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PRIONS AND PLATELETS: A POSSIBLE ROLE FOR CELLULAR PRION
PROTEIN
BY
CATHERINE ROBERTSON
A Thesis Submitted to the Faculty of Graduate Studies
In Partial Fulfillment of the Requirements for the Degree of
MASTER OF SCIENCE
Department of Oral Biology University of Manitoba
Winnipeg, Manitoba
Catherine Robertson, March 2005
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TABLE OF CONTENTS
ABSTRACT.......................................................................................................................5
DEDICATION....................................................................................................................6
ACKNOWLEDGEMENTS..............................................................................................7
ABBREVIATIONS ...........................................................................................................8
LIST OF TABLES............................................................................................................9
LIST OF FIGURES ........................................................................................................10
1. INTRODUCTION .......................................................................................................11
1.1 Cellular prion protein structure .......................................................................................................................11
1.2 Biosynthesis, cellular location and trafficking of PrPc...............................................................................14
1.3 Interaction of PrPc with other proteins ..........................................................................................................14
1.4 Putative functions of PrPc ..................................................................................................................................17 1.4.1 Copper Binding............................................................................................................................................... 17 1.4.2 Signal Transduction....................................................................................................................................... 18 1.4.3 Cell Adhesion.................................................................................................................................................. 19
1.5 Transmissible Spongiform Encephalopathies ...............................................................................................19
1.6 The Role of PrPc in TSEs ...................................................................................................................................22
1.7 Pathogenesis of TSE.............................................................................................................................................22
1.8 PrPc and Blood......................................................................................................................................................24
1.9 Platelets ....................................................................................................................................................................27 1.9.1 Plasma membrane proteins........................................................................................................................... 27 1.9.2 Platelet Granules ............................................................................................................................................. 27 1.9.3 Tubular Systems ............................................................................................................................................. 31 1.9.4 Platelet Function............................................................................................................................................. 31
1.10 Objectives ..............................................................................................................................................................34
2. MATERIALS AND METHODS................................................................................35
2.1 Antibodies and reagents ......................................................................................................................................35 2.1.1 Anti-PrP antibodies ........................................................................................................................................ 35
3
2.1.2 Other antibodies.............................................................................................................................................. 35 2.1.3 Reagents........................................................................................................................................................... 36 2.1.4 Buffers .............................................................................................................................................................. 36
2.2 Preparation of washed platelets ........................................................................................................................36
2.3 Preparation of whole platelet homogenates ...................................................................................................37
2.4 Fractionation of platelets ....................................................................................................................................37
2.5 Protein Determination.........................................................................................................................................38
2.6 Electrophoresis and Immunoblotting ..............................................................................................................38
2.7 Immunofluorescence ............................................................................................................................................39
2.8 Mepacrine labeling of dense granules .............................................................................................................40
2.9 Flow cytometry ......................................................................................................................................................40
2.10 Platelet microvesicle release.............................................................................................................................41
2.11 Microvesicle and Exosome preparation........................................................................................................41
2.12 Slot Blots................................................................................................................................................................42
2.13 Electron microscopy and immunocytochemistry.......................................................................................43 2.13.1 Resin sections of whole platelets............................................................................................................... 43 2.13.2 Whole mounts of sucrose gradient fractions........................................................................................... 43
2.14 Antibody staining ................................................................................................................................................43
2.15 Platelet Aggregation Studies ............................................................................................................................44
2.16 Platelet Adhesion Assays...................................................................................................................................44
2.17 Immunoprecipitations .......................................................................................................................................45
3 RESULTS.....................................................................................................................46
3.1 Cellular prion protein is present on platelet alpha granule membranes ...............................................46
3.2 Cellular prion protein translocates to the surface, and is released, on platelet activation...............52
3.3 Thrombin causes the release of microvesicles from platelets ..................................................................52
3.4 Cellular prion protein is present in microvesicles and exosomes ............................................................56
3.5 Cellular prion protein is not involved in platelet aggregation ..................................................................66
3.6 Cellular prion protein may be involved in platelet adhesion ....................................................................66
3.7 Cellular prion protein co-precipitates with calreticulin.............................................................................66
4
4. DISCUSSION .............................................................................................................72
5. LIMITATIONS.............................................................................................................78
6. CONCLUSIONS.........................................................................................................80
7. APPENDICES............................................................................................................81
APPENDIX A. BUFFER SOLUTIONS.......................................................................81
APPENDIX B. PROCESSING SCHEDULE FOR TRANSMISSION ELECTRON MICROSCOPY...............................................................................................................87
8. REFERENCES...........................................................................................................88
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ABSTRACT
Cellular prion protein (PrPc) is a GPI–anchored protein, of unknown function,
found in a various cell types throughout the body. It is now widely believed that a mis-
folded, protease resistant form of this protein is responsible for a group of fatal
neurodegenerative diseases called transmissible spongiform encephalopathies (TSE),
including Creutzfeldt-Jakob disease (CJD) and kuru in humans, scrapie in sheep, chronic
wasting disease (CWD) in deer and elk and bovine spongiform encephalopathy (BSE) in
cattle. Although the exact function of PrPc is unknown it binds copper and has been
implicated in copper homeostasis, signal transduction and cell adhesion.
The pathogenesis of prion diseases is poorly understood, however the expression
of PrPc in target cells is an absolute requirement for disease progression. Platelets have
been shown to be the largest reservoir of PrPc in peripheral blood cells and studies in
animal models have suggested platelets may also be involved in TSE infectivity.
In this study, we determine the exact location of PrPc within human platelets,
examine the mobilization and release of PrPc from activated platelets on both
microvesicles and exosomes and suggest a possible role for platelets in prion infectivity.
In addition we examine the role of PrPc within normal platelet functions including
aggregation, signal transduction and adhesion.
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DEDICATION
It is the duty of the working class to educate themselves to their fullest potential. - D.H. Lawrence 1885-1930
This work is dedicated to the memory of my Grandmother,
Catherine Quinn
A woman who recognized the value of a good education.
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ACKNOWLEDGEMENTS This project would never have been completed without the help and support of a large number of people. First of all, I would like to express my sincere gratitude to my supervisor Dr. Archie McNicol for his patience and guidance throughout this project. You have not only been an excellent supervisor but a true friend. To Dr. Stephanie Booth who is a member of my committee, but who also acted as co-supervisor and advisor, I thank you not only for your advice and input into this study, but for not laughing out loud at my wild hypotheses. I wish to thank Dr. Catalina Birek, and Dr. Elliot Scott for agreeing on short notice to be members of my committee. I am indebted to Dr. Michael Coulthart of the National Laboratory for Prion Diseases for allowing me both the time and laboratory resources to carry out this project, and for the many insightful conversations. I wish to thank Dr. Daniel Beniac from the National Laboratory for Bloodborne Pathogens, and Lynne Burton from the Canadian food Inspection Agency for their excellent technical assistance with the samples for electron microscopy as well as Dr. Tim Booth from the National Laboratory for Bloodborne Pathogens for not only allowing me the use of the electron microscope but for providing helpful suggestions regarding electron microscopy. To Deborah Godal, Susan Shead and Elke Jackson who willingly bled all of the volunteer donors, I thank you from the bottom of my heart, and to all the volunteers, I can honestly say I could not have done this project without you. Thank you all so much. I want to thank Rhonda Mogk not only for her friendship and support but for also showing me the correct way to end a telephone call. To everyone in HGPD and alsoViral Exanthema, thank you all for making the 5th floor such a fun place to work. To my husband John, and daughters Suzanne and Kayleigh whose unfailing support and understanding throughout this endeavour mean more to me than I can possibly put into words. We’ll soon be able to do all the things I had promised to do “after Mum finishes her thesis”. I wish to thank my Aunt Cathie and Uncle Archie not only for their support and encouragement during this time but for always being such a positive influence throughout my childhood. To my good friends and “adopted family” the Tennents, I am so glad you decided to come to Canada. Finally, a big thank you to “the Captain, for keeping my spirits up over the last few years.
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ABBREVIATIONS
Ab Antibody
ACD Acid citrate dextrose
BSA Bovine serum albumin
BSE Bovine spongiform encephalopathy
CJD Creutzfeldt-Jakob Disease
CK Casein kinase
EDTA Ethylenediaminetetraacetic acid
GPI Glycophosphatidylinositol
kDa Kilodalton
PBS Phosphate buffered saline
PILPC Phosphatidylinositol-specific phospholipase C
PMSF Phenyl methyl sulphonyl fluoride
PrP Prion protein
PrPc Cellular prion protein
PrPres Protease resistant prion protein
SDS Sodium dodecyl sulfate
SOD Superoxide dismutase
TBS Tris buffered saline
TBS-T Tris buffered saline with 0.05% tween20
TME Transmissible mink encephalopathy
TSE Transmissible spongiform encephalopathy
vCJD variant Creutzfeldt-Jakob disease
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LIST OF TABLES
Table 1. Protein molecules interacting with cellular prion protein 16
Table 2. Distribution of cellular prion protein in blood 26
Table 3. Main glycoprotein receptors on platelet plasma membrane 29
Table 4. Platelet Adhesion Assays 71
10
LIST OF FIGURES
Figure 1. Structure of prnp gene 12
Figure 2. Structure of PrPc and PrPres 13
Figure 3. Histology and immunohistochemistry of TSEs 21
Figure 4. Normal resting platelet 28
Figure 5. Electron microscopy: Platelet Activation 33
Figure 6. Immunofluorescent staining of PrPc in quiet and activated intact platelets47
Figure 7. Double immunofluorescent staining of PrPc, CD62 and CD63 in resting
permeabilised platelets 48
Figure 8. Double immunofluorescent staining of dense granules, PrPc and CD62 positive
granules in resting permeabilised platelets 50
Figure 9. Western blot of platelet fractions 51
Figure 10. Western blot of platelet pellets and supernatants 53
Figure 11. Densitometry on activated platelets and supernatants 54
Figure 12. Flow cytometry of activated platelets 55
Figure 13. Flow cytometry of activated platelets 57
Figure 14. Immunogold labeling of activated platelets 60
Figure 15. Slot blot of sucrose gradient fractions 62
Figure 16. CD63 Immuno electron microscopy of isolated exosomes 63
Figure 17. PrPc Immunolabeling of sucrose gradient pellet 65
Figure 18. Platelet aggregation 68
Figure 19. Immunoprecipitation with antibody to PrPc and calreticulin 70
Figure 20. Double labeling with antibodies to calreticulin and prion protein 71
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1. INTRODUCTION
In the course of studies to determine the causative agent of the group of diseases
known as Transmissible Spongiform Encephalopathies (TSE), an insoluble protein,
termed prion for “Proteinaceous Infectious Particle”, was partially purified from the
brains of affected animals (Prusiner, S. B., 1998).
Two forms of the prion protein (PrP) were identified, the non- infectious cellular
form (PrPc), which was expressed in a variety of normal neuronal and non-neuronal
tissues including tonsils, spleen and blood cells, and a protease-resistant, form (PrPres)
which is believed to be the agent responsible for TSEs (Martins, V. R. et al., 2002;
Barclay, G. R. et al., 1999).
1.1 Cellular prion protein structure
PrPc is a glycophosphatidyinositol (GPI) anchored protein of 254 amino acids
with a molecular mass of 33-40 kilodaltons (kDa) or, in its unglycosylated form, of
27kDa. Structurally, the protein has 5 glycine/proline rich octapeptide repeat regions, two
N-linked glycosylation sites, a single disulfide bond and signaling peptides at both the
amino and carboxyl terminii (Stahl, N. et al., 1987) (Figure1). PrPc is composed of about
40% alpha helix formation with less than 10% being in the beta sheet formation.
However the beta sheet content increases to about 50% in the infectious form (Martins,
V. R., 1999) (Figure 2). The prnp gene is located on chromosome 20 and the protein is
highly conserved among species, with a similarity of 85-97% among mammals (Gabriel,
J. M. et al., 1992).
12
Figure 1. Structure of prnp gene (Vostal JG et al 2001)
254 Signal peptide
Octapeptide repeats
N-linked oligosaccharides
GPI-anchor
Cell membrane
S-S 1
13
Figure 2. Structure of Normal cellular Prp and infectious isoform
The normal form of cellular prion protein on the left is composed of mainly alpha helical
formation whereas the infectious misfolded isoform is predominantly beta sheet
formation (http://www.cmpharm.ucsf.edu/cohen/media/pages/gallery.html)
PrPc PrPres
14
1.2 Biosynthesis, cellular location and trafficking of PrPc
PrPc is synthesized in the rough endoplasmic reticulum, and passes through the
golgi en route to the plasma membrane. During biosynthesis it undergoes a number of
post-translational modifications including cleavage of the N-terminal signal peptide,
addition of N-linked oligosaccharide chains, formation of a single disulphide bond,
cleavage of C-terminal peptide and attachment of the GPI anchor (Turk, E. et al., 1988;
Haraguchi, T. et al., 1989).
The majority of PrPc is found in detergent resistant domains, called lipid rafts, at
the cell membrane of neuronal and non-neuronal cells, as well as of cultured cell lines
(Vey, M. et al., 1996). However PrPc does not remain on the cell surface but cycles
between the membrane and an endocytic compartment (Shyng, S. L. et al., 1993).
Clathrin coated pits are the main structures responsible for endocytic uptake of PrPc
(Shyng, S. L. et al., 1994).
PrPc has been demonstrated in the secretory granules of epithelial cells in the
stomach (Fournier, J. G. et al., 1998), as well as on the surface of both quiet and activated
platelets, (Barclay, G. R. et al., 1999; Holada, K. et al., 1998; MacGregor, I. et al., 2000).
1.3 Interaction of PrPc with other proteins
PrPc interacts with a large number of proteins (Table 1). The first interacting
proteins identified were a pair of prion protein ligands Pli45 and Pli 110 (Oesch, B. et al.,
1990); the former being glial fibrillary acidic protein, a marker for astrocytes which
proliferate in response to TSE infections (DeArmond, S. J. et al., 1992).
15
Subsequently, a number of other PrPc- interacting proteins have been identified
using a yeast two-hybrid system; these include the anti-apoptotic protein Bcl-2
(Kurschner, C. et al., 1996), the cellular chaperone heat shock protein 60 (Hsp60)
(Edenhofer, F. et al., 1996), the 37kDa laminin receptor precursor (Rieger, R. et al.,
1997), the synaptic vesicle marker synapsin1b, the adaptor protein Grb2 and a protein
named prion interaction protein (Pint 1), for which no function has been determined
(Spielhaupter, C. et al., 2001). Capellari et al co- immunoprecipitated PrPc from a M17
neuroblastoma cell line with antibodies to grp94, protein disulphide isomerase, calnexin,
and calreticulin suggesting that PrPc interacts, with these proteins (Capellari, S. et al.,
1999).
PrPc also binds to laminin in PC12 cells and rodent primary neurons, and this
interaction promotes neurite outgrowth in these cells (Graner, E. et al., 2000). A number
of additional cell surface proteins interact with PrPc including neuronal cell adhesion
molecules (N-CAMs) (Schmitt-Ulms, G. et al., 2001), apolipoprotein 1 (Aplp1) an
amyloid precursor protein which has been implicated in Alzheimers disease (Yehiely, F.
et al., 1997), the 67 kDa laminin receptor (Gauczynski, S. et al., 2001) and
glycosaminoglycans (GAGS) (Priola, S. A. et al., 1994; Pan, T. et al., 2002).
Finally, complementary hydropathy, a technique in which cDNA is used to
generate a molecule which is a complementary mirror image of the target protein,
identified a 66 kDa protein, the stress inducible protein STI-1, which bound to PrPc and
which may be involved in neuroprotection (Zanata, S. M. et al., 2002).
16
Table 1. Protein molecules interacting with cellular prion protein
Molecule Function Possible binding site Reference Synapsin 1b Regulation of Intracellular vesicles Spielhaupter(2001)
neurotransmitter Grb2 Adapter protein Pint 1 Unknown Unknown Caveolin 1 Signaling Caveolae raft Mouillet-Richard (2000) CK2 Ser/Thr Caveolae raft Meggio (2000)
phosphorylation STI1 HSP rela ted Cell surface Zanata (2002) Bcl-2 Apoptosis Unknown Kurschner (1996) Laminin Adhesion Cell surface Graner (2000) GAG Biomolecular Cell surface Priola (1994)
transport Pan (2002) N-CAM Adhesion Caveolae-like domain Schmitt-Ulms (2001) GFAP Cell repair Unknown Oesch (1990) Pli 110 Unknown Unknown Hsp60 Chaperone Unknown Edenhofer (1996) Stockel (1998) Nrf2 Apoptosis Unknown Yehiely (1997)
inhibitor Aplp1 Regulation of Cell surface
neurite outgrowth Laminin laminin Cell surface Gauczynski(2001) receptor binding
Grp94 protein disulphide Unknown Capellari (1999) isomerase Calnexin Chaperone Calreticulin Chaperone
17
1.4 Putative functions of PrPc
As outlined above, PrPc binds to a large number of proteins, however its
function(s) remains unclear. Early attempts to determine a function for PrPc using prnp
knockout mice (PrP-/-) were disappointing, since there were no apparent differences
between the knockout and wild type mice (Bueler, H. et al., 1992). To date, four different
strains of PrP-/- mice have been developed (Manson, J. C. et al., 1994; Sakaguchi, S. et
al., 1996). Two of the later strains developed ataxia which suggested a neuroprotective
role for PrPc (Moore, R. C. et al., 1999).
1.4.1 Copper Binding
In 1992, Pan et al. demonstrated that PrPc from hamster brain bound copper (Pan,
K. M. et al., 1992), and subsequent studies showed that PrPc can bind other metal ions,
such as zinc, manganese and nickel, but with a much lower affinity than copper (Stockel,
J. et al., 1998; Brown, D. R. et al., 2000). The main copper binding site in human PrPc is
an N-terminal octapeptide repeat region encompassing residues 60-91 (Hornshaw, M. P.
et al., 1995; Brown, D. R. et al., 1997a; Viles, J. H. et al., 1999). PrP-/- mice have 50%
lower copper concentrations in their synaptosomal fractions than wild type mice,
suggesting that PrPc may play a role in copper regulation in neuronal synapses (Herms, J.
et al., 1999). Similarly, there are higher levels of copper in mice over-expressing PrPc
(Brown, D. R. et al., 1998). Interestingly the association with copper gives PrPc similar
enzymatic activities to that of Cu2+/Zn2+ superoxide dismutase (SOD-1) (Brown, D. R.
et al., 1999) and SOD-1 activity in cultured neurons from PrP-/- mice is 50% lower than
that from normal mice, while activity from PrPc overexpressing mice is about 20%
18
higher than control mice (Brown, D. R. et al., 1997b). Taken together these studies
suggested a role for PrPc in the protection against oxidative stress.
These studies, however, have not been substantiated. Waggoner and colleagues
failed to detect any differences in copper levels between PrP-/-, wild type and PrPc over-
expressing mice. In addition, no effect of PrPc on SOD-1 activity was seen, suggesting
that PrPc may simply act as a carrier for copper ions (Waggoner, D. J. et al., 2000). In
addition, recent in vivo studies using PrP-/- and PrP over-expressing mice crossed with
SOD-1 over-expressing mice failed to detect any effects of PrPc on SOD-1 activity
(Hutter, G. et al., 2003).
1.4.2 Signal Transduction
The role of PrPc in signal transduction has also been investigated. As outlined
above, two putative surface receptors for PrPc have been identified; the 37kDa/66 kDa
laminin receptor (Gauczynski et al 2001) and STI-1 (Zanata et al 2001). A yeast two-
hybrid system demonstrated that PrPc also interacts with the cytoplasmic adaptor protein
Grb2, which acts as a link between extracellular receptors and intracellular signaling
molecules (Koch, C. A. et al., 1991).
In cultured neuronal cells, PrPc indirectly increases the phosphorylation of the
tyrosine kinase fyn and it has been suggested that caveolin–1 acts as the intermediate
signaling molecule between PrPc and fyn (Mouillet-Richard, S. et al., 2000).
In addition, recombinant bovine PrPc has been shown to increase the
phosphotransferase activity of the pleiotropic protein kinase, casein kinase 2 (CK-2),
which is abundant in the brain and is involved in a number of signal transduction, as well
as gene expression, pathways (Meggio, F. et al., 2000).
19
1.4.3 Cell Adhesion
Graner et al. (2000) demonstrated that, in addition to an interaction with the
laminin receptor, PrPc is directly associated with laminin, and have suggested a role for
PrPc in neuronal cell adhesion and neurite outgrowth.
Studies using the neuroblastoma cell line, N2A, demonstrated that cells over-
expressing PrPc showed increased aggregation when compared to wild type N2A cells.
Furthermore, when surface PrPc was removed by incubation with phosphatidylinositol-
specific phospholipase C (PILPC), which cleaves GPI- linked proteins, aggregation levels
returned to that of wild type cells (Mange, A. et al., 2002). Cross-linking PrPc with
formaldehyde in N2A cell lines, revealed that PrPc was present in high molecular weight
complexes and that these complexes were three splice variants of N-CAMs (Schmitt-
Ulms, G. et al., 2001). These studies strengthened the argument for the involvement of
PrPc in neuronal cell adhesion.
1.5 Transmissible Spongiform Encephalopathies
TSEs are a group of fatal neurodegerative diseases that affect both humans and
animals. The animal TSEs include BSE in cattle, scrapie in sheep, CWD in mule deer
and elk, transmissible mink encephalopathy in farmed mink and feline spongiform
encephalopathy in zoological and domestic cats (Sigurdson, C. J. et al., 2003).
The human TSEs are typically diseases of late middle age and can be either
inherited or acquired (Palmer, M. S. et al., 1992). The inherited forms include Familial
CJD, Fatal Familial Insomnia or Gerstmann-Straussler-Schienker syndrome, and can all
be attributed to a germline prnp gene mutation (Ironside, J. W., 1996). The acquired
20
forms can be iatrogenic, as in the cases of transplantation of dura matter (Liscic, R. M. et
al., 1999), or the use of pituitary hormones from previously infected patients (Collins, S.
et al., 1996), or can also be acquired through ingestion of PrPres. For example, in Papua,
New Guinea, the prion disease Kuru was attributed to ritualistic cannibalism (Pedersen,
N. S. et al., 2002), and the outbreak of variant CJD (vCJD) in the United Kingdom has
been linked to ingestion of beef from cattle infected with BSE (Bruce, M. E. et al., 1997).
There have also been cases of sporadic CJD or atypical CJD, which have an
unknown cause but have been thought to be due to a somatic mutation in the prnp gene
(Poser, S. et al., 2000).
The physical symptoms of prion diseases may include visual disturbances,
cerebellar ataxia (gait difficulties), myoclonus (muscle spasms) and progressive
dementia. These symptoms are not present in all TSE cases. For example, vCJD differs
from sporadic CJD in that it usually affects patients at a much younger age, and the
duration of the disease is longer. However all of these forms of prion diseases are fatal
and, at the present time, there is no treatment (Will, R. G., 2002).
At the tissue level, the family of diseases is characterized by the spongiform
appearance of brains of affected individuals, due to vacuolation of neurons and
surrounding neuropil. This vacuolation is accompanied by deposition of protease-
resistant protein (Wells, G. A., 1993) (Figure 3).
21
Figure 3. Histology and Immunohistochemistry of TSEs
Example of TSE infected brain. A: Haemotoxylin and Eosin stained section showing vacuolation and degeneration of neurons (n) and
surrounding neuropil (arrows). B: Ab to PrP demonstrates presence of protease resistant protein deposits in neuronal cell bodies (n) as
well as along the neuronal processes (arrows). (Micrographs courtesy of Dr. Y Robinson, Canadian Food Inspection Agency)
A B
n
n
n
22
1.6 The Role of PrPc in TSEs
The role of PrPc in TSEs has been widely studied and it is now widely
accepted that PrPres is the agent responsible. This has led to the prion hypothesis
which states that PrPc undergoes a conformational change to PrPres, which acts as a
template for the transformation of additional PrPc to PrPres (Telling, G. C. et al.,
1996). The mechanism of this conversion is poorly understood, although another
protein, termed “Protein X”, has been implicated (Telling, G. C. et al., 1995; Kaneko,
K. et al., 1997).
Studies using PrP-/- mice suggest that the PrPc must be present in order for
the disease to progress (Brandner, S. et al., 2000), PrP-/- mice inoculated with murine
scrapie were still healthy 500 days post inoculation compared to the wild-type mice
which developed scrapie in less than 165 days post inoculation (Prusiner, S. B. et al.,
1993). Furthermore, PrP-/- mice grafted with PrPc expressing neuroectodermal tissue
tissue showed only neurodegeneration in the area of the graft after inoculation with
scrapie (Brandner, S. et al., 1996)
1.7 Pathogenesis of TSE
The pathogenesis of prion diseases is also poorly understood. It has been
proposed that ingested infectious prions are absorbed through Payers patches in the
gut and accumulate in the spleen. The spleen is highly innervated and PrPres travels
to the brain via peripheral nerves (Glatzel, M. et al., 2001).
Although this is the most likely route for neuroinvasion, it has been suggested
that blood may also play a role in the pathogenesis of prion diseases (Ramasamy, I. et
23
al., 2003; Radebold, K. et al., 2001). For example, a number of studies in animals
have produced prion infections from inoculation of buffy coats and other blood
components (Bons, N. et al., 2002; Holada, K. et al., 2002; Cervenakova, L. et al.,
2003). In 2000, Hunter et al used sheep, previously infected with BSE or scrapie, as
blood donors to TSE-free sheep. Due to the long incubation period of BSE, these
studies are expected to take 5 years to complete; however in 2002 the authors
reported that 17% of the recipient animals were showing clinical signs of BSE and
19% were positive for scrapie, providing evidence for the transmission of BSE and
scrapie via blood transfusion (Hunter, N. et al., 2002; Houston, F. et al., 2000).
Of importance there have been two recent reports of transmission of variant
CJD by blood transfusion of blood from a donor later found to have been infected
with vCJD (Llewelyn, C. A. et al., 2004; Peden, A. H. et al., 2004). The mechanism
of transmission is currently not clear.
Recently the effectiveness of standard leucoreduction for removing TSE
infectivity from whole blood was investigated (Gregori, L. et al., 2004). The removal
of all white cells reduced infectivity by only 42%, suggesting that other blood
components, cells or plasma, carry TSE infectivity. These data, when considered
along with the animal studies of prion infectivity in blood, have highlighted both the
significance of PrPc and the role of individual blood components in the pathogenesis
of TSEs. Therefore in order to understand the transmission of the disease, it is
important to determine the physiological behaviour and function of the normal
cellular form of this protein in blood cells and plasma.
24
1.8 PrPc and Blood
Several flow cytometric studies on whole blood have shown PrPc to be
present on the surface of a number of peripheral blood cells (Vostal, J. G. et al.,
2001). There is a wide variation, however, between these studies in the number of
PrPc molecules found on each type of blood cell (Table 2). For example, two studies
reported that the majority of PrPc was associated with platelets with minimal levels in
red blood cells (Barclay, G. R. et al., 1999; MacGregor, I. et al., 2000). In contrast,
Holada et al reported only 44.9% associated with platelets and over 50 % to be
associated with red blood cells (Holada, K. et al., 2000) These differences have been
attributed to a number of factors including choice of antibody(Ab), method of
detection and the type of reference beads used for calibration (Vostal, J. G. et al.,
2001).
There are documented similarities between platelet secretion and neuronal
exocytosis (Reed, G. L. et al., 2000). Neurons have both small and large dense core
granules, which bear some resemblance to the alpha and dense granules of platelets.
Indeed, dense granules have previously been used as a model to study serotonin
uptake in neurons (Pletscher, A., 1986; Da Prada, M. et al., 1988). Since human
platelets have been reported to contain high proportions of PrPc (Table 2),
understanding PrPc function in platelets may also give insight into its function in
neurons.
Previous studies have shown that PrPc is present on both internal and external
platelet membranes, and that it relocates to the surface, along with other granule
membrane proteins, when the platelet is activated (Holada, K. et al., 1998).
25
Although the exact location of internal PrPc has yet to be determined, the alpha
granule membrane seem to be a good candidate as PrPc, like the alpha granule
membrane protein P-selectin, is expressed on the surface of activated platelets.
26
Table 2. Distribution of cellular prion protein in blood (% whole blood)
Barclay et al(1999) MacGregor et al (2000) Holada et al (2000) Platelets 96.3 84.2 44.9 RBCs 0 5.7 53.7 Mononuclear 2.8 7.6 1.3 cells Granulocytes 0.9 2.5 0.1
27
1.9 Platelets
Platelets are anucleate blood cells produced by differentiation of bone marrow
derived megakaryocytes (Italiano, J. E., Jr. et al., 2003b). Circulating platelets have a
lifespan of around 10-12 days and are removed primarily by the spleen. Resting
platelets are discoid in shape with an equatorial diameter of 2-3 microns (Figure 4).
1.9.1 Plasma membrane proteins
The platelet plasma membrane contains a number of membrane spanning
proteins including members of the serpentine, integrin, the leucine rich glygoprotein,
the selectin, the quadraspan and the immunoglobulin supergene families. These
proteins have a variety of functions including acting as receptors for various platelet
agonists and adhesion molecules (Table 3) (Kunicki, T. J., 1989).
The open canallicular system (OCS) is continuous with the plasma membrane
and is thought to be the channel through which internal platelet granules release their
contents upon cell activation.
1.9.2 Platelet Granules
Alpha granules are the largest, and the most numerous, of the platelet
granules. They contain a number of adhesive molecules, coagulation factors and
growth factors. Fibrinogen, fibronectin and thrombospondin are also present in alpha
granules, as are platelet factor 4, platelet derived growth factor and β-
thromboglobulin. The alpha granule membrane contains a number of adhesive
receptors which are expressed on the platelet surface after activation, consistent with
a role for alpha granule membrane receptors in platelet adhesion (Wencel-Drake, J.
D. et al., 1986).
28
Figure 4. Normal resting platelet
Normal resting platelet. PM – plasma membrane, MT – microtubules,
AG – alpha granules, DG – dense granules, OCS – open canallicular system. (bar =
1µM)
PM
DG
MT
OCS
AG
29
Table 3. Main glycoprotein receptors on platelet plasma membrane Agonist Receptor Integrin Fibronectin GPIc/Iia α2β5
Collagen GPIa/IIa α2β1
Thrombospondin GpIV
Thrombin GpV GPIb
Fibrinogen GPIIb/IIIa αΙΙββ3
Laminin GpIIa/Ic α6β1
von Willebrand factor GPIb/IX
30
These receptors include CD62 (P-selectin, GMP-140,), which plays a role in
the interaction of platelets with neutrophils (McEver, R. P. et al., 1984), GMP-33,
which has been reported to be an N-terminal proteolytic fragment of thrombospondin
(Damas, C. et al., 2001) and Gn24, a GTP binding protein thought to be involved in
exocytosis (van der, Meulen J. et al., 1991).
Dense granules are smaller, and less numerous (3-8 per platelet), than alpha
granules and contain serotonin, ATP, calcium and ADP (Rendu, F. et al., 2001). The
presence of calcium in these granules imparts an electron dense appearance when
viewed with a transmission electron microscope. The release of serotonin and ADP
following platelet activation promotes further platelet aggregation in a positive
feedback manner. The membranes of dense granules have been shown to contain P-
selectin (Israels, S. J. et al., 1992), the tyrosine kinase pp60src (Rendu, F. et al.,
1989), as well as two almost identical proteins granulophysin (Gerrard, J. M. et al.,
1991; Nishibori, M. et al., 1993) and CD63 (a lysosomal membrane protein)
(Metzelaar, M. J. et al., 1991). Each of these proteins has been shown to relocate to
the surface of activated platelets.
Lysosomal granules contain a number of lysosomal enzymes which have been
suggested to play a role in clot lysis, and clearing of thrombi (Van Oost, B. A. et al.,
1985). In addition to CD63 mentioned above, lysosomal granule membranes also
contain the lysosomal-associated membrane proteins LAMP-1 and LAMP-2
(Silverstein, R. L. et al., 1992). Interestingly LAMP-2 has also been identified to be
present in platelet dense granule membranes (Israels, S. J. et al., 1996).
31
The presence of peroxisomes in platelets remains contentious. Parmley et al
suggest that they are in fact dense granules (Parmley, R. T. et al., 1974), whereas
Breton-Gorius and Guichard demonstrated that they were distinct granules and
unrelated to dense granules (Breton-Gorius, J. et al., 1975).
1.9.3 Tubular Systems
In addition to the open canalicular system, there are two other tubular systems
within platelets. The first is a band of microtubules that encircles the platelet just
below the plasma membrane. Formed from a single coiled microtubule from the
megakaryocyte (Italiano, J. E., Jr. et al., 2003a), it is thought to be involved in platelet
shape change in response to agonists.
The second system is the dense tubular system which is thought to be the main
calcium sequestration and thromboxane synthesising organelle within platelets
(Ebbeling, L. et al., 1992).
1.9.4 Platelet Function
The function of platelets is to form a primary hemostatic plug at the site of
vessel injury (Gerrard, J. M. et al., 1976). This is a multi-step process involving
platelet adhesion to the damaged vessel wall and subsequent aggregation to form the
haemostatic plug. The initial stimulus is collagen which is exposed by the injured
vessel wall and binds to receptors on the platelet membrane (Gerrard, J. M., 1988).
This interaction of collagen with its receptor triggers a series of events including
platelet shape change, associated with centralization of organelles and extension of
pseudopods, the exposure of adhesive receptors, granule exocytosis, and the synthesis
32
and secretion of the pro-aggregatory eicosanoid thromboxane A2 (Gerrard, J. M. et
al., 1993).
Furthermore activated platelets release microvesicles (Figure 5).
Microvesicles are derived from the plasma membrane, range in size from 100nm to 1
micron and are enriched in the anionic phospholipid phosphatidylserine which serves
as the platform for both the VIIIa/IXa and the Va/Xa complexes in the coagulation
cascade (Heijnen, H. F. et al., 1999). It is therefore believed that microvesicles play a
procoagulant function (Siess, W., 1989; Fox, J. E. et al., 1990).
There is some evidence suggesting that, in addition to microvesicles, platelets
also release exosomes (Heijnen, H. F. et al., 1999). Ranging in size between 40-
100nm, exosomes are smaller than microvesicles and are thought to be derived from
both alpha granules and multivesicular bodies; the latter of which are present in
platelets and megakaryocytes and are believed to represent an intermediate stage in
alpha granule formation (Heijnen, H. F. et al., 1998).
The precise function of platelet-derived exosomes remains to be determined
although the low binding of factor X, prothrombin and annexin V to their surface
suggests that they do not have the same pro-coagulant activity as microvesicles. The
presence of the tetraspan protein CD63 on their surface however points to a possible
role for exosomes in cell adhesion, since CD63 molecules are involved in adhesion of
neutrophils to endothelial cells (Toothill, V. J. et al., 1990).
33
Figure 5. Electron Microscopy: Platelet Activation
p
Granule centralisation, extrusion of contents and formation of microvesicles (mvs) (Bar=500nm)
Quiet platelet – discoid alpha granule (a) dense granule (d) open canallicular system (ocs)
Activated platelet – shape change, formation of pseudopods (p)
a
d
ocs mvs
34
1.10 Objectives
As outlined above, PrPc is present on both internal and external platelet
membranes and relocates to the surface following platelet activation (Holada, K. et al.,
1998). However the role PrPc, if any, in normal platelet function has not been addressed.
The purpose of this work is therefore twofold:
• to determine the internal location of PrPc within platelets.
• to determine if PrPc is involved in normal platelet function mechanisms.
35
2. MATERIALS AND METHODS
2.1 Antibodies and reagents
2.1.1 Anti-PrP antibodies
The following anti-PrP Abs were used. The final concentrations or dilutions used
are indicated in brackets.
• Monoclonal Ab 308 (3 µg/ml), raised against amino acids (aa)106-126 of human PrP,
was purchased from Cayman Chemical Co. Ann Arbor, MI
• Monoclonal Ab 6H4 (1/1000 dilution), raised against aa 144-152 human PrP was
purchased from Prionics, Switzerland
• Monoclonal Ab 3F4 (1/1000 dilution), raised against aa 108-111 human and hamster
PrP, was purchased from DakoCytomation, Mississauga, Ontario
• Monoclonal Ab F89/160.1.5 (5µg/ml) raised against aa 142-145 ovine PrP was a
generous gift from Dr. K. O’ Rourke , USDA, Pullman, WA
• Monoclonal Ab F99/97.6.1 (5µg/ml)raised against aa 220-225, was a generous gift
from Dr. K. O’ Rourke USDA, Pullman, WA
• Polyclonal Ab 703 (1/2000 dilution), raised against full length bovine PrP, was
purchased from Abcam Inc. Cambridge, MA
2.1.2 Other antibodies
• Anti-CD62P and anti CD63 Abs (1:100 dilution) were purchased from Abcam Inc.
Cambridge, MA
• Anti calreticulin (1:100 dilution) was purchased from Stressgen Biotechniologies,
Victoria B.C.
36
• Anti fyn (1:100 dilution) was purchased form BD Biosciences, Mississauga,Ont
• Anti- laminin receptor (1:100 dilution)was purchased form Neomarkers, Fremont, CA
• CD9-FITC mouse monoclonal (1:50 dilution) was purchased from DakoCytomation,
Mississauga, Ontario
2.1.3 Reagents
Alexa fluor 488 (10ug/ml) was obtained from Molecular Probes, Eugene,
OR.,Texas Red conjugated secondary Ab (10ug/ml) was from Vector laboratories,
Burlingame,CA, and horseradish peroxidase conjugated secondary Ab (1:1000 dilution)
from DakoCytomation, Mississauga, Ontario
Thromboxane analogue U46619 was from Cayman Chemical Ann Arbor, MI, and
equine tendon collagen was from Helena Laboratories, Beaumont, TX.
The “Seize” primary immunoprecipitation kit and the “Super Signal West
Pico”chemiluminescence kit were obtained from Pierce Chemical Company, Rockford,
IL. Hyperfilm was purchased from Amersham, Piscataway, NJ, and nitrocellulose
membranes, and protein assay kits, were purchased from Bio-Rad, Mississauga, ON.
Secondary Abs conjugated to gold, bovine thrombin, metrizamide, protease inhibitors
and all other reagents were from Sigma-Aldrich Canada Ltd, Oakville, ON, and were of
the highest grade available.
2.1.4 Buffers
All buffers used are listed in appendix A.
2.2 Preparation of washed platelets
Blood from healthy donors, who had refrained from taking aspirin in the previous
two weeks, was drawn into 20 ml syringes containing ACD anticoagulant in the ratio of
37
8.1ml blood to 1.9 ml of ACD (Aster, R. H., 1972). The whole blood was transferred to
15 ml polypropylene tubes and centrifuged at 800 x g for 5 minutes at room temperature.
The resultant platelet rich plasma was transferred to a fresh tube and centrifuged at 800 x
g for a further 15 minutes to pellet the platelets. The supernatant was removed and the
platelet pellet re-suspended in an appropriate buffer (McNicol, A. et al., 1997).
2.3 Preparation of whole platelet homogenates
Platelet rich plasma, prepared as outlined above, was transferred to a fresh
polypropylene tube, 2mM EDTA was added and centrifuged at 800 x g for 15 minutes at
4oC to pellet the platelets. The pellet was re-suspended in Hepes/Tyrodes buffer and
centrifuged at 800 x g for 15 minutes at 4oC. The washed platelet pellet was re-suspended
in TBS to a volume of 1ml of buffer per 10ml of platelet rich plasma. After the addition
of protease inhibitors (10µg/ml leupeptin, 1mM PMSF, 1mM benzamidine), the platelet
suspension was dispensed into 200µl aliquots and stored at –80oC until use.
2.4 Fractionation of platelets
Platelet rich plasma prepared from 60ml blood, as outlined above, was transferred
to a fresh tube, 2mM EDTA added, and centrifuged at 800 x g for 15 minutes at room
temperature to pellet the platelets. The platelet pellet was re-suspended in wash buffer,
and any remaining red blood cells were removed by a brief (10 second) centrifugation at
800 x g. The supernatant was transferred to a fresh tube and the red blood cell pellet was
discarded. This step was repeated until the platelet preparation was free of red blood cell
contamination. The platelets were centrifuged at 800 x g for 15 minutes and the resultant
pellet re-suspended in homogenizing buffer containing protease inhibitors (1mM PMSF,
10µg/ml leupeptin, 1mM benzamidine). The platelets were sonicated for three 15-second
38
pulses at 4°C and the lysate layered on to a 40% metrizamide gradient (Rendu, F. et al.,
1982). The gradient was centrifuged at 54,000 x g for 30 minutes at 4oC and the alpha
granule enriched layer above the gradient was removed to a fresh tube. The metrizamide
was removed and the dense granule-enriched pellet was washed with homogenizing
buffer and finally re-suspended in Tris buffer pH 7.4. Both granule-enriched fractions
were stored at –80oC until use.
2.5 Protein Determination
The protein concentrations of the samples for electrophoresis were determined by
a Bio-Rad protein assay, following the manufacturer’s instructions. An aliquot (4µl) of
the sample was mixed with 800µl of water and 200µl of Bio-Rad protein assay reagent
added. The samples were mixed and absorbance at a wavelength of 595nM measured on
a Gemini max spectrophotometer. Protein concentrations were determined from a
standard curve made of varying concentrations of BSA.
2.6 Electrophoresis and Immunoblotting
Samples (10 –15 µg of protein) were mixed with 3X SDS reducing sample buffer
in the ratio 2:1 sample:buffer, denatured by incubation at 100°C for 5 minutes and the
proteins separated on a 12% SDS polyacrylamide gel (175 volts for 1 hour.).
The proteins were transferred to 0.45µΜ? nitrocellulose using a semi-dry transfer
system (Invitrogen; 20 milliamps per gel for 1 hour). The transferred proteins were
visualised by staining in 0.1% ponceau S for 30 seconds followed by rinsing in water.
The nitrocellulose was incubated in non-fat powdered milk (5% in TBS-T) for 30
minutes at room temperature, to block non-specific binding, and subsequently incubated
in the appropriate Ab for 60 minutes at room temperature. Following 3 x 5 minute washes
39
in TBS-T, the nitrocellulose was incubated with an appropriate, horseradish peroxidase-
conjugated secondary Ab for 30 minutes at room temperature. The nitrocellulose was
washed (6 x 5 minutes in TBS-T) and incubated for 5 minutes in Super Signal West Pico
chemiluminescence system. Following drying, the bands were visualised by exposure to
Hyperfilm or by digital imaging on the Fluor–S Max multimager (Bio-Rad). Multiple
exposures were made to optimise the intensity.
2.7 Immunofluorescence
Platelets were washed into Hepes/Tyrodes buffer, as described above, and
aliquots (700µl) dispensed into eppendorf tubes. The platelets were stimulated with 1
unit/ml of thrombin (Sigma) over a time course of 0, 30, 60, and 120 seconds. The
reaction was stopped, and fixed, by the addition 700µl of 4% paraformaldehyde in
phosphate buffered saline (PBS) for 30 minutes. Some unstimulated platelet samples
were permeabilised, after fixation, by incubation with 0.2% Triton-X 100 for 3 minutes
prior to the addition of excess PBS.
In all case, samples were washed x 3 in PBS with 0.1% BSA, incubated in the
appropriate primary Ab diluted in PBS/0.1% BSA overnight at 4oC, re-washed x 3 in
PBS/0.1% BSA and incubated in the appropriate secondary Ab conjugated to either
Alexa Fluor 488nm or Texas red for 30 minutes at room temperature in the dark.
Following a further washing x 3 in PBS the samples were re-suspended in 90%
glycerol/ 10% PBS. A drop was placed on a slide and allowed to settle for 30 minutes
before being visualised on an Olympus IX70 microscope with confocal attachment
(Carsen Group, ON).
40
For double labeling experiments, the samples were washed in PBS with 0.1%
BSA after labeling with the first fluorochrome then incubated in the second Ab, washed
x3 in PBS with BSA then labeled with the second fluorochrome.
2.8 Mepacrine labeling of dense granules
Platelet dense granules were labeled with mepacrine according to a method by
McNicol et al (McNicol, A. et al., 1994) Briefly, 1µl of 55mM mepacrine was added to
every 1ml of platelet rich plasma and incubated at 37oC. ACD was added in the ratio of
1.9ml ACD: 8.1 ml platelet rich plasma and the platelets were pelleted by centrifugation
as above. The platelet pellet was re-suspended in Hepes/Tyrodes buffer.
For double labeling experiments, aliquots of the mepacrine labeled platelets were
fixed by addition of an equal amount of 4% paraformaldehyde in phosphate buffer and
allowed to fix for 10 minutes. After fixation the platelets were washed in PBS /0.1% BSA
then incubated in the appropriate primary Ab as described in section 2.7, re-washed in
PBS/0.1% BSA then incubated in the appropriate secondary Ab conjugated to Texas Red.
The samples were further washed in PBS/0.1% BSA and re-suspended in 90%
glycerol/105 PBS and visualized as above.
2.9 Flow cytometry
Platelets were washed into Hepes/Tyrodes buffer as described above. The washed
platelets were incubated with 1 unit/ml of thrombin, or saline control, for various times
(0, 30, 60, 120 seconds) and the reaction terminated by addition of an equal volume of
4% paraformaldehyde in PBS. The platelets were allowed to fix at room temperature for
30 minutes, then centrifuged at 10,000 x g for 2 minutes and the paraformaldehyde
removed. The pellet was washed x 3 in PBS with 0.1% BSA and centrifuged as above.
41
The platelets were re-suspended in an appropriate primary Ab and incubated for 60
minutes at room temperature (or overnight at 4oC), washed x 3 in PBS/0.1% BSA as
above, and incubated in a flourescein isothiocyanate (FITC)-conjugated secondary Ab
(30 minutes; room temperature). All FITC incubations were performed in the dark. The
samples were finally washed x 3 in PBS and re-suspended in PBS. Flow cytometry was
carried out on a Becton Dickinson FacsCalibur flow cytometer with forward and side
scatter set to a logarithmic scale.
2.10 Platelet microvesicle release
Following flow cytometric analysis an area was drawn around the unstimulated
platelet sample according to the forward and side scatter properties, and labeled platelet
region (P). A second region was drawn to gate on smaller, more granular particles and
labeled microvesicle (MVS) region. The percentage of samples from each region was
calculated and the number of fluorescent labeled events in each region determined
(Heijnen, H. F. et al., 1999)
2.11 Microvesicle and Exosome preparation
Microvesicles and exosomes were prepared according to a method by Heijnen
and colleagues (Heijnen, H. F. et al., 1999). Briefly, platelets were washed into
Hepes/Tyrodes buffer and divided into 2ml aliquots. Pefabloc (4µM) was added to the
buffer to prevent endogenous protease digestion, and 5?µM RGDS peptide added to
prevent platelet clumping. Platelets were incubated with either 1unit/ml thrombin or
saline control, for 120 seconds and the reaction stopped by the addition of an equal
amount of ice-cold ACD. Samples were centrifuged at 800 x g for 15 minutes to obtain
the platelet pellet fraction (P). The supernatant was further centrifuged at 10,000 x g for
42
30 minutes to obtain the microvesicle fraction (MV). The supernatant from the MV
fraction was centrifuged at 100,000 x g for 60 minutes to pellet the exosomes (EX). All
pellets were re-suspended in PBS. Samples were used for either immunoblotting or
electron microscopy.
To ensure that exosomes were being produced, the supernatant from the 10,000 x
g spin was mixed with 2M sucrose, and a 0.8-0.25M sucrose gradient was layered on top.
This gradient was centrifuged at 65,000 x g for 16 hours and 500µl fractions were
collected. These fractions were diluted in 4.5ml PBS and centrifuged at 100,000 x g for
60 minutes at 4oC. The pellets were resuspended in PBS for slot blots or electron
microscopy.
2.12 Slot Blots
Pellets from the sucrose gradient fractions were mixed with 30 µl of PBS and
10µl was bound to a nitrocellulose membrane using a Schleicher and Schuell slot blot
apparatus. The membrane was incubated in 5% non-fat powdered milk in TBS-T to block
non-specific binding then incubated in the appropriate Ab diluted in TBS for 60 minutes
at room temperature or overnight at 4oC. Following washing in TBS-T (3x5 minutes),
the membrane was incubated in the appropriate secondary Ab conjugated to horseradish
peroxidase for 30 minutes at room temperature. The membranes were washed in TBS-T
(6 x 5 minutes), incubated in Super Signal West Pico chemiluminescence substrate and
visualised on a Bio-Rad Fluoro S Max imaging system. Densitometry was performed
using Quantity One software (Bio-Rad). The presence of exosomes in these fractions was
confirmed by staining with a monoclonal Ab to CD63.
43
2.13 Electron microscopy and immunocytochemistry
2.13.1 Resin sections of whole platelets
Platelet pellets were re-suspended in Hepes/Tyrodes buffer as described above.
The washed platelets were incubated with 1 unit/ml of thrombin, or saline control, for
various times (0, 30, 60, 120 seconds) and the reaction terminated at by the addition of an
equal amount of 0.1% glutaraldehyde in White’s Saline. The samples allowed to fix for
10 minutes at room temperature then processed for electron microscopy according to the
protocol in Appendix B.
Sections (80 nm) were prepared on a Leica ultramicrotome and picked up on 300
mesh copper grids. Grids were stained in lead citrate for 10 minutes, washed in distilled
water and dried before being examined in a Tecnai electron microscope.
2.13.2 Whole mounts of sucrose gradient fractions
Pellets from the sucrose gradient fractions were re-suspended in 30µl of PBS as
described above and adsorbed onto 300 mesh formvar coated nickel grids for 10 minutes.
Excess PBS was blotted with filter paper and the grids were fixed with 2%
paraformaldehyde in PBS for 10 minutes. After fixation, the grids were washed in PBS
with 1% BSA and stained for immunocytochemistry.
2.14 Antibody staining
Grids for immunocytochemistry were incubated in PBS with 1% BSA for 15
minutes then incubated in the appropriate Ab for 30 minutes. The grids were washed 3 x
5 minutes in PBS with 0.1% BSA and incubated in the secondary Ab conjugated to 10nm
gold for 30 minutes. After gold labeling, the grids were washed 3 x 5minutes in PBS
44
followed by washing x 3 in deionised water. The grids were then stained in 3% uranyl
acetate, dried and examined in a Tecnai transmission electron microscope.
2.15 Platelet Aggregation Studies
Platelet pellets were re-suspended in Hepes/Tyrodes buffer as described above.
Aggregation studies were carried out on a Peyton Aggregometer (McNicol, A. et al.,
1993) Briefly, platelet aliquots (400µl) were stimulated with the thromboxane A2
analogue U46619 or collagen. In some studies anti-PrP Abs were added to the platelet
samples and incubated at 37oC for 5 minutes before addition of agonist.
2.16 Platelet Adhesion Assays
Flat bottomed microtitre plates (96 well) were coated with fibrinogen, laminin,
collagen or fibronectin (each at 10mg/ml). Coated wells were incubated with 1%BSA in
calcium free Hepes/Tyrodes buffer for 90 minutes at room temperature before being used
for adhesion assays. Platelets, washed into Hepes/Tyrodes buffer as described above,
were incubated with various concentrations of anti-PrP Abs for 5 minutes. Platelets
(200µl) were added to the wells and allowed to settle for 15 minutes. The wells were
washed with PBS, incubated with anti-CD9, a platelet plasma membrane marker,
conjugated to FITC for 30 minutes. The wells were washed twice in PBS and
fluorescence determined on a Gemini Max fluorospectrophotometer with excitation and
emission set to 495nm and 530nm respectively. BSA was used as a negative control and
the background counts were subtracted from the samples. Platelets incubated with anti-
PrP Abs were compared controls that had no Ab added.
45
2.17 Immunoprecipitations
Immunoprecipitations with anti-PrP monoclonal Ab 6H4 were carried out using a
“Seize” primary immunoprecipitation kit following the manufacturer’s instructions.
Platelets, in Hepes /Tyrodes buffer, were lysed using the mammalian cell lysis buffer
provided. The lysates were then added to a column containing 6H4 Ab bound to
sepharose beads, rocked overnight at 4oC and the proteins in the eluate separated on a
10% SDS PAGE gel, transferred to nitrocellulose and immunoblotted with Abs raised
against to laminin receptor, fyn and calreticulin.
46
3 RESULTS
3.1 Cellular prion protein is present on platelet alpha granule membranes
Immunofluorescence staining of quiescent, non-permeabilised platelets with anti-
PrP Abs demonstrated the presence of PrPc on the plasma membrane (Figure 6A, 6B).
Following platelet stimulation with 1 unit/ml of thrombin there was an apparent increase
in the surface levels of PrPc (Figure 6C, 6D).
Similar studies in permeabilised platelets which allowed access of detecting Abs
to internal organelles showed the presence of PrPc on internal membranes (Figure 7A,
7D). Double immunolabeling, confocal microscopic studies of permeabilised platelets co-
localised PrPc with CD62 (Figure 7A, 7B, 7C), an acknowledged alpha granule
membrane marker (Furie, B. et al., 2001) but not CD63 (Figure 7D, 7E, 7F), which is
present on the membranes of dense granules and lysosomes (Nishibori, M. et al., 1993)
Platelets were incubated with mepacrine which is actively and selectively
accumulated by dense granules, and subsequently double stained with anti-PrP or anti
CD62 Abs. Visualization, using a secondary Ab conjugated to Texas Red, confirmed the
absence of PrPc on the membranes of dense granules (Figure 8).
Immunoblotting of platelet sub-cellular preparations with an anti-PrP Ab, was
consistent with the presence of PrPc in whole platelet homogenates and in the alpha
granule-enriched fraction, but not in the dense granule-enriched fraction (Figure 9).
Taken together these data are consistent with the presence of PrP in alpha granule, but not
dense granule, membranes.
47
Figure 6. Immunofluorescent staining of PrPc in quiet and activated intact platelets
Immunofluorescent staining of quiet (panels A and B) and activated (panels C and D)
platelets with Ab 308 showed presence of PrPc on the plasma membrane (large arrow).
Following activation with 1 unit/ml of thrombin, the internal stores of PrPc relocated to
the outside of the cell (small arrow). Panels A and C – light microscopy, panels B and D
Ab 308 followed by secondary antibody conjugated to FITC.
A B
C D
48
Figure 7. Double immunofluorescent staining of PrPc, CD62 and CD63 in resting permeabilised platelets
PrPc CD62 Merge
PrPc CD63 Merge
B C
D E F
A
49
Figure 7. Double immunofluorescent staining of PrPc, CD62 and CD63 in resting permeabilised platelets Top row: Permeabilised platelets stained with anti-PrP Ab 308 and visualized using a
secondary Ab conjugated to AF488(A), anti-CD62 Ab , visualized using a secondary Ab
conjugated to Texas Red (B), overlay of A and B (C). Bottom row: Stained with the
anti-PrPc Ab 308 and visualized using a secondary Ab conjugated to AF488 (D), stained
with anti-CD63 Ab and visualized using a secondary Ab conjugated to Texas Red
(E),overlay of D and E (F) (n=3).
50
Figure 8. Double immunofluorescent staining of dense granules, PrPc and CD62
positive granules in resting permeabilised platelets
A: Platelets labeled with mepacrine were subsequently stained with anti-PrP Ab3F4 and
visualized using a secondary Ab conjugated to Texas Red. The dense granules are
stained bright green (arrows), and the PrPc labeled granules are stained red. No co-
localization was seen with dense granules and PrPc.
B: Platelets labeled with mepacrine were subsequently stained with anti-CD62 Ab and
visualized using a secondary Ab conjugated to Texas Red. The dense granules are
stained bright green (arrows), and the CD62 positive alpha granules are stained red. No
co-localization was seen with dense granules and CD62
A B
51
.Figure 9. Western blot of platelet fractions
(A) Western blotting with anti-PrP Ab 308 revealed a 35-40 kDa band in the whole
homogenate (H) and alpha granule enriched fraction (A) but not the dense granule
fraction (D)
(B) Blotting with anti-CD63 Ab showed enrichment of this protein in both the alpha
granule enriched fraction and the dense granule fraction. The presence of CD 63 in the
alpha granule enriched fraction could be due to the presence of lysosomal granules in this
fraction or the retention of some dense granules above the metrizamide gradient. (n=2
- PrP
H A D
H A D
- CD63
A
B
52
3.2 Cellular prion protein translocates to the surface, and is released, on platelet
activation.
Immunoblotting of lysates from thrombin stimulated platelets with an anti-PrP Ab
showed a decrease in PrPc levels over time (Figure 10), although there was significant
donor variability in the time course of the loss of PrPc. Densitometric analysis of the
representative blot showed that, prior to thrombin stimulation, the density of the pellet
sample was 1.75 ODunits/mm2. This fell to 0.8 ODu/mm2 at 120 secs following
activation. This decrease in platelet-associated PrPc from these activated platelets was
accompanied by a corresponding accumulation of PrPc in the releasate (Figure 11).
3.3 Thrombin causes the release of microvesicles from platelets
Thrombin-activated platelets caused the release of microvesicles, determined by
flow cytometry, as has been previously reported (Heijnen, H. F. et al., 1999). Under
resting conditions most of the events fall within the platelet region, however the number
of events in the microvesicle region increased over time (Figure 12). There was
significant donor variability in the time course with maximum release observed between
60 and 300 sec.
Analysis of the flow cytometry event numbers in the representative experiment
(Figure 12) showed that in resting platelets, 0.8% of the total events corresponded to
microvesicles. Following stimulation by thrombin the percentage of microvesicles
increased to 12% of total events at 60 seconds, and to 43.6% of total events at 120
seconds. There was a corresponding decrease in the number of events in the platelet
region falling from 98% at 0 seconds thrombin to 80% at 60 seconds and finally 45.5% at
120 seconds of thrombin stimulation.
53
Figure 10. Western Blot of platelet pellets and supernatants
0 60 30 120 0 30 60 120
PrPc -
Pellet Supernatant
Platelet pellets and supernatants from a timecourse of thrombin stimulation. Blotting with anti-PrP Ab 308 confirmed that
PrPc was being released from activated platelets in a time-dependent manner. The supernatants from the activated platelet
samples were spun at 200,000 x g and the pellets blotted with Ab308. By 120 seconds stimulation, the intensity of the PrPc
band from the platelet pellet had diminished to about 50% of the other samples. PrPc was detected in the corresponding
supernatant at 120 seconds. (n=3)
54
Figure 11. Densitometry on activated platelets and supernatants.
prion protein densitometry
0
0.5
1
1.5
2
2.5
0 30 60 120
Time thrombin (seconds)
Den
sity
OD
u/m
m2
platelet pellet
supernatant
Densitometry was performed on immunoblots of pellets and supernatants from the timecourse of thrombin stimulated
platelets in figure 9
There was a slight decrease in density from the 60second pellet sample, but by 120 seconds the density of the pellet had
fallen from 1.75 ODu/mm2 in the unstimulated sample to 0.8 ODu/mm2. There was a corresponding rise in the density
of the 120second supernatant sample, showing that PrPc was being released to the supernatant.
55
0s thrombin 60s thrombin 120s thrombin
P
MVS
A timecourse of thrombin stimulation was analysed using a forward scatter vs. side scatter dot plot. Gates
were drawn around the platelet (P) and microvesicle (MVS) fractions. At 0 seconds thrombin the MVS
region constituted 0.8% of the total events counted. By 60 seconds this percentage has risen to 12%, and by
120 seconds had increased to 43.6% of total events. (representative of 8 experiments)
Figure 12. Flow cytometry of activated platelets
56
3.4 Cellular prion protein is present in microvesicles and exosomes
Flow cytometry using an anti-PrP Ab demonstrated that the fluorescence intensity
of PrPc was 37.7% in unstimulated platelets, consistent with the presence of PrPc on the
platelet surface. Following thrombin stimulation this increased to 67.8 % at 30 seconds,
then fell to 42.9% at 60 seconds (Figure 13A). As outlined above, thrombin stimulated
the release of microvesicles although the level of PrPc on the surface of the microvesicles
remained relatively constant (Figure 13A). This suggests that the PrPc remains on the
surface of the microvesicles.
A similar pattern of results were seen in platelet samples labeled with Abs
directed against CD62 and CD63 (Figure 13B,C). As expected the there was cons iderably
lower surface levels of CD62 and CD63, than of PrPc, in the unstimulated platelets.
Immunogold labeling of a whole mount of thrombin-stimulated platelets
demonstrated the presence of PrPc around the periphery of the cell and at the tips of
pseudopods. In addition there is some evidence for the presence of PrPc on small vesicles
released from platelets (Figure14).
Previous studies have shown that, in addition to microvesicles, thrombin
stimulates the release of exosomes from platelets (Heijnen, H. F. et al., 1999). Given the
relatively low levels of PrPc on the surface of released microvesicles (Figure 13A), the
possible association of PrPc with exosomes was therefore examined.
57
Figure 13A. Flow cytometry of activated platelets
At 0 seconds of thrombin, 98.6% of total events fell within the platelet gate. This number
decreased over time until at 60 seconds thrombin only 56.8% of events fell within this
gate. There was a corresponding increase in events in the microvesicle gate, rising from
2% at 0 seconds thrombin to 37.5% of total events at 60 seconds thrombin.
Fluorescence intensity of PrPc was at 37.7% in unstimulated platelets.This amount
increased to just below 67.8% at 30 seconds, but fell to 42.9% at 60 seconds of thrombin
stimulation. Surface PrPc on microvesicles remained relatively constant between 3.9-
4%. (Representative of 3 experiments)
PrPc in platelets and microvesicles
0
20
40
60
80
100
120
0 20 40 60 80
Time(seconds)
Per
cent
platelets
PrPc positiveplateletsMVS
PrPc positiveMVS
58
Figure 13 B. Flow cytometry of activated platelets
CD62 in platelets and microvesicles
0
20
40
60
80
100
120
0 20 40 60 80 100 120 140
Time(seconds)
per
cen
t platelets
CD62 positive platelets
MVSCD62 positive mvs
97.5% of total events fell into the platelet fraction in the unstimulated sample. By
120seconds stimulation this had fallen to 51.7%. Events in the MVS fraction rose from
1.6% in the unstimulated sample to 27.2% after 120 seconds of thrombin stimulation.
Surface CD62 intensity on the unstimulated platelet fraction was 13.9%, ris ing to 62.9%
by 30 seconds then falling to 51.7% by 120 seconds. CD62 on microvesicles remained
relatively constant at between 2.4 and 3.6% (Representative of 3 experiments)
59
Figure 13C. Flow cytometry of activated platelets
CD 63 in platelets and microvesicles
0
20
40
60
80
100
120
0 50 100 150
Time (seconds)
Per
cen
t
platelets
MVS
CD63 positive platelets
CD63 positive MVS
98.3% of total events fell into the platelet fraction in the unstimulated sample. By
120seconds stimulation this had fallen to 50.6%. Events in the MVS fraction rose from
1.7% in the unstimulated sample to 49.4% after 120 seconds of thrombin stimulation.
Surface CD63 intensity on the unstimulated platelet fraction was 25.5%, rising to 87.7%
by 30 seconds then falling to 58% by 120 seconds. CD63 on microvesicles rose to only
0.38%. (Representative of 3 experiments)
60
Figure 14. Immunogold labeling of activated platelets
A: Immunogold labelling with anti-PrP MoAb 6H4 demonstrated the presence of PrPc at the periphery of the activated platelet and at
the end of a pseudopod (P)
B: Release of membrane vesicles from an activated platelet. Gold- labelled PrPc is seen on these vesicles as well as on a smaller
vesicle in the top right area of the picture (arrows).
C: Another example of PrPc seen in smaller vesicles released from activated platelets. These vesicles ranged from 40-100nm
suggesting that they are exosomes. (Bar =200nm)
P
A B C
61
Exosomes were prepared by differential centrifugation, and separation through a
sucrose gradient, of the releasate of thrombin-stimulated platelets. Slot blot analysis, and
associated densitometry, with an anti-CD63 Ab confirmed the presence of exosomes in
fractions 3 and 4, as previously reported (Heijnen, H. F. et al., 1999).
Immunoblotting using an anti-PrP Ab was consistent with the presence of PrPc in
the exosome fraction (Figure 15 A and B). There was also significant PrPc present in
fraction 1, likely representing soluble PrPc.
Transmission electron immunomicroscopy identified exosomes, which were
considerably smaller than released microvesicles, stained with CD63 (Figure 16A).
Subsequent staining with an ant-PrP Ab was consistent with PrPc on exosomes (Figure
16B).
Fraction 10 was also examined for the presence of PrPc by transmission electron
immunomicroscopy. This fraction contained PrPc on both various membrane fragments
and larger granules which, by size, are possibly microvesicles. PrPc seemed to be
polarized to one end of these larger granules suggesting a possible lipid raft association
(Figure 17).
62
Figure 15. Slot blot of sucrose gradient fractions
10000
8000
6000
4000
2000
0
PrPc
OD
U/m
m2
1 2 3 4 5 6 7 8 9 10
Fraction
22000
1000
800
600
400
200
0
CD
63O
D U
/mm
2
Fractions from the sucrose gradient in figure 13A were bound to nitrocellulose and
blotted with antip PrP and anti-CD63 MoAbs. Fractions 3 and 4, which contained large
amounts of both PrPc and CD63 corresponded to the density of exosomes and were
subsequently collected and used for immunoelectron microscopy.
Fraction 1 is most likely a soluble form of PrPc. Most of the remaining PrPc was found
in fraction 10 (pellet). Although there was a substantial amount of CD63 in the exosome
fraction, most of the CD63 was found in the membranes in fraction 10.
63
Figure 16A. CD63 Immuno electron microscopy of isolated exosomes
Presence of exosomes in fractions 3 and 4 was confirmed by staining with anti-CD63 MoAb. (A and B). Structures between 40-
100nm are exosomes labelled with 10nm gold (Larger structure in panel B is most likely a microvesicle).
A B
64
Figure 16B. PrPc Immuno electron microscopy of isolated exosomes
Immunolabelling of whole mounts of exosomes from fraction 3 and 4 with anti-PrP MoAb 3F4. PrPc was seen on released
exosomes(A). Higher magnification of single exosome (B)
50nm
A B
65
Figure 17. PrPc Immunolabeling of sucrose gradient pellet
200nm
Fraction 10 from the sucrose gradient was seen to contain various membrane fragments
and larger granules which ranged from 150-350nm in size consistent with microvesicles.
PrPc staining seemed to be polarized to one end of these granules (arrows).
66
3.5 Cellular prion protein is not involved in platelet aggregation
Platelets were incubated with several antibodies to PrP prior to stimulation by
either U46619 or collagen, each at the minimal concentration required to produce
aggregation. Initial studies suggested that anti-PrP Abs 6H4 and 3F4, but not 308,
inhibited aggregation in response to both agonists (Figure18). However, both 6H4 and
3F4 contained sodium azide which is a powerful inhibitor of platelet aggregation. Thus it
is unlikely that blocking PrPc had any effect on platelet aggregation. Furthermore neither
6H4 nor 3F4 were used in subsequent platelet func tion studies.
3.6 Cellular prion protein may be involved in platelet adhesion
Platelet adhesion to collagen, fibrinogen, fibronectin or laminin was measured
using an anti-CD42 Ab directly conjugated to FITC. Pre-incubation of platelets with
anti- PrP antibodies 308, 8G8, F89 and F99 each inhibited platelet adhesion to all four
matrices to some degree, with inhibition ranging between 15-23% (n=3) (Table 4).
3.7 Cellular prion protein co-precipitates with calreticulin
Platelet eluates from an anti-PrP Ab (6H4) column were separated by SDS-PAGE
and immunoblotted with Abs to calreticulin, the laminin receptor and fyn. Calreticulin,
but neither the laminin receptor nor fyn, was present in the eluate of both unstimulated
and thrombin-stimulated platelets (Figure 19A). Furthermore, immunoblotting of a
platelet eluate from a corresponding anti-calreticulin column demonstrated the presence
of both calreticulin and PrP (Figure 19B). Taken together these data are consistent with
PrPc associating with calreticulin in platelets.
Immunofluorescence, using anti-PrP and anti-calreticulin Abs, of permeabilised
unstimulated platelets showed the significant co- localization of PrPc and calreticulin
67
(Figure 20). However there was additional, non-PrPc-associated calreticulin present in
the platelet cytoplasm.
68
Figure 18. Platelet Aggregation
1µM U46619 3F4 6H4 308
1µM U46619 caused complete aggregation of platelets. Monoclonal antibodies to prion
protein were added at a 1/400 dilution 5 minutes before aggregation. 3F4 and 6H4
completely blocked U46619 aggregation while 308 had no effect. (n=3)
69
Table 4. Platelet Adhesion Assays
Percentage Adhesion
Antibody
Matrix
Control 308 8G8 F89 F99
Collagen 100% 81.3±5.4 80.8±6.3 81.0±4.6 85.9±6.5
Fibrinogen 100% 82.0±2.5 77.9±1.7 78.8±4.3 80.5±4.4
Fibronectin 100% 87.4±5.6 86.1±6.1 83.0±3.0 85.4±2.9
Laminin 100% 81.± 9.5 82.7± 9.8 81.8±8.0 79.8±6.5
Platelelet adhesion to collagen, fibrinogen, fibronectin or laminin was measured +/-
preincubation with various antibodies to PrP. All antibodies inhibited adhesion to some
degree. Inhibition to the various matrices ranged from 15-23% with Ab 8G8 the most
effective in inhibiting adhesion to fibronectin at 23%. (n=3)
70
Figure 19. Immunoprecipitation with antibody to PrPc and Calreticulin
A B
A. Eluates, of quiet (Q) and thrombin stimulated platelets (A), from an anti -PrP 6H4 Ab
column were blotted with anti-calreticulin (CR), -Laminin receptor (LRc) and -Fyn Abs.
Only calreticulin gave a positive signal in both quiet and activated platelets.
B. Platelets immunoprecipitated with anti-calreticulin Ab and blotted with anti -
calreticulin Ab and anti-PrP Ab 308 . Both gave a positive signal. (n=3)
Q A Q A Q A
C R LRc Fyn CR PrPc
CR -
- PrPc
- CR
71
Figure 20. Double labelling with antibodies to calreticulin and prion protei
PrPc Calreticulin Merge
Permeabilised platelets were labelled with anti- PrP Ab 308 with the secondary antibody conjugated to FITC (A), and anti-calreticulin
Ab with the secondary antibody conjugated to Texas Red (B). The yellow areas in the overlay of both images showed some co-
localization of PrPc and calreticulin (C) ( n=2)
A B C
72
4. DISCUSSION
Historically TSEs were relatively rare and obscure fatal neurodegenerative
diseases of humans and animals. However they were highlighted in the mid 1980s with
the outbreak of BSE in the United Kingdom, the cause of which was suggested to be due
to a combination of feeding meat and bone meal to cattle coupled with a change in
rendering practices. (Taylor, D. M. et al., 2003; Taylor, D. M. et al., 1997). Between
1988 and 2004 a total of 179,085 cattle were diagnosed as positive for BSE
http://www.defra.gov.uk/animalh/bse/statistics/weeklystats.html#pass. with 214,080
being slaughtered to date. This disease is not unique to the U.K.; a total of 25 countries
worldwide have reported at least 1 or more positive cases of BSE since 1987
http://home.hetnet.nl/~mad.cow/. The finding of BSE in cattle causes an immediate and
great impact on the economy of affected countries. For example, the single Canadian case
in May of 2003 caused the immediate slaughter of 2000 herd-mates and offspring, and
the closure of various borders to the import of Canadian beef had a devastating effect on
the Canadian beef industry with an estimated loss of $5billion. (Canadian Cattlemens
Association http://www.cattle.ca/ ) The recent discovery of two more cases in Western
Canada may hamper negotiations with the United States to reopen the border to
importatin of live cattle.
Another result of the BSE outbreak in the U.K. was the emergence of a variant
form of CJD which was first identified in 1996 . It was linked to the digestion of beef
contaminated with BSE (Lasmezas, C. I. et al., 2001). The CJD Surve illance Unit n the
U.K. was set up in 1990 to monitor all new cases of CJD (Ironside, J. W., 1996), and to
73
date there have been a total of 147 human deaths from vCJD
(http://www.cjd.ed.ac.uk/figures.htm).
Although the pathogenesis of CJD is still uncertain, the presence of PrPres in
peripheral lymphoid tissue led to the fear that TSEs may be transmitted though blood
transfusion (Lee, C. A. et al., 1998). In 1998, in an effort to prevent blood-borne
transmission, the U.K. adopted the practice of universal leucodepletion. The decision
arose from findings that described B lymphocytes as playing a major role in TSE
pathogenesis (Klein, M. A. et al., 1997). However the effectiveness of leucodepletion to
completely remove TSE infectivity has still to be established. Indeed, Klein et al. later
downplayed the role of B lymphocytes in TSE pathogenesis and suggested a stronger role
for follicular dendritic cells of the spleen (Klein, M. A. et al., 1998).
Although there have been no reports of classical CJD being transmitted by blood
transfusion, there have recently been two reports linking the acquisition of vCJD to blood
transfusions (Peden, A. H. et al., 2004). These reports have brought into question the
effectiveness of leucodepletion and the possibility that PrPres may be carried on blood
components other than leucocytes (Gregori L. et al., 2004).
Pathogenesis studies using PrP-/- mice have demonstrated that propagation of
PrPres cannot occur in the absence of PrPc (Brandner S.et al., 2000) and, since a number
of studies have shown PrPc to be present on platelets as well as leucocytes (Holada, K. et
al., 2000), the objective of the current study was to further investigate the role of PrPc
within platelets.
Previous studies have shown PrPc to be present on both external and internal
platelet membranes; furthermore the internal store relocates to the surface on platelet
74
activation. In addition flow cytometric studies suggested an alpha granule location for
PrPc since it relocated to the surface in a similar fashion to P-selectin, however the exact
internal location had not been determined (Holada, K. et al., 1998). In the current study,
by using a combination of imuno-blotting of platelet granule fractions and
immunofluorescent co- localization studies, we have unequivocally shown that PrPc is
located on the alpha granule membrane, but not on either dense granule or lysosomal
granule membranes. Furthermore, we have shown that, following thrombin stimulation,
PrPc relocates from these granule membranes to the plasma membranes, and is
eventually released from the platelet surface. This is, in part, agreement with an earlier
study which described a soluble form of PrPc being released from stored platelet packs
(Bessos, H. et al., 2001).
It has been shown that on activation, platelets release two types of membranous
packages; microvesicles, which are plasma membrane derived and range from 100nm to
1 µm, and smaller exosomes, which range between 40-100nm and are derived from alpha
granules and multivesicular bodies (Heijnen, H. F. et al., 1999). We therefore
investigated the possibility that PrPc was being released on exosomes and/or
microvesicles.
We initially used flow cytometry to determine the amount of PrPc that was
released on the microvesicles from thrombin-activated platelets, and although the number
of microvesicles being released from activated platelets increased, the amount of PrPc in
the microvesicle fraction did not rise above 4%. This suggested that the released PrPc
was either in a soluble form or attached to membranes or vesicles that were much smaller
than microvesicles. Since exosomes are smaller than 100nm they are beyond the limits
75
of detection by flow cytometry and it was therefore possible that some PrPc may be
released on these vesicles.
Using a sucrose gradient, we isolated exosomes from thrombin-activated platelets
and found increased amounts of PrPc present in both soluble and exosome containing
fractions. Immunoelectron microscopy confirmed the presence of PrPc in the exosome
fraction. Larger granules with smaller internal vesicles were also observed in the
releasate from activated platelets. PrPc was located on the both the external and internal
membranes of these structures. The size of these structures (approximately 500µm) and
the presence of many internal vesicles are consistent with them being multivesicular
bodies.
Therefore the current study has shown, for the first time that PrPc is released from
activated platelets on both exosomes and microvesicles.
Recent bioassays using mouse models, demonstrated infectivity in both platelets
and platelets poor plasma (Cervenakova et al., 2003). Although this study did not isolate
exosomes from plasma it is possible that exosomes were present in the platelet poor
plasma fraction and may be contributing to infectivity. Of interest, Fevrier and
colleagues have recently demonstrated that cultured neuronal cells can release PrPres on
exosomes (Fevrier, B. et al., 2004), suggesting that exosomes may be a likely source of
prion infectivity. While the presence of PrPc on exosomes released from platelets does
not necessarily indicate the presence of PrPres, the involvement of plasma exosomes in
the transmission of TSE infection from blood cannot be ruled out and is an area which
obviously merits further investigation.
76
It is widely acknowledged that the spleen is the major site for accumulation of
PrPres (Jeffrey, M. et al., 2000). The spleen is also the main sequestration organ for
senescent platelets (Berger, G. et al., 1998). It is therefore possible that any PrPres that
may be carried on platelets or released granules could contribute to the accumulated
PrPres in the spleen.
The role of PrPc in normal platelet function was also investigated. Initially the
role of PrPc in platelet aggregation was examined. Blocking PrPc with a number of
antibodies initially showed promise as two of the antibodies used completely blocked
aggregation in response to the thromboxane analogue U46619. However, further
investigation revealed that these antibodies contained sodium azide, which is itself, a
potent inhibitor of platelet aggregation. Subsequent experiments using sodium azide-free
antibodies had no effect on aggregation suggesting that PrPc was not involved in this
function.
PrPc had been previously shown to be invo lved in cell adhesion in neuronal
cells. We therefore investigated the possibility that it could also be involved in platelet
adhesion. Blocking PrPc with a number of antibodies inhibited platelet adhesion to
fibrinogen, collagen, fibronectin and laminin matrices by up to 25%. Although apparently
a low level of inhibition, it should be noted that platelets contain a number of other
receptors on their surface that are involved in adhesion imparting a degree of high
adhesive capacity and a degree of receptor redundancy. These studies suggest a possible
role for PrPc in platelet adhesion. However further studies are required to ensure that the
inhibition of adhesion is a result of the specific inhibition of PrPc rather than a non-
specific steric interference by the antibodies. In addition the effects of anti-bodies against
77
individual regions of PrPc should be used to determine the specific domain responsible
for adhesion.
A number of studies in neuronal cells have suggested that PrPc is associated with
a number of protein-protein interactions important for signaling pathways. These studies
have implicated interactions between PrPc and calreticulin, laminin receptor and fyn. In
our hands, calreticulin, but neither the laminin receptor nor fyn, co-precipitated with
PrPc. Furthermore immunofluorescence double labeling studies suggested that PrPc may
also co- localise with calreticulin. This is the first time that an association between PrPc
and calreticulin has been described in platelets.
Interestingly calreticulin has been shown to be present on the surface of platelets
(Elton, C. M. et al., 2002), and earlier studies have suggested the dense tubular system as
a possible location for calreticulin (Arber, S. et al., 1992), although the ring like structure
in activated platelets suggests a granule membrane location. In the current study PrPc
was found to be mainly present in alpha granule membrane. To date there are no reports
of a granular location for calreticulin although this could be addressed by applying the
same fractionation and double labeling techniques that we have used in this study to
characterise the location of calreticulin within platelets.
It may be also possible to elucidate a role for PrPc within platelets through its
association with calreticulin. Calreticulin is known to be a multi functional calcium
binding protein and an endoplasmic reticulum chaperone protein (Michalak, M. et al.,
2002). It has been implicated in a number of processes including glycoprotein folding
(Ellgaard, L. et al., 2003), cell adhesion and integrin dependent calcium signaling
(Coppolino, M. G. et al., 1997). In platelets, it has been associated with collagen
78
receptors (Elton, C. M. et al., 2002), and thrombospondin signaling (Goicoechea, S. et
al., 2002).
The tyrosine kinase fyn is also involved in the collagen activation pathway in
platelets (Briddon, S. J. et al., 1999), and PrPc has been shown to phosphorylate fyn
through caveolin-1 signaling in a neuronal cell line (Mouillet-Richard, S. et al., 2000). It
may also be possible that both PrPc and calreticulin are associated during collagen
signaling in platelets although investigation of this possibility warrants a more intensive
approach than this project allows.
As this work progressed it became increasingly clear that further investigation
into the various roles for PrPc within platelet function were necessary. The role that this
protein plays in cell adhesion or signal transduction alone, warrants much further
research than could be carried out here and we can only hope that this work may be done
in future projects.
5. LIMITATIONS The initial objective of these studies was to pinpoint the exact location of PrPc
within platelets and, if possible, determine a role for the protein within normal platelet
function. The localization of PrPc to platelet-derived exosomes raises the possibility that
there may be a role for platelet derived exosomes in TSE pathogenesis. Investigating this
role however is outside the scope of this project for a number of reasons:
79
§ One approach would be to isolate exosomes from the plasma of previously
infected animals and inoculate a new set of animals with these isolates.
However the long incubation times of TSE infection put this type of
experiment outside the time frame for this project. For example ME7 strain of
scrapie has an incubation time of 171 days in the C57/BL strain of mice
(Bruce, M. E. et al., 1991).
§ There is also the possibility that the exosomes, which carry infectivity in plasma
may be derived from other blood cells or endothelial cells. Like platelets,
endothelial cells have also been shown to release microvesicles which contain
PrPc (Simak, J. et al., 2002), although there have been no reports of
endothelial cells releasing exosomes.
The antibodies used for the functional studies were all raised against different
areas of the PrP peptide and therefore may have had varying degrees of effectiveness.
Since PrPc is a GPI anchored protein, an alternative approach would be to remove the
protein by incubation in phosphatidylinositol-specific phospholipase C (PILPC), which
breaks the GPI anchor. However Holada et al found that the PrPc expressed on activated
platelets was resistant to PILPC (Holada K. et al., 1998), so an alternative method of
blocking PrPc would have to be found. A possibility is to use a peptide that would bind to
PrPc thus rendering it inactive.
80
6. CONCLUSIONS
The experiments carried out in this study have enabled us to reach a number of
conclusions about the location and possible function of PrPc in human platelets.
In addition to being located on the plasma membrane, we have shown PrPc to be
located internally on the alpha granule membrane but not on dense granule or lysosomal
granule membranes. Furthermore, this internal store of PrPc relocates to and is
subsequently released from the plasma membrane when the platelet is activated with
thrombin.
The PrPc from activated platelets is released in a soluble form as well as on
microvesicles and exosomes.
As to a role for PrPc in normal platelet function, blocking PrPc with antibodies
has shown that PrPc is not involved in platelet aggregation however it may be involved in
platelet adhesion since blocking PrPc with antibodies inhibited adhesion to laminin,
collagen, fibronectin and fibrinogen.
PrPc may also play a role in signal transduction since it co-precipitates
calreticulin. However it did not co-precipitate fyn or the laminin receptor as had been
reported in other cell lines.
81
7. APPENDICES
Appendix A. Buffer solutions
ACID CITRATE DEXTROSE ANTICOAGULANT (ACD)
Trisodium citrate 86mM
Citric Acid 70mM
Dextrose 200mM
HEPES /TYRODES BUFFER
Sodium Chloride 134mM
Sodium hydrogen carbonate 12mM
Potassium chloride 2.9mM
Di-sodium hydrogen phosphate 0.6mM
Magnesium chloride 1mM
Hepes 8.4mM
Adjust pH to 7.4. Add 0.09% dextrose and 0.3% BSA before use.
TRIS BUFFERED SALINE (for whole platelet homogenate preparation)
Tris 20mM
NaCl 130mM
Adjust pH to 7.0. Add 1mM PMSF, 10µg/ml leupeptin and 1mM benzamidine before
use.
WASH BUFFER
Sodium chloride 130mM
Potasium chloride 5mM
Sodium dihydrogen orthophosphate 1mM
82
Sodium hydrogen carbonate 24mM
Dextrose 10mM
Sucrose 2.5mM
EDTA 2mM
BSA 0.7%
Adjust pH to 7.4
HOMOGENIZING BUFFER
Hepes 25mM
Potassium chloride 500mM
Magnesium sulphate 2mM
Dextrose 10mM
Sodium citrate 20mM,
Adenosine triphosphate 5mM
Sucrose 100mM
Adjust to pH 7.0. Add 1mM PMSF, 10µg/ml leupeptin, and 1mM benzamidine before
use.
METRIZAMIDE GRADIENT
Metrizamide 40%
KCl 0.35M
150mM TRIS/HCl BUFFER pH 8.8
Tris 3.3g
Deionised water 80ml
Adjust pH to 8.8 with hydrochloric acid and bring volume to 200ml with deionised water
83
50mM TRIS/HCl BUFFER pH 6.8
Tris 6g
Deionised water 80ml
Adjust pH to 6.8 with hydrochloric acid and bring volume to 100ml with deionised water
12% POLYACRYLAMIDE RESOLVING GEL ( (Laemmli, U. K., 1970)
Deionised water 3.3 ml
30% Acrylamide/bis-acrylamide 29:1 4.0 ml
150mM Tris/HCl buffer pH 8.8 2.5 ml
10% SDS 0.1 ml
1% Ammonium persulfate 0.1 ml
Temed 0.004 ml
4%POLYACRYLAMIDE STACKING GEL
Deionised water 6.8ml
30% Acrylamide/bis-acrylamide 29:1 1.7ml
50mM Tris/HCl buffer pH 6.8 1.25ml
10% SDS 0.1ml
Ammonium persulfate 0.1ml
Temed 0.01ml
4X SDS SAMPLE BUFFER
1M Tris /HCL 2.4ml
SDS 8g
80% Glycerol 2.4ml
Bromophenol blue 4mg
84
Deionised water 2.1 ml
TRIS/GLYCINE/SDS RUNNING BUFFER
Tris 25mM
Glycine 192mM
SDS 0.5.%
TRIS/GLYCINE TRANSFER BUFFER
Tris 25mM
Glycine 192mM
Methanol 20%
TRIS BUFFERED SALINE (TBS)
Tris 50mM
Sodium Chloride 138mM
Adjust pH to 8.0
TBS-T
As above but with the addition of 0.05% Tween-20
PHOSPHATE BUFFERED SALINE (PBS)
Sodium Chloride 145mM
Sodium dihydrogen orthophosphate 1.45mM
di-sodium hydrogen phosphate 17mM
Adjust pH to7.0
85
WHITE’S SALINE SOLUTION A
Sodium Chloride 240mM
Potassium Chloride 100mM
Magnesium sulphate 45mM
Calcium nitrate 60mM
Adjust pH to 7.4
WHITE’S SALINE SOLUTION B
Sodium hydrogen carbonate 12.5mM
Potasium phosphate 3mM
Phenol Red 0.002mM
Dissolve in 100ml deionised water and adjust pH to 7.4
0.1% GLUTARALDEHYDE IN WHITES SALINE
Deionised water 8.875ml
White’s saline solution A 0.5ml
White’s saline solution B 0.5ml
8% Glutaraldehyde 0.125ml
3% GLUTARALDEHYDE IN WHITES SALINE
Deionised water 5.24ml
White’s saline solution A 0.5ml
White’s saline solution B 0.5ml
8% Glutaraldehyde 3.74ml
86
1% OSMIUM TETROXIDE
Deionised water 7.0ml
0.1M Calcium Chloride 5.0ml
Potassium ferricyanide 2.5ml
4% osmium tetroxide 2.5ml
87
Appendix B. Processing schedule for transmission electron microscopy
1. Fix platelet samples by addition of an equal volume of 0.1% glutaraldehyde in
Whites saline; 10 minutes.
2. Centrifuge at 2000 x g ; 5 minutes.
3. Remove supernatant and incubate pellet in 3% glutaraldehyde in Whites saline;
60 minutes.
4. Centrifuge at 2000 x g; 5 minutes.
5. Discard supernatant and incubate pellet in 1% osmium tetroxide solution; 90
minutes.
6. Remove osmium tetroxide and wash pellet in deionised water; 2 x 5 minutes.
7. Incubate pellet in 3% uranyl acetate overnight at 4oC.
8. Wash pellet in deionised water; 2 x 5 minutes.
9. Incubate in 70% ethanol; 30 minutes.
10. Incubate in 90% ethanol; 30 minutes.
11. Incubate in 100% ethanol; 2 x 30 minutes.
12. Incubate in propylene oxide; 2 x 10 minutes.
13. Incubate in a 1:1 mixture of propylene oxide and epon /araldite resin; 60 minutes.
14. Incubate in epon/araldite resin; 2 x 60 minutes.
15. Embed pellet in fresh epon /araldite resin and cure at 60oC; 24 hours.
88
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