Cellular and Molecular Events Regulating Factor V Endocytosis by Megakaryocytes Jacqueline Michelle Gertz University of Vermont

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Graduate College Dissertations and Theses Dissertations and Theses

2015 Cellular And Molecular Events Regulating Factor V Endocytosis By Megakaryocytes Jacqueline Michelle Gertz University of Vermont

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This Dissertation is brought to you for free and open access by the Dissertations and Theses at ScholarWorks @ UVM. It has been accepted for inclusion in Graduate College Dissertations and Theses by an authorized administrator of ScholarWorks @ UVM. For more information, please contact [email protected]. CELLULAR AND MOLECULAR EVENTS REGULATING FACTOR V ENDOCYTOSIS BY MEGAKARYOCYTES

A Dissertation Presented

Jacqueline Michelle Gertz

The Faculty of the Graduate College

The University of Vermont

In Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy Specializing in Biochemistry

October, 2015

Defense Date: June 5, 2015

Dissertation Examination Committee: Beth A. Bouchard, PhD., Advisor Anthony Morielli, Ph.D., Chairperson Stephen J. Everse, Ph.D. Christopher S. Francklyn, Ph.D. Robert J. Kelm, Ph.D. Paula B. Tracy, Ph.D. Cynthia J. Forehand, Ph.D., Dean of the Graduate College

Abstract

Platelet- and plasma-derived factor Va are absolutely essential for thrombin generation catalyzed by the prothrombinase complex, a 1:1 stoichiometric complex of the serine protease factor Xa and the nonenzymatic cofactor, factor Va, assembled on an appropriate membrane surface in the presence of calcium ions. Two whole blood pools of the procofactor, factor V, exist: approximately 75% circulates in the plasma as a single chain inactive molecule, while the other 25% resides in platelet α-granules in a partially proteolytically-activated state. Our laboratory demonstrated that the platelet-derived cofactor originates following endocytosis of plasma-derived factor V by megakaryocytes, the platelet precursor cells, via a two receptor system including an uncharacterized, specific factor V receptor and low density lipoprotein receptor related protein-1. Following endocytosis factor V is physically and functionally modified and trafficked to the platelet α-granule from where it is released upon platelet activation at sites of vascular injury. The first goal of this dissertation was to define how factor V endocytosis changes over the course of megakaryocyte development. Hematopoietic multipotential stem cells were isolated from human umbilical cord blood and subjected to ex vivo differentiation into megakaryocytes. Megakaryocyte differentiation was assessed by flow cytometry using fluorescently-labeled antibodies against megakaryocyte- and platelet-specific markers and factor V directly conjugated to a fluorophore over 12 days. Differentiation was confirmed by a decrease in a stem cell marker (CD34) and an increase in a mature megakaryocyte marker (CD42) and coincident with factor V endocytosis. Live cell imaging verified differentiation and permitted the observation of proplatelet formation, the precursor to circulating platelets. Analogous experiments verified the trafficking of factor V into proplatelet extensions. Factor V is a highly glycosylated protein: potential roles of these glycans may be endocytosis and trafficking by megakaryocytes. We previously demonstrated that factor V endocytosis is mediated by the light chain region of the procofactor. This region of factor V contains three glycans – one high mannose and two complex N-linked glycans. In the second part of this dissertation, a role for the complex N-linked glycans at Asn1675 and Asn2181 of the factor V light chain in factor V endocytosis by megakaryocytes was assessed. Exoglycosidases were used to selectively trim the complex N-linked glycans on human factor V under native conditions. Treatment with neuraminidase removed 100% of the sialic acid residues on the factor V light chain as demonstrated by gel electrophoresis and mass spectrometry. Treatment with β-1,4-galactosidase removed 69% of the galactose residues at Asn1675 and 100% at Asn2181. Glycosidase-treated factor Va behaves similarly to untreated factor Va in thrombin generation assays suggesting that cofactor activity is unaltered by glycan trimming. In addition, glycan removal had no effect on factor V endocytosis by megakaryocyte-like cells. These observations suggest that complex N-linked glycans on the factor V light chain are not important for factor Va cofactor activity or factor V endocytosis by megakaryocyte-like cells, which strongly suggests that they have a role in trafficking.

Citations

Material from this dissertation has been submitted for publication to the Journal of Thrombosis and Haemostasis on May 9th, 2015 in the following form:

Gertz, J.M., Schwertz, H., McLean, K., Bouchard, B.A.. Expression of CD42 is a phenotypic characteristic of megakaryocytes capable of factor V endocytosis and its trafficking to proplatelet extensions

Material from this dissertation has been submitted for publication to Cytometry Part A on May 19th, 2015 in the following form:

Gertz, J.M., Meuser, M., Bouchard, B.A.. Simultaneous flow cytometric analysis of megakaryocyte ploidy and a labile intracellular protein using zinc-based fixation.

Acknowledgements

First and foremost I would like to thank my advisor, Dr. Beth Bouchard, for her guidance, support, and encouragement during the entire graduate school process. I would like to thank the following faculty for being a part of my dissertation committee: Drs.

Anthony Morielli, Stephen Everse, Christopher Francklyn, Robert Kelm, and Paula

Tracy. Many thanks to our collaborators: Drs. Andrew Weyrich and Hanzjorg Schwertz for inviting me to visit their lab and teaching me techniques that were essential for my dissertation and Dr. Mark Jennings for performing mass spectrometry analysis; without them these studies would not have been possible. I would also like to thank all of the members of the Tracy and Bouchard labs, past and present, for moral support, helpful discussion and performing numerous factor V purifications, without which these studies would not have been possible. In particular the lab members I would like to thank are Dr.

Paula Tracy, Dr. Jay Silveira, Dr. Laura Haynes, Dr. Jeremy Wood, Sarah Abdalla and

Francis Ayombil. I would also like to thank Dr. Kenneth Mann for allowing me to participate in the thrombosis and hemostasis training program.

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Table of Contents

Citations ...... ii Acknowledgements ...... iii List of Tables ...... vi List of Figures ...... vii Chapter 1. Introduction ...... 1 1.1. Overview of Megakaryocyte Differentiation and Platelet Production...... 1 1.2. Hematopoietic Stem Cells...... 2 1.3. Megakaryocyte development ...... 6 1.3.1. Signals regulating megakaryocyte development ...... 10 1.4. Platelet production ...... 11 1.4.1. Proplatelets ...... 11 1.4.2. Platelet release ...... 13 1.5. Ex vivo-derived megakaryocytes ...... 13 1.6. Overview of hemostasis ...... 15 1.7. The prothrombinase complex ...... 20 1.8. Factor V: biochemistry and physiology ...... 21 1.9. Structural features of factor V/Va ...... 23 1.10. Platelet versus plasma-derived factor V/Va ...... 28 1.11. Acquisition of platelet-derived factor V/Va ...... 31 1.12. Summary ...... 36 1.13. References ...... 37 Chapter 2. Expression of CD42 is a phenotypic characteristic of megakaryocytes capable of factor V endocytosis and its trafficking to proplatelet extensions ...... 61 2.1. Summary ...... 62 2.2. Introduction ...... 64 2.3. Materials and methods ...... 67 2.4. Results ...... 72 2.5. Discussion ...... 85 2.6. Authorship details ...... 89 2.7. Acknowledgements ...... 89 2.8. References ...... 91 Chapter 3. Complex N-linked glycans in the factor V light chain are not involved in endocytosis by megakaryocytes ...... 100 3.1. Summary ...... 101 3.2. Introduction ...... 102 3.3. Materials and methods ...... 106 3.4. Results ...... 110

3.5. Discussion ...... 116 3.6. Authorship details ...... 121 3.7. Acknowledgements ...... 122 3.8. References ...... 123 Perspectives and Future Directions ...... 132 References ...... 139 Appendix I. Simultaneous flow cytometric analysis of megakaryocyte ploidy and a labile intracellular protein using zinc-based fixation ...... 145 Summary ...... 147 Introduction ...... 149 Materials and methods ...... 150 Results ...... 153 Discussion ...... 160 Acknowledgements ...... 163 References ...... 164 Appendix II. Galectin-8 is not a component of the two receptor system that mediates factor V endocytosis by megakaryocytes ...... 168 Introduction ...... 169 Materials and methods ...... 171 Results ...... 174 Discussion ...... 176 References ...... 180

List of Tables

Table 1. Nomenclature of megakaryocyte differentiation markers...... 3

Table 2. Median fluorescence intensities of differentiation markers in the super factor V+

megakaryocyte population...... 80

Table 3. Glycan composition of glycosidase-treated factor V...... 113

Table 4. Densitometric analysis of the factor Va light chain in CMK lysates following

endocytosis...... 118

List of Figures

Figure 1. Hematopoietic stem cell development...... 4

Figure 2. Megakaryocyte maturation from hematopoietic stem cells...... 7

Figure 3. Platelet production and release...... 12

Figure 4. Enzyme complexes of the coagulation cascade...... 18

Figure 5. The importance of the prothrombinase components to thrombin generation. ... 22

Figure 6. Human factor V structure and activation by thrombin...... 24

Figure 7. Potential N-linked glycosylations in human factor V...... 26

Figure 8. Hypothetical model describing the megakaryocyte cell surface events mediating

factor V endocytosis...... 33

Figure 9. Isolation and analysis of megakaryocytes cultured ex vivo from umbilical cord

blood-derived CD34+ cells...... 69

Figure 10. Factor V endocytosis and surface expression of megakaryocyte lineage markers.

...... 73

Figure 11. Nearly all factor V+CD42+ cells express CD41...... 77

Figure 12. CD42 expression increases over time on super factor V+ cells...... 79

Figure 13. Live cell imaging of proplatelet formation by megakaryocytes derived ex vivo

from human umbilical cord blood...... 81

Figure 14. CD42+ proplatelets contain endocytosed factor V...... 83

Figure 15. N-linked glycans on the human, plasma-derived factor V light chain...... 105

Figure 16. Mobility of glycosidase-treated and untreated factor V on SDS-PAGE...... 112

vii

Figure 17. Time course of factor Va activity after glycosidase treatment...... 115

Figure 18. Endocytosis of glycosidase-treated factor V by CMKs...... 117

Figure 19. Standard protocols are not appropriate to assess factor V endocytosis in

conjunction with DNA content...... 155

Figure 20. ZBF will allow simultaneous analysis of DNA content, an integral membrane

protein and/or a labile intracellular protein...... 158

Figure 21. Simultaneous flow cytometric analysis of DNA content and endocytosed factor

V in polyploidal CMK cells...... 161

Figure 22. Expression of galectin-8 by megakaryocytes...... 175

Figure 23. Inhibition of galectin-8 does not alter factor V endocytosis...... 177

viii

Chapter 1. Introduction

1.1. Overview of Megakaryocyte Differentiation and Platelet Production.

Platelets are small anucleate cell fragments that reside in the blood (Osler 1874,

Tocantins 1938). When an injury occurs platelets adhere to the site to form a platelet plug

(Brass 2003), become activated, and release proteins that are essential for clot formation

(Vicic, Lages et al. 1980, Tracy and Mann 1983, Harrison and Cramer 1993, McNicol and

Israels 1999, Brass 2003, Blair and Flaumenhaft 2009, Fager, Wood et al. 2010). This leads to formation of a fibrous mesh network that will stabilize the platelet plug to form the clot and prevent further bleeding (Peerschke, Zucker et al. 1980).

Platelets are generated by megakaryocytes (Wright 1906) in the bone marrow where they differentiate from hematopoietic stem cells (Long, Williams et al. 1982, Ogawa

1993). Once the cell has completely differentiated, platelet production can begin (Long,

Williams et al. 1982, Ogawa 1993, Kondo, Wagers et al. 2003). During this process the megakaryocyte will develop a unique multi-nuclei phenotype (Ebbe and Stohlman 1965,

Odell and Jackson 1968, Ebbe 1976). In addition, they will begin to generate all necessary platelet proteins and organelles in addition to their own (Zimmet and Ravid 2000, Machlus and Italiano 2013). Finally a long, thin, tubular proplatelet extension is extended into the blood stream (Italiano, Lecine et al. 1999) where the functional platelets are sheared off by the flowing blood (Dunois-Larde, Capron et al. 2009).

1.2. Hematopoietic Stem Cells

The hematopoietic stem cell, expressing cluster of differentiation (CD) 34 (Table

1) (Figure 1), is derived from the mesoderm layer (Mikkola and Orkin 2006) to give rise to all blood cells (hematopoiesis) through a series of differentiation steps (Long, Williams et al. 1982, Ogawa 1993, Kondo, Wagers et al. 2003) in the bone marrow. As with all stem cells, hematopoietic cells are multipotent, allowing them to self-renew and replenish all types of blood cells (Mikkola and Orkin 2006). Thus, a small number of hematopoietic stem cells can expand into a very large number of daughter cells comprising many cell types. Hematopoietic stem cells initially give rise to multi-potent progenitor cells which no longer have the capacity to self-renew yet maintain full lineage differentiation potential

(Morrison and Weissman 1994, Christensen and Weissman 2001). Subsequently, multipotent progenitor cells advance to the oligopotent progentors cells: the common lymphoid progenitor (which will develop into T-cells, B-cells) and the common myeloid progenitor

(becoming monocytes, macrophages, neutrophils, erythrocytes, and megakaryocytes)

(Seita and Weissman 2010). Differentiation along either of these lineages involves a loss of self-renewal capacity, which requires the original hematopoietic stem cell population to self-renew to maintain the population. However, lineage differentiation is not always equivalent; some hematopoietic stem cells repopulate one lineage more frequently than the other (Muller-Sieburg, Cho et al. 2002, Muller-Sieburg, Cho et al. 2004).

Table 1. Nomenclature of megakaryocyte differentiation markers.

Cluster of Glycoprotein Integrin Complex Role Differentiation

- - Cell adhesion CD34 Stem cell marker Fibrinogen GPIIb α CD41 GPIIb/IIIa IIb receptor GPIb-V-IX von Willebrand GPIb - CD42 complex Factor receptor Fibrinogen GPIIIa β3 CD61 GPIIb/IIIa receptor

Figure 1. Hematopoietic stem cell development.

Multipotent hematopoietic stem cells are capable of self-renewal, allowing a small number of stem cells to generate a very large number of daughter cells. In addition, they can give rise to common myeloid and lymphoid progenitor cells. The myeloid lineage will produce erythrocytes, megakaryocytes and platelets, basophils, eosinophils, monocytes and macrophages, as well as neutrophils. Common lymphoid progenitors will develop into B and T lymphocytes as well as natural killer cells.

1.3. Megakaryocyte development

Megakaryocytes are highly specialized cells responsible for assembling and releasing platelets. These cells develop from CD34+ hematopoietic stem cells (Figure 2) into common myeloid progenitor cells expressing CD41 (glycoprotein (GP) IIb, αIIb)

(Figure 2) (Long, Williams et al. 1982, Ogawa 1993). As the cell differentiates from stem cell to megakaryocyte (Figure 2) they increase in size and DNA content, develop both granules and an invaginated membrane system, and express the megakaryocyte-specific marker, and CD42 (GPIb) (Table 1) (Zimmet and Ravid 2000). During this process the megakaryocyte tailors its cytoplasm and membrane systems for platelet biogenesis (Long,

Williams et al. 1982). By growing to a size of 100 µm with a high concentration of ribosomes, megakaryocytes are able to increase metabolic output to allow for production of platelet proteins (Zimmet and Ravid 2000, Machlus and Italiano 2013). This enlargement is mediated through endomitosis, a process which allows the chromosomes to replicate without devoting extra energy to the other aspects of cell division, resulting in a polyploidal (Nagata, Muro et al. 1997, Vitrat, Cohen-Solal et al. 1998) cell with multiple nuclei and DNA contents ranging from 4N to 128N per megakaryocyte (Ebbe and

Stohlman 1965, Odell and Jackson 1968, Odell, Jackson et al. 1970, Ebbe 1976). A single

32N megakaryocyte can yield approximately 3000 platelets, while several precursor 2N cells would be required to produce a similar number of platelets (Winkelmann, Pfitzer et al. 1987).

Figure 2. Megakaryocyte maturation from hematopoietic stem cells.

CD34+ hematopoietic stem cells (left) in the bone marrow develop into immature megakaryocytes with decreased CD34 (red), increased CD41 (purple) expression, and transition to an endomitotic cell cycle leading to a polyploidal phenotype with many nuclei

(black). The cells will continue to develop into mature megakaryocytes that generate proplatelet extensions (right) and express to the mature megakaryocyte marker, CD42

(yellow), in addition to CD41.

In addition to a polyploidal nucleus and a high concentration of ribosomes, megakaryocytes develop an expansive and interconnected membranous network called the demarcation membrane system (Yamada 1957). These membranes are connected to the plasma membrane (Nakao and Angrist 1968, Behnke 1969) and are thought to function as membrane reservoirs for formation of proplatelet extensions, the platelet precursor bodies that are formed through extrusion of the internal membrane system (Radley and Haller

1982). Platelet-specific proteins, such as von Willebrand factor and fibrinogen receptors, are synthesized. Others proteins, are endocytosed from the plasma by megakaryocytes

(Heijnen, Debili et al. 1998, Bouchard, Williams et al. 2005, Suehiro, Veljkovic et al.

2005). Once the necessary proteins have been synthesized or acquired they accumulate in small vesicles that bud off from the trans-Golgi network [24-26] and travel to the multivesicular body (Heijnen, Debili et al. 1998, Youssefian and Cramer 2000). Here they are sorted to their final destination: the cell surface (Heijnen, Debili et al. 1998), or platelet- specific α- and dense-granules (Handagama, Scarborough et al. 1993, Youssefian, Masse et al. 1997, Heijnen, Debili et al. 1998, Youssefian and Cramer 2000, Gould, Simioni et al.

2005, Suehiro, Veljkovic et al. 2005). It is unclear if proteins are specifically sorted by granule type (Italiano, Richardson et al. 2008, Kamykowski, Carlton et al. 2011,

Jonnalagadda, Izu et al. 2012), as recent studies have identified heterogeneity within the α- granule population of circulating platelets with respect to morphology and membrane protein composition (van Nispen tot Pannerden, de Haas et al. 2010).

1.3.1. Signals regulating megakaryocyte development

Thrombopoietin is the primary cytokine for megakaryocyte differentiation and is essential for megakaryocytes to maintain platelet production (Bartley, Bogenberger et al. 1994, de

Sauvage, Hass et al. 1994, Kaushansky 1994, Kaushansky, Lok et al. 1994, Kuter, Beeler et al. 1994, Wendling, Maraskovsky et al. 1994, Kaushansky 1995). However, mice lacking either thrombopoietin or its receptor, c-Mpl, are still able to produce functional platelets, although in a limited quantity, indicating a role for other regulators in the process of megakaryocyte differentiation and platelet formation (Choi, Hokom et al. 1995, Ito, Ishida et al. 1996). In in vitro culture systems stem cell factor, expressed by bone marrow endothelial cells, encourages proliferation of all hematopoietic stem cells (Broxmeyer,

Cooper et al. 1991). Megakaryocyte specific proliferation is stimulated by interleukin (IL)-

3 (Emerson, Yang et al. 1988, Briddell and Hoffman 1990) yielding increased total cell number and an immature megakaryocyte population (Schattner, Lefebvre et al. 1996, Sun,

Tan et al. 2004). In contrast, IL-6 affects the later stages of megakaryocyte differentiation

(Kimura, Ishibashi et al. 1990, Koike, Nakahata et al. 1990, Burstein, Mei et al. 1992).

Interestingly, IL-6 (Bruno and Hoffman 1989, Koike, Nakahata et al. 1990) and stem cell factor (Brandt, Briddell et al. 1992, Imai and Nakahata 1994) only show an effect when used in conjunction with other cytokines, such as IL-3. Together they lead to an increase in megakaryocyte number, size, and DNA content (Bruno and Hoffman 1989, Kimura,

Ishibashi et al. 1990, Brandt, Briddell et al. 1992). The use of these cytokines has allowed ex vivo differentiation of megakaryocytes from hematopoietic stem cells by which to study

the mechanisms that regulate megakaryocyte differentiation and platelet production (Choi,

Hokom et al. 1995, Cramer, Norol et al. 1997, Lecine and Shivdasani 1998).

1.4. Platelet production

1.4.1. Proplatelets

Once megakaryocytes reach maturity (Behnke 1969, Becker and De Bruyn 1976,

Junt, Schulze et al. 2007) a pseudopod forms at a single site on the cell, elongates and tapers into a tubule called the proplatelet (Figure 3) (Italiano, Lecine et al. 1999). This process is initiated through signaling by the megakaryocyte specific markers (Dunois-

Larde, Capron et al. 2009), CD42 and CD41/CD61 expressed on the cell surface.

Proplatelet formation is a microtubule-driven process (Tablin, Castro et al. 1990) in which continuous polymerization of tubulin in conjunction with motor-driven sliding of overlapping microtubules leads to elongation (Hartwig and Italiano 2006). When a portion of the proplatelet is bent, some of the microtubules will separate from the bundle to form a branch point (Italiano, Lecine et al. 1999). If the microtubules continue to bend forming a loop, they may reenter the shaft leading to formation of a bud at the proplatelet tip. The same microtubule bundles within the microtubule shaft also serve as a conduit by which mitochondria, granules, and other organelles can travel bidirectionally to the platelet bud

(Richardson, Shivdasani et al. 2005).

Figure 3. Platelet production and release.

Mature megakaryocytes will develop long, thin proplatelet extensions that extend into the small blood vessels of the bone marrow by breaking down extracellular matrix proteins.

Once the proplatelet is in the blood vessel, shear forces from the passing blood flow remove preplatelet particles from the proplatelet extensions. Preplatelets will undergo a fission process to yield individual functional platelets.

1.4.2. Platelet release

Proplatelets extend into the small bone marrow sinusoidal blood vessels (Figure 3) by breaking down extracellular matrix proteins through the use of an unidentified matrix metalloproteases (Schachtner, Calaminus et al. 2013). As the proplatelet enters the vessel, shear forces from the passing blood flow separate preplatelet particles from the cell body

(Junt, Schulze et al. 2007, Thon, Montalvo et al. 2010). Preplatelets will convert to a barbell shape in a microtubule dependent process before undergoing fission into individual platelets (Thon, Montalvo et al. 2010). Once the cytoplasm has been completely converted into platelets the remaining megakaryocyte cell body, including nuclei, is extruded and degraded by macrophages in the bone marrow (Italiano, Lecine et al. 1999).

1.5. Ex vivo-derived megakaryocytes

Most studies to assess the mechanisms of platelet formation have been performed using murine bone marrow cells. Purified low density bone marrow cells were allowed to mature for 4-6 days to yield large cells with up to 128N DNA content, expression of the megakaryocyte lineage markers CD41, CD61, and CD42 up to 90% (Shiraga, Ritchie et al.

1999). However, using this culture model few cells, if any, proceed to proplatelet formation. In contrast, mature megakaryocytes isolated from murine bone marrow rapidly produce proplatelets when cultured on a fibrinogen matrix (Larson and Watson 2006).

However, the yields are generally low using this method and require that mice receive thrombopoietin injections for four days prior to bone marrow collection. A further

improvement is the use of primary megakaryocytes obtained from murine fetal livers

(Italiano, Lecine et al. 1999). Live cells are cultured in the presence of thrombopeotin for

4-6 days prior to separation using an albumin step gradient to obtain an enriched megakaryocyte population. Within this population large polyploidal cells are observed and proplatelet formation occurs rapidly.

Megakaryocytes-derived from human tissues take longer to mature (13-17 days in culture), and often yield fewer proplatelets than murine cell cultures. The first report of the generation of human megakaryocytes and platelets in culture was published by Choi et al. in 1995 (Choi, Hokom et al. 1995). Megakaryocytes, larger proplatelets, and smaller- platelet sized particles were present when CD34+ peripheral blood progenitor cells where cultured in the presence of thrombopoietin. Recent studies have identified additional sources of human CD34+ cells including umbilical cord blood (Tao, Gaudry et al. 1999), fetal liver (Ma, Sun et al. 2000), or bone marrow (Guerriero, Testa et al. 1995). Human

CD34+ cells isolated from bone marrow can be cultured into large polyploidal megakaryocytes expressing markers of megakaryocyte maturation, CD41 and CD42.

However, they do not form proplatelet extensions (Debili, Coulombel et al. 1996, Tao,

Gaudry et al. 1999, Miyazaki, Ogata et al. 2000). Ex vivo expansion studies using umbilical cord blood-derived CD34+ stem cells suggest that they have a higher proliferative capacity in response to stimulation by stem cell factor (Emerson 1996), leading to increased cell number and low ploidy levels (Hagiwara, Kodama et al. 1998, Tao, Gaudry et al. 1999,

Lepage, Leboeuf et al. 2000, Miyazaki, Ogata et al. 2000, Sun, Tan et al. 2004, Denis,

Tolley et al. 2005, Hofmeister, Zhang et al. 2007, Robert, Cortin et al. 2012). Interestingly, despite low ploidy levels they express the mature megakaryocyte marker, CD42, and generate proplatelet extensions (Lepage, Leboeuf et al. 2000, Denis, Tolley et al. 2005,

Hofmeister, Zhang et al. 2007) .

Ex vivo differentiation of megakaryocytes has proved a useful tool by which to study platelet formation, however, it is not without limitations. Without fail, ex vivo- derived cultures contain a mix of hematopoietic stem cells, immature megakaryocytes, proplatelet-producing megakaryocytes, as well as other cells from both the myeloid and lymphoid lineages (Tao, Gaudry et al. 1999, Proulx, Boyer et al. 2003, Cortin, Garnier et al. 2005, Matsunaga, Tanaka et al. 2006, Boyer, Robert et al. 2008), requiring further purification by techniques such as fluorescence activated cell sorting (Tao, Gaudry et al.

1999) in order to achieve a pure megakaryocyte culture at the expense of cell number. In addition, techniques have only recently been able to isolate the intermediate stages of platelet release (Thon, Montalvo et al. 2010). As such, most studies have focused on the qualitative aspects of megakaryocyte maturation. Therefore many questions remain, particularly about what signals regulate each step of megakaryocyte maturation and platelet formation.

1.6. Overview of hemostasis

Blood is comprised of a heterogeneous mixture of platelets, red blood cells, and white blood cells suspended in a fluid, known as plasma. In addition to these cellular

components, plasma also contains carbohydrates, salts, fats, proteins, hormones, nucleic acids, and other small molecules. The functions of blood include: oxygen delivery to tissues, transportation of nutrients, and protection against infectious agents (Alberts 2008).

In addition, blood plays a role in hemostasis, a highly regulated process in which blood coagulation is rapidly initiated and terminated in order to maintain a balance between the procoagulant and anticoagulant states and prevent uncontrolled bleeding or thrombosis at the site of vascular injury.

Hemostasis occurs in three stages following vascular damage: primary hemostasis, secondary hemostasis, and termination. During primary hemostasis, the endothelial cell lining of the blood vessel wall is damaged to expose subendothelial proteins including collagen, fibronectin, tissue factor, and von Willebrand factor as well as subendothelial cells such as macrophages, foam cells, and smooth muscle cells. Generally, these proteins are sequestered within the intact vasculature, preventing interactions with the blood and unwanted clot formation (Savage, Saldivar et al. 1996, Farndale, Sixma et al. 2004). Exposure of these proteins to the bloodstream allows platelets to adhere to the site of injury using integrins and glycoprotein receptors (Brass 2003). Interactions with the subendothelium leads to platelet activation (Moroi, Jung et al. 1996, Gross, Lee et al.

1999), a dramatic shape change involving cytoskeletal protein rearrangement and release of hemostatically-relevant factors, such as factor V/Va and fibrinogen, from the platelet α- and dense- granules (Jandrot-Perrus, Lagrue et al. 1997, Kehrel, Wierwille et al. 1998).

Platelet to platelet bridges are formed through interactions between CD41/CD61 on the

platelet and released fibrinogen to form a platelet plug (Bennett and Vilaire 1979,

Peerschke, Zucker et al. 1980).

While the platelet plug will stop the flow of bleeding, it can easily be dislodged due to pressure from the blood leading additional blood loss. Stabilization of the platelet plug is achieved through fibrin crosslinking in a process known as secondary hemostasis.

This is initiated through exposure of the transmembrane protein tissue factor to the blood

(Wilcox, Smith et al. 1989) (Figure 4). Tissue factor binds to plasma factor VIIa to form the extrinsic tenase complex (Mackman 2006). This complex activates factor IX to IXa and factor X to factor Xa (Komiyama, Pedersen et al. 1990, Lawson and Mann 1991,

Krishnaswamy, Field et al. 1992). Factor IXa forms a complex with the circulating non- enzymatic cofactor factor VIIIa known as the intrinsic tenase complex (Barton 1967,

Hemker and Kahn 1967, Hultin and Nemerson 1978), which also activates factor X to Xa

(Silverberg, Nemerson et al. 1977, van Dieijen, Tans et al. 1981). Factor Xa, together with factor Va, calcium, and an appropriate membrane surface, such as the activated platelet surface (Walsh 1974), form the prothrombinase complex (Barton, Jackson et al. 1967) to convert prothrombin to thrombin (Barton, Jackson et al. 1967). Thrombin activates platelets (Davey and Luscher 1967), and propagates its own generation through activation of factor V (Esmon 1979, Nesheim and Mann 1979), factor VII (Radcliffe and Nemerson

1975), factor VIII (Osterud, Rapaport et al. 1977, Vehar and Davie 1980) and factor XIII

(Takagi and Doolittle 1974, Greenberg, Miraglia et al. 1985). Thrombin also cleaves fibrinogen to fibrin (Bailey, Bettelheim et al. 1951, Doolittle 1973) giving rise to a fibrin

Figure 4. Enzyme complexes of the coagulation cascade.

The extrinsic tenase complex, comprised of the serine protease factor VIIa (FVIIa) and tissue factor (TF) expressed on subendothelial cells forms at the site of vascular injury and activates factor X (FX) to factor Xa (FXa) and factor IX (FIX) to factor IXa (FIXa). FIXa and the non-enzymatic cofactor FVIIIa form the intrinsic tenase complex, which also activates FX to FXa. Together, FXa and factor Va (FVa, shown as consisting of a heavy chain and light chain) form the prothrombinase complex which activates prothrombin (II) to thrombin (IIa). Thrombin activates fibrinogen to fibrin to form a fibrin clot. Thrombin will also bind thrombomodulin to activate protein C to inhibit the intrinsic tenase and prothrombinase complexes. Tissue factor pathway inhibitor (TFPI) inhibits the extrinsic tenase complex while antithrombin inhibits thrombin and factor Xa. The extrinsic tenase, intrinsic tenase and prothrombinase complexes all require Ca2+ and a membrane surface for optimal activity.

clot (Ferry 1952). The fibrin clot is further stabilized through factor XIIIa, which covalently cross-links fibrin molecules together (Doolittle, Chen et al. 1971, Kanaide and Shainoff

1975).

In order to prevent excessive thrombosis, the coagulation response must be terminated. Once factor Xa concentrations reach a critical level it will bind to tissue factor pathway inhibitor (TFPI) and the complex will inhibit the extrinsic tenase, to form a tissue factor-factor VIIa-factor Xa-TFPI inhibitory complex (Broze, Warren et al. 1988, Girard,

Warren et al. 1989). In addition, thrombin will bind to thrombomodulin on endothelial cells

(Esmon, Esmon et al. 1982) to activate protein C to activated protein C (Esmon and Owen

1981, Owen and Esmon 1981, Comp, Jacocks et al. 1982, Esmon, Owen et al. 1982) leading to inactivation of the cofactors factor Va (Kisiel, Canfield et al. 1977, Walker,

Sexton et al. 1979) and factor VIIIa (Vehar and Davie 1980).

1.7. The prothrombinase complex

As previously mentioned, hemostasis requires the assembly of many membrane bound protein-enzyme complexes (Figure 4) (Mann, Nesheim et al. 1990). For example, thrombin is an essential enzyme involved in numerous procoagulant and anticoagulant events. In order to perform these duties, it must first be activated from the inactive precursor, prothrombin, by the prothrombinase complex. Together the serine protease factor Xa, the non-enzymatic cofactor factor Va, along with calcium and a phospholipid membrane surface, such as the platelet form the prothrombinase complex (Barton, Jackson

et al. 1967, Walsh 1974, Mann, Nesheim et al. 1990). Complex formation increases the catalytic rate of thrombin formation by 300,000-fold when compared with the enzyme, factor Xa, alone (Figure 5). When the cofactor, factor Va, is removed from the complex, the thrombin generation rate is decreased by 10,000 fold, suggesting a critical role for the cofactor in the regulation of thrombin generation via this complex.

1.8. Factor V: biochemistry and physiology

There are two pools of factor V in the blood: 75-80% is found in the plasma and

20-25% is found inside the platelets (Breederveld, Giddings et al. 1975, Osterud, Rapaport et al. 1977, Vicic, Lages et al. 1980, Tracy, Eide et al. 1982). The platelet-derived pool is stored within the platelet α-granules (Chesney, Pifer et al. 1981, Wencel-Drake, Dahlback et al. 1986) and released upon platelet activation. Factor V is synthesized by the liver and circulates in plasma as a large single chain 330kDa protein (Rapaport, Ames et al. 1960,

Kupfer, Gee et al. 1964). The circulating plasma concentration of factor V is 21-30 nM

(Tracy, Eide et al. 1982), and the half-life is 12-15 hours (Borchgrevink and Owren 1961).

During its synthesis, factor V undergoes several post-translational modifications, including phosphorylation (Kalafatis, Rand et al. 1993, Rand, Kalafatis et al. 1994, Gould, Silveira et al. 2004), N- and O-linked glycosylation (Jenny, Pittman et al. 1987, Fernandez,

Hackeng et al. 1997, Silveira, Kalafatis et al. 2002, Gould, Silveira et al. 2004), and sulfation (Hortin 1990, Pittman, Tomkinson et al. 1994, Pittman, Tomkinson et al. 1994).

Additionally, a 28 residue signaling peptide is removed prior to its secretion as a 2196

Figure 5. The importance of the prothrombinase components to thrombin generation.

The relative rates of thrombin generation by components of the prothrombinase complex: factor Xa (green oval), the non-enzymatic cofactor factor Va (red oval) shown as a heavy chain (HC) and light chain (LC), calcium ions (Ca2+), and a membrane surface (blue and yellow phospholipid vesicles). While factor Xa is capable of activating thrombin alone, it occurs at a very slow rate. With the incorporation of the other prothrombinase components, this rate is increased almost 300,000-fold. Removal of factor Va from the complex results in a four order of magnitude decrease in the reaction rate, while removal of the membrane surface decreases the reaction rate by three orders of magnitude. Relative rates are from

Nesheim et al. (Nesheim, Taswell et al. 1979).

amino acid protein (Kane and Davie 1986). Thrombin cleavage of the zymogen, factor V, yields the active form of the protein, factor Va.

1.9. Structural features of factor V/Va

The domain structure of factor V (Figure 6) is identical to that observed in the homologous protein, factor VIII, consisting of A1-A2-B-A3-C1-C2 (Gitschier, Wood et al. 1984, Toole, Knopf et al. 1984, Vehar, Keyt et al. 1984). In the thrombin-activated cofactor, the A1 and A2 domains comprise the heavy chain, while the A3, C1 and C2 domains comprise the light chain (Smith and Hanahan 1976). Both the A and C domains of factor V and VIII are ~40% homologous, while the B domains differ greatly, with only

14% identity (Jenny, Pittman et al. 1987). In addition to the human factor V sequence, the complete amino acid sequence has been determined for bovine (Guinto, Esmon et al. 1992), porcine (Grimm, Colter et al. 2001), and murine (Yang, Cui et al. 1998) factor V proteins.

In addition, proposed amino acid sequences are available in the NCBI protein database for chimpanzee, dog, horse, opossum, platypus, rhesus monkey, and zebra fish factor V based on the DNA sequence from each animal. In all cases, the greatest homology is seen between the A and C domains, with the lowest homology observed between B domains.

Crystal structures have identified membrane binding sites within the C1 and C2 domain of the light chain (Macedo-Ribeiro, Bode et al. 1999, Adams, Hockin et al. 2004), providing support for several previously published studies suggesting that the light chain

Figure 6. Human factor V structure and activation by thrombin.

The procofactor, factor V, is a single chain molecule with A1-A2-B-A3-C1-C2 domain structure. Upon activation by thrombin cleavage, the B-domain is removed and the active factor Va molecule consists of a heavy (A1-A2) and light chain (A3-C1-C2) associated in a Ca2+-dependent manner. Cleavage sites for thrombin activation of factor V, Arg709,

Arg1018, and Arg1545, are shown are. The residues defining each domain are indicated above their respective locations.

is involved in binding to both phospholipid vesicles and activated platelets (Higgins and

Mann 1983, Tracy and Mann 1983, van de Waart, Bruls et al. 1983, Pusey and Nelsestuen

1984, Krishnaswamy and Mann 1988, Kalafatis, Jenny et al. 1990, Ortel, Devore-Carter et al. 1992). The ability of the factor Va light chain to bind the phospholipid membrane is also affected by an N-linked glycosylation at Asn2181 (Figure 7) in the C2 domain, located adjacent to the C2 membrane binding region. This differential glycosylation is responsible for the two observed isoforms of the factor Va light chain, known as factor Va1 (~74kDa) and factor Va2 (~71kDa) when visualized by SDS-PAGE (Kim, Ortel et al. 1999, Nicolaes,

Villoutreix et al. 1999). Depending on the specific conditions used for analysis, factor Va2 has a 3-45 fold increased affinity for the phospholipid membrane compared to factor Va1

(Rosing, Bakker et al. 1993, Koppaka, Talbot et al. 1997, Kim, Ortel et al. 1999). This decreased membrane binding of factor Va1 also leads to decreased cofactor activity as part of the prothrombinase complex (Rosing, Bakker et al. 1993, Hoekema, Nicolaes et al. 1997,

Koppaka, Talbot et al. 1997, Kim, Ortel et al. 1999, Nicolaes, Villoutreix et al. 1999), reduced susceptibility to inactivation by activated protein C (Hoekema, Nicolaes et al.

1997), and a reduced ability to act as a cofactor for activated protein C in the inactivation of factor VIIIa (Varadi, Rosing et al. 1996).

Including Asn2181, there are 37 potential N-linked glycosylation sites on human factor V (Figure 7) (Jenny, Pittman et al. 1987). Three are found in the light chain, nine in the heavy chain, and the remaining 25 are in the B domain. With the exception of Asn2181, few of these sites have been characterized. Nicolaes et al. used endoglycosidase treatment

Figure 7. Potential N-linked glycosylations in human factor V.

Factor V contains 37 potential N-linked glycosylation sites. Nine are found in the heavy chain, three in the light chain, and 25 in the B domain. Four of these have been identified as glycosylated (red circles), and one (Asn2181, green circle) has been identified as being partially glycosylated. Glycosylations at the remaining sites have yet to be confirmed (blue circles).

to show that both Asn1675 and Asn1982 are likely fully glycosylated (Nicolaes, Villoutreix et al. 1999). Recent proteomic studies identified glycopeptides containing Asn269 and

Asn432 (Lewandrowski, Moebius et al. 2006). In addition, Liu et al. used a lectin affinity approach to identify Asn269 as a high mannose glycan (Liu, Qian et al. 2005). The complex

N-linked glycosylations of factor Va have been recently characterized with respect to location and composition (Tracy, Jemings et al. 2013). This is of particular interest as deglycosylated factor V has been shown to have a reduced rate of thrombin catalyzed activation (Bruin, Sturk et al. 1987), and once activated, the deglycosylated form of factor

Va may have slightly increased clotting activity (Gumprecht and Colman 1975, Bruin,

Sturk et al. 1987, Pittman, Tomkinson et al. 1994) and be more susceptible to inactivation by activated protein C (Fernandez, Hackeng et al. 1997, Silveira, Kalafatis et al. 2002). In addition, high mannose glycans in the B domain of factor V are required for its trafficking and secretion by hepatocytes (Moussalli, Pipe et al. 1999). In addition to the N-linked glycosylations, human factor V contains additional post translational modifications, including O-linked glycans within the B domain (Fernandez, Hackeng et al. 1997, Silveira,

Kalafatis et al. 2002), tyrosine sulfation (Hortin, Folz et al. 1986, Hortin 1990, Pittman,

Tomkinson et al. 1994, Pittman, Tomkinson et al. 1994, Gould, Silveira et al. 2004), and phosphorylation (Kalafatis, Rand et al. 1993, Rand, Kalafatis et al. 1994, Kalafatis 1998,

Gould, Silveira et al. 2004).

1.10. Platelet versus plasma-derived factor V/Va

As stated earlier, factor V approximately 25% of the factor V pool is located within the platelet α-granule while the other 75% circulates in the plasma. Initial studies using animal models suggested that megakaryocytes synthesize platelet-derived factor V and traffic it to the α-granule. While murine megakaryocytes do synthesize factor V (Chiu,

Schick et al. 1985, Gewirtz, Keefer et al. 1986, Yang, Pipe et al. 2003, Rowley, Schwertz et al. 2012), it has been shown that human platelet-derived factor V originates exclusively through endocytosis of the plasma-derived procofactor by megakaryocytes (Camire, Pollak et al. 1998, Bouchard, Williams et al. 2005, Gould, Simioni et al. 2005, Suehiro, Veljkovic et al. 2005, Bouchard, Meisler et al. 2008).

Although platelet- and plasma-derived factor V are identical in their biosynthetic origin, they are structurally and functionally distinct. For example, studies have shown that the plasma-derived molecule exists solely as a single chain, and exhibits no cofactor activity. In contrast, platelet-derived factor V is stored and released as a mixture of intact single chain and partially proteolytically-activated factor V (Viskup, Tracy et al. 1987,

Monkovic and Tracy 1990). Compared with thrombin-activated factor Va, this partially active cofactor exhibits 5-50% of the activity (Kane, Lindhout et al. 1980, Vicic, Lages et al. 1980, Kane, Mruk et al. 1982, Viskup, Tracy et al. 1987, Monkovic and Tracy 1990).

While factor Xa and thrombin activate the plasma-derived factor V at equivalent rates, the platelet-derived cofactor is activated 50-100 times more effectively by factor Xa than by

thrombin once released at the site of vascular injury (Monkovic and Tracy 1990, Monkovic and Tracy 1990).

Proteolysis of the platelet-derived form occurs within the megakaryocyte. The proteolytic state of the active platelet-derived factor V molecule demonstrates cleavage consistent with thrombin-activated factor V in addition to peptide fragments that are most likely generated by a separate, unidentified, protease (Ayombil, Abdalla et al. 2013). The platelet-derived factor V light chain has a unique cleavage site at Tyr1543 (Gould, Silveira et al. 2004) and is partially resistant to inactivation by activated protein C (Camire,

Kalafatis et al. 1995, Camire, Kalafatis et al. 1998). Plasmin, a serine protease which is known to inactivate plasma factor V in the presence of a membrane surface (Lee and Mann

1989), leads to increased and sustained activity of the platelet-derived cofactor (Conlon,

Camire et al. 1997), allowing the platelet-derived cofactor to remain active much longer compared with the plasma-derived counterpart. In addition, At the site of vascular injury, the platelet-derived factor Va concentration is 100 fold higher than its plasma counterpart.

As platelets contain 20% of the total factor V pool in whole blood and maintain it in such a small volume, the concentration of platelet-derived factor V at the site of injury is able to greatly surpass that of the plasma-derived molecule (Nesheim, Nichols et al. 1986)

Comparisons between platelet and plasma-derived factor V have demonstrated several differences in the post-translational modifications between the molecules. A unique

O-linked glycosylation is seen at Thr402 on the heavy chain of platelet-derived factor V

(Gould, Silveira et al. 2004). The high mannose glycan located at Arg1982 of the light

chain of the platelet-derived molecule has been trimmed (Tracy, Jemings et al. 2013). A phosphorylation site is present at Ser692 in both molecules, however, phosphorylation by casein kinase-2, is only seen in the plasma-derived factor V (Rand, Kalafatis et al. 1994,

Kalafatis 1998, Gould, Silveira et al. 2004). Studies have also demonstrated that a portion of the platelet-derived factor V is non-dissociable and bound to the membrane of the platelet α-granule by a glycophosphatidylinositol anchor (Wood 2008)..

The clinical importance of platelet-derived factor V is evident in individuals with

Quebec platelet disorder, a hemorrhagic disorder involving reduced platelet factor V levels

(Tracy, Giles et al. 1984) due to the enhanced proteolysis of all α-granule proteins by overexpression of urokinase-type plasminogen activator (Kahr, Zheng et al. 2001). An essential role for platelet factor V is further highlighted by the severe bleeding problems associated with factor V New York (Weiss, Lages et al. 2001), a disorder characterized by defective platelet-derived factor V and normal plasma factor V levels. In addition, an individual with an inhibitor to plasma-derived factor V, but whose platelet-derived factor

V is unaffected, demonstrated no adverse bleeding and demonstrated adequate hemostasis during surgery (Nesheim, Nichols et al. 1986). Hemostatic improvement has also been observed in a patient receiving platelet transfusion for a complete factor V deficiency

(Borchgrevink and Owren 1961).

A protective role for factor V in bleeding pathologies has recently been identified, although its significance in hemostasis has been known for decades. Individuals with congenital factor V deficiency often present with hemorrhagic diatheses of variable

severity that correlate poorly with plasma factor V levels (Lak, Sharifian et al. 1998). Two different studies have explained this variation in phenotype by demonstrating that patients deficient in plasma factor V still have residual factor V in their platelets to protect against major bleeding events (Bouchard, Chapin et al. 2005, Bouchard, K. E. Brummel Ziedins et al. 2009, Duckers, Simioni et al. 2010, Castoldi, Duckers et al. 2011). Studies by

Bouchard et al. showed that in a patient completely devoid of factor V, transfusion of the plasma protein allowed for generation of a platelet-derived factor V/Va pool providing hemostatic competence despite a total absence of plasma derived factor V (Bouchard, K.

E. Brummel Ziedins et al. 2009).

1.11. Acquisition of platelet-derived factor V/Va

As stated previously platelet-derived factor V/Va is generated through endocytosis of the plasma-derived molecule by the megakaryocyte. Endocytosis occurs through a two receptor system involving low density lipoprotein receptor related protein-1

(LRP-1) and an uncharacterized factor V receptor (Figure 8) (Bouchard, Meisler et al.

2008). LRP-1 is a member of the low desnsity lipoprotein receptor family consisting of a

515-kDa α-subunit containing several extracellular ligand binding domains, and a non- covalently associated 85-kDa β-subunit, which includes a transmembrane domain and a short cytoplasmic tail (Herz and Strickland 2001). LRP-1 is found on the cells of several different tissue types where it binds to and delivers a variety of ligands to the lysosome for degradation, including lipoproteins, and coagulation and fibrinolytic proteins (Strickland,

Gonias et al. 2002). Similar to endocytosis of LRP-1 ligands (Li, Marzolo et al. 2000), endocytosis of factor V is calcium-(Bouchard, Meisler et al. 2008) and clathrin-dependent

(Bouchard, Williams et al. 2005). Factor V also displays a high level of homology to a known LRP-1 ligand, factor VIII (Gitschier, Wood et al. 1984, Toole, Knopf et al. 1984,

Vehar, Keyt et al. 1984, Jenny, Pittman et al. 1987, Bouchard, Meisler et al. 2008). In order to examine a role for LRP-1 in factor V binding to megakaryocytes, the effect of receptor- associated protein (RAP) on factor V binding was assessed (Bouchard, Meisler et al. 2008).

In vivo, RAP functions as an LRP-1 chaperone and has been used extensively to determine a role for LRP-1 in various cellular processes (Bouchard, Meisler et al. 2008). Under conditions precluding endocytosis (i.e. in the absence of Ca2+), 125I-factor V binding is reversible and time-dependent, with binding reaching steady state at ~20 min (Bouchard,

Meisler et al. 2008).

RAP displaced ~50% of the bound 125I-factor V from the megakaryocyte cell surface when added 10 minutes after the initiation of binding (Bouchard, Meisler et al.

2008). Addition of excess factor V or factor V plus RAP displaced ~90% of the bound radiolabeled factor V. In a similar experiment, anti-LRP-1 antibodies displaced ~40% of the bound 125I-factor V from the cell surface. In contrast, factor V endocytosis was inhibited nearly 100% in the presence of excess RAP. Concentration-dependent binding of radiolabeled factor V to megakaryocytes was sigmoidal, suggesting cooperative binding

(Bouchard, Meisler et al. 2008). These data were subjected to regression analysis by fitting to the Hill equation. Factor V binding was seen to be saturable and positively cooperative

Figure 8. Hypothetical model describing the megakaryocyte cell surface events mediating factor V endocytosis.

Two receptors, including LRP-1 and the currently unidentified factor V receptor (FV Rc), mediate factor V endocytosis by megakaryocytes. Factor V (FV) initially binds to the FV receptor which facilitiates subsequent binding of a second molecule of FV to LRP-1.

Endocytosis is initiated by LRP-1 in clathrin- and Ca2+- dependant manners. Once inside the megakaryocyte, factor V is phsyically- and functionally-modified before trafficking to the α-granule. LRP-1 is hypothesized to recycle back to the cell surface; however, the fate of the factor V receptor is unknown.

with a Hill coefficient of 1.92 (Bmax = 1038 ± 60 pM). The addition of excess RAP inhibited factor V binding by 50% (Bmax = 618 ± 91 pM) and reduced the Hill coefficient to approximately 1. Cell surface expression of LRP-1 by ex vivo-derived megakaryocytes was confirmed using anti-LRP-1 antibodies and was consistent with the detection of LRP-1 message in these cells (Bouchard, Meisler et al. 2008, Lambert, Wang et al. 2009). RT-

PCR for other low density lipoprotein family members was negative (Bouchard, Meisler et al. 2008).

Based on these observations, a model for factor V endocytosis has been developed

(Figure 8) (Bouchard, Meisler et al. 2008), in which megakaryocytes that can endocytose factor V express both a specific factor V receptor and LRP-1. A molecule of factor V will bind to the factor V receptor causing a second molecule of factor V to bind to LRP-1. LRP-

1 will initiate endocytosis in calcium- and clathrin- dependent manners. Once internalized factor V is physically and functionally modified prior to trafficking to the α-granule.

Traditionally LRP-1 ligands are trafficked to the lysosome for degradation and the receptor is recycled back to the surface. Thus, generation of the platelet-derived factor V pool represents a novel role for LRP-1 in the endocytosis of a ligand not destined for degradation.

While the region of LRP-1 that binds factor V has yet to be identified, it is known that the light chain of factor V is important for endocytosis (Bouchard, Abdalla et al. 2013).

Purified regions of factor V were incubated with CMK cells and their effect on factor V binding was assessed. Endocytosis was inhibited by human and bovine factor V and Va by

nearly 100%. Additionally, B-domainless factor V inhibited endocytosis by 86%. Further, purified factor V heavy chain, did little to inhibit factor V binding, demonstrating that the heavy chain and B-domain are not involved. However, the purified light chain prevented binding of factor V to the megakaryocyte cell surface by almost 100%. Additionally, antibodies directed at the factor V light chain inhibited factor V binding, while the heavy chain antibodies did not, further suggesting that it is the light chain that is important for factor V endocytosis by megakaryocytes.

It has been suggested that the specific factor V receptor is the glycan binding protein, galectin-8 (Zappelli, van der Zwaan et al. 2012). Traditionally cytosolic proteins, galectin family members were recently shown to be secreted (Hughes 1999) from cells and tethered back to the cell membrane to aid in endocytosis (Ochieng, Furtak et al. 2004,

Rabinovich, Toscano et al. 2007). They act by cross-linking specific glycoproteins which triggers an intracellular signaling cascade (Rabinovich, Toscano et al. 2007) to alter cell differentiation, cellular adhesion, growth regulation, and apoptosis (Leffler 2001). Platelets were recently shown to be activated by galectins-1 and -8 (Pacienza, Pozner et al. 2008,

Romaniuk, Tribulatti et al. 2010). Additionally factor V was identified as a potential ligand of galectin-8 (Romaniuk, Tribulatti et al. 2010) using in vitro techniques including immunoaffinity purification of platelet lysates and solid phase binding assays. Presumably these interactions take place between galectin-8 and the complex N-linked glycans containing β-galactoside sugar units on factor V. The megakaryocyte-like cell line, DAMI, was shown to express galectin-8 on the cell surface (Zappelli, van der Zwaan et al. 2012).

Endocytosis of factor V by these cells could be inhibited through the use of purified β- galactoside sugars or α-galectin-8 antibodies (Zappelli, van der Zwaan et al. 2012). Further, factor V endocytosis was decreased in cells treated with galectin-8 siRNA (Zappelli, van der Zwaan et al. 2012). Together, these results suggest that galectin-8 plays a role in factor

V endocytosis at the surface of the megakaryocyte-like cell line, DAMI.

1.12. Summary

Studies aimed at understanding the mechanism by which factor V is internalized by megakaryocytes are important given the significance of platelet factor V in maintaining normal hemostasis, and the fact that this cellular process could be manipulated into a potential therapy hemostatic maintenance. Presently, the endocytic processes is not fully understand, and key questions including the identity of the unknown factor V receptors remain to be addressed. The goal of the studies detailed in this dissertation was to develop a cellular model by which to study factor V endocytosis and assess a potential role for the glycans in the factor V light chain in endocytosis by megakaryocytes. In

Chapter 2, factor V endocytosis and expression of megakaryocyte specific markers will be assessed over the course of megakaryocyte differentiation. Chapter 3 will use glycosidases to selectively trim the complex N-linked glycans in factor V and determine if they play a role in factor V endocytosis by the megakaryocyte.

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Chapter 2. Expression of CD42 is a phenotypic characteristic of megakaryocytes

capable of factor V endocytosis and its trafficking to proplatelet extensions

J.M. Gertz*, H. Schwertz†, K. McLean‡, and B.A. Bouchard*

*Department of Biochemistry and ‡Department of Obstetrics, Gynecology and

Reproductive Sciences, University of Vermont, Burlington, VT; and Department of

Surgery, University of Utah.†

Address correspondence to: Beth A. Bouchard, Ph.D.

Department of Biochemistry

University of Vermont College of Medicine

89 Beaumont Avenue, Given C440

Burlington, VT 05405-0068, USA

Tel.: +1 (802)656-6529; Fax: +1 (802)656-8220

E-mail: [email protected]

Running title: Factor V endocytosis and trafficking to proplatelets

Total figure number: 6

Total table number: 1

2.1. Summary

Background: Plasma- and platelet-derived factor Va are essential for thrombin generation via the prothrombinase complex. Both cofactors function efficiently in thrombin generation; however, several observations demonstrate that the platelet-derived cofactor, which is formed following megakaryocyte endocytosis and modification of the plasma procofactor, factor V, is more hemostatically relevant. Objective: The goal of the current study was to relate factor V endocytosis to megakaryocyte differentiation and proplatelet formation. Methods: CD34+-derived hematopoietic progenitor cells were isolated from human umbilical cord blood and cultured for 12 days in the presence of cytokines to induce ex vivo differentiation into megakaryocytes. Endocytosis of AlexaFluor488-labeled factor

V, expression of megakaryocyte-specific markers, and proplatelet formation were assessed by flow cytometry and microscopy. Results: The percentage of cells that endocytosed factor V and expressed an early marker of megakaryocyte differentiation, CD41, remained constant (days 6 -12). In contrast, the expression of the stem cell marker CD34 decreased while expression of CD42, a late marker of megakaryocyte differentiation, increased. By day 12 nearly all CD42+ cells were capable of factor V endocytosis, and expressed CD41.

In addition, the density of CD41 and CD42 was greatest on megakaryocytes that endocytosed increased amounts of factor V on day 12. Following its endocytosis, factor V localized in granules within megakaryocytes and was trafficked to proplatelet extensions, which expressed CD42 on their surface. Conclusion: In megakaryocytes derived ex vivo from umbilical cord blood, expression of CD42, a late marker of megakaryocyte

differentiation, appears to define cells capable of factor V endocytosis and trafficking to proplatelet extensions.

Keywords: Blood Platelets, Factor V, Megakaryocytes, Platelet Glycoprotein GPIb-IX

Complex, Umbilical Cord

2.2. Introduction

Platelet- and plasma-derived factor Va are absolutely essential for physiological thrombin generation catalyzed by the prothrombinase complex, a 1:1, Ca2+-dependent complex of the serine protease factor Xa and the non-enzymatic cofactor factor Va bound to the surface of activated platelets adhered to sites of vascular injury (Mann, Nesheim et al. 1990). Although platelet-derived factor Va represents only 20-25% of the total whole blood pool of the cofactor (Tracy, Eide et al. 1982), its storage in and release from platelet

α-granules (Breederveld, Giddings et al. 1975, Vicic, Lages et al. 1980, Chesney, Pifer et al. 1981) allows for an increased local concentration at sites of vascular injury as compared with the plasma-derived pool (Nesheim, Nichols et al. 1986). Indeed, several clinical observations suggest that the platelet-derived cofactor plays a more hemostatically relevant role in clot formation at sites of vascular injury (Tracy, Giles et al. 1984, Nesheim, Nichols et al. 1986, Camire, Kalafatis et al. 1995, Camire, Kalafatis et al. 1998, Weiss, Lages et al.

2001, Bouchard, Chapin et al. 2015). Furthermore, the platelet-derived factor Va pool is physically distinct (Monkovic and Tracy 1990, Kalafatis, Rand et al. 1994, Rand, Kalafatis et al. 1994, Camire, Kalafatis et al. 1995, Conlon, Camire et al. 1997, Gould, Silveira et al.

2004, Gould, Simioni et al. 2005, Wood 2008, Tracy, Jennings et al. 2013), and functions as a more procoagulant cofactor (Lee and Mann 1989, Monkovic and Tracy 1990, Conlon,

Camire et al. 1997, Camire, Kalafatis et al. 1998, Gould, Silveira et al. 2004) as compared to its plasma-derived counterpart. Despite these differences, the platelet-derived factor V pool is generated through endocytosis of the procofactor, factor V, from plasma by

megakaryocytes, platelet precursor cells (Bouchard, Williams et al. 2005, Suehiro,

Veljkovic et al. 2005).

Megakaryocytes arise from multipotent CD34+ hematopoietic stem cells residing in the bone marrow (Lu, Briddell et al. 1988). Megakaryocyte differentiation is controlled by numerous cytokines and growth factors including thrombopoietin (TPO) (Bruno,

Briddell et al. 1988, Bartley, Bogenberger et al. 1994, de Sauvage, Hass et al. 1994,

Gurney, Carvermoore et al. 1994, Kaushansky 1994, Kaushansky, Lok et al. 1994, Kuter,

Beeler et al. 1994, Wendling, Maraskovsky et al. 1994, Kaushansky 1995) , stem cell factor

(SCF) (Ebbe, Phalen et al. 1973) and interleukin-3 (IL-3) (Emerson, Yang et al. 1988,

Briddell and Hoffman 1990), and is characterized by acquisition of a polyploidal DNA content (up to 128N) through an endomitotic cell cycle (De Leval 1964, Marianogarcia

1964) and expression of proteins important for platelet function (Rabellino, Nachman et al. 1979). Expression of these proteins is differentially regulated and includes: CD41

[glycoprotein (GP) IIb; integrin αIIb], a cell surface protein involved in platelet aggregation

(Phillips, Charo et al. 1988) during primary hemostasis that is expressed early in megakaryocyte differentiation (Debili, Robin et al. 2001), and CD42 (GPIbα), a cell surface protein expressed at later stages of megakaryocyte differentiation (Debili, Robin et al. 2001) that functions as a receptor for von Willebrand factor (vWF) (Kroll, Harris et al.

1991). Megakaryocytes also synthesize (Heijnen, Debili et al. 1998) (e.g. vWF and P- selectin) and endocytose (e.g. fibrinogen (Handagama, Scarborough et al. 1993) and factor

V (Bouchard, Williams et al. 2005, Suehiro, Veljkovic et al. 2005)) a variety of hemostatic

proteins that are trafficked to and stored in α-granules to be released from activated platelets at sites of vascular injury during the hemostatic response (Jandrot-Perrus, Lagrue et al. 1997). Mature megakaryocytes are large (up to 60 μm) and undergo extensive remodeling to produce proplatelet extensions that are shed to form functional platelets

(Bunting 1909). Platelets are anucleate and contain α-granules, dense granules, lysosomes, and other organelles derived from the megakaryocyte (Richardson, Shivdasani et al. 2005).

While the synthesis of various platelet-specific proteins as a function of megakaryocyte differentiation is well defined, little is known about how factor V endocytosis is developmentally regulated. In the current study, factor V endocytosis was correlated with expression of megakaryocyte/platelet specific markers and proplatelet formation using megakaryocytes derived ex vivo from CD34+ cells isolated from human umbilical cord blood.

2.3. Materials and methods

Materials: Ficoll-Paque was purchased from GE Healthcare (Pittsburgh, PA, USA) and

EasySep human CD34 selection kit was purchased from Stem Cell Technologies

(Vancouver, British Columbia, Canada). X-Vivo 20 serum-free media was obtained from

BioWhittaker (Walkersville, Maryland, USA) and the hematopoietic cytokines TPO, SCF and IL-3 were acquired from Life Technologies (Grand Island, NY, USA). The following antibodies were used in these studies: mouse anti–human CD34 and mouse anti-human

CD42 (BD Pharmingen, San Jose, Ca, USA); mouse anti-human CD41 clone 5B12 conjugated to phycoerythrin (PE) and mouse IgG1-PE (Dako, Carpinteria, CA, USA); and goat anti–mouse IgG conjugated to AlexaFluor647 (Life Technologies). Human factor V, purified from freshly frozen plasma as described (Katzmann, Nesheim et al. 1981), and human factor IX (Haematologic Technologies, Inc., Essex Junction, VT, USA) were conjugated to AlexaFluor488 using the manufacturer’s (Life Technologies) instructions

(Bouchard, Williams et al. 2005, Bouchard, Meisler et al. 2008). Eight well covered borosilicate chamber slides were purchased from Nunc Lab-Tek (Rochester, NY, USA) and coated with sterile human plasma fibrinogen (100 μg/mL; 30 min at 37°C or overnight at 4°C) (Calbiochem, Billerica, MA, USA). Prior to their use, non-specific binding sites were blocked by incubating fibrinogen-coated slides with 1% bovine serum albumin (30 min, 37°C).

Ex vivo-differentiation of megakaryocytes: Cord blood from 2-4 full-term deliveries by elective cesarean section was obtained at The University of Vermont Medical Center

(formerly Fletcher Allen Healthcare) after verbal consent using a protocol approved by the

University of Vermont Committee on Human Research. On day 0 (Figure 9)

CD34+ hematopoietic progenitor cells were isolated and cultured for differentiation into megakaryocytes as previously described (Denis, Tolley et al. 2005, Foulks, Marathe et al.

2009, Shi, Smith et al. 2014). Briefly, mononuclear cells were isolated using centrifugation through Ficoll (400xg, 30 min). From the mononuclear cells, CD34+ cells were isolated by immunomagnetic separation with anti-CD34-coupled beads according to the manufacturer’s protocol. Isolated cells were cultured (37°C, 5% CO2) in X-Vivo 20 serum free media in the presence of TPO (50 ng/mL), SCF (40 ng/mL), and IL-3 (10 ng/mL).

Cell culture media was changed every 2-3 days when cells were at a concentration of ~4 x

106 cells/mL. Cultures were reseeded at a concentration of 0.5 x 106 cells/mL in fresh media. From day 5 onward, the cells were cultured in X-Vivo 20 and TPO (50 ng/mL) and

SCF (40 ng/mL) alone (Figure 9).

Flow cytometric analyses: On day 5 (Figure 9) an aliquot of the cells (1x106 cells/mL) was removed and cultured with AlexaFluor488-labeled factor V or IX (30nM, 16 hr) in culture media. The following day (day 6) factor V endocytosis and cell surface expression of CD34, CD41, and CD42 by these cells was assessed by flow cytometry using the following protocol. The cells were removed from the culture dish, washed two times by

Figure 9. Isolation and analysis of megakaryocytes cultured ex vivo from umbilical cord blood-derived CD34+ cells.

CD34+ multipotent hematopoietic stem cells were isolated from human umbilical cord blood and cultured in serum free media containing thrombopoietin (TPO), stem cell factor

(SCF), and interleukin-3 (IL-3) (days 0 -5) as described. On the days indicated by the arrows (days 5, 7, 9, and 11), cells (1x106/mL) were incubated (16 hr, 37°C) with 30 nM

AlexaFluor488-labeled factor V or factor IX in culture media. The next day (days 6, 8, 10, or 12), factor V endocytosis and cell surface expression of CD34, CD41, and CD42 were assessed by flow cytometry (X).

centrifugation (500xg, 5 min) followed by resuspension in 20 mM Hepes, 0.15 M NaCl, pH 7.4 (HBS) to remove cell surface–associated factor V- and factor IX-AlexaFluor488.

Non-specific binding sites were then blocked using 1% bovine serum albumin. Cells were immunostained for 1 hr on ice with the following monoclonal antibodies: anti-CD34 (1

μg/mL), anti-CD41-phycoerythrin (PE) (7.4 μg/mL), or anti-CD42 (20 μg/mL). Cells were washed to remove excess antibodies as described above. Reactions containing anti-CD34 or anti-CD42 were incubated for 1 hr on ice with Alexa647-conjugated goat anti-mouse

IgG (4 μg/mL) then washed to remove excess antibodies. Samples were resuspended in 4% paraformaldehyde and incubated for at least 30 min before flow cytometry analysis. For three color flow cytometry experiments CD42 immunostaining was performed as described above. Subsequently, cells were washed and immunostained with anti-CD41-PE as described above prior to fixation with 4% paraformaldehyde.

Cells (10,000) were analyzed on a BD LSR II flow cytometer (BD Biosciences,

San Jose, CA). The cell population of interest was first selected based on forward and side scatter. Analysis regions were defined such that ~2% of the cells cultured in the presence of AlexaFlour488-factor IX, or immunostained in the absence of primary antibodies (anti-

CD34 or anti-CD42) or using a non-immune antibody conjugated to PE (anti-CD41). All data were collected as binary FCS files and analyzed using FlowJo version 10 data analysis software (FlowJo, LLC, Ashland, OR, USA). For some analyses, the 10% of megakaryocytes with the highest fluorescence intensity corresponding to factor V-

AlexaFluor488 endocytosis were identified (super factor V+). Expression of CD34, CD41

and CD42 was assessed within this population and recorded as the median fluorescence intensity (mean ± SEM). The y-axis on all histograms is modal. The percent cells (mean ±

SEM) that endocytosed factor V and/or expressed CD34, CD41, and CD42 on days 8, 10 and 12 were compared to that on day 6 using a student’s T-test. p≤0.05 was considered significant.

Assessment of proplatelet formation using live cell imaging: Day 12 ex vivo-derived megakaryocytes were seeded on fibrinogen-coated slides and cultured for 24 hr to induce proplatelet formation. The cells were visualized on an Olympus upright BX50 light microscope every 30 min to assess proplatelet formation. Once proplatelet formation was observed, the slides were transferred to a Zeiss LSM 510 Laser Scanning Confocal

Microscope with an incubation chamber maintained at 37°C and 5% CO2. Differential interference contrast (DIC) images (40X) were taken every 15 min until no change in proplatelet formation was seen.

Confocal microscopy: Day 12 cells were incubated with 30 nM factor V-AlexaFluor488 or factor IX-AlexaFluor488 for 8 hr at 37°C, washed three times as described above in X-

Vivo 20 serum free media, and cultured on fibrinogen-coated chamber slides (48 hr, 37°C) to induce proplatelet formation (Lindemann, Tolley et al. 2001, Galt, Lindemann et al.

2002, Denis, Tolley et al. 2005, Schwertz, Tolley et al. 2006, Weyrich, Denis et al. 2007).

Cells and proplatelet extensions were subjected to fixation with 4% paraformaldehyde (30 min, ambient temperature), rinsed three times with HBS, permeabilized with 0.1% triton

X (5 min), and rinsed three times with HBS. Non-specific binding sites were blocked by incubation with 10% goat serum for 1 hr. Following incubation with anti-CD42 (10 μg/mL,

1 hr), the cells were rinsed three times with HBS to remove excess antibody and incubated with goat anti-mouse IgG conjugated to AlexaFluor647 (4 μg/mL, 1 hr, ambient temperature). After rinsing the cells with HBS to remove excess antibodies, they were incubated with Hoechst (5 μg/mL, 5 min). The cells were viewed by confocal microscopy using a 63X objective lens with a numerical aperture of 1.4 on a Zeiss LSM 510 Laser

Scanning Confocal Microscope (Carl Zeiss Microimaging, Thornwood, NY, USA) with photo multiplier tube values adjusted to no signal with IgG controls.

2.4. Results

Correlation of factor V endocytosis with markers of megakaryocyte differentiation.

Previous observations by Weyrich et al. demonstrate that megakaryocytes derived ex vivo from human umbilical cord blood express early markers of megakaryocyte differentiation, CD41 and CD61, and store P-selectin in α-granules (Denis, Tolley et al.

2005). This cell system was used to examine the relationship of factor V endocytosis to markers of megakaryocyte differentiation by flow cytometry (Figure 10). In our initial experiments, propidium iodide staining demonstrated that the DNA content of the cells does not exceed 2N/4N over the culture period (data not shown), confirming that the cells 72

Figure 10. Factor V endocytosis and surface expression of megakaryocyte lineage markers.

Factor V endocytosis was assessed in the total cell population (A) (n=20) and in conjunction with cell surface markers (B, C, and D, gray circles, gray lines) (n=5) on days 6, 8, 10 and 12 of differentiation by flow cytometry as described in Materials &

Methods. Cell surface expression of (B) CD34, (C) CD41, and (D) CD42 was also assessed in the total cell population on days 6, 8, 10 and 12 (open circles, dashed lines)

(n=5) as described. The percentage of total cells that endocytosed factor V and also express (B) CD34, (C) CD41, and (D) on their membrane surface are also shown (black circles, black line). All data are expressed as the mean ± SEM. *p≤0.002 compared to day 6; **p≤0.02 compared to day 6; ***p≤0.05 compared to day 6.

do not become polyploidal under the culture conditions (Denis, Tolley et al. 2005) used in this study. Therefore, DNA content was not used as a maturation marker. The percentage of cells that endocytosed AlexaFluor488-labeled factor V remained nearly constant over the course of megakaryocyte differentiation (Figure 10A) (34.6 ± 2.8% vs. 40.8 ± 3.5% on days 6 and 12, respectively) and did not vary substantially among each experimental group

(Figures 10B, C and D, gray circles, gray line). On day 6 of differentiation, 82.2 ± 3.6% of the cells expressed CD34, a multipotent hematopoietic stem cell marker (Figure 10B, open circles, dotted line), 36.6 ± 8.8% of the cells expressed CD41, a marker of the multipotent megakaryocyte precursor cell (Basch, Zhang et al. 1999, Proulx, Boyer et al. 2003, Boyer,

Robert et al. 2008) (Figure 10C, open circles, dotted line), and 12.6 ± 1.1% of the cells expressed CD42, a megakaryocyte marker that is expressed late in differentiation (Debili,

Robin et al. 2001) (Figure 10D, open circles, dotted line). A statistically significant decrease in the number of CD34+ cells was observed over the course of megakaryocyte differentiation (Figure 10B, open circles, dotted line). On days 10 and 12, only 26.9 ± 4.1% and 18.6 ± 3.6% of the cells, respectively, expressed CD34 (p≤0.002, days 10 and 12).

While the expression of CD41 only slightly increased over the course of differentiation

(44.8 ± 6.0% on day 12) (Figure 10C, open circles, dotted line), a statistically significant increase in expression of CD42 was observed over this time such that, by day 10, 20.5 ±

2.4% of the cells were CD42+ (p≤0.02), and by day 12 25.4 ± 5.8% of the cells were positive for expression of this marker (p≤0.05) (Figure 10D, open circles, dotted line).

Multiparameter flow cytometric analyses were used to assess CD34, CD41 and

CD42 expression on cells capable of factor V endocytosis. Over the course of megakaryocyte differentiation, the percentage of cells that endocytosed factor V and expressed CD34 decreased two-fold from 27.2 ± 5.0% on day 6 to 8.1 ± 1.9% on day 10 and 11.7 ± 3.4% on day 12 (Figure 10B, black circles, black line) (p≤0.02, days 10 and

12). Although the total numbers of cells that endocytosed factor V and expressed CD41 were similar over the course of differentiation, two color flow cytometric analyses clearly demonstrated that the CD41+ and factor V+ cell populations do not overlap. While the percentage of factor V+CD41+ cells nearly doubled from 9.0 ± 2.7% on day 6 to 17.7 ±

4.0% at day 12, this represents less than half of the total CD41+ cell population (Figure

10C, black line vs. dotted line). The percentage of factor V+CD42+ cells increased nearly three-fold over the course of differentiation from 8.6 ± 2.0% (day 6) to 21.0 ± 6.5% (day

12) (Figure 10D, black circles black line) (p=0.053). Comparison of these data with the increase in the total CD42+ cell population over the course of megakaryocyte differentiation (Figure 10D, open circles dotted line) demonstrated that on day 12 the percentage of factor V+CD42+ cells and the percentage of all CD42+ cells were not statistically different (p=0.324), suggesting that nearly 100% of the cells that express CD42 endocytose factor V. Three color flow cytometric analyses confirmed that nearly all of the factor V+CD42+ cells expressed CD41 on their cell surface (Figure 11). These studies also demonstrated the presence of a population of factor V+ cells that do not express either megakaryocyte marker (data not shown).

Figure 11. Nearly all factor V+CD42+ cells express CD41.

Endocytosis of factor V and cell surface expression of CD41 and CD42 were assessed on day 12 by tricolor flow cytometry as described in Materials and Methods. Immunostaining of the factor V+CD42+ cell population with a non-immune antibody conjugated to PE is depicted by the gray histogram. The overlay is the population of factor V+CD42+ cells that express CD41+ (black histogram) following immunostaining with anti-CD41-PE.

Further analyses were done to determine if the population of cells containing the most endocytosed factor V (super factor V+) also expressed different amounts of megakaryocyte differentiation markers on their membrane surfaces. A three-fold increase was observed in the median fluorescence intensity of CD42+ cells in the super factor V+ population between days 6 and 12 (Figure 12, black histograms) (Table 2). A similar trend was seen for expression of CD41 on the super factor V+ population (Table 2). In contrast, the density of the stem cell marker, CD34, decreases in the super factor V+ population over time (Table 2).

Factor V is trafficked to proplatelet extensions.

Approximately 20% of the megakaryocytes derived ex vivo from human umbilical cord blood generate proplatelet extensions upon culture on fibrinogen (Proulx, Boyer et al.

2003). Live cell imaging and time lapse photography confirmed formation of proplatelet extensions under the present culture conditions (Figure 13). Additional experiments used confocal microscopy to determine if endocytosed factor V is trafficked to proplatelet extensions, and to define the co-distribution of endocytosed factor V and CD42 (Figure

14). Endocytosed AlexaFluor488-factor V (white pixels) was observed in the cell bodies and proplatelet extensions suggesting that following its packaging in megakaryocyte α- granules (Bouchard, Taatjes et al. 2006) factor V is trafficked to proplatelet extensions

(Figure 14A and B). The same cells and proplatelets that endocytosed factor V also immunostained positively for CD42 (Figure 14B). In contrast to factor V, which was

Figure 12. CD42 expression increases over time on super factor V+ cells.

Factor V endocytosis and CD42 expression were assessed in the total cell population on days 6, 8, 10 and 12 as described in Materials and Methods. CD42 expression on each day is depicted by the gray histogram. CD42 expression by the populations of cells with the highest 10% median fluorescence intensities with regards to factor V-AlexaFluor488 endocytosis (super factor V+) (black histogram) are shown as overlays.

Table 2. Median fluorescence intensities of differentiation markers in the super factor

V+ megakaryocyte population.

Day 6 8 10 12 CD34a 1266.2 ± 348.9 942.8 ± 285.6 426.7 ± 142.5 561.6 ± 206.3 CD41a 852.0 ± 294.8 795.4 ± 366.5 4605.3 ± 2336.0 10814.0 ± 8420.5 CD42a 707.4 ± 681.2 1191.3 ± 604.3 1345.4 ± 772.3 5284.2 ± 2175.6

a Surface expression of CD34, CD41, or CD42 was assessed on super factor V+ cells as described in Materials & Methods. The median fluorescence intensities corresponding to marker expression (mean ± SEM) are shown.

Figure 13. Live cell imaging of proplatelet formation by megakaryocytes derived ex vivo from human umbilical cord blood.

Day 12 cells were cultured on fibrinogen-coated slides and proplatelet formation was assessed and documented as described in Materials & Methods. The differential interference contrast images (40X) are shown. The white arrows are used to indicate the formation of two different proplatelet extensions. Scale bar = 10 µm. For each individual image, the time of image acquisition is shown in seconds.

Figure 14. CD42+ proplatelets contain endocytosed factor V.

A) Day 12 cells were incubated (8 hr, 37°C) with 30 nM AlexaFluor488-factor V in culture media. Cells were subsequently cultured on fibrinogen-coated slides as described to induce the formation of proplatelet extensions (black arrow). Endocytosed factor V in the cell bodies and proplatelet extensions is pseudocolored white; nuclei are stained using Hoechst

(blue). Scale bar = 10 μm. B) Following factor V endocytosis and formation of proplatelet extensions, immunostaining for CD42 was performed as described in Materials and

Methods. Top: Overlay of a DIC and fluorescence image. Proplatelet extension, black arrow; nuclei, blue. Scale bar = 10 μm. Bottom: A fluorescence image of the same proplatelet extension demonstrating the presence of endocytosed factor V (white) and

CD42 immunostaining (red). Scale bars = 10 μm.

localized in the cytoplasm in a punctate pattern, CD42 was localized on the membrane surface.

2.5. Discussion

Megakaryocytes derived ex vivo from CD34+ cells isolated from human umbilical cord blood endocytose human factor V and traffic it to proplatelet extensions. Following its endocytosis from plasma by megakaryocytes, factor V undergoes retrograde transport through the Golgi apparatus (Bouchard, Taatjes et al. 2006) and is modified to form the physically and functionally unique platelet-derived cofactor (Monkovic and Tracy 1990,

Kalafatis, Rand et al. 1994, Rand, Kalafatis et al. 1994, Camire, Kalafatis et al. 1995,

Conlon, Camire et al. 1997, Gould, Silveira et al. 2004, Gould, Simioni et al. 2005, Wood

2008, Tracy, Jennings et al. 2013). It is subsequently trafficked to and stored in α-granules

(Suehiro, Veljkovic et al. 2005). Endocytosed factor V was observed in CD42+ cells and proplatelet extensions. It was distributed in the cytoplasm in a punctate pattern consistent with its localization in α-granules (Suehiro, Veljkovic et al. 2005). Granules are derived from vesicles containing granule cargo from the trans-Golgi network (Jones 1960, Cramer,

Harrison et al. 1990, Hegyi, Heilbrun et al. 1990, Blair and Flaumenhaft 2009) in a process that appears to be dependent upon proteins involved in regulation of vesicle trafficking

(Blair and Flaumenhaft 2009, Kahr, Hinckley et al. 2011, Urban, Li et al. 2012, Kahr, Lo et al. 2013). These vesicles mature into multivesicular bodies and granules where the granule proteins are sorted (Heijnen, Debili et al. 1998). It is not clear at what stage of

megakaryocyte differentiation α-granule formation commences; however, two studies suggest that immature (2N and 4N) megakaryocytes contain no detectable α-granules

(Bretongorius and Reyes 1976, Hegyi, Heilbrun et al. 1990). In contrast, a more recent study using fresh bone marrow aspirates suggests that the α-granule protein vWF is a sensitive and distinctive marker for 2N and 4N megakaryocytes (Tomer 2004). Proplatelets are formed following remodeling of the microtubule cytoskeleton (Tablin, Castro et al.

1990, Hartwig and Italiano 2006). Granules and organelles are transported along these microtubules to the proplatelet ends (Richardson, Shivdasani et al. 2005). Large preplatelets are released from proplatelets into the vasculature where they undergo fission events to become mature platelets (Thon, Montalvo et al. 2010). The appearance of endocytosed factor V in the cell bodies suggests that factor V endocytosis and packaging into α-granules occurs prior to proplatelet formation.

While those cells capable of factor V endocytosis cannot be defined by a single phenotype, nearly all CD41+CD42+ megakaryocytes endocytose factor V by day 12.

Furthermore, the population of megakaryocytes that endocytose more factor V express higher densities of CD41 and CD42 on the membrane surface. These observations are consistent with previous studies using megakaryocytes derived ex vivo from human bone marrow (Bouchard, Williams et al. 2005). In these experiments, days 4 and 7 cells were incubated with a plasma concentration of plasma-derived factor V followed by flow cytometric analyses at days 7 and 10, respectively, to correlate factor V endocytosis with expression of CD41 and DNA content. Only the more mature CD41+ day 10 cells were

capable of substantial uptake of factor V. Furthermore, only CD41+ cells with >4N DNA content endocytosed factor V (Bouchard, Williams et al. 2005). On day 12, a population of

CD41-CD42- cells capable of factor V endocytosis was also observed. On the same day,

~12% of the cells that endocytosed factor V also expressed CD34. Expression of CD34 by immature megakaryocytes derived ex vivo from human bone marrow (Debili, Issaad et al.

1992, Norol, Vitrat et al. 1998) or mobilized blood cells (Norol, Vitrat et al. 1998, Debili,

Robin et al. 2001) has been described previously. Therefore, it is likely that these cells represent a small population of CD34+ megakaryocyte progenitors capable of factor V endocytosis. Under the conditions used in this study, the cell numbers obtained were inadequate to allow for flow cytometric analyses prior to day 5/6 of culture. Furthermore, at day 6 of culture a small, but not unsubstantial fraction of the cells expressed CD42, a megakaryocyte/platelet specific marker that is expressed during later stages of differentiation. Therefore, we were unable to assess factor V endocytosis at the earliest stages of megakaryocyte expansion and differentiation.

Interestingly, not all CD41+ cells are capable of factor V endocytosis. This observation is consistent with previous studies using megakaryocyte-like cell lines and megakaryocytes derived ex vivo from human bone marrow (Bouchard, Williams et al.

2005). These observations can be interpreted in one of two ways. One possibility is that only a subpopulation of CD41+ cells expresses the machinery necessary for factor V endocytosis. Factor V endocytosis is mediated by a two receptor system consisting of low- density lipoprotein receptor related protein-1 (LRP-1), a ubiquitous endocytic receptor, and

an unidentified, specific factor V receptor (Bouchard, Meisler et al. 2008). As cell surface expression of LRP-1 is unchanged over the course of differentiation (data not shown), these data suggest that only some megakaryocytes express the unidentified factor V receptor.

Another possibility is that while these cells express CD41, they are not all destined to become megakaryocytes. Indeed, not all cells cultured under the conditions of this study developed into mature megakaryocytes and generated proplatelets. At day 12 of culture we observed that only ~25% of the cells expressed CD42, which is consistent with previous observations (Debili, Robin et al. 2001). Although the conditions of culture are designed to favor megakaryocyte development, several studies have reported that ex vivo culture of

CD34+ cells can lead to expansion of not only megakaryocytes, but also cells with myeloid and lymphoid properties including those that express CD41 (Tao, Gaudry et al. 1999,

Proulx, Boyer et al. 2003, Cortin, Garnier et al. 2005, Matsunaga, Tanaka et al. 2006,

Boyer, Robert et al. 2008). This observation would also suggest that the ability to endocytose factor V may be a useful megakaryocyte-specific marker in a mixed cell population.

While megakaryocytes derived ex vivo from CD34+ human umbilical cord blood appear to be a suitable megakaryocyte model to study factor V endocytosis and trafficking to proplatelet α-granules, it is not without its limitations. As knowledge regarding the cellular and molecular events regulating megakaryocyte differentiation and platelet formation is gained, it is likely that improved culture conditions for expansion and differentiation of human megakaryocytes ex vivo will be identified which will be useful for

further studies of factor V endocytosis and packaging in α-granules. These studies are essential as even small changes in the level or function of the highly procoagulant cofactor that predominates at sites of vascular injury will have a profound impact on clot formation.

2.6. Authorship details

J.M. Gertz was responsible for the experimental design; data acquisition, analyses and interpretation; and preparation of the manuscript. H. Schwertz provided instruction on cell isolation from umbilical cord blood and culture of ex vivo-derived megakaryocytes. K.

Mclean assisted with collection of umbilical cord blood at the University of Vermont

Medical Center, Burlington, VT, USA. B.A. Bouchard conceived of the research; provided general guidance on experimental design and data acquisition, analyses and interpretation; and preparation of the manuscript.

2.7. Acknowledgements

We would like to thank Andrew S. Weyrich, PhD (University of Utah, Salt Lake City, UT,

USA) for assistance with cell isolation from umbilical cord blood and ex vivo differentiation of megakaryocytes. In addition, we would like to thank Paula B. Tracy,

PhD, Jay R. Silveira, PhD, and Laura M. Haynes, PhD (University of Vermont, Burlington,

VT, USA) for helpful discussions. This work was supported by grants from the NIH (K02

HL91111) (BB) and T32 HL007894 (JG), a grant from the American Heart Association

(SDG 0635048N) (BB), and the University of Vermont College of Medicine. Hansjörg 89

Schwertz was supported by a Post-Doctoral fellowship (0625098Y), a Beginning-Grant- in-Aid (09BG1A 2250381) from the American Heart Association Western States Affiliate, and a Lichtenberg-Professorship from the Volkswagen Foundation.

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47. Lee, C. D. and K. G. Mann (1989). "Activation/inactivation of human factor V by plasmin." Blood 73(1): 185-190.

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49. Lu, L., R. A. Briddell, C. D. Graham, J. E. Brandt, E. Bruno and R. Hoffman (1988). "Effect of recombinant and purified human haematopoietic growth factors on in vitro colony formation by enriched populations of human megakaryocyte progenitor cells." Br J Haematol 70(2): 149-156.

50. Mann, K. G., M. E. Nesheim, W. R. Church, P. Haley and S. Krishnaswamy (1990). "Surface-dependent reactions of the vitamin K-dependent enzyme complexes." Blood 76(1): 1-16.

51. Marianogarcia, A. (1964). "Feulgen-DNA Values in Megakaryocytes." J Cell Biol 20: 342-345.

52. Matsunaga, T., I. Tanaka, M. Kobune, Y. Kawano, M. Tanaka, K. Kuribayashi, S. Iyama, T. Sato, Y. Sato, R. Takimoto, T. Takayama, J. Kato, T. Ninomiya, H. Hamada and Y. Niitsu (2006). "Ex vivo large-scale generation of human platelets from cord blood CD34+ cells." Stem Cells 24(12): 2877-2887.

53. Monkovic, D. D. and P. B. Tracy (1990). "Functional characterization of human platelet-released factor V and its activation by factor Xa and thrombin." J Biol Chem 265(28): 17132-17140.

54. Nesheim, M. E., W. L. Nichols, T. L. Cole, J. G. Houston, R. B. Schenk, K. G. Mann and E. J. Bowie (1986). "Isolation and study of an acquired inhibitor of human coagulation factor V." J Clin Invest 77(2): 405-415.

55. Norol, F., N. Vitrat, E. Cramer, J. Guichard, S. A. Burstein, W. Vainchenker and N. Debili (1998). "Effects of cytokines on platelet production from blood and marrow CD34+ cells." Blood 91(3): 830-843.

56. Phillips, D. R., I. F. Charo, L. V. Parise and L. A. Fitzgerald (1988). "The platelet membrane glycoprotein IIb-IIIa complex." Blood 71(4): 831-843.

57. Proulx, C., L. Boyer, D. R. Hurnanen and R. Lemieux (2003). "Preferential ex vivo expansion of megakaryocytes from human cord blood CD34+-enriched cells in the presence of thrombopoietin and limiting amounts of stem cell factor and Flt-3 ligand." J Hematother Stem Cell Res 12(2): 179-188.

58. Rabellino, E. M., R. L. Nachman, N. Williams, R. J. Winchester and G. D. Ross (1979). "Human megakaryocytes. I. Characterization of the membrane and cytoplasmic components of isolated marrow megakaryocytes." J Exp Med 149(6): 1273-1287.

59. Rand, M. D., M. Kalafatis and K. G. Mann (1994). "Platelet coagulation factor Va: the major secretory platelet phosphoprotein." Blood 83(8): 2180-2190.

60. Richardson, J. L., R. A. Shivdasani, C. Boers, J. H. Hartwig and J. E. Italiano (2005). "Mechanisms of organelle transport and capture along proplatelets during platelet production." Blood 106(13): 4066-4075.

61. Schwertz, H., N. D. Tolley, J. M. Foulks, M. M. Denis, B. W. Risenmay, M. Buerke, R. E. Tilley, M. T. Rondina, E. M. Harris, L. W. Kraiss, N. Mackman, G. A. Zimmerman and A. S. Weyrich (2006). "Signal-dependent splicing of tissue factor pre-mRNA modulates the thrombogenicity of human platelets." J Exp Med 203(11): 2433-2440.

62. Shi, D. S., M. C. Smith, R. A. Campbell, P. W. Zimmerman, Z. B. Franks, B. F. Kraemer, K. R. Machlus, J. Ling, P. Kamba, H. Schwertz, J. W. Rowley, R. R. Miles, Z. J. Liu, M. Sola-Visner, J. E. Italiano, Jr., H. Christensen, W. H. Kahr, D. Y. Li and A. S. Weyrich (2014). "Proteasome function is required for platelet production." J Clin Invest.

63. Suehiro, Y., D. K. Veljkovic, N. Fuller, Y. Motomura, J. M. Masse, E. M. Cramer and C. P. Hayward (2005). "Endocytosis and storage of plasma factor V by human megakaryocytes." Thromb Haemost 94(3): 585-592.

64. Tablin, F., M. Castro and R. M. Leven (1990). "Blood platelet formation in vitro. The role of the cytoskeleton in megakaryocyte fragmentation." J Cell Sci 97 ( Pt 1): 59-70.

65. Tao, H., L. Gaudry, A. Rice and B. Chong (1999). "Cord blood is better than bone marrow for generating megakaryocytic progenitor cells." Exp Hematol 27(2): 293-301.

66. Thon, J. N., A. Montalvo, S. Patel-Hett, M. T. Devine, J. L. Richardson, A. Ehrlicher, M. K. Larson, K. Hoffmeister, J. H. Hartwig and J. E. Italiano, Jr. (2010). "Cytoskeletal mechanics of proplatelet maturation and platelet release." J Cell Biol 191(4): 861-874.

67. Tomer, A. (2004). "Human marrow megakaryocyte differentiation: multiparameter correlative analysis identifies von Willebrand factor as a sensitive and distinctive marker for early (2N and 4N) megakaryocytes." Blood 104(9): 2722-2727.

68. Tracy, P. B., L. L. Eide, E. J. Bowie and K. G. Mann (1982). "Radioimmunoassay of factor V in human plasma and platelets." Blood 60(1): 59-63.

69. Tracy, P. B., A. R. Giles, K. G. Mann, L. L. Eide, H. Hoogendoorn and G. E. Rivard (1984). "Factor V (Quebec): a bleeding diathesis associated with a qualitative platelet Factor V deficiency." J Clin Invest 74(4): 1221-1228.

70. Tracy, P. B., M. E. Jennings, 2nd, K. Stringer, J. R. Silveira, J. P. Wood and A. Blanchard (2013). "Site-specific glycan trimming in megakaryocyte-endocytosed factor V." J Thromb Haemost 11(Suppl. s2): AS36.32.

71. Urban, D., L. Li, H. Christensen, F. G. Pluthero, S. Z. Chen, M. Puhacz, P. M. Garg, K. K. Lanka, J. J. Cummings, H. Kramer, J. D. Wasmuth, J. Parkinson and W. H. Kahr (2012). "The VPS33B-binding protein VPS16B is required in megakaryocyte and platelet α-granule biogenesis." Blood 120(25): 5032-5040.

72. Vicic, W. J., B. Lages and H. J. Weiss (1980). "Release of human platelet factor V activity is induced by both collagen and ADP and is inhibited by aspirin." Blood 56(3): 448-455.

73. Weiss, H. J., B. Lages, S. Zheng and C. P. Hayward (2001). "Platelet factor V New York: a defect in factor V distinct from that in factor V Quebec resulting in impaired prothrombinase generation." Am J Hematol 66(2): 130-139.

74. Wendling, F., E. Maraskovsky, N. Debili, C. Florindo, M. Teepe, M. Titeux, N. Methia, J. Breton-Gorius, D. Cosman and W. Vainchenker (1994). "cMpl ligand is a humoral regulator of megakaryocytopoiesis." Nature 369(6481): 571-574.

75. Weyrich, A. S., M. M. Denis, H. Schwertz, N. D. Tolley, J. Foulks, E. Spencer, L. W. Kraiss, K. H. Albertine, T. M. McIntyre and G. A. Zimmerman (2007). "mTOR- dependent synthesis of Bcl-3 controls the retraction of fibrin clots by activated human platelets." Blood 109(5): 1975-1983.

76. Wood, J. P., Fager, A.M., Silveira, J.R., Tracy, P.B. (2008). "Platelet-derived factor Va expressed on the surface of the activated platelet is GPI-anchored. ." Blood 112(11): 219a.

Chapter 3. Complex N-linked glycans in the factor V light chain are not involved in

endocytosis by megakaryocytes

J.M. Gertz*, M.E. Jennings II†, Cody Couperus*, JR Silveira*, PB Tracy* and B.A. Bouchard*

*Department of Biochemistry and †Department of Chemistry, University of Vermont, Burlington, VT

Address correspondence to: Beth A. Bouchard, Ph.D. Department of Biochemistry University of Vermont College of Medicine 89 Beaumont Avenue, Given C440 Burlington, VT 05405- 0068, USA Tel.: +1 (802)656-6529; fax: +1 (802)656-8220 E-mail: [email protected]

Running title: The role of glycans in factor V endocytosis

Total Figure Number: 6

Total Table Number: 2

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3.1. Summary

Background: Platelet-derived factor V/Va forms the hemostatically relevant cofactor in thrombin generation via the prothrombinase complex at sites of vascular injury. It is generated through endocytosis of plasma-derived factor V by megakaryocytes via a two receptor system including a specific, unidentified factor V receptor and the low density lipoprotein receptor related protein-1. Recent studies have demonstrated that the light chain of factor V is important for its binding and endocytosis by megakaryocytes. One high mannose glycan (Asn1982) and two complex N-linked glycans (Asn1675, Asn2181) have been identified in this domain. Objective: The goal of this study is to determine if complex N-linked glycans in the factor V light chain play a role in endocytosis. Methods:

The factor V glycans were selectively trimmed using exoglycosidases. Glycan removal was assessed using gel electrophoresis and mass spectrometry. The effect of glycan trimming on factor V cofactor function and endocytosis by megakaryocytes were subsequently determined. Results: Treatment of native, human factor V with neuraminidase and β-1,4-galactosidase gave increased mobility by gel electrophoresis suggesting removal of sugar residues. Mass spectrometry analysis confirmed removal of

100% of the sialic acid residues at Asn1675 and Asn2181 following treatment with neuraminidase. Treatment with β-1,4-galactosidase removed approximately 69% of the galactose residues from Asn1675 and 100% of the galactose residues from Asn2181.

Removal of these sugars did not alter the ability of factor V to function as a cofactor in the generation of thrombin. Glycan removal also had no effect on factor V endocytosis by

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megakaryocytes. Conclusions: These studies indicate that the complex N-linked glycans at Asn1675 and Asn2181 are not important for factor V endocytosis by megakaryocytes and suggest that the unidentified factor V receptor is not a lectin.

Keywords: Blood Platelets, Endocytosis, Factor V, Glycosylation, Megakaryocytes.

3.2. Introduction

The prothrombinase complex, assembled on the activated platelet membrane at sites of vascular injury, is essential for physiological thrombin generation. It is comprised of a 1:1 stoichiometric complex of the serine protease factor Xa and the non-enzymatic cofactor factor Va assembled on an appropriate membrane surface in the presence of Ca2+

(Mann, Nesheim et al. 1990). Approximately 20-25% of the total pool of the procofactor, factor V (Breederveld, Giddings et al. 1975, Vicic, Lages et al. 1980), is stored in platelet

α-granules in a partially, proteolytically-activated state and is released at sites of vascular injury upon platelet activation leading to an increased localized concentration of active platelet-derived cofactor (Nesheim, Nichols et al. 1986). This is supported by several clinical observations that highlight the importance of the platelet-derived cofactor in normal hemostasis (Tracy, Giles et al. 1984, Nesheim, Nichols et al. 1986). Furthermore, studies have established that the platelet-derived pool is physically distinct (Monkovic and Tracy 1990, Kalafatis, Rand et al. 1994, Rand, Kalafatis et al. 1994, Camire,

Kalafatis et al. 1995, Conlon, Camire et al. 1997, Gould, Silveira et al. 2004, Gould,

Simioni et al. 2005, Wood 2008, Tracy, Jemings et al. 2013) and functions as a more

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procoagulant molecule (Lee and Mann 1989, Monkovic and Tracy 1990, Conlon, Camire et al. 1997, Camire, Kalafatis et al. 1998, Gould, Silveira et al. 2004) as compared with its plasma-derived counterpart.

Due to the predominance of the platelet-derived cofactor at sites of vascular injury, its structural and functional properties will have a distinct advantage over plasma- derived factor Va during thrombin generation and clot formation. Platelets acquire

(Bouchard, Meisler et al. 2008) and modify (Ayombil, Abdalla et al. 2013) factor V following endocytosis of the plasma procofactor by platelet precursor cells, megakaryocytes. Endocytosis is clathrin-dependent and mediated via a two receptor system consisting of low density lipoprotein receptor related protein-1 (LRP-1), a ubiquitous endocytic receptor, and an uncharacterized, specific factor V binding site

(Hughes 1999). Endocytosis is mediated by the factor V light chain (Bouchard, Abdalla et al. 2013) although the specific factor V residues and/or features involved are unknown.

More recently, Zappelli et al. suggested that a glycan binding protein, galectin-8, may function as the unidentified factor V receptor on the megakaryocyte cell surface

(Zappelli, van der Zwaan et al. 2012). In these studies, it was reported that inhibition of galectin-8 decreased factor V endocytosis by megakaryocytes. Other studies demonstrated a clear interaction between factor V and galectin-8 in vitro using mass spectrometry (Romaniuk, Tribulatti et al. 2010) and surface plasmon resonance (Zappelli, van der Zwaan et al. 2012). Several other platelet glycoproteins were found to interact

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with galectin-8 including von Willebrand factor, multimerin 1, and glycoprotein Ibα

(CD42) (Romaniuk, Tribulatti et al. 2010).

Galectin-8 has two carbohydrate recognition domains (CRD) (Hirabayashi,

Hashidate et al. 2002). The N-terminal CRD binds complex N-linked glycans containing a sialic acid (Neu5Ac) linked to a galactose (Gal) residue (Stowell, Arthur et al. 2010).

The C-terminal CRD binds to N-acetylglucosamine (GlcNAc) linked to a galactose residue (N-acetyllactosamine) in complex N-linked glycans (Stowell, Arthur et al. 2010).

The human factor V cDNA sequence predicts 37 potential N-linked glycosylation sites in human factor V (Jenny, Pittman et al. 1987). Nine are found in the heavy chain, three in the light chain (Rosing, Bakker et al. 1993, Hoekema, Nicolaes et al. 1997, Nicolaes,

Villoutreix et al. 1999), and the remaining 25 are in the B-domain of plasma-derived factor V. Of the three proposed N-linked glycan sites in the factor V light chain, one has been identified as a high mannose glycan on Asn1982 (Tracy, Jemings et al. 2013), while the other two were identified as being complex N-linked glycans on Asn1675 and

Asn2181 (Figure 15) (Tracy, Jemings et al. 2013). All N-linked glycans are comprised of a common biantennary core composed of two GlcNAc and three mannose (Man) residues to which additional sugar units are added (Kornfeld and Kornfeld 1985). In addition to the common core, the complex N-linked glycans on the factor V light chain contain two

GlcNAc residues, two Gal residues, and sometimes one Neu5Ac (Figure 15).

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Figure 15. N-linked glycans on the human, plasma-derived factor V light chain.

The light chain of factor V contains complex N-linked glycans at Asn1675 and 2181 as well as a high mannose-type glycan at Asn1982. In all cases, heterogeneous glycan species are found. Complex N-linked glycans always contain a common core of N- acetylglucosamine (GlcNAc, ■) and mannose (Man, ●) residues. GlcNAc residues are connected directly to the common core forming either bi- or tri-antennary structures. The external sugars consist of galactose (Gal, ●) residues, sometimes followed by a sialic acid residue (Neu5Ac, ♦). Asn2181 is also observed with no glycan present.

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Based on these data and the galectin-8 ligand preference (Jenny, Pittman et al. 1987,

Stowell, Arthur et al. 2010), it is possible that galectin-8 could interact with factor V via these glycans.

The goal of the current study was to determine if complex N-linked glycans are involved in factor V receptor endocytosis by megakaryocytes. These studies used exoglycosidase treatment in conjunction with mass spectrometry to selectively trim the complex N-linked glycans on factor V. These methods will determine if the complex N- linked glycans in the factor V light chain are important for endocytosis by megakaryocytes.

3.3. Materials and methods

Materials: CMK cells were a generous gift from Dr. Hava Avraham (The Division of

Experimental Medicine, Harvard Medical School, Boston, MA) and cultured as described previously (Bouchard, Williams et al. 2005). Human α-thrombin and factor Xa were purchased from Haematologic Technologies, Inc. (Essex Junction, VT, USA). Human prothrombin and factor V were purified from freshly frozen plasma as described (Bajaj and Mann 1973, Katzmann, Nesheim et al. 1981). PNGaseF, endoglycosidase H, α2-3,6,8 neuraminidase, β-N-acetylglucosaminidase and β-1,4-galactosidase were purchased from

New England BioLabs (Ipswich, MA, USA). Endoglycosidase F1, F2, and F3 were purchased from QA-Bio (San Mateo, CA, USA). Protease Inhibitor Cocktail and Surfact-

Amps® NP-40 were purchased from Thermo Scientific (Rockford, IL, USA). NuPage

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gels (4–12%) were obtained from Life Technologies (Carlsbad, CA, USA). Western

Lightening Plus enhanced chemiluminescence substrate was purchased from PerkinElmer

(Waltham, MA, USA). Horse α-mouse IgG-horseradish peroxidase was obtained from

Vector Laboratories, Inc. (Burlingame, CA, USA). Mouse anti-human FV #9 (α-FV#9) directed against the light chain (Foster, Tucker et al. 1983) and vesicles composed of

75% phosphatidylcholine and 25% phosphatidylserine (PCPS) were generously provided by Dr. Kenneth Mann (Department of Biochemistry, University of Vermont, Burlington,

VT, USA). Spectrozyme TH was purchased from American Diagnostica (Lexington,

MA, USA). Hirudin was purchased from Calbiochem (Darmstadt, Germany).

ProteaseMax surfactant and trypsin were from Promega Corporation (Madison, WI,

USA).

Enzymatic removal of Neu5Ac and Gal residues from factor V: Enzymatic deglycosylation of human, plasma-derived factor V (4 µM) was performed under non- denaturing conditions. The following glycosidases were used: PNGaseF (1.26 µM), endoglycosidase F1 (1.28 µM), endoglycosidase F2 (0.33 µM), endoglycosidase F3 (0.25

µM), endoglycosidase H (0.23 µM), neuraminidase (0.29 µM) and β-1,4-galactosidase

(0.38 µM). Enzymes were diluted in 20 mM HEPES, 0.15 M NaCl, pH 7.4 (HBS)

2+ containing 5 mM CaCl2 (HBS/Ca ) or enzyme buffer (20 mM Tris, 50 mM NaCl, 1 mM

EDTA). After incubation, aliquots were removed for endocytosis and activity assays and stored on ice. Gel samples were made by activating factor V (1 µM) with 2 U/mL α-

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thrombin for 10 min at 37°C. The reaction was quenched by dilution with sample preparation buffer (SPB) (62.5 mM Tris, pH 6.8, 0.07 M sodium dodecyl sulfate, 1.09 M glycerol, 0.15 mM bromophenol blue) (Laemmli 1970). Protein (2.5 μg) was resolved by

SDS-PAGE under reducing conditions (Laemmli 1970). The gel was stained using

Coomassie blue for 30 min followed by destaining in 18% methanol/9% acetic acid in water.

LC-MS/MS analysis of deglycosylated factor V: Mass spectrometry samples were prepared according to the directions included with ProteaseMAX Surfactant, trypsin enhancer. Briefly, Coomasie blue-stained proteins were extracted from the gel, washed with water, and destained twice with 50% methanol containing 25 mM NH4HCO3.

Samples were then dehydrated twice using 50% acetonitrile in 25 mM NH4HCO3 and dried in a Speed vacuum. Proteins were rehydrated in 25 mM dithiothreitol in 50 mM

NH4HCO3 and then incubated with 55 mM iodoacetamide in 25 mM NH4HCO3.

Washing, dehydration and drying steps were repeated prior to digestion using

ProteaseMAX and trypsin. LC-MS/MS analyses were performed as previously described

(Krudysz-Amblo, Jennings et al. 2010, Tracy, Jemings et al. 2013). Observed peak heights (Xcalibur software NL values, Thermo Scientific, Rockford, IL, USA) from the extracted ion currents of the glycopeptides mass to charge were used to generate relative abundance to assess the change in the glycan composition.

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Effect of glycosidase treatment on factor Va cofactor activity: Untreated and glycosidase-treated factor V (1 µM) were activated to factor Va (2 U/mL thrombin, 10 min, 37°C). Thrombin activity was quenched using a 1.5-fold molar excess of hirudin.

The prothrombinase complex was assembled by incubating dansylarginine N-3(ethyl-1,5- pentanediyl)amide (DAPA, 30 µM), and prothrombin (1.4 µM) with PCPS vesicles (20

μM) for 5 min in HBS containing 0.1% polyethylene glycol 8000 (PEG) and 2 mM CaCl2. Factor Va (20 nM) was added and the prothrombinase reaction was initiated with the addition of factor Xa (0.3 nM) (Haynes, Bouchard et al. 2012). At 15 s, 30 s, 45 s, 60 s, 75 s, 90 s, 2 min, and 5 min, the reaction mixtures were quenched with HBS containing 50 mM EDTA and 0.1% PEG-8000. Thrombin activity was assessed using the chromogenic substrate Spectrozyme TH (200 μM) as described previously (Haynes,

Bouchard et al. 2012).

Effect of glycosidase treatment on factor V endocytosis by megakaryocytes: CMK cells (1×106/mL) were cultured with glycosidase-treated factor V (30 nM) for 3 hr at

37°C. The cells were washed twice with HBS by centrifugation (260xg, 7 min, 25°C) to remove excess and cell surface bound protein. Cell viability was assessed by trypan blue exclusion before the final cell pellets were solubilized in lysis buffer (25 mM Tris, 0.15

M NaCl, 1% NP-40, pH 7.4) containing protease inhibitor cocktail and 5 mM EDTA at equivalent cell concentrations for 30 min on ice. The lysate was centrifuged (14,000xg,

32 min, 4°C) and the supernatant containing endocytosed factor V was incubated for 10

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min at 37°C with 2 U/mL α-thrombin to activate factor V to Va. The reaction was quenched by dilution with SPB. The factor Va present in the cell lysates was resolved by

SDS-PAGE under reducing conditions (Laemmli 1970), transferred to nitrocellulose (Towbin, Staehelin et al. 1979), and subsequently probed with anti-factor V

#9 using standard Western blotting techniques (Foster, Tucker et al. 1983). Factor Va

Western blot bands were quantified using Quantity One software (Bio-Rad Laboratories,

Hercules, CA). The density of the band was measured in intensity units per mm2 and the background for each lane was subtracted.

3.4. Results

Electrophoretic analyses of glycosidase-treated factor V.

Experiments were performed to define the appropriate conditions for deglycosylation of the light chain of native, human plasma-derived factor V. Initially, several endoglycosidases such as, PNGaseF, endoglycosidase H, and a cocktail of endoglycosidases F1, F2, F3 were incubated with factor V for 4, 8, 24, and 48 hr under native and denaturing conditions in an attempt to remove the glycan at the core; however, treatment with these enzymes did not lead to an increased electrophoretic mobility corresponding with removal of a large glycan under any of the conditions tested (data not shown). As a control, in gel deglycosylation was also attempted without success (data not shown). Therefore, specific exoglycosidases were employed to selectively trim the glycans. Neuraminidase was used to remove terminal Neu5Ac residues (Figure 15)

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(Schauer 1982), β-1,4-galactosidase to remove Gal residues (Figure 15) (Huber, Kurz et al. 1976). Attempts to remove GlcNAc (Wong-Madden and Landry 1995) residues were also made using β-N-acetylglucosaminidase (0.83 µM, 2, 4, or 12 hr incubation time) without success (data not shown). Following conversion to factor Va, deglycosylation was assessed by SDS-PAGE. The differential glycosylation on Asn2181 is known to cause the light chain to display as a doublet by SDS-page (Nicolaes, Villoutreix et al.

1999). When compared with a factor Va standard (Figure 16, lanes 1 and 2) collapse of the light chain doublet can be seen following treatment with neuraminidase (Figure 16, lane 3) or neuraminidase plus β-1,4-galactosidase (Figure 16, lane 4) suggesting that the glycans have been trimmed. Trimming of the glycans in the heavy chain was also observed as evident by an increased electrophoretic mobility.

Mass spectrometry analysis of factor V deglycosylation.

Deglycosylation was confirmed using LC-MS/MS analysis (Table 1). It is important to note that mass spectrometry analyses can determine the type of sugar residues present, but not the order in which they are connected. However, based on what is known about other complex N-linked glycans we can determine the residue order

(Kornfeld and Kornfeld 1985). The complex N-linked glycans on Asn1675 and Asn2181 of human, plasma-derived factor V consist of the common N-linked biantennary glycan core followed by two GlcNAc residues connected to two Gal residues, with one or two

Neu5Ac as the outermost residue (Figure 15 and Table 3, No enzyme). Neuraminidase

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Figure 16. Mobility of glycosidase-treated and untreated factor V on SDS-PAGE.

Factor V was treated with glycosidases and its electrophoretic mobility assessed following activation to factor Va by thrombin as described in Materials and Methods. Factor V was incubated for 2 hr (37°C) under the following conditions: HBS + 5 mM Ca2+ (lane 1);

Enzyme buffer (lane 2); Enzyme buffer + Neuraminidase (lane 3); Enzyme buffer +

Neuraminidase + β-1,4-galactosidase (lane 4). The factor V heavy chain (FV HC) and light chain (FV LC) are indicated on the left side of the gel. The data are representative of 4 experiments.

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Table 3. Glycan composition of glycosidase-treated factor V.

Relative Abundance‡ Glycopeptide Site Composition§ m/z (Da) No enzyme* Treated†

(Core)GlcNAc2 1266 3798 0.00 0.69

(Core)GlcNAc2Gal1 1306.5 3919.5 0.00 0.31

Asn1675 (Core)GlcNAc2Gal2 1347.5 4042.5 0.09 0.00 (Core)GlcNAc Gal 2 2 1493.9 4481.7 0.53 0.00 Neu5Ac1 (Core)GlcNAc Gal 2 2 1420.8 4262.4 0.38 0.00 Neu5Ac2

(Core)GlcNAc2 1194 2388 0.00 1.00

(Core)GlcNAc2Gal1 1274.7 2549.4 0.00 0.00

Asn2181 (Core)GlcNAc2Gal2 1356.6 2713.2 0.00 0.00 (Core)GlcNAc Gal 2 2 1098.4 3295.2 0.37 0.00 Neu5Ac1 (Core)GlcNAc Gal 2 2 1001.4 3004.2 0.63 0.00 Neu5Ac2 ______

Core, Common N-linked glycan core; GlcNAc, N-acetyl glucosamine; Gal, galactose; Neu5Ac, Sialic acid. §Glycan composition was determined using tandem mass spectrometry. ‡The relative abundance of each glycan species was determined using Xcalibur NL values. *Native factor V incubated in HBS containing 5 mM Ca2+. †Native factor V incubated in enzyme buffer with neuraminidase and β-1,4-galactosidase as described in the Materials and Methods.

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treatment quantitatively removed the Neu5Ac residues on both Asn1675 and Asn2181

(Table 1, Treated). Treatment with β-1,4-galactosidase removed 69% of the Gal residues from Asn1675 and 100% of the Gal residues at Asn2181 (Table 1, Treated). The remaining glycan species on Asn1675 consist of the common core with two additional

GlcNAc residues; in some cases one Gal residue is present. In the case of Asn2181, all the Neu5Ac and Gal residues were removed leaving only two GlcNAc residues in addition to the common core.

Glycosidase treatment has no effect on factor Va cofactor activity.

The cofactor activity of deglycosylated factor Va was assessed via the prothrombinase complex assembled on PCPS vesicles. For these experiments, factor V was deglycosylated as described above and activated to factor Va. No difference could be seen between the rate of thrombin generation using glycosidase-treated factor Va and untreated factor Va suggesting that glycosidase treatment does not affect prothrombinase assembly, or the ability of factor Va to participate as a cofactor in thrombin generation

(Figure 17).

Endocytosis of glycosidase-treated factor V by CMKs.

Glycosidase-treated factor V was cultured with a megakaryocyte like cell line,

CMK, to determine if factor V glycans play a role in its endocytosis by megakaryocytes.

Following endocytosis, the cells were lysed and treated with thrombin to activate the

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800 700 600 500 400 nM IIa nM 300 200 100 0 0 10 20 30 40 50 60 70 Time(s)

Figure 17. Time course of factor Va activity after glycosidase treatment.

The ability of glycosidase-treated factor Va to participate in prothrombinase complex assembly and function on PCPS vesicles was measured as described in Materials and

Methods. Factor V treatments include: HBS (●), enzyme buffer (▲), enzyme buffer and neuraminidase (■), enzyme buffer with neuraminidase and β-1,4-galactosidase (○), and untreated factor V (×). The dotted line is fitted to the enzyme buffer with neuraminidase and β-1,4-galactosidase (○) time course. These data are representative of 4 experiments.

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factor V and its partial activation products (Ayombil, Abdalla et al. 2013) to factor Va, and the light chain visualized by Western blotting (Figure 18). Glycosidase treatment appeared to have no effect on factor V endocytosis (lane 3) by megakaryocytes as compared to untreated factor V (lane 4) or factor V that was incubated in the absence of enzyme (lanes 1 and 2) as similar amounts of factor V light chain were detected. The relative densities of the factor Va light chain in each sample were quantified and were not substantially different (Table 4).

3.5. Discussion

The goal of this study was to determine if complex N-linked glycans play a role in factor V endocytosis. Unsuccessful attempts were made to remove this type of glycan from the factor V light chain using endoglycosidases. Instead, exoglycosidases were used to selectively trim the complex N-linked glycans. Treatment with glycosidases did not alter the ability of factor Va to participate as a cofactor in the prothrombinase complex.

Previous studies assessing the role of glycans in factor Va cofactor activity have given contrasting results (Gumprecht and Colman 1975, Elder and Alexander 1982, Bruin,

Sturk et al. 1987, Ortel, Devore-Carter et al. 1992, Rosing, Bakker et al. 1993, Ortel,

Quinn-Allen et al. 1994, Varadi, Rosing et al. 1996, Hoekema, Nicolaes et al. 1997,

Nicolaes, Villoutreix et al. 1999). Differential glycosylation at Asn2181 leads to the appearance of a light chain doublet by gel electrophoresis (Ortel, Devore-Carter et al.

1992, Rosing, Bakker et al. 1993, Ortel, Quinn-Allen et al. 1994, Nicolaes, Villoutreix et al. 1999). The deglycosylated form has been shown to have decreased procoagulant

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Figure 18. Endocytosis of glycosidase-treated factor V by CMKs.

Treated factor V was cultured with CMKs and lysates were activated using thrombin.

Endocytosis was assessed by Western blotting using α-FV#9, a monoclonal antibody directed against the factor V light chain (N=2). All samples were normalized to cell number prior to lysis. Lane 1: HBS + 5 mM Ca2+, lane 2: Enzyme Buffer, lane 3: Enzyme Buffer

+ Neuraminidase + β-1,4-galactosidase, lane 4: Untreated Control, lane 5: No factor V.

The factor V light chain (FV LC) is indicated on the left side of the gel.

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Table 4. Densitometric analysis of the factor Va light chain in CMK lysates following endocytosis.

Density

(Intensity/mm2)*

HBS + 5 mM CaCl2 8848

Enzyme buffer 10571

Enzyme buffer + Neuraminidase+ 12300 β-1,4-galactosidase

Untreated 9583

* Endocytosis was visualized using Western blotting and quantified using densitometry as described in the Materials and Methods. All samples were normalized to cell number prior to lysis.

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activity (Rosing, Bakker et al. 1993, Varadi, Rosing et al. 1996, Hoekema, Nicolaes et al.

1997) through reduced membrane binding when compared with the glycosylated form.

Similarly, removal of the outermost Neu5Ac residues with neuraminidase leads to a 1.5 to 2-fold increase in factor V activity as measured by a clotting assay using bovine

(Gumprecht and Colman 1975) and human (Bruin, Sturk et al. 1987) factor V. In contrast, incubation of human factor V with endoglycosidase F, which hydrolyses the bond between GlcNAc residues in the core giving release to the majority of the glycan

(Elder and Alexander 1982), demonstrated increased electrophoretic mobility, but had no effect on factor V clotting activity. At the time these results were published, purification of glycosidases was rudimentary with a large amount of contamination. It is likely that some of these results may be due to contaminating enzymes.

Frequently glycoproteins can interact with lectin receptors on the cell surface

(Rice and Lee 1993, Sheikh, Yarwood et al. 2000, Szolnoky, Bata-Csorgo et al. 2001,

Weigel and Yik 2002). One example of this is the asialoglycoprotein receptor (Weigel and Yik 2002). Upon removal of external Neu5Ac residues the internal Gal residues are exposed signaling the need for clearance by the asialoglycoprotein receptor in the liver

(Morell, Gregoriadis et al. 1971, Yu and Gan 1977, Park, Manzella et al. 2003, Grewal,

Uchiyama et al. 2008, Steirer, Park et al. 2009). Through his mechanism, desialyation of circulating platelets (Greenberg, Packham et al. 1975, Steiner and Vancura 1985,

Sorensen, Hoffmeister et al. 2008, Sorensen, Rumjantseva et al. 2009), erythrocytes

(Bratosin, Mazurier et al. 1995) and glycoproteins (Van Baelen and Mannaerts 1974,

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Weigel and Yik 2002, Park, Manzella et al. 2003, Bovenschen, Rijken et al. 2005, Steirer,

Park et al. 2009) regulates their clearance (Van Den Hamer, Morell et al. 1970, Morell,

Gregoriadis et al. 1971, Zijderhand-Bleekemolen, Schwartz et al. 1987, Ii, Kurata et al.

1990, Monroe and Huber 1994, Mu, Fallon et al. 1994) by the liver (Morell, Gregoriadis et al. 1971, Yu and Gan 1977, Hubbard, Wilson et al. 1979, Weigel and Oka 1983, Park,

Manzella et al. 2003, Grewal, Uchiyama et al. 2008, Steirer, Park et al. 2009). A second example is the HIV-1 envelope glycoprotein 120 complex which initiates virus entry by binding to extracellular lectin receptors on target cells (de Witte, Bobardt et al. 2007).

Another important lectin in human pathology is influenza hemagglutinin, which initiates influenza virus entry by binding to cells with Neu5Ac on the membranes (Russell, Kerry et al. 2008). With the precedent of lectin-based endocytosis, studies were performed to determine if the complex N-linked glycans at Asn1675 and Asn2181 on the factor V light chain are involved in its endocytosis by megakaryocytes. Removal of the external

Neu5Ac and Gal residues did not alter megakaryocyte endocytosis, suggesting that

Asn2181 and Asn1675 do not play a role in endocytosis of factor V at the cell surface. In addition, both the glycosylated and unglycosylated forms of Asn2181 are found within the platelet (Hayward, Furmaniak-Kazmierczak et al. 1995), suggesting that factor V is endocytosed by the megakaryocyte regardless of the glycosylation status of Asn2181.

A search of the megakaryocyte transcriptome (A. Weyrich, unpublished data) indicates that megakaryocytes do not possess asialylglycoprotein receptor mRNA. They do contain mRNA for the mannose-binding lectin which will bind to high mannose

120

glycans (Fraser, Koziel et al. 1998). Proteins from the sialic acid binding protein family which bind to external sialic acid residues were also found. Additionally, no receptors capable of binding terminal Gal or GlcNAc residues, as found in glycosidase treated factor V, could be identified in the transcriptome. It can be hypothesized that no glycan binding proteins exist on the megakaryocyte cell surface that are capable of endocytosing glycosidase-treated factor V via complex N-linked glycans in the light chain. Thus, endocytosis of treated factor V must be via the same mechanism as untreated factor V.

It was previously suggested that galectin-8 may serve as an extracellular factor V receptor (Zappelli, van der Zwaan et al. 2012). However, as shown in Appendix II, galectin-8 is not found on the megakaryocyte cell surface. Additionally, blocking galectin-8 did not decrease factor V endocytosis (Appendix II). Removal of all the β- galactosides, the galectin-8 ligand (Carlsson, Oberg et al. 2007), from factor V using exoglycosidases did not have an effect of factor V endocytosis. Therefore, while it may play an intracellular role in factor V endocytosis it cannot function as a factor V receptor.

Additionally, through the glycan trimming used here, the sugar recognition sequence of any lectin would be destroyed. However, based on the siRNA it is possible that galectin-

8 may play an intracellular role in factor V endocytosis or trafficking.

3.6. Authorship details

J.M. Gertz was responsible for the experimental design; data acquisition, analyses and interpretation; and preparation of the manuscript. M.E. Jennings acquired and analyzed

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mass spectrometry data. C. Couperus performed prothrombinase assays. J.R. Silveira assisted with preparation of mass spectrometry samples. B.A. Bouchard conceived of the research; provided general guidance on experimental design and data acquisition, analyses and interpretation; and prepared the manuscript.

3.7. Acknowledgements

The authors would like to thank Andrew S. Weyrich, PhD (University of Utah) for providing access to the unpublished megakaryocyte transcriptome and Laura M. Haynes,

PhD (University of Vermont) for helpful discussions. This work was supported by grants from the NIH (K02 HL91111) (BB), (PHS P01 HL46703) (MJ) and T32 HL007894 (JG), a grant from the American Heart Association (SDG 0635048N) (BB), and the University of Vermont College of Medicine.

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Perspectives and Future Directions

Factor Va functions as the essential cofactor for the prothrombinase complex consisting of factor Xa, calcium, and a phospholipid membrane surface, such as the activated platelet, to convert the zymogen prothrombin to the active form, thrombin (Mann,

Nesheim et al. 1990). Once formed, thrombin cleaves fibrinogen to form a stable fibrin clot and modulates its own formation through activation of platelets as well as coagulation factors V, VIII, XI and the potent anticoagulant, activated protein C. In addition to these roles in coagulation, thrombin activates cellular protease-activated receptors (PARs)

(Coughlin 2000), leading to mitogenic and chemotactic properties and participation in cellular growth, brain development, and inflammatory processes (Narayanan 1999).

Thrombin generation can occur through cleavage of prothrombin by factor Xa alone; however, assembly of the prothrombinase complex is 300,000 times more efficient

(Nesheim, Taswell et al. 1979), highlighting the importance of factor Va in this process.

This is further confirmed through clinical observations that deficiencies of factor Va can lead to severe bleeding in the affected individual (Camire 2011). In contrast, thrombosis can arise through inefficient inactivation of the cofactor by activated protein C, as is seen with factor VaLeiden (Dahlback, Hillarp et al. 1996, Dahlback, Zoller et al. 1996, Hillarp,

Zoller et al. 1996, Kujovich 2011).

The procofactor, factor V, exists in two pools found within the whole blood: 75-

80% circulates in the plasma as a single chain procofactor while the other 20-25% is stored within the platelet α-granule as a partially proteolytically activated form that is released

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upon platelet activation at the site of vascular injury (Tracy, Eide et al. 1982). Platelet- derived factor V/Va is acquired via endocytosis of the plasma derived procofactor by the platelet precursor cell, the megakaryocyte. Endocytosis is mediated by a two-receptor system including an uncharacterized factor V receptor, and low density lipoprotein receptor related protein-1 (LRP-1). Upon binding of factor V to its receptor, LRP-1 initiates endocytosis in a clathrin- and calcium-dependent manner (Bouchard, Williams et al. 2005,

Bouchard, Meisler et al. 2008). Studies are being performed to narrow the list of candidate factor V receptors. It was previously demonstrated that the light chain of factor V is important for endocytosis (Bouchard, Abdalla et al. 2013). Located within this region are three N-linked glycans, two complex type at Asn1675 and 2181, and one high mannose type at Asn1892 (Jenny, Pittman et al. 1987, Tracy, Jemings et al. 2013). In this dissertation, the complex N-linked glycans at Asn1675 and Asn2181 in the factor V light chain were assessed for a potential role in factor V endocytosis by megakaryocytes

(Chapter 3). A search of the megakaryocyte transcriptome identified one lectin family, the galectins, that could participate in factor V endocytosis by interacting with the complex N- linked glycans in the factor Va light chain. Through the use of exoglycosidases, the glycans were trimmed to a level that would destroy the binding ability of all galectin family members. In addition, no lectins capable of binding the complex N-linked glycans in glycosidase treated factor V were identified in the megakaryocyte transcriptome. Trimming of the complex N-linked glycans in the factor V light chain did not affect factor V endocytosis or thrombin generation, suggesting that these processes are not dependent on

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these glycans. In other studies using similar techniques it was shown that the high mannose glycan at Asn1982 does not play a role in factor V endocytosis or thrombin generation

(Ayombil and Tracy, unpublished data). The high mannose binding lectin, ER-Golgi intermediate compartment 53 kDa protein (ERGIC-53)(Arar, Carpentier et al. 1995, Itin,

Roche et al. 1996), was identified in the megakaryocyte transcriptome, and is known to play a role in factor V trafficking in hepatocytes (Segal, Zivelin et al. 2004); however, its role in megakaryocytes was not assessed. It is possible that the glycans at Asp1675, 1982, and 2181 play a role in factor V trafficking to the α-granule. This could be confirmed through co-distribution of glycosidase-treated and untreated factor V with markers of the endoplasmic reticulum, Golgi apparatus, lysosome and α-granule in order to determine if there is any alteration in the trafficking pathway.

The other major objective of this dissertation was to establish and characterize a primary cell model of megakaryocyte differentiation using umbilical cord blood-derived

CD34+ cells by which to study factor V endocytosis through platelet production. While umbilical cord blood-derived cells have been used to study megakaryocyte production of platelets (Denis, Tolley et al. 2005, Foulks, Marathe et al. 2009, Shi, Smith et al. 2014), no study has thoroughly characterized megakaryocyte development using this model.

Experiments to characterize megakaryocyte development were performed on cells that had been cultured for 6-12 days, and the results demonstrated that factor V endocytosis and expression of the early megakaryocyte differentiation marker, CD41 (Zimmet and Ravid

2000), did not change during this time. In contrast, expression of the mature megakaryocyte

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marker (Zimmet and Ravid 2000), CD42, increased nearly two-fold during this time period.

The number of cells was not sufficient for analysis prior to day 6 of culture; therefore, nothing is known about the process before this time. It is likely that factor V endocytosis and CD41 expression increase from days 0 to 5. It is important to note that cells are isolated based on expression of the hematopoietic stem cell marker, CD34 (Long, Williams et al.

1982, Ogawa 1993), with a high level of purity. At the time of isolation some of these cells may also express megakaryocyte specific markers. If further synchronization of the culture was required, cell sorting could be utilized to select the population that is CD34+ and devoid of any myeloid or lymphoid lineage markers to yield an immature hematopoietic stem cell population. Alternatively, proplatelet production and expression of the mature megakaryocyte marker, CD42, occurs in approximately 20% of the cells in culture. Cell sorting could be utilized to select the population that is CD42+, yielding a pure mature megakaryocyte culture that will likely lead to nearly 100% proplatelet formation.

While factor V was visualized in proplatelet extensions, localization to the α- granule has not been confirmed using this cell model. Congruent endocytosis with fluorescently-labeled fibrinogen could provide evidence for this; however, as fibrinogen is also endocytosed and trafficked through the cell (Broekman, Handin et al. 1975,

Handagama, Shuman et al. 1989, Handagama, Rappolee et al. 1990, Handagama,

Scarborough et al. 1993), it would likely be detected in other vesicles and organelles, such as the endosomes and Golgi apparatus. In contrast to factor V, endocytosed fibrinogen is not modified by the megakaryocyte and would not be expected to localize to the

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endoplasmic reticulum. A second α-granule marker, P-selectin (Wagner 1993), could be used, but because it is synthesized by the megakaryocyte (Schick, Konkle et al. 1993) it is expected that staining would also be visualized in the endoplasmic reticulum, the Golgi apparatus, and cargo vesicles. Dual co-distribution with both of these α-granule markers would provide improved confidence. Unequivocal localization of factor V to the α-granule could be achieved using electron microscopy.

Live cell imaging techniques were also developed in order to observe the process of proplatelet formation in human ex vivo-derived megakaryocytes. Attempts were made to visualize factor V trafficking in the proplatelet extension using this technique, but were unsuccessful due to photobleaching of the fluorophore conjugated to factor V. In order to remedy this problem, factor V could be conjugated to quantum dots, which are extremely bright and resistant to photo bleaching (Ness, Akhtar et al. 2003, Wu, Liu et al. 2003,

Alivisatos, Gu et al. 2005), even over the 24 hours required to visualize proplatelet formation in its entirety. Only ~20% of the cells in culture will generate proplatelet extensions and nearly all of these cells express the mature megakaryocyte marker, CD42.

Cell sorting could be used to produce a pure mature megakaryocyte population, with nearly

100% proplatelet formation, increasing the chances of successfully filming this process.

Identification of the cellular and molecular mechanisms by which factor V is endocytosed by megakaryocytes would also be useful to the hemophilia A community.

This disorder is caused by a deficiency in factor VIII, and traditionally protein replacement therapy is used to treat severe bleeding episodes but it has been confounded by the

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formation of inhibitory antibodies to transfused human FVIII in 30% of patients (Viel,

Ameri et al. 2009, Hay, Palmer et al. 2011). As activated platelets mediate the primary response to vascular injury by adhering to a wound site and secreting biologically active proteins (Nurden and Caen 1975), storage of FVIII within platelets may be an ideal strategy for providing a continuous, locally inducible treatment for maintaining hemostasis for hemophilia A (Ortel, Devore-Carter et al. 1992, Shi, Wilcox et al. 2003, Yarovoi, Nurden et al. 2005, Shi, Wilcox et al. 2006, Shi, Wilcox et al. 2007, Gewirtz, Thornton et al. 2008,

Neyman, Gewirtz et al. 2008, Doering, Denning et al. 2009, Greene, Lambert et al. 2011,

Siner, Iacobelli et al. 2013, Greene, Lyde et al. 2014). As factor VIII demonstrates a high level of structural homology to factor V (Jenny, Pittman et al. 1987), the knowledge gained about generation of the platelet-derived factor V/Va pool could be applied to produce a novel platelet-derived factor VIII pool. Megakaryocyte endocytosis and trafficking of factor VIII to the platelet α-granule could create designer platelets containing a protected pool that is released only at the site of vascular injury. Designer platelets could be infused into hemophilia A patients instead of purified factor VIII, leading to prolonged hemostatic competence.

In conclusion, while the studies presented in this dissertation have provided much insight into the mechanisms involved in megakaryocytes endocytosis of factor V, there are many unanswered questions remaining. Much work needs to be done to identify the specific factor V receptor present on the megakaryocyte cell surface and to determine signaling mechanisms by which factor V is trafficked to the α-granule. The studies and

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techniques presented in this dissertation provide the groundwork for such future research endeavors.

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Appendix I. Simultaneous flow cytometric analysis of megakaryocyte ploidy and a

labile intracellular protein using zinc-based fixation

This work was submitted for publication, in part, to Cytometry Part A.

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Simultaneous flow cytometric analysis of megakaryocyte ploidy and a labile

intracellular protein using zinc-based fixation1

J.M. Gertz, M. Meuser, and B.A. Bouchard

Department of Biochemistry, University of Vermont, Burlington, VT

Running headline: DNA and protein analysis in megakaryocytes.

Address correspondence to: Beth A. Bouchard, Ph.D.

Department of Biochemistry

University of Vermont College of Medicine

89 Beaumont Avenue, Given C440

Burlington, VT 05405- 0068, USA

Tel.: +1 (802)656-6529; Fax: +1 (802)656-8220

E-mail: [email protected]

1 This research was supported by grants from the NIH (K02 HL91111) (BB) and T32

HL007894 (JG), a grant from the American Heart Association (SDG 0635048N) (BB), and the University of Vermont College of Medicine.

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Summary

Megakaryocytes undergo a unique endomitotic cell cycle leading to large polyploidal cells.

Simultaneous assessment of DNA content and cellular markers would be a useful tool by which to study megakaryocyte development. Traditional fixatives, paraformaldehyde and methanol, only allows for analysis of one of these parameters at a time. Aldehyde-based fixatives preserve extracellular epitopes as well as labile endocytosed proteins. However, the cross-linking mechanisms of these fixatives is known to fragment DNA and pervert intercalation of nucleic acid stains between the bases. In contrast, methanol fixation gives excellent preservation of the DNA but alters the cellular morphology of proteins, destroying the antibody epitope. Recently, zinc salt-based fixatives (ZBF) have been developed as a non-toxic, non-crosslinking method of cell fixation for flow cytometry that may be appropriate for the study of megakaryocyte development. ZBF preserves extracellular and intracellular epitopes and forward scatter/side scatter parameters as well as low concentrations of aldehyde-based fixatives, and was shown to be adequate for simultaneous analyses of extracellular epitopes and DNA content (Cytometry A 77(8):798-

804, 2010). The current study demonstrates that analysis of ZBF cells yields preservation of proteins similar to paraformaldehyde fixation, and simultaneously preserves DNA content in a manner similar to methanol fixation. This is highlighted by experiments in which polyploidal megakaryocytes were analyzed for both endocytosis of a labile protein and DNA content. As megakaryocytes naturally become polyploidal, this could be a valuable tool in correlating megakaryocyte function with DNA content. Together these

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results indicate that ZBF is an appropriate fixation method for simultaneous analyses of a labile intracellular epitope and DNA content in megakaryocytes by flow cytometry.

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Introduction

Megakaryocytes, derived from hematopoietic stem cells found in the bone marrow

(Long, Williams et al. 1982, Ogawa 1993), are responsible for production of all circulating platelets (Wright 1906). Initially they undergo a traditional cell cycle (Zimmet and Ravid

2000) in which they begin with 2N DNA content, duplicate the DNA to 4N, and then perform cytokinesis, yielding two identical cells. Upon completion of this proliferation stage the megakaryocyte will transition to an endomitotic cell cycle in which they continue to duplicate the DNA but do not undergo cytokinesis leading to a polyploidal phenotype

(Nagata, Muro et al. 1997, Vitrat, Cohen-Solal et al. 1998). Later, the mature polyploidal megakaryocyte will cease DNA synthesis, undergo cytoplasmic maturation, and fragment into functional circulating, blood platelets (Long, Williams et al. 1982, Zimmet and Ravid

2000, Machlus and Italiano 2013). Platelets are small anucleate cell fragments that circulate within the blood (Osler 1874, Tocantins 1938). Upon vascular injury, they adhere and aggregate at the site to form a platelet plug (Brass 2003), and provide an appropriate membrane surface for clot formation (Mann, Nesheim et al. 1990).

The aim of this study was to develop a simple, non-toxic protocol that would allow for simultaneous flow cytometric analysis of polyploidal DNA content in conjunction with a labile intracellular protein and an extracellular epitope. The ability to analyze intra and extracellular proteins in conjunction with DNA content in megakaryocytes will be useful for understand megakaryocyte biology and the development of megakaryocyte and platelet focused treatments.

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Materials and methods

Materials: CMK cells were a generous gift from Dr. Hava Avraham (The Division of

Experimental Medicine, Harvard Medical School, Boston, MA). Human factor V, purified from freshly frozen plasma as described (Katzmann, Nesheim et al. 1981), and human factor IX (Haematologic Technologies, Inc., Essex Junction, VT) were conjugated to

AlexaFluor488 using the manufacturer’s (Life Technologies, Grand Island, NY) instructions (Bouchard, Williams et al. 2005, Bouchard, Meisler et al. 2008). Anti-CD61 conjugated to FITC was purchased from BioLegend (San Diego, CA) and an isotype- matched non-immune antibody conjugated to FITC (NI-FITC) was purchased from Becton

Dickenson and Company (Franklin Lakes, NJ). Propidium iodide and RNase A were purchased from Sigma-Aldrich (St. Louis, MO). Phorbol 12-myristate-13-acetate (PMA) was a generous gift from Dr. Benjamin Surratt (University of Vermont College of

Medicine, Burlington, VT). Superfrost Plus Micro Slides were purchased from VWR

(Radnor, PA).

Cell culture: CMK cells were cultured as described previously (Taniguchi, London et al.

1999). For some experiments, CMK cells (2.5 x 105 cells/mL) were cultured for eleven days in the presence of 10 nM PMA or DMSO (Komatsu, Suda et al. 1989) on non-treated, tissue culture wells. On days 2, 5, 7, and 9 fresh media containing PMA or DMSO was added in a volume equivalent to the original culture volume. Prior to each experiment, cell viability was assessed by trypan blue exclusion.

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Fixation of cells: Cells were washed three times using HBS and resuspending at 1 x 106 cells/mL.

Paraformaldehyde (PFA)-fixation: Washed cells were incubated with three volumes of 1 or 4% PFA (30 min, ambient temperature).

Methanol fixation: Three volumes of chilled 100% methanol were added to washed cells while vortexing. The cells were subsequently incubated at 4°C for up to 6 hr.

Zinc-based fixation (ZBF)-fixation: Two volumes of ZBF-buffer (0.1 M Tris-HCl, pH 7.8,

0.05% calcium acetate, 0.5% zinc acetate, 0.5% zinc chloride) were added while vortexing to washed cells. The cells were subsequently incubated at 4°C 16-48 hr.

All cells subjected to fixation were washed three times with HBS to remove the fixative.

Factor V endocytosis: CMK cells were washed twice by centrifugation (250xg, 7 min) followed by resuspension in serum-free media containing 0.1% endotoxin-free bovine serum (Bouchard, Williams et al. 2005, Bouchard, Meisler et al. 2008) and incubated with

30 nM AlexaFluor488-conjugated factor V (3 hr, 37°C). Control reactions were incubated with 30 nM AlexaFluor488-conjugated factor IX (Bouchard, Williams et al. 2005,

Bouchard, Meisler et al. 2008). Cells were subsequently washed by centrifugation (250xg,

7 min) followed by resuspension in 20 mM Hepes, 0.15 M NaCl, pH 7.4 (HBS), and subjected to fixation as described above in “Fixation of cells”.

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Cell surface immunostaining: Cells were washed by centrifugation (250xg, 7 min) followed by resuspension in HBS. Cells (2.5 x 106 cells/mL, 100 µL) were incubated (1 hr, ambient temperature) with anti-CD61-FITC (4 µg/mL) in 1% BSA. Control reactions contained NI-FITC. Cells were washed as described above to remove unbound antibody and subjected to fixation by one of the three methods describe above.

Assessment of DNA ploidy: Fixed cells were resuspended in 200 μL of a solution containing 10 U/mL RNase A, 0.05% saponin, 0.01 mg/mL propidium iodide, 2 mM

MgCl2, and phosphate-buffered saline (0.15 M sodium chloride, 4 mM sodium phosphate monobasic monohydrate, 6 mM sodium phosphate dibasic heptahydrate, pH 7.4) (PBS).

Samples were also prepared in the absence of propidium iodide.

Flow cytometric analyses: Cells were analyzed for DNA content and either CD61-FITC or endocytosed AlexaFluor488-factor V. Fluorescence from 10,000 cells was analyzed on a BD LSRII using a 488 laser with a 530/30 emission filter. Cells were analyzed using a

150-200 μm diameter nozzle. Cells were selected based on forward scatter and side scatter, and a singlet population was selected based on the area and width of the propidium iodide signal. All data were collected as binary FCS files and analyzed using FlowJo version 10 data analysis software (FlowJo, LLC, Ashland, OR).

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Fluorescence microscopy: Following flow cytometry analysis, cells were adhered to microscopy slides through cytocentrifugation and visualized using an Olympus BX50

Upright Fluorescence Microscope. The propidium iodide was excited with a krypton/argon laser. The individual fluorescent images were acquired using Q-Capture pro image analysis software (Q-imaging, Surrey, BC, Canada).

Results

Paraformaldehyde and methanol fixation will not allow for simultaneous analysis of a labile endocytosed protein and DNA ploidy.

A megakaryocyte-like cell line, CMK, was used to determine an appropriate protocol to assess DNA ploidy in conjunction with expression of a labile endocytosed protein and a cell surface glycoprotein by flow cytometry. Most flow cytometry protocols utilize formaldehyde-based crosslinking fixatives for flow cytometric analysis; however, these fixatives are known to fragment DNA and prevent intercalation of nucleic acid stains between the bases (Clevenger, Bauer et al. 1985, Gillio-Tos, De Marco et al. 2007, Jensen,

Owens et al. 2010) in addition to being toxic (National Toxicology 2011). Precipitating fixatives (e.g. ethanol, methanol) are also widely used despite disruption of protein structure and antibody epitopes (Mann, Dyne et al. 1987, Morkve and Hostmark 1991).

ZBF has been developed as a non-toxic alternative to crosslinking and precipitating fixatives (Wester, Asplund et al. 2003, Lykidis, Van Noorden et al. 2007, Jensen, Owens et al. 2010). It has been shown to have improved structure preservation of RNA, DNA and

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proteins while maintaining cellular morphology (Wester, Asplund et al. 2003) and enzymatic activity (Hadler-Olsen, Kanapathippillai et al. 2010) when compared with crosslinking fixatives allowing for DNA, cell cycle, and immunophenotypic analysis using flow cytometry. Initially, the scatter properties (forward scatter or size and side scatter or granularity) of the cells following fixation with 1% PFA, 4% PFA, or methanol were compared (Figure 19A, B, and C, respectively). Small changes in CMK cell forward and side scatter were observed between the different fixatives and as compared to unfixed cells

(data not shown). Fixation with increasing concentrations of PFA leads to decreased side scatter and forward scatter, while fixation with methanol appears to slightly increase the side scatter or granularity of the cells. In contrast to these results, only 4% PFA fixation led to preservation of endocytosed AlexaFluor488-labeled factor V (Figure 19D, E and F) as evidenced by a substantial increase in the fluorescent signal (Figure 19E, black histogram) as compared to cells incubated in the presence of AlexaFluor488-labeled factor

IX (Figure 19E, shaded histogram). Little or no fluorescence was observed following fixation with 1% PFA or methanol. Immunostaining of a megakaryocyte-specific, cell surface, integral membrane protein, CD61, was similar for all three fixatives (Figure 19G,

H, and I, black histogram) although the greatest increase in fluorescence as compared to cells stained with NI-FITC (shaded histograms) was observed in cells subjected to 4% fixation (see Figure 19H).

Substantially different results were obtained when the cells were analyzed for

DNA content using propidium iodide. When the cells were subjected to fixation with 1%

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Figure 19. Standard protocols are not appropriate to assess factor V endocytosis in conjunction with DNA content.

Forward and side scatter (A, B, C), endocytosis of AlexaFluor488-factor V (D, E, F; black histogram), anti-CD61-FITC immunostaining (G, H, I; black histogram) and DNA ploidy

(J, K, L) were assessed by flow cytometry following fixation of CMK cells with 1% PFA

(A, D, G, J), 4% PFA (B, E, H, K), or methanol (C, F, I, L). The grey histograms represent

CMK cells incubated with AlexaFluo488-factor IX (D, E, F) or immunostained with NI-

FITC (G, H, I). The 2N and 4N populations are indicated with arrows (J, L).

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PFA (Figure 19J) or methanol (Figure 19L), two distinct peaks representing the 2N and 4N populations were observed. These low ploidy numbers are not surprising as CMK cells must be treated with cytokines or phorbol diesters such as PMA to induce further differentiation along the megakaryocyte lineage (Ogura, Morishima et al. 1988, Fugman,

Witte et al. 1990, Nagano, Kishimoto et al. 1993) even though they closely recapitulate megakaryocyte biology. In marked contrast, 4% PFA fixation did not allow for assessment of DNA content (Figure 19K).

Factor V endocytosis and DNA content can be assessed simultaneously by flow cytometry in polyploidal megakaryocytes using zinc-based fixation

Previous studies have demonstrated that ZBF allows for the simultaneous analysis of DNA and intracellular and cell surface proteins by flow cytometry (Jensen, Owens et al.

2010, Zhao, Li et al. 2011). The forward and side scatter of CMK cells subjected to ZBF fixation (Figure 20A) were also similar to unfixed cells (data not shown). Side scatter was most similar to cells subjected to fixation with 1% PFA (see Figure 19A), while the forward scatter was most like that of cells subjected to fixation with 4% PFA (see Figure 19B). In addition, ZBF preserved endocytosed AlexaFluor488-labeled FV (Figure 20B), as well

CD61 immunostaining (Figure 20C), and distinct 2N and 4N populations were observed following propidium iodide staining (Figure 20D). Both AlexaFluor488-factor V endocytosis (Figure 20E) and CD61 expression (Figure 20F) were slightly increased in cells with a 4N DNA content.

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Figure 20. ZBF will allow simultaneous analysis of DNA content, an integral membrane protein and/or a labile intracellular protein.

(A) Forward and side scatter, (B) endocytosis of AlexaFluor488-factor V (black histogram), (C) anti-CD61-FITC immunostaining (black histogram) and (D) DNA ploidy were assessed by flow cytometry following ZBF of CMK cells. AlexaFluor488-factor V endocytosis (E) and CD61 expression (F) are also displayed as a function of DNA content.

The grey histograms represent CMK cells incubated with AlexaFluo488-factor IX (B) or immunostained with NI-FITC (C). The 2N and 4N populations are indicated with arrows

(D, E, F).

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159

CMK cells were stimulated with PMA for nine days to induce megakaryocyte differentiation and DNA polyploidization (Komatsu, Suda et al. 1989), and factor V endocytosis and DNA ploidy were assessed following ZBF by flow cytometry. Similar to what was observed in unstimulated CMK cells, ZBF preserved endocytosed AlexFluor488- factor V (Figure 21A, black histogram). In addition, distinct populations of cells could be distinguished that had a DNA content that ranged from 2N to 32N DNA (Figure 21B), which corresponds to cells with 1 – 5 nuclei the presence of which could be confirmed by fluorescence microscopy (Figure 21D). A strong correlation between increased

AlexaFluor488-factor V endocytosis and increasing DNA content was also observed

(Figure 21C).

Discussion

Chemical fixation has been used for decades to preserve tissue and subcellular components to allow for analysis at a later time (Paavilainen, Edvinsson et al. 2010). This process is often used in conjunction with immunochemistry techniques such as microscopy and flow cytometry where preservation of a cellular epitope is necessary. Formaldehyde- based crosslinking fixatives have become the predominant fixative for these types of analyses as it is inexpensive and provides irreversible fixation through crosslinking of amino acids side chains or nucleic acids (Puchtler and Meloan 1985, Helander 1994); however, fixation in this manner can fragment DNA and inhibit analysis of nucleic acids

(Gillio-Tos, De Marco et al. 2007, Jensen, Owens et al. 2010). In addition, there are well known hazards associated with the use of formaldehyde as a fixative through skin or eye

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Figure 21. Simultaneous flow cytometric analysis of DNA content and endocytosed factor V in polyploidal CMK cells.

(A) Endocytosis of AlexaFluor488-factor V

(black histogram) and (B) DNA ploidy were assessed by flow cytometry following ZBF of

CMK cells. The grey histogram represents

CMK cells incubated with AlexaFluo488-factor

IX (A). In (B), the 2N, 4N, 8N, 16N and 32N populations are indicated with arrows. (C)

Endocytosis of AlexaFluor488-factor V is also displayed as a function of DNA content. (D)

Propidium iodide (red) staining of a polyploidal megakaryocyte visualized by fluorescence microscopy. The white arrow indicates a single nucleus in a polyploidal megakaryocyte.

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contact or via the respiratory tract (National Toxicology 2011). Because of these reasons other fixation methods have been developed.

The most common alternatives, ethanol and methanol, replace water molecules in the tissue environment thereby disrupting hydrophobic and hydrophilic interactions to precipitate the proteins (Lillie and Fullmer 1976). Precipitants are known to alter the cellular morphology and tertiary structure of proteins, disrupting some antibody epitopes, which can be reversed through fixation removal and rehydration (Mann, Dyne et al. 1987,

Morkve and Hostmark 1991). In contrast, much less is known about the mechanism of

ZBF. Zinc ions are thought to stabilize the tertiary structures of proteins and have been used in conjunction with formaldehyde-based fixatives to improve morphology. The acetate ions in ZBF are thought to act as a chaotropic agent (Selevsek, Rival et al. 2009), disrupting non-covalent interactions including van der Waals forces, hydrophobic interactions, and hydrogen bonds. Some have suggested that when combined, the zinc and acetate ions introduce structural changes to the protein which cause it to form a more rigid structure (Jensen, Owens et al. 2010, Zhao, Li et al. 2011). Despite not knowing its mechanism of fixation, ZBF provides exceptional conservation of DNA, RNA, and protein morphology as shown by our present findings and others (Lykidis, Van Noorden et al.

2007, Jensen, Owens et al. 2010, Zhao, Li et al. 2011), and can be used to assess DNA content in polyploidal megakaryocytes in conjunction with a labile endocytosed protein.

Furthermore, an increase in AlexaFluor488-factor V endocytosis by higher ploidy cells was observed suggesting that factor V endocytosis occurs as a function of megakaryocyte

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differentiation and is consistent with our observations that factor V endocytosis by megakaryocytes derived ex vivo from umbilical cord blood correlates with expression of a late marker of megakaryocyte differentiation (Chapter 2).

Acknowledgements

We would like to thank Paula B. Tracy, PhD, Jay R. Silveira, PhD, and Laura M. Haynes,

PhD (University of Vermont) for helpful discussions.

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2. Bouchard, B. A., J. L. Williams, N. T. Meisler, M. W. Long and P. B. Tracy (2005). "Endocytosis of plasma-derived factor V by megakaryocytes occurs via a clathrin- dependent, specific membrane binding event." J Thromb Haemost 3(3): 541-551.

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6. Gillio-Tos, A., L. De Marco, V. Fiano, F. Garcia-Bragado, R. Dikshit, P. Boffetta and F. Merletti (2007). "Efficient DNA extraction from 25-year-old paraffin-embedded tissues: study of 365 samples." Pathology 39(3): 345-348.

7. Hadler-Olsen, E., P. Kanapathippillai, E. Berg, G. Svineng, J. O. Winberg and L. Uhlin- Hansen (2010). "Gelatin in situ zymography on fixed, paraffin-embedded tissue: zinc and ethanol fixation preserve enzyme activity." J Histochem Cytochem 58(1): 29-39.

8. Helander, K. G. (1994). "Kinetic studies of formaldehyde binding in tissue." Biotech Histochem 69(3): 177-179.

9. Jensen, U. B., D. M. Owens, S. Pedersen and R. Christensen (2010). "Zinc fixation preserves flow cytometry scatter and fluorescence parameters and allows simultaneous analysis of DNA content and synthesis, and intracellular and surface epitopes." Cytometry A 77(8): 798-804.

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13. Long, M. W., N. Williams and S. Ebbe (1982). "Immature megakaryocytes in the mouse: physical characteristics, cell cycle status, and in vitro responsiveness to thrombopoietic stimulatory factor." Blood 59(3): 569-575.

14. Lykidis, D., S. Van Noorden, A. Armstrong, B. Spencer-Dene, J. Li, Z. Zhuang and G. W. Stamp (2007). "Novel zinc-based fixative for high quality DNA, RNA and protein analysis." Nucleic Acids Res 35(12): e85.

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Appendix II. Galectin-8 is not a component of the two receptor system that mediates

factor V endocytosis by megakaryocytes

This work was presented in abstract form at the XXIV Congress of the International

Society on Thrombosis and Haemostasis

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Introduction

Platelet-derived factor V is produced through endocytosis of the plasma-derived molecule by the platelet precursor cell, the megakaryocyte (Bouchard, Williams et al. 2005,

Suehiro, Veljkovic et al. 2005). Our research to date has established a two receptor model of this process (Bouchard, Meisler et al. 2008). A molecule of factor V initially binds to a specific, unidentified factor V receptor on the surface of the megakaryocyte and a second molecule of factor V will bind to low density lipoprotein receptor related protein-1 (LRP-

1). LRP-1 mediates endocytosis in calcium- and clathrin-dependent manners. Once internalized factor V is physically (Monkovic and Tracy 1990, Kalafatis, Rand et al. 1994,

Rand, Kalafatis et al. 1994, Camire, Kalafatis et al. 1995, Conlon, Camire et al. 1997,

Gould, Silveira et al. 2004, Gould, Simioni et al. 2005, Wood 2008, Tracy, Jemings et al.

2013) and functionally (Kane, Lindhout et al. 1980, Vicic, Lages et al. 1980, Kane, Mruk et al. 1982, Viskup, Tracy et al. 1987, Monkovic and Tracy 1990, Monkovic and Tracy

1990) modified before trafficking to the platelet α-granule (Breederveld, Giddings et al.

1975, Vicic, Lages et al. 1980, Chesney, Pifer et al. 1981). Studies are currently underway in our laboratory to characterize the cellular and molecular mechanisms by which these events occur.

Recently, Zappelli et al. suggested that the specific factor V receptor is galectin-8

(Zappelli, van der Zwaan et al. 2012), a carbohydrate-binding protein. The galectin protein family plays a role in a wide range of processes including: cell differentiation, growth regulation, and apoptosis (Leffler 2001). They were originally identified as cytosolic

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proteins but have more recently been observed to be secreted and tethered to the cell via glycoproteins expressed on the cell surface (Hughes 1999). Galectin-8 has two carbohydrate recognition domains (CRD) (Hirabayashi, Hashidate et al. 2002). The N- terminal CRD has a preference for complex N-linked glycans containing a sialic acid linked to a galactose residue (Stowell, Arthur et al. 2010) while the C-terminal CRD will bind to

N-acetyllactosamine, consisting of a modified glucose residue, N-acetylglucosamine, linked to a galactose residue, in complex N-linked glycans (Stowell, Arthur et al. 2010).

Factor V, which is heavily glycosylated (Jenny, Pittman et al. 1987), was shown to interact with galectin-8 using galectin-8 affinity chromatography of platelet lysates followed by mass spectrometric analysis (Romaniuk, Tribulatti et al. 2010). Zappelli et al. used a megakaryocyte like cell line, DAMI, to demonstrate that known galectin-8 ligands block factor V endocytosis (Zappelli, van der Zwaan et al. 2012). In addition, siRNA knockdown of galectin-8 in these cells lead to decreased factor V endocytosis.

Previous studies have identified the light chain of factor V as the region that is important for binding and endocytosis (Bouchard, Abdalla et al. 2013). Interestingly, the glycans at Asn1675 and Asn2181 in this region contain galactose residues linked to sialic acid residues as well as N-acetyllactosamine, both of the galectin-8 binding sites (Tracy,

Jemings et al. 2013) and may mediate these interactions. Thus, the goal of this study was to confirm the results published Zappelli et al. (Zappelli, van der Zwaan et al. 2012) and demonstrate a role for galectin-8 in factor V endocytosis. Flow cytometric analyses were used to evaluate intracellular and extracellular expression of galectin-8 by megakaryocytes.

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The effect of galectin-8 inhibition on factor V endocytosis was also monitored. In these experiments, we were not able to confirm that galectin-8 functions as the specific factor V receptor on megakaryocytes.

Materials and methods

Materials: CMK cells were a generous gift of Dr. Hava Avraham (The Division of

Experimental Medicine, Harvard Medical School, Boston, MA). DAMI cells were a generous gift of Dr. Walter Kahr (Program in Cell Biology, Research Institute, The

Hospital for Sick Children, Toronto, Ontario, Canada) and Dr. Robert Handin

(Hematology Division, Brigham and Women's Hospital, Boston, MA) (Greenberg,

Rosenthal et al. 1988). Human factor V, purified from freshly frozen plasma (Katzmann,

Nesheim et al. 1981), and human factor IX (Haematologic Technologies, Essex Junction,

VT) were conjugated to AlexaFluor488 using the manufacturer’s (Life Technologies,

Grand Island, NY) instructions (Bouchard, Williams et al. 2005, Bouchard, Meisler et al.

2008). Alexa647-conjugated goat α-mouse IgG was also from Life Technologies. Anti- galectin-8 antibody was purchased from Santa Cruz Biotechnology (Bergheimer

Heidelberg, Germany). D-lactose monohydrate was acquired from Fisher Scientific (Fair

Lawn, NJ) while D-Mannose was from Alfa Aesar (Ward Hill, MA).

Megakaryocyte cell culture: The megakaryocyte-like cell lines, CMK (Taniguchi,

London et al. 1999) and DAMI (Zappelli, van der Zwaan et al. 2012), were cultured as

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previously described. DAMI cell lots were maintained as separate cultures. Ex vivo megakaryocytes derived from human umbilical cord blood were isolated and cultured as described in serum free media containing thrombopoietin, interleukin-3, and stem cell factor (Denis, Tolley et al. 2005, Foulks, Marathe et al. 2009, Shi, Smith et al. 2014,

Gertz, Schwertz et al. Submitted). Umbilical cord blood was collected with approval of the internal review board and patient consent at University of Vermont Medical Center from full term elective cesarean sections. For each experiment using CMKs and DAMIs, cells were washed twice by centrifugation (250xg, 7 min) followed by resuspension in serum-free media containing 0.1% endotoxin-free bovine serum albumin. Ex vivo- derived megakaryocytes were cultured for 11 days and then washed once (500xg, 5 min) with fresh culture media. Prior to each experiment, cell viability was assessed by trypan blue exclusion.

Flow cytometry analysis of galectin-8 expression on megakaryocytes: CMK, DAMI or ex vivo-derived megakaryocytes were washed two times by centrifugation (250xg, 7 min) followed by resuspension in 20 mM Hepes, 0.15 M NaCl, pH 7.4 (HBS). To assess intracellular galectin-8 expression, cells were subjected to fixation using 4% paraformaldehyde (30 min). Non-specific binding sites were blocked using 1% bovine serum albumin. Cells were incubated for 1 hr on ice in the presence or absence of an anti- galectin-8 antibody (0.4 μg/mL). For some experiments, saponin (0.1%) was included in the immunostaining reactions to assess intracellular galectin-8 expression. Cells were

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washed as described above to remove excess antibodies, and incubated for 1 hr on ice with

Alexa647-conjugated goat anti-mouse IgG (4 μg/mL) then washed to remove excess antibodies. Finally, non-permeabilized cells were subject to 4% paraformaldehyde fixation.

The effects of galectin-8 inhibition on factor V endocytosis: CMK, DAMI or ex vivo- derived megakaryocytes (1x106 cells/mL) were cultured for 30 min at 37°C in the presence or absence of lactose (5 mM), mannose (5 mM), or α-galectin-8 antibodies (4 µg/mL) prior to addition of AlexaFluor488-labeled factor V or IX (30 nM, 3 hr) The cells were removed from the culture dish, washed two times by centrifugation (250xg, 7 min) followed by resuspension in HBS to remove cell surface–associated factor V- and factor IX-

AlexaFluor488. Subsequently, cells were fixed using 4% paraformaldehyde.

Flow cytometry analysis: Cells (10,000) were analyzed on a BD LSR II flow cytometer

(BD Biosciences, San Jose, CA). The cell population of interest was first selected based on forward and side scatter. Analyses regions were defined such that approximately 2% of the negative control cells (cultured with AlexaFluor488-FIX or stained in the absence of primary antibody) were positive. All data were collected as binary FCS files and analyzed using FlowJo version 10 data analysis software (FlowJo, LLC, Ashland, OR).

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Results

Galectin-8 is not expressed on the surface of megakaryocytes.

In an attempt to reproduce the results published by Zappelli et al. (Zappelli, van der Zwaan et al. 2012), our studies examined the effects of galectin-8 inhibitors on factor

V endocytosis by megakaryocyte-like cell lines CMK and DAMI, and megakaryocytes derived ex vivo from human umbilical cord blood to more closely mimic the in vivo scenario. It is thought that after their submission to ATCC the DAMI cell line became contaminated with HeLa cells (MacLeod, Dirks et al. 1997). Therefore, for this report

DAMI cells from ATCC, as used by Zappelli et al. (Zappelli, van der Zwaan et al. 2012), and provided by Dr. Kahr, will be referred to as DAMI A. DAMI cells provided to us by the ATCC depositor, Dr. Handin (Greenberg, Rosenthal et al. 1988), will be referred to as

DAMI B. As no differences were observed with the results obtained between DAMI A and DAMI B cells, only the data using DAMI A cells will be presented. Intracellular

(top) and cell surface (bottom) expression of galectin-8 was assessed by flow cytometry on CMK cells, DAMI A cells, and ex vivo-derived megakaryocytes (Figure 22). When compared to reactions stained in the absence of primary antibody (Figure 22, red histogram) a large quantity of galectin-8 appears to be located intracellularly (Figure 22, blue histogram) as demonstrated by flow cytometry on permeabilized megakaryocyte- like cell lines and ex vivo-derived megakaryocytes. In contrast, no expression of galectin-

8 (Figure 22, blue histogram) could be detected in the absence of cell permeabilization

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Figure 22. Expression of galectin-8 by megakaryocytes.

The megakaryocyte-like cell lines CMK and DAMI, or megakaryocytes derived ex vivo from CD34+ umbilical cord blood cells were immunostained in the presence (blue histogram) or absence (red histogram) of an α-galectin-8 antibody as described in

Materials and Methods. Intracellular galectin-8 staining (top) was performed subsequent to fixation and permeabilization. Extracellular galectin-8 staining (bottom) was performed prior to fixation and in the absence of permeabilization.

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when compared with controls (Figure 22, red histogram) suggesting that these cells do not express galectin-8 on their membrane surface.

Inhibition of Galectin-8 does not alter factor V endocytosis.

The ability of known galectin-8 ligands and an inhibitory anti-galectin-8 antibody (Zappelli, van der Zwaan et al. 2012) to inhibit factor V endocytosis (Figure 23, black histogram) was assessed by flow cytometry on CMK, DAMI, and ex vivo-derived megakaryocytes. The known galectin-8 ligand, lactose (Figure 23, green histogram), which consists of a galactose connected to a glucose, did not alter factor V endocytosis when compared to factor V alone (Figure 23, pink histogram) or a negative control, mannose (Figure 23, purple histogram), which does not bind galectin-8.

Discussion

Results described in the current study indicate that galectin-8 is not expressed on the megakaryocyte cell surface and does not mediate factor V endocytosis. Furthermore, studies described in this thesis (Chapter 3) demonstrate that selective trimming of the complex N-linked glycans in factor V to remove the galectin-8 binding site does not have an effect on factor V endocytosis. Taken together, these observations suggest that galectin-8 does not function as part of the two receptor system mediating factor V endocytosis by megakaryocytes. These results are in contrast to those published by

Zappelli et al. in which galectin-8 was shown to be expressed on the DAMI cell surface 176

Figure 23. Inhibition of galectin-8 does not alter factor V endocytosis.

CMK, DAMI and ex vivo-derived megakaryocytes were preincubated for 30 min at 37°C in the presence of lactose (green histogram), mannose (purple histogram), or an inhibitory

α-galectin-8 antibody (black histogram) prior to the addition of AlexaFluor488-factor V

(30 nM, 3 hr, 37°C). Endocytosis of AlexaFluor488-factor V by these cells was compared to endocytosis of AlexaFluor488-factorV by untreated cells (pink histogram) or by cells incubated with AlexaFluor488-factor IX (red histogram).

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(Zappelli, van der Zwaan et al. 2012). In our experiments, cells were subjected to fixation with paraformaldehyde following galectin-8 immunostaining and washing away of excess antibody. Duplication of the immunostaining technique used by Zappelli et al.

(Zappelli, van der Zwaan et al. 2012) in which cells were subjected to fixation prior to immunostaining lead to galectin-8 detection at a level similar to permeabilized cells (data not shown) suggesting that paraformaldehyde fixation may cause unwanted cell permeabilization and intracellular access of antibodies. This observation is well known within the flow cytometry community, although no published reference could be found. It is also possible that under the conditions of our study, galectin-8, although secreted from the cell and associated with the cell membrane in culture, did not remain bound to the cell surface during experimental processing and therefore could not be detected by immunostaining. However, as we still observed factor V endocytosis despite the absence of cell surface expressed galectin-8, this is a less likely explanation for the contrasting results.

Not surprisingly, and consistent with our inability to observe cell surface galectin-8 expression, factor V endocytosis performed in the presence of lactose or an inhibitory anti-galectin-8 antibody had no effect on its uptake by megakaryocytes in contrast to observations made by Zappelli et al. (Zappelli, van der Zwaan et al. 2012). In addition, they also observed reduced factor V endocytosis following knockdown of galectin-8 with galectin-8 siRNA. It is of interest to note that LRP-1 was recently identified as a galectin-8 binding partner in osteoblasts (Vinik, Shatz-Azoulay et al.

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2015). Additional experiments demonstrated that in this system, LRP-1 is a negative regulator of galectin-8 function. While LRP-1’s primary function is endocytosis of ligands destined for lysosomal degradation (Goldstein and Brown 1974), its partnering with other cell surface or integral membrane proteins as well as cytoplasmic signaling molecules to modulate various cellular functions is well known (Czekay, Kuemmel et al.

2001, Loukinova, Ranganathan et al. 2002). As we did not verify the effect of galectin-8 knockdown on factor V endocytosis, the possibility remains that cytoplasmic galectin-8, in conjunction with LRP-1, modulates factor V endocytosis by megakaryocytes.

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