et al. Mechanisms of thrombocytopenia in platelet-type von Willebrand disease. Haematologica.
22. Nurden P, Gobbi G, Nurden A, et al. Abnormal VWF modifies megakaryocytopoiesis: studies of platelets and megakaryocyte cultures from patients with von Willebrand disease type 2B. Blood.
23. McGrath RT, McRae E, Smith OP, et al. Platelet von Willebrand factor--structure, function and biological importance. Br J Haematol.
24. Ebaugh FG Jr, Bird RM. The normal megakaryocyte concentration in aspirated human bone marrow. Blood.
25. Voorberg J, Fontijn R, Calafat J, et al. Biogenesis of von Willebrand factor-containing organelles in heterologous transfected CV-1 cells. EMBO J.
26. Bonthron DT, Handin RI, Kaufman RJ, et al. Structure of pre-pro-von Willebrand factor and its expression in heterologous cells. Nature.
27. Voorberg J, Fontijn R, van Mourik JA, et al. Domains involved in multimer assembly of von willebrand factor (vWF): multimerization is independent of dimerization. EMBO J.
28. Berber E, Ozbil M, Brown C, et al. Functional characterisation of the type 1 von Willebrand disease candidate VWF gene variants: p.M771I, p.L881R and p P1413L. Blood Transfus.
29. Hommais A, Stepanian A, Fressinaud E, et al. Mutations C1157F and C1234W of von Willebrand factor cause intracellular retention with defective multimerization and secretion. J Thromb Haemost.
30. Huang RH, Wang Y, Roth R, et al. Assembly of Weibel-Palade body-like tubules from N-terminal domains of von Willebrand factor. Proc Natl Acad Sci U S A.
31. Michaux G, Abbitt KB, Collinson LM, et al. The physiological function of von Willebrand's factor depends on its tubular storage in endothelial Weibel-Palade bodies. Dev Cell.
32. Groeneveld DJ, Wang JW, Mourik MJ, et al. Storage and secretion of naturally occurring von Willebrand factor A domain variants. Br J Haematol.
33. Wang JW, Groeneveld DJ, Cosemans G, et al. Biogenesis of Weibel-Palade bodies in von Willebrand's disease variants with impaired von Willebrand factor intrachain or interchain disulfide bond formation. Haematologica.
34. Wang JW, Valentijn KM, de Boer HC, et al. Intracellular storage and regulated secretion of von Willebrand factor in quantitative von Willebrand disease. J Biol Chem.
35. de Jong A, Dirven RJ, Oud JA, et al. Correction of a dominant-negative von Willebrand factor multimerization defect by small interfering RNA-mediated allele-specific inhibition of mutant von Willebrand factor. J Thromb Haemost.
36. Blagoveshchenskaya AD, Hannah MJ, Allen S, et al. Selective and signal-dependent recruitment of membrane proteins to secretory granules formed by heterologously expressed von Willebrand factor. Mol Biol Cell.
37. Haberichter SL, Fahs SA, Montgomery RR. von Willebrand factor storage and multimerization: 2 independent intracellular processes. Blood.
38. Wagner DD, Saffaripour S, Bonfanti R, et al. Induction of specific storage organelles by von Willebrand factor propolypeptide. Cell.
39. Hannah MJ, Williams R, Kaur J, et al. Biogenesis of Weibel-Palade bodies. Semin Cell Dev Biol.
40. Haberichter SL. von Willebrand factor propeptide: biology and clinical utility. Blood.
41. Lin Y, Weisdorf DJ, Solovey A, et al. Origins of circulating endothelial cells and endothelial outgrowth from blood. J Clin Invest.
42. Medina RJ, Barber CL, Sabatier F, et al. Endothelial progenitors: a consensus statement on nomenclature. Stem Cells Transl Med.
43. van den Biggelaar M, Bouwens EA, Kootstra NA, et al. Storage and regulated secretion of factor VIII in blood outgrowth endothelial cells. Haematologica.
44. Starke RD, Paschalaki KE, Dyer CE, et al. Cellular and molecular basis of von Willebrand disease: studies on blood outgrowth endothelial cells. Blood.
45. Wang JW, Bouwens EA, Pintao MC, et al. Analysis of the storage and secretion of von Willebrand factor in blood outgrowth endothelial cells derived from patients with von Willebrand disease. Blood.
46. Groeneveld DJ, van Bekkum T, Dirven RJ, et al. Angiogenic characteristics of blood outgrowth endothelial cells from patients with von Willebrand disease. J Thromb Haemost.
47. Bowman ML, Pluthero FG, Tuttle A, et al. Discrepant platelet and plasma von Willebrand factor in von Willebrand disease patients with p.Pro2808Leufs∗24. J Thromb Haemost.
48. Hawke L, Bowman ML, Poon MC, et al. Characterization of aberrant splicing of von Willebrand factor in von Willebrand disease: an underrecognized mechanism. Blood.
49. Selvam SN, Casey LJ, Bowman ML, et al. Abnormal angiogenesis in blood outgrowth endothelial cells derived from von Willebrand disease patients. Blood Coagul Fibrinolysis.
50. Sadler JE. von Willebrand factor in its native environment. Blood.
51. Starke RD, Ferraro F, Paschalaki KE, et al. Endothelial von Willebrand factor regulates angiogenesis. Blood.
52. de Jong A, Weijers E, Dirven R, et al. Variability of von Willebrand factor-related parameters in endothelial colony forming cells. J Thromb Haemost.
2019;(In press; DOI: 10.1111/jth.14558).
53. Smadja DM, Melero-Martin JM, Eikenboom J, et al. Standardization of methods to quantify and culture endothelial colony-forming cells derived from peripheral blood: Position paper from the International Society on Thrombosis and Haemostasis SSC. J Thromb Haemost.
54. Kolbe M, Dohle E, Katerla D, et al. Enrichment of outgrowth endothelial cells in high and low colony-forming cultures from peripheral blood progenitors. Tissue Eng Part C Methods.
55. Liew A, Barry F, O’Brien T. Endothelial progenitor cells: diagnostic and therapeutic considerations. Bioessays.
56. O’Neill CL, O’Doherty MT, Wilson SE, et al. Therapeutic revascularisation of ischaemic tissue: the opportunities and challenges for therapy using vascular stem/progenitor cells. Stem Cell Res Ther.
57. Paschalaki KE, Randi AM. Recent advances in endothelial colony forming cells toward their use in clinical translation. Front Med (Lausanne).
58. Geti I, Ormiston ML, Rouhani F, et al. A practical and efficient cellular substrate for the generation of induced pluripotent stem cells from adults: blood-derived endothelial progenitor cells. Stem Cells Transl Med.
59. Takahashi K, Tanabe K, Ohnuki M, et al. Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell.
60. Aasen T, Raya A, Barrero MJ, et al. Efficient and rapid generation of induced pluripotent stem cells from human keratinocytes. Nat Biotechnol.
61. Staerk J, Dawlaty MM, Gao Q, et al. Reprogramming of human peripheral blood cells to induced pluripotent stem cells. Cell Stem Cell.
62. Dambrot C, van de Pas S, van Zijl L, et al. Polycistronic lentivirus induced pluripotent stem cells from skin biopsies after long term storage, blood outgrowth endothelial cells and cells from milk teeth. Differentiation.
63. Raab S, Klingenstein M, Liebau S, et al. A comparative view on human somatic cell sources for iPSC generation. Stem Cells Int.
64. Ormiston ML, Toshner MR, Kiskin FN, et al. Generation and culture of blood outgrowth endothelial cells from human peripheral blood. J Vis Exp.
65. Orlova VV, Drabsch Y, Freund C, et al. Functionality of endothelial cells and pericytes from human pluripotent stem cells demonstrated in cultured vascular plexus and zebrafish xenografts. Arterioscler Thromb Vasc Biol.
66. Hu S, Zhao MT, Jahanbani F, et al. Effects of cellular origin on differentiation of human induced pluripotent stem cell-derived endothelial cells. JCI Insight.
67. Kim M, Costello J. DNA methylation: an epigenetic mark of cellular memory. Exp Mol Med.
68. Wilson HK, Canfield SG, Shusta EV, et al. Concise review: tissue-specific microvascular endothelial cells derived from human pluripotent stem cells. Stem Cells.
69. Ikuno T, Masumoto H, Yamamizu K, et al. Efficient and robust differentiation of endothelial cells from human induced pluripotent stem cells via lineage control with VEGF and cyclic AMP. PLoS One.
70. Orlova VV, van den Hil FE, Petrus-Reurer S, et al. Generation, expansion and functional analysis of endothelial cells and pericytes derived from human pluripotent stem cells. Nat Protoc.
71. Halaidych OV, Freund C, van den Hil F, et al. Inflammatory responses and barrier function of endothelial cells derived from human induced pluripotent stem cells. Stem Cell Reports.
72. Rufaihah AJ, Huang NF, Kim J, et al. Human induced pluripotent stem cell-derived endothelial cells exhibit functional heterogeneity. Am J Transl Res.
73. McCracken IR, Taylor RS, Kok FO, et al. Transcriptional dynamics of pluripotent stem cell-derived endothelial cell differentiation revealed by single-cell RNA sequencing. Eur Heart J.
74. De Bruyn C, Delforge A, Martiat P, et al. Ex vivo expansion of megakaryocyte progenitor cells: cord blood versus mobilized peripheral blood. Stem Cells Dev.
75. Nakamura S, Takayama N, Hirata S, et al. Expandable megakaryocyte cell lines enable clinically applicable generation of platelets from human induced pluripotent stem cells. Cell Stem Cell.
76. Tao H, Gaudry L, Rice A, et al. Cord blood is better than bone marrow for generating megakaryocytic progenitor cells. Exp Hematol.
77. Heremans J, Garcia-Perez JE, Turro E, et al. Abnormal differentiation of B cells and megakaryocytes in patients with Roifman syndrome. J Allergy Clin Immunol.
78. Choi KD, Yu J, Smuga-Otto K, et al. Hematopoietic and endothelial differentiation of human induced pluripotent stem cells. Stem Cells.
79. French TW, Fox LE, Randolph JF, et al. A bleeding disorder (von Willebrand's disease) in a Himalayan cat. J Am Vet Med Assoc.
80. Hogan AG, ME Muhrer ME, Bogart R. A hemophilia-like disease in swine. Proc Soc Exp Biol Med.
81. Olsen EH, McCain AS, Merricks EP, et al. Comparative response of plasma VWF in dogs to up-regulation of VWF mRNA by interleukin-11 versus Weibel-Palade body release by desmopressin (DDAVP). Blood.
82. Pendu R, Christophe OD, Denis CV. Mouse models of von Willebrand disease. J Thromb Haemost.
2009;7 (Suppl 1):61–64.
83. Fass DN, Bowie EJ, Owen CA Jr, et al. Inheritance of porcine von Willbrand's disease: study of a kindred of over 700 pigs. Blood.
84. Brinkhous KM. Animal models: importance in research on hemorrhage and thrombosis. Adv Exp Med Biol.
85. Wu QY, Drouet L, Carrier JL, et al. Differential distribution of von Willebrand factor in endothelial cells. Comparison between normal pigs and pigs with von Willebrand disease. Arteriosclerosis.
86. Dodds WJ. Canine von Willebrand's disease. J Lab Clin Med.
87. Johnson GS, Turrentine MA, Kraus KH. Canine von Willebrand's disease. A heterogeneous group of bleeding disorders. Vet Clin North Am Small Anim Pract.
88. De Meyer SF, Vanhoorelbeke K, Chuah MK, et al. Phenotypic correction of von Willebrand disease type 3 blood-derived endothelial cells with lentiviral vectors expressing von Willebrand factor. Blood.
89. Denis C, Methia N, Frenette PS, et al. A mouse model of severe von Willebrand disease: defects in hemostasis and thrombosis. Proc Natl Acad Sci U S A.
90. Marx I, Christophe OD, Lenting PJ, et al. Altered thrombus formation in von Willebrand factor-deficient mice expressing von Willebrand factor variants with defective binding to collagen or GPIIbIIIa. Blood.
91. Sweeney JD, Novak EK, Reddington M, et al. The RIIIS/J inbred mouse strain as a model for von Willebrand disease. Blood.
92. Nichols WC, Cooney KA, Mohlke KL, et al. von Willebrand disease in the RIIIS/J mouse is caused by a defect outside of the von Willebrand factor gene. Blood.
93. Golder M, Pruss CM, Hegadorn C, et al. Mutation-specific hemostatic variability in mice expressing common type 2B von Willebrand disease substitutions. Blood.
94. Rayes J, Hollestelle MJ, Legendre P, et al. Mutation and ADAMTS13-dependent modulation of disease severity in a mouse model for von Willebrand disease type 2B. Blood.
95. Adam F, Casari C, Prevost N, et al. A genetically-engineered von Willebrand disease type 2B mouse model displays defects in hemostasis and inflammation. Sci Rep.
96. Sanda N, Suzuki N, Suzuki A, et al. Vwf K1362A resulted in failure of protein synthesis in mice. Int J Hematol.
97. Hockemeyer D, Jaenisch R. Induced pluripotent stem cells meet genome editing. Cell Stem Cell.
98. Park CY, Kim DH, Son JS, et al. Functional correction of large factor VIII gene chromosomal inversions in hemophilia A patient-derived iPSCs using CRISPR-Cas9. Cell Stem Cell.
99. Mayadas TN, Wagner DD. Vicinal cysteines in the prosequence play a role in von Willebrand factor multimer assembly. Proc Natl Acad Sci U S A.
Copyright © 2019 the Author(s). Published by Wolters Kluwer Health, Inc. on behalf of the European Hematology Association.
100. Hop C, Fontijn R, van Mourik JA, et al. Polarity of constitutive and regulated von Willebrand factor secretion by transfected MDCK-II cells. Exp Cell Res.