Prion-infection and Cellular Signaling

Influence of scrapie-infection on lipid raft-associated proteins

Hanna Gyllberg

Stockholm University

Doctoral thesis

©Hanna Gyllberg, Stockholm 2007

ISBN: 978-91-7155-518-2

Printed in Sweden by US-AB, Stockholm 2007

Distributor: Department of Biochemistry and Biophysics, Stockholm

University

Cover page: GT1-1 cells at approximately 20% confluency. Modified

by Eva Gyllberg and Jenny Wikstrand

Till Mamma och Pappa

List of Publications

This thesis is based on the following papers, which are referred to by Roman numerals (I-IV) in the text:

I Increased Src kinase level results in increased protein ty-

rosine phosphorylation in scrapie-infected neuronal cell

lines. Hanna Gyllberg, Kajsa Löfgren, Heléne Lindegren, Katarina

Bedecs. FEBS Lett. 580, 2603-2608 (2006)

II Altered membrane distribution and increased specific ty-

rosine kinase activity of Fyn in scrapie-infected

neuronal cells. Hanna Gyllberg, Kajsa Löfgren, Ana Almeida, Katarina

Bedecs. Manuscript November (2007)

III Increased levels of insulin and insulin-like growth factor-1

hybrid receptors and decreased glycosylation of the insu- lin receptor α- and β-subunits in scrapie-infected neuro-

blastoma N2a cells. Daniel Nielsen, Hanna Gyllberg, Pernilla Östlund, Tomas

Bergman, Katarina Bedecs. Biochem J. 380, 571-579 (2004)

IV Loss of LPS-induced nitric oxide production and iNOS

expression in scrapie-infected N2a cells. Heléne Lindegren, Pernilla Östlund, Hanna Gyllberg,

Katarina Bedecs. J Neurosci Res. Jan 15;71(2): 291-9 (2003)

Contents

List of Publications .......................................................................................... v

Abbreviations................................................................................................viii

Introduction .....................................................................................................1 1. Prion diseases and the prion protein .........................................................................2

1.1 Prion diseases ...................................................................................................2 1.2 The prion protein isoforms .................................................................................3 1.3 Putative functions of PrPC ..................................................................................7

2. Lipid rafts – specialized plasma membrane domains ..............................................10 2.1 The lipid raft hypothesis ...................................................................................10 2.2 Lipid composition of rafts .................................................................................11 2.3 Proteins associated with rafts ..........................................................................12 2.4 Raft function ....................................................................................................13 2.5 Distribution and trafficking of rafts....................................................................14 2.6 The lipid raft debate .........................................................................................14

3. Protein tyrosine kinases ..........................................................................................16 3.1 Non-receptor tyrosine kinases (NRTKs)...........................................................16 3.1.1 Src family kinases (SFKs).............................................................................16 3.2 Receptor tyrosine kinases (RTKs) ...................................................................21 3.2.1 Insulin and insulin-like growth factor-1 receptors ..........................................22 3.3 Protein tyrosine phosphatases.........................................................................26

4. LPS mediated production of nitric oxide ..................................................................28 4.1 The bacterial endotoxin LPS............................................................................28 4.2 LPS receptor complex and LPS signaling ........................................................28 4.3 Nitric oxide and inducible nitric oxide synthase................................................30

Methodology..................................................................................................32 Cell lines .....................................................................................................................32

GT1, N2a and BV-2 ...............................................................................................32 Scrapie-infected cell lines ......................................................................................32 Generation of scrapie-infected cell lines; ScGT1a/b and ScN2a............................33 Detection of PrPSc..................................................................................................33

Phenotypic differences between GT1 and ScGT1a/b..................................................34 Src and Fyn antibodies ...............................................................................................35 Isolation of lipid rafts ...................................................................................................36

Protein deglycosylation ...............................................................................................37 Determination of nitrite production ..............................................................................37

Results and Discussion.................................................................................38 Paper I and II ..............................................................................................................38

Increased level of active Src kinase in scrapie-infected cells (paper I)...................39 Increased specific tyrosine kinase activity of Fyn in scrapie-infected cells

(paper II) ................................................................................................................42 Increased tyrosine phosphorylation in scrapie-infected cells (paper I and II) .........43 Plasma membrane distribution of PrPC, PrPSc and active Fyn (paper II) ................45 Physiological implications (paper I and II) ..............................................................45

Paper III ......................................................................................................................47 Localization of IR in ScN2a cells and elevated expression of IR/IGF-1R hybrid

receptors in ScN2a ................................................................................................48 Altered molecular mass of the IRβ-subunit in ScN2a.............................................49

Paper IV......................................................................................................................49 Lack of LPS-induced iNOS expression in ScN2a...................................................50

Sammanfattning på svenska – Hur reagerar nervceller på prioninfektion?..52

Acknowledgements .......................................................................................54

References....................................................................................................56

Abbreviations

AG aminoguanidine BSE bovine spongiform encephalopathy CD14 cluster of differentiation 14 CJD Creutzfeldt-Jacobs disease CNS central nervous system CTXB cholera toxin fragment B DRM detergent resistant membrane ERK extracellular signal regulated kinase GPI glycosylphosphatidylinositol IGF-1R insulin-like growth factor-1 receptor iNOS inducible nitric oxide synthase IR insulin receptor LBP lipopolysaccharide-binding protein LPS lipopolysaccharide LRP/LR laminin receptor precursor/laminin receptor MAPK mitogen activated protein kinase NFκB nuclear factor κB NMDA N-methyl-D-aspartate NO nitric oxide NRTK non-receptor tyrosine kinase PK proteinase K PLC phospholipase C PP2 4-amino-5-(4-chlorophenyl)-7-(t-

butyl)pyrazolo(19)pyrimidine prnp prion protein gene PrPC cellular prion protein PrPSc scrapie prion protein PTK protein tyrosine kinase RTK receptor tyrosine kinase SH Src homology SFK Src family kinase SOD superoxide dismutase TSE transmissible spongiform encephalopathy TLR toll-like receptor WGA wheat germ agglutinin

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Introduction

Prion diseases are a group of neurodegenerative conditions occurring amongst humans and other mammals. All forms of prion diseases are to date incurable and fatal. The first prion disease to be recognized was scrapie in sheep, a problem discussed in the British parliament already in 1755 since it had great economical effects in the era of the industrial revolution. The in-fectious agent was originally considered a “slow virus”, but treatments known to inactivate viruses did not abolish infectivity (Alper, et al., 1966, Alper, et al., 1967). Surprisingly, scrapie infectivity proved to be sensitive to irradia-tion at 237 nm, the wavelength at which proteins are inactivated (Latarjet, et

al., 1970). Hence, the infective agent was named prion, a word derived from proteinaceous infectious particle (Prusiner, 1982). Today, there is convincing evidence that prions consist exclusively of the disease-causing scrapie pro-tein, PrPSc. In the 1980´s it was discovered that PrPSc is the misfolded form of an endogenously expressed cellular prion protein, PrPC.

Infectious cases of prion disease are termed transmissible spongiform en-cephalopathies (TSEs), which captured the public attention with the emer-gence of the bovine spongiform encephalopathy (BSE) or “Mad-cow dis-ease” epidemic in the 1990´s, and later with the appearance of variant Creutzfeldt-Jacob disease (vCJD) in humans, a novel form of CJD linked to dietary exposure of BSE. Prion diseases can also occur spontaneously or be inherited. All inheritable cases are with no exception results from mutations in the gene coding for PrPC.

The aim of this project has been to characterize putative cytopathological changes induced by prion infection, using in vitro cell culture models of chronically prion-infected neuronal cells.

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1. Prion diseases and the prion protein

Prion diseases are a group of fatal neurodegenerative diseases that can strike almost all mammals. Until the 1980´s, the pathogenic agent was unknown, although there were speculations of a “slow virus”. Today, it is well estab-lished that prion diseases are caused by the conformational change of the cellular prion protein (PrPC) into the disease causing isoform the scrapie prion protein (PrPSc). The diseases are characterized by accumulation of PrPSc in the brain, astrogliosis, activation of microglia cells and vacuoliza-tion of nerve cell processes resulting in the typical spongiform neurodegen-eration. This chapter covers novel research about prion diseases, the sug-gested functions of the prion protein and the models suggested for the con-version of PrPC into PrPSc.

1.1 Prion diseases

Spongiform encephalopathies are the only known group of diseases contain-ing infectious as well as genetic and sporadic cases (Prusiner, 1991) where in-fectious diseases are termed transmissible spongiform encephalopathies (TSEs). Human prion diseases include Kuru, affecting the Fore people at New Guinea through ritualistic cannibalism, iatrogenic (i), variant (v) and familiar (f) CJD, Gerstmann-Sträussler-Scheinker disease (GSS), fatal famil-ial insomnia (FFI) and fatal sporadic insomnia (FSI) (Prusiner, 1998). Iatro-genic cases, which are a subtype of an infectious form of CJD, have been documented to derive from blood transfusions, corneal- or other tissue trans-plantations or administration of pituitary gland hormones. The source of infectivity in all such cases has been individuals suffering from a non-identified human TSE (Aguzzi, 2006). GSS, FFI and fCJD are always caused by point mutations in the PRNP gene e.g. P102L, D178N and E200K respec-tively. More than 20 different point mutations in the PRNP gene have been shown to cause various types of prion diseases. Sporadic cases of CJD are the most common of all the human prion diseases however, the incidence of all different types of CJD cases combined is around 1.3/106 per year. The protein-only hypothesis of prion diseases postulates that PrPSc is the only constituent of the prion and this isoform is very resistant to cellular degrada-tion. Thus, in a person carrying such a PRNP mutation, misfolded PrPSc is continuously expressed and PrPSc accumulation in the brain is considered to be the main cause of the disease (Prusiner, 1998).

Prion diseases occurring amongst other mammals include scrapie in sheep, with one of the symptoms being to scrape off their wool (Greig, 1950). BSE in cattle and chronic wasting disease (CWD) in deer and elk popula-tions are also well documented TSEs.

Clinically, transmitted TSE pathogenesis is divided into three distinct phases: (i) infection and peripheral replication of prions, (ii) neuroinvasion

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and replication in the central nervous system (CNS), followed by (iii) neu-rodegeneration (Prusiner, 1998). Still the question remains what actually causes this neuronal cell death. A number of possible scenarios causing the neu-rodegeneration can be offered; gain of function – direct neuronal toxicity of PrPSc, loss of function – consequence of loss of PrPC function, microglial and astroglial production of neurotoxic proinflammatory cytokines as a response to PrPSc, indirect disturbance of the synthesis, processing and/or action of other proteins due to neuronal accumulation of PrPSc or a combination of some or all suggested mechanisms.

1.2 The prion protein isoforms

The prion protein (PrP) is well conserved amongst species displaying overall structural similarities. The human PRNP gene is located on chromosome 20 and the mouse prnp gene on chromosome 2 (Sparkes, et al., 1986) encoding the 253 amino acid (254 in mouse) unprocessed PrPC precursor protein. In mammals, PrP is highly expressed in the brain, but although the amount of PrP in the brain is very high in contrast to other tissues, it represents less than 0.1% of the total CNS proteins (Riesner, 2003). PrP mRNA in human and mouse is detected mostly in neurons of the hippocampus, cortex, thalamus, cerebellum and medulla but is also present in non-neuronal astrocytes and ependymal cells (Brown, et al., 1990, Makrinou, et al., 2002, McLennan, et al., 2001, Tanji, et al., 1995).

1.2.1 PrPC – the cellular prion protein

PrPC (Fig. 1) is synthesized in the rough endoplasmatic reticulum (ER) and transits to the Golgi on its way to the cell surface. It is a plasma membrane-associated protein and the final processed form of PrPC contains amino acids 23-231 from the original translation product, resulting from an N-terminal cleavage of a hydrophobic signal sequence (residues 1-22) and cleavage of 22 amino acids in the C-terminal prior to glycosylphosphatidylinositol (GPI) modification (Prusiner, 1998). Mature PrPC can be divided in two distinct re-gions; one flexible N-terminal region that is essentially unstructured and comprises amino acids 23-125; and a C-terminal region comprising amino acids 126-231, composed of three α-helical structures and a short β-sheet motif (Abid, et al., 2006). PrPC contains an eight-residue repeating sequence (octarepeat) in the N-terminal (Cohen, et al., 1998) and during its biosynthesis in the ER, PrPC undergoes several post-translational modifications. During folding, a disulfide-bridge is formed between cysteine residues 179 and 214, stabilizing the C-terminal protein region (Zahn, et al., 2000). Overall, the PrPC structure contains 42% α-helix and 3% β-sheet (Pan, et al., 1993). PrPC is a glycoprotein that may contain two N-linked oligosaccharide chains at aspar-agine residues 181 and 197 for human and 180 and 197 for mouse PrPC. In Syrian hamster, more than 50 different sugar chain combinations have been shown to be attached to PrPC (Endo, et al., 1989, Lawson, et al., 2005). Oligosac-

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charide chains are added in the ER, further modified and extended to contain sialic acid in the Golgi. Although PrPC possesses two glycosylation sites, the protein is found as a mixture of di- mono- or unglycosylated forms, depend-ing on the neuronal region and species (Russelakis-Carneiro, et al., 2002). How-ever, the physiological significance of these differences remains unknown.

Figure 1. Schematic structure and post-translational modification of the prion pro-tein. PK resistant fragment concerns PrPSc and is described in chapter 1.2.2. OR, octarepeat; PK, proteinase K.

After passage through Golgi, vesicles containing PrPC bud off and fuse with the cell surface, localizing PrPC to the plasma membrane within one hour after synthesis. The association of PrPC to lipid rafts has been suggested to occur in the Golgi, although recently, an immature precursor to PrPC was found to be associated to cholesterol rich rafts in ER (Taylor, et al., 2006).

The majority of PrP biosynthesis leads to a GPI-anchored form, but at the ER membrane PrP can also be synthesized in two other alternative forms designated CtmPrP and NtmPrP which may be produced with their C- or N-terminus respectively towards the lumen. Mature CtmPrP has been found to cause neurodegenerative effects in mice similar to those of genetic prion disease (De Fea, et al., 1994, Hegde, et al., 1998).

PrPC has a half-life of six hours and is degraded through a two-step mechanism (Caughey, et al., 1989, Taraboulos, et al., 1992, Taraboulos, et al., 1995). The first step occurs in rafts where the polypeptide is N-terminally cleaved yield-ing a 17 kDa polypeptide. The second step occurs in endosomes where the 17 kDa peptide is completely degraded (Taraboulos, et al., 1995).

1.2.2 PrPSc

- the scrapie prion protein

The infectious agent of prion diseases, PrPSc, is also referred to solely as the prion. Although PrPC and PrPSc share the same primary structure and post-translational modifications, they differ dramatically in their secondary struc-ture. In contrast to PrPC, PrPSc is enriched in β-sheets. PrPSc consists of ap-proximately 40% β-sheets and 30% α-helices (Pan, et al., 1993) (Fig. 2). PrPSc is also differently metabolized e.g. it is N-terminally trimmed in an acidic compartment in cultured cells which is most likely an endosomal compart-ment whereas PrPC is not (Caughey, et al., 1991, Taraboulos, et al., 1992). In addi-

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tion, the two PrP isoforms possess very different biochemical properties. In contrast to PrPC, PrPSc is insoluble in non-denaturing detergents, possesses a core of approximately 142 amino acids (designated PrP 27-30) that is par-tially resistant to proteinase K (PK) treatment (Fig. 1) and polymerizes into very large amyloid structures (prion rods) under combined action of prote-ases and detergents (McKinley, et al., 1991, Prusiner, et al., 1983). PrPSc cannot readily be degraded by neuronal cells and appears to accumulate in late en-dosomes, lysosomes and on the cell surface (Caughey, et al., 1991, Naslavsky, et

al., 1997, Shyng, et al., 1993). Studies have shown that PrPSc also accumulates in extracellular spaces in brain tissue as amorphous deposits, diffuse fibrils or dense amyloid plaques (DeArmond, et al., 1995).

Figure 2. Plausible models for the tertiary structures of PrPC and PrPSc. Figures are adopted from The Cohen group homepage; http://www.cmpharm.ucsf.edu/cohen/.

1.2.3 Models for prion propagation

The transition of PrPC into PrPSc involves a conformational change whereby a portion of PrPC´s α-helical and coil structure is refolded into β-sheets (Pan,

et al., 1993). Two different mechanisms for conversion have been proposed. The “template assisted/refolding” model proposes that PrPSc contains the refolding instruction that is applied to PrPC upon interaction between the two isoforms. The process is catalyzed by an unknown protein or co-factor des-ignated protein X and mediated by the formation of a conformational inter-mediate (Cohen, et al., 1998). This model thus argues that the infections unit consists of a single PrPSc monomer functioning as a folding chaperone (Abid,

et al., 2006). However, it was recently shown that small oligomers composed of less than six units of PrPSc were noninfectious in Syrian hamsters (Silveira,

et al., 2005). The particles harboring the highest infectious potential were non-fibrilar structures composed of 14-28 units of PrPSc. Thus, growing evidence

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supports the hypothesis that small aggregates of PrPSc rather than monomers or large fibrilar structures can catalyze the conversion (Abid, et al., 2006). The “seeding/nucleation” model proposes that monomeric PrPSc exists in equilib-rium with PrPC (Jarrett, et al., 1993, Kocisko, et al., 1994). In this scenario, mono-meric PrPSc would represent a minor and transient isoform of PrP and would be stabilized only when forming ordered aggregates. The stabilized oli-gomers act as nuclei to recruit monomeric PrPSc into the polymer in a proc-ess that is much faster than the initial formation of the seed.

PrPSc has been found to co-localize with PrPC in lipid rafts in the plasma membrane (lipid rafts are described in more detail in chapter 2.) and this co-localization has been shown to be necessary for the conversion of PrPC to PrPSc (Kaneko, et al., 1997). In addition, disruption of lipid rafts by lovastatin-mediated cholesterol depletion diminishes PrPSc formation (Taraboulos, et al., 1995). Furthermore, metabolic studies indicate that PrPSc formation occurs after PrPC reaches the cell surface and release of nascent PrPC from the cell surface by phosphatidylinositol-specific phospholipase C (PIPLC) prevents PrPSc synthesis (Borchelt, et al., 1992, Caughey, et al., 1991). These results support that lipid rafts play an important role in the conversion of PrPC to PrPSc.

Presumably, the endocytic pathway and/or recycling of PrPC are involved in the synthesis of PrPSc. The endosomes are the proposed site of conversion and the gradually lowered pH of endocytic pathways is believed to be a driv-ing force (Borchelt, et al., 1992). Several compounds have been tested as cofac-tors of conversion by application in in vitro conversion assays. This method has shown that presence of RNA molecules as well as heparan sulfate (HS) promotes formation of PrPSc from PrPC (Supattapone, 2004). Many reports show that HS play a role in the life cycle of the prion protein and possibly in the pathogenesis of the disease. Recent studies indicate that cell surface HS may serve both as a cofactor for PrPC to PrPSc conversion and also as a receptor for the uptake of prions (Diaz-Nido, et al., 2002). Sulfated glycans stimulate cell-free conversion of PrPC (Wong, et al., 2001) and are associated with amyloid plaques or PrPSc deposits (McBride, et al., 1998, Snow, et al., 1990) in the brain during prion disease, pointing to a role of these molecules in the pathogene-sis of these diseases.

Divergent results have been obtained regarding the role of divalent metal ions for conversion in vitro. It was recently shown that PrPSc amplification by the protein-misfolding cyclic amplification (PMCA) technique is inhib-ited by CuCl2 and ZnCl2. Conversely, another report identified a number of transition metal ions (manganese, copper and iron) to promote PMCA con-version of PrPC into PrPSc (Kim, et al., 2005, Orem, et al., 2006).

How prions move from one cell to another during infection and/or disease propagation is not clarified. One suggestion is that secreted cellular vesicles called exosomes carry prions between cells. Exosomal secretion can be used to eject molecules targeted for degradation, but some cell types use exosomes for intercellular communication by proteins and lipid transfer (Fevrier, et al., 2004). Often, endocytosed raft-associated components, including

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PrPC, transit through multivesicular bodies, the specific type of endosomal organelles where exosomes are formed. Indeed, PrPC is found in exosomes and recent findings also show PrPSc association with exosomes (Fevrier, et al., 2004, Vella, et al., 2007). In addition, transfer of prion infectivity has been shown to occur by cell-cell contact in culture (Kanu, et al., 2002). It has also been found that conditioned cell free medium from infected cells could transfer prions to uninfected GT1 cells (Schätzl, et al., 1997).

The most convincing evidence that PrPSc formation and propagation is re-sponsible for neurodegeneration in prion diseases is that PrP-deficient mice are resistant to prion infection and fail to replicate prions (Bueler, et al., 1993, Prusiner, et al., 1993). This means that if PrPSc cannot be formed because the substrate PrPC is lacking, the mice will not become ill.

1.3 Putative functions of PrPC

The physiological role of PrPC remains elusive. However, in the past decade, several efforts have been made to elucidate the function of PrPC, with a num-ber of hypotheses being proposed.

1.3.1 Molecular interactions of PrPC

Several proteins are found to bind and interact with PrPC. Ligands identified mainly belong to heat-shock proteins, membrane-bound receptors and HS (Lasmezas, 2003). A yeast two-hybrid system was used to screen for PrPC-interacting proteins from a cDNA expression library. This led to the identifi-cation of the anti-apoptotic molecule Bcl-2 (Kurschner, et al., 1995, Kurschner, et

al., 1996), the chaperone Hsp60 (Edenhofer, et al., 1996), the 37 kDa laminin re-ceptor precursor LRP (Rieger, et al., 1997) as well as synapsin Ib, the adaptor protein Grb2 and the unknown protein Pint 1 (protein interactor 1) (Spielhaupter, et al., 2001). PrPC binds to the extracellular matrix-protein laminin, promoting neurite outgrowth in a rat hypothalamic cell line and in primary neurons of rodents (Graner, et al., 2000). The laminin receptor (LR) has been shown to act as a cell surface receptor for PrPC and play a role in PrPSc propagation in cultured mammalian cells. LRP/LR mediates the binding and internalization of purified PrPC to cells in culture, suggesting a role in the endocytic pathway of PrPC (Lasmezas, 2003). The finding that PrPC and neu-ronal cell adhesion molecule (N-CAM) interacts is consistent with a role of PrPC as a signaling receptor (Schmitt-Ulms, et al., 2001). Another study sug-gested a possible role of PrPC as a lipid raft signaling protein as antibody-mediated cross-linking of PrPC in murine 1C11 differentiated neuronal cells induce a caveolin-1 dependent activation of the tyrosine kinase Fyn (Mouillet-Richard, et al., 2000). An ongoing debate however, is whether caveolin-1 is expressed in neurons or not (Gorodinsky, et al., 1995). It has been shown that ligation of PrPC is able to induce phosphorylation of extracellular regulated kinase 1/2 (ERK1/2) without the expression of caveolin-1 in GT1-7 hypotha-lamic cells (Schneider, et al., 2003). In addition, in enterocytes, PrPC and Src

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kinase were shown to co-localize in raft domains in cell-cell junctional com-plexes and a direct physical interaction with Src was also demonstrated by co-immunoprecipitation of PrPC with the Src kinase (Morel, et al., 2004). Also, PrPC ligation in T-cells induces lipid raft clustering and activation of the Src family kinase members Fyn and Lck (Stuermer, et al., 2004).

1.3.2 Copper metabolism, oxidative stress and neuronal survival

PrPC is a metal ion-binding protein as it binds copper with affinity in the femtomolar range (Brown, et al., 1997, Jackson, et al., 2001). Zinc, manganese and nickel also bind to PrPC, but with lower affinities than does copper (Brown, et

al., 2000, Jackson, et al., 2001, Pan, et al., 1992). The binding is cooperative and occurs via histidines in the N-terminal region of PrPC. In this region, six conserved His residues have been identified (four in the octarepeats, the others at position 96 and 111) (Jackson, et al., 2001). The fact that mice devoid of PrPC harbor 50% lower copper concentrations in synaptosomal fractions than wild type mice suggests that PrPC could regulate the copper concentra-tion in the synaptic region of the neuron. PrPC may play a role in the re-uptake of copper into the pre-synaptic cell (Kretzschmar, et al., 2000). Copper added to cultured neuroblastoma cells has been shown to stimulate PrPC endocytosis (Pauly, et al., 1998). In addition, deletion of the octarepeats or sub-stitution of the histidine residues (His68 and His76) in the central two re-peats abolished copper-induced endocytosis of PrPC expressed in human neuroblastoma SH-SY5Y cells (Perera, et al., 2001). Hence, it can be hypothe-sized that the transport of copper from the extra- to the intracellular com-partment is operated through PrPC internalization.

Recombinant as well as immunoprecipitated murine PrPC have been shown to harbor the activity of a copper/zink-dependent superoxide dismu-tase (SOD1) that endows PrPC with anti-oxidant activity (Brown, et al., 1999). SOD1 expression in cells is known to alter cell survival and resistance to oxidative stress. In accordance with these results, another study showed a reduced activity of cytosolic SOD1 in the brains of PrP0/0 mice (Brown, et al., 1997) and an increased SOD1 activity and copper loading in mice that over-express PrPC (Brown, et al., 1998). Moreover, SOD activity in brain lysates from wild-type mice was reduced after PrPC depletion (Wong, et al., 2000). In con-trast to these results, others failed to detect significantly differing amounts of copper in subcellular fractions of brain from PrP0/0 mice, wild-type mice and mice overexpressing PrPC 10 times as well as any effect of PrPC expression level on SOD activity (Waggoner, et al., 2000).

Additional neuroprotective effects of PrPC have been shown. Immortal-ized hippocampal cells from PrP0/0 were found to be more sensitive to apop-tosis induced by serum-withdrawal than wild-type neurons. Transfection of a PrPC or Bcl-2 expressing construct into the PrP null neurons protected them from apoptosis induced by serum deprivation (Kuwahara, et al., 1999). In addi-tion, overexpression of PrPC has been shown to inhibit Bax-induced apop-tosis in primary human neurons (Bounhar, et al., 2001). These studies suggest

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that depletion of PrPC is involved in apoptosis and increased susceptibility to oxidative stress. The role of PrPC in modulating neuronal antioxidant ho-meostasis is further indicated by evidence of oxidative stress in both scrapie-infected brains and neuronal cells (Guentchev, et al., 2000, Milhavet, et al., 2000).

1.3.3 PrPC knock-out mice

Several lines of mice devoid of PrPC have been generated by homologous recombination in embryonic stem cells, using either of two strategies. One in which the disruptive modifications is restricted to the open reading frame (ORF) (Bueler, et al., 1992, Manson, et al., 1994). Mice homozygous for the inacti-vated gene develop normally, show no striking pathology and are resistant to prion infection. The other strategy involves deletion of not only the reading frame, but also of its flanking regions. These mice also develop normally, but exhibit severe ataxia and Purkinje cell loss later in life (Moore, et al., 1999, Rossi, et al., 2001, Sakaguchi, et al., 1996, Silverman, et al., 2000). In the ORF-modified mice behavioral studies revealed no significant differences to wild-type (Bueler, et al., 1992), except for alterations in circadian activity (Tobler, et al., 1997, Tobler, et al., 1996). Changes in the synaptic behavior in brain slices from ORF-modified mice have been observed (Collinge, et al., 1994), although such changes have not been confirmed by others (Lledo, et al., 1996).

In the mice where also flanking regions of the PrP gene was ablated, Pur-kinje cell loss and ataxia were shown to be due to ectopic expression of a prion protein paralog, doppel (Li, et al., 2000, Moore, et al., 1999). Doppel (Ger-man for double) or Dpl (downstream of the prnp locus) was expressed be-cause of an intergenic splicing event that placed the Dpl gene under the con-trol of the prnp promoter. Thus, ectopic expression of Dpl in the absence of PrPC, rather than absence of PrPC itself caused Purkinje cell loss. To circum-vent the problems involved in the interpretation of PrPC-null mice, two lines of conditional knock-out mice have been generated, with an intact PrPC ex-pression during embryogenesis, which can be knocked down in the adult mouse. Unfortunately, these mice did not reveal the function of PrPC as no obvious effects on neuronal viability, neuropathological abnormalities or neurological status were observed (Mallucci, et al., 2002, Tremblay, et al., 1998).

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2. Lipid rafts – specialized plasma membrane domains

The fluid mosaic concept (Singer, et al., 1972) is still the most established model of how biological membranes are organized in which the proteins are uniformly dispersed in the lipid solvent. However, this theory has been ques-tioned since it oversimplifies the complexity of the membrane and cannot explain many of its important functions. Research now indicates that mem-brane lipids are not randomly distributed, but instead show local heteroge-neities.

There is evidence for the existence of more than one lipid bilayer state, a liquid-ordered (Lo) and a liquid-disordered (Ld). Instead of random mixing as postulated by the fluid mosaic model, differential packing abilities of sphin-golipids and phospholipids probably lead to phase separation in the mem-brane.

2.1 The lipid raft hypothesis

The existence of lipid raft domains was proposed by Simons and co-workers almost 20 years ago and originated from studies on epithelial cell polarity (Simons, et al., 1988). The hypothesis postulates the existence of dynamic as-semblies of cholesterol and sphingolipids in the exoplasmic leaflet of the bilayer. The high content of saturated hydrocarbon chains in cell sphingolip-ids allows cholesterol to be tightly intercalated, similar to the organization of the liquid-ordered state in model membranes (Brown, et al., 1998, Simons, et al., 1988, Simons, et al., 1997). The membrane surrounding lipid rafts is more fluid, as it consists mostly of phospholipids with unsaturated, and therefore kinked, fatty acyl chains and cholesterol.

The existence of rafts has been discussed for many years and evidence for rafts has been presented in numerous studies e.g. by electron microscopy (Wilson, et al., 2000), photonic force microscopy (Pralle, et al., 2000) and antibody cross-linking of raft proteins into patches segregating them from non-raft proteins (Harder, et al., 1998).

An important early indication that rafts exist, came from the observation that cell membranes are not fully solubilized by non-ionic detergents at low temperature. Instead, Lo-phase detergent resistant membranes (DRMs) re-main and can be isolated on a sucrose gradient (Brown, et al., 1992, Schroeder, et

al., 1994). The existence of rafts is further discussed in chapter 2.6 in this the-sis.

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Table 1. Alternative names for DRMs

DRM Detergent-resistant membrane

CEM cholesterol-encriched membrane

GEM glycosphingolipid-enriched membrane DIG detergent-insoluble glycosphingolipid-enriched membrane TIM Triton-insoluble membranes

TIFF Triton-insoluble floating fraction

2.2 Lipid composition of rafts

The most familiar state in which saturated acyl chains are highly ordered is the gel phase. However, raft lipids do not exist in this phase because of the high concentration of cholesterol present since cholesterol has important effects on phase behavior contributing to membrane properties between gel and Ld phases. Rafts probably exist in the Lo phase or a state with similar properties (Brown, et al., 2000). In vitro reconstitution with defined lipids has shown that, at the relative concentrations found in the plasma membrane, sphingolipids can indeed form a Lo phase in the lipid bilayer (Ahmed, et al., 1997).

Cholesterol is thought to serve as a spacer between the hydrocarbon chains of the sphingolipids and to function as a dynamic glue that keeps the raft assembly together (Simons, et al., 2000). Cholesterol partitions between the raft and the non-raft phase with higher affinity to raft sphingolipids than to unsaturated phospholipids (Fig. 3). Recent studies showed important features required for cholesterol’s capability to induce Lo phases in sphingolipid con-taining membranes; a small polar headgroup, the planarity of the steroid ring system and optimal length and structure of the aliphatic site chain at the D7 ring of cholesterol (Xu, et al., 2000). In spite of their dense packing, lipids in Lo phases retain lateral and rotational mobility (Almeida, et al., 1992). Rafts are also thicker than non-raft domains since the lipid acyl chains are more ex-tended (Brown, 2006).

In a study of 1% Triton X-100-insoluble, low-density fractions from Madin-Darby canine kidney (MDCK) cells, Brown and Rose (Brown, et al., 1992) reported that the isolated membranes contained 32 mol% cholesterol and 14 mol% sphingomyelin compared to 12 mol% cholesterol and 1 mol% sphingomyelin in whole cells. These fractions were enriched, about 5-fold in glycolipids such as gangliosides and sulfatides compared to non-fractioned membranes (Fig. 3).

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Figure 3. Schematic picture of lipid rafts.

2.3 Proteins associated with rafts

Membrane proteins can be assigned to three categories: those that are mainly found in the rafts, those that are present in the Ld phase and those that repre-sent an intermediate state, moving in and out of rafts. Some of the proteins targeted to rafts are modified by saturated chain lipid groups that pack well into an ordered lipid environment (Brown, et al., 2000). Therefore, proteins with raft affinity include GPI-anchored proteins (Brown, et al., 1998), doubly acy-lated proteins such as some Src-family kinases or the α-subunits of hetero-trimeric G-proteins (Resh, 1999), cholesterol-linked and palmitoylated proteins such as Hedgehog (Rietveld, et al., 1999) and transmembrane proteins, particu-larly palmitoylated ones such as influenza virus hemagglutinin and β-secretase (BACE) (Simons, et al., 2000). GPI-anchored proteins or proteins that carry hydrophobic modifications probably partition into rafts owing to pref-erential packing of their saturated membrane anchors. In contrast, both membrane-spanning proteins and prenyl groups (which are bulky and branched) are difficult to accommodate in highly ordered lipid rafts (Melkonian, et al., 1999).

Some membrane proteins are regulated raft residents and have weak affin-ity for rafts in the unliganded state. After binding to a ligand, they undergo a conformational change and/or become oligomerized which increases their raft affinity (Harder, et al., 1998). A peripheral membrane protein, such as the non-receptor tyrosine kinase Lyn, can be reversibly palmitoylated and can lose its raft association after depalmitoylation (Zacharias, et al., 2002). Thus, the partitioning of proteins in and out of rafts can be tightly regulated.

This variety of components suggests that, if these fractions represent ac-tual domains, they could serve as signal transduction platforms (Jacobson, et

al., 1999). In fact, the association of proteins with ordered lipid domains is specific and of functional importance since disruption of these domains with

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cholesterol depleting agents can affect signal transduction and receptor-ligand interactions (Dobrowsky, 2000).

2.4 Raft function

Several approaches have been taken to investigate raft function. One ap-proach is to study proteins after experimental clustering of lipid rafts since several proteins must interact and/or co-localize to function. Another ap-proach is to show that disrupting the association of a protein to rafts also disrupts its function. Finally, raft disruption by depletion of cholesterol or occasionally sphingolipids can affect function of a raft resident protein (Brown, et al., 2000).

Rafts were first proposed to mediate protein sorting in the trans-Golgi network, especially in polarized epithelial cells and neurons (Benting, et al., 1999, Brown, et al., 1998, Rodriguez-Boulan, et al., 1999, Simons, et al., 1997). Rafts may also be important in the endocytic pathway (Mukherjee, et al., 1999).

Lipid rafts are thought to function as platforms for the dynamic associa-tion of signaling molecules. Gathering of the receptors for interaction with ligands and effectors on both sides of the membrane can allow efficient and rapid coupling of activated receptors to the effector system (Kasahara, et al., 2000, Lai, 2003). These lipid domains may also create a microenvironment that is locally protected from other transmembrane negative regulators of signal-ing, such as phosphatases (Lai, 2003, Simons, et al., 2000). Some of the strongest cases for involvement of lipid domains in signaling come from studies in hematopoietic cells. Cross-linking of many GPI-anchored proteins or trans-membrane receptors in T- and B-cells by various means induces clustering of components of signal transduction, and often correlates with an increase in the raft association for relevant signaling components (Wilson, et al., 2000, Viola, et al., 1999). The first signaling process shown to involve lipid rafts was immunoglobulin E (IgE) signaling during the allergic immune response (Field, et al., 1995, Sheets, et al., 1999).

The glial cell derived neurotrophic factor (GDNF) family of ligands is important for the development and maintenance of the nervous system. GDNF binds to a multi-component receptor complex that is composed of the GPI-linked GDNF receptor-α (GFRα) and the transmembrane tyrosine kinase, RET. The receptor subunits are not associated in the absence of ligand, but after extracellular GDNF stimulation, RET moves into rafts, where it associates with GFRα. Signal transduction depends on the co-localization of RET and GFRα in lipid rafts, as cholesterol depletion with methyl-β-cyclodextrin decreases GDNF signaling (Tansey, et al., 2000).

Caveolae are functionally specialized lipid rafts that are invaginations in the plasma membrane coated with the cholesterol-binding protein caveolin-1 (Taylor, et al., 2006). Several functions have been attributed to these structures e.g. an involvement in endocytosis (Parton, 1996), transcytosis (Schnitzer, et al., 1994) and regulation of signaling cascades (Anderson, 1998), although they are

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not absolutely required as several cell types that lack caveolin, such as lym-phocytes and neurons can nevertheless signal through rafts (Simons, et al., 2000).

2.5 Distribution and trafficking of rafts

Rafts are likely to be most abundant in membranes that are rich in choles-terol and sphingolipids including the plasma membrane, the late secretory pathway and endocytic compartments (Brown, et al., 2000). Whereas cholesterol is synthesized in the ER, sphingolipid synthesis and head group modification are completed largely in the Golgi (van Meer, 1989). This means that choles-terol-sphingolipid rafts first assemble in the Golgi (Brown, et al., 1998) and move out mainly towards the plasma membrane, as vesicles going back to the ER contain little sphingomyelin and cholesterol (Brugger, et al., 2000).

The distribution of lipid rafts over the cell surface varies between cell types. In polarized epithelial cells and neurons, lipid rafts accumulate in the apical and axonal plasma membrane, respectively. In lymphocytes and fi-broblasts, rafts are distributed over the cell surface, without obvious polarity (Simons, et al., 1997).

Rafts are continuously endocytosed (Mukherjee, et al., 2000) from the plasma membrane. From early endosomes, rafts either recycle directly back to the cell surface or return indirectly through recycling endosomes, which also can deliver rafts to the Golgi (Puri, et al., 1999). Lipid rafts have also been shown to serve as docking sites for certain pathogens and toxins (Fivaz, et al., 1999).

2.6 The lipid raft debate

Membrane fragments that are insoluble in non-ionic detergents (DRMs), can be isolated from most mammalian cells (Brown, et al., 1992). DRMs appear to be derived from rafts, they are rich in cholesterol and sphingolipids and are in the Lo phase when isolated from cells (Ge, et al., 1999). Thus, there is a close relation between rafts and DRMs.

This method of isolation of lipid rafts involves cooling before detergent extraction in order to stabilize the Lo phase and enhance its detergent resis-tance. However, the phase of the membrane is certainly affected by the de-creasing temperature, and probably, more of the membrane is in the Lo phase at 0°C than at 37°C (Brown, et al., 2000). On the other hand, in some cases de-tergent may partially solubilize raft lipids and proteins even after cooling, leading to an underestimation of the fraction of these molecules in rafts (Arni, et al., 1998, Field, et al., 1999, Ostermeyer, et al., 1999). In contrast to these argu-ments, model membrane studies showed that Lo-Ld phase separation could occur at 37°C in lipid mixtures with physiological levels of each of these lipid classes, making phase separation in cell membrane plausible (Ahmed, et

al., 1997, Silvius, 2003).

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If rafts are present, they should be possible to detect by microscopy. Never-theless, molecules such as GPI-anchored proteins and gangliosides, taken as putative raft markers often appear uniformly distributed on the cell surface when detected microscopically (Brown, et al., 1998, Jacobson, et al., 1999). One interpretation of this is that the rafts are too small to be seen by light micros-copy (Simons, et al., 2000). However, upon clustering with antibodies or other agents, the distribution of these markers can change dramatically. In some cases, clustering of one marker can cause redistribution of another, although the two are unlikely to interact directly (Brown, et al., 1998, Harder, et al., 1998,

Janes, et al., 2000, Viola, et al., 1999). However, it is not proven that clustering of lipid rafts occurs in vivo. If rafts do not exist, a continuous model could be that the outer leaflet of the plasma membrane would be an essentially ho-mogenous phase rich in cholesterol and sphingolipids that would provide a permeability barrier to cells that remained highly fluid in the plane of the bilayer, thereby allowing proteins to move freely by lateral diffusion and participate in protein:protein interactions. The high levels of cholesterol and sphingolipids are also likely to cause the bilayer to be thicker, analogous to lipid raft compartments, than those of the earlier compartments of the secre-tory pathway (Bretscher, et al., 1993).

Another way of investigating the existence of lipid rafts would be to modulate membrane cholesterol. However, this method would not be con-clusive since agents that modulate membrane cholesterol are likely to affect cell function in ways that are unrelated to raft formation. Sterols have pro-found effects on bilayer structure, even in single-phase membranes (Huang, et

al., 1999, McConnell, et al., 2003), and their removal would greatly affect even membranes that do not contain rafts. Thus, inhibition of a process by choles-terol modulators suggests that the process requires cholesterol but does not prove that it requires rafts (Brown, 2006).

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3. Protein tyrosine kinases

Protein tyrosine kinases (PTKs) are a group of enzymes that catalyzes the transfer of the terminal phosphate group (γ) from ATP to the hydroxyl group of the amino acid tyrosine on target proteins. Activation of PTKs elicits nu-merous different intracellular events such as proliferation, differentiation and motility. There are mainly two different types of PTKs, the receptor and the non-receptor tyrosine kinases, both of which will be described in more de-tail.

3.1 Non-receptor tyrosine kinases (NRTKs)

A large group of PTKs is the NRTKs. Examples of these proteins are the family of Janus kinases (JAKs), the family of focal adhesion kinases (FAKs) and the Src family kinases (SFKs). The SFKs have been studied in this thesis and will therefore be discussed in chapter 3.1.1.

Most NRTKs are found in the cytoplasm (Neet, et al., 1996), although some are anchored to the cell membrane through amino-terminal modification, such as myristoylation and/or palmitoylation. In addition to the tyrosine kinase domain, NRTKs also possess domains that mediate protein:protein, protein:lipid and protein:DNA interactions, such as Src homology 2 (SH2) and SH3 domains.

3.1.1 Src family kinases (SFKs)

Almost a century ago, Peyton Rous described that injection of cell-free ex-tracts from chicken tumors could cause tumors in other animals, giving rise to the hypothesis that cancer could be caused by a transmissible agent, a virus that now bears his name, Rous Sarcoma Virus (RSV) (Rous, 1911). By 1977 the gene responsible for this virus’s oncogenic properties as well as its protein product, pp60v-Src were identified (Martin, 2001). Over the next 20 years, studies on RSV yielded an incredibly rich medley of new and startling information; cellular transformation was caused by a single gene, (Src); Src was a protein tyrosine kinase; Src was derived from a cellular gene (the proto-oncogene, c-Src); Src activity was regulated by intramolecular interac-tions controlled by tyrosine phosphorylation; and domains of Src (SH2 and SH3) mediated protein:protein interactions. Concurrent with the studies on Src, it was recognized that Src was a member of a large family of structur-ally related kinases, SFKs (Thomas, et al., 1997).

There are eleven SFKs in humans including Src, Fyn, Yes, Frk, Lyn, Hck, Fgr, Blk, Brk, Srm and Lck (Table 1). The members Src, Fyn, Yes and Frk are expressed in most tissues although different levels of the proteins are found in different tissues (Roskoski, 2004). For example, Src is ubiquitously

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expressed, however, platelets, neurons and osteoclasts express 5-200-fold higher levels of this protein (Brown, et al., 1996).

Table 2. SFK members in humans

Src family member

Pattern of expression Isoforms

Src Ubiquitous Neuron-specific isoforms Fyn Ubiquitous T cell-specific isoform (Fyn T) Yes Ubiquitous Frk Ubiquitous Mainly in primary epithelial cells Blk B cells Fgr Myeloid cells, B cells Hck Myeloid cells Two different translational starts Lyn Brain, B cells, myeloid cells Two alternatively spliced forms Lck T cells, NK cells, brain

Srm Keratinocytes

Brk Colon, prostate, small intestine, breast cells

Structure of SFKs

SFKs are 52-62 kDa proteins composed of six distinct functional regions (Fig. 4); the SH4 domain, the unique region, the SH3 domain, the SH2 do-main, the catalytic domain and a short negative regulatory tail (Brown, et al., 1996). The SH4 domain is a 15-amino acid sequence that contains signals for lipid modification of SFKs (Resh, 1993). The glycine at position 2 is important for addition of a myristic acid moiety (Kamps, et al., 1985) and cysteine residues in the SH4 domain are subject to palmitoylation. These cysteines are present in all members except Src and Blk (Resh, 1993).

The unique domain has been proposed to be important for mediating in-teractions with receptors or proteins that are specific for each family member (Thomas, et al., 1997).

SH3 domains bind short contiguous amino acid sequences rich in proline residues (Cohen, et al., 1995) and examples of proteins shown to interact with SFK SH3 domains either in vitro or in vivo include p68sam, p85 phosphatidy-linositol-3 kinase (PI3-K) and paxillin (Thomas, et al., 1997). The SH3 domain of SFKs is composed of 50 amino acids (Cohen, et al., 1995, Pawson, 1995). How-ever, alternatively spliced forms of Src have been found, which contain 6- or 11-amino acid insertions in the SH3 domain in CNS neurons (Brugge, et al., 1985, Martinez, et al., 1987, Pyper, et al., 1990, Sugrue, et al., 1990). However, there is little evidence for a functional difference among Src splice variants (Thomas,

et al., 1997). The SH2 domains bind to short contiguous amino acid sequences contain-

ing phosphotyrosine, and the specificity of individual SH2 domains lies in the 3-5 residues following the phosphotyrosine (Pawson, 1995). Examples of

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proteins shown to interact with the Src SH2 domain in vivo include the focal adhesion kinase (FAK), p130cas, p85 PI3-K and p68sam (Fukui, et al., 1991, Petch,

et al., 1995, Schaller, et al., 1994). Binding interactions mediated by the SH2 do-main are regulating the catalytic activity of SFKs as well as the localization of the enzyme and also its binding proteins.

Figure 4. Domain structure of Src family kinases

Regulation of SFKs

The C-terminal tail of SFKs bears an autoinhibitory phosphorylation site (Y527 in Src) (Cooper, et al., 1986). SFKs require phosphorylation within a segment of the kinase domain termed the activation loop for full catalytic activity. In Src, this autophosphorylation site is Y416 (Smart, et al., 1981). In

vivo, SFKs are phosphorylated on either Y416 (in their active state) or Y527 (in their inactive state). The inactivating phosphorylation on Y527 is carried out by the SFK-specific kinase, Csk (Nada, et al., 1991). Phosphorylation of the C-terminal tail promotes binding to the SH2 domain leading to autoinhibi-tion of the kinase activity (Sicheri, et al., 1997, Williams, et al., 1997).

The SH2 and SH3 domains have four important functions. First, these domains constrain the activity of the enzyme via intramolecular contacts. Second, proteins that contain SH2 or SH3 ligands can bind to the domains and attract Src to specific cellular locations. Third, as a result of displacing the intramolecular SH2 or SH3 domains, interacting proteins can activate Src kinase activity, and fourth, proteins containing SH2 or SH3 ligands can en-hance their ability to function as substrates for SFKs (Brown, et al., 1996).

Csk homology kinase (Chk) is another enzyme that catalyzes the phos-phorylation of the inhibitory tyrosine, Y527, of SFKs (Zrihan-Licht, et al., 1997). Csk is expressed in all mammalian cells, whereas Chk is limited to certain cells in breast and testes as well as hematopoietic cells and neurons (Brown, et

al., 1996). An important mechanism for SFK activation involves Y527 dephosphory-

lation. Candidate phosphatases include cytoplasmic protein tyrosine phos-phatase 1B (PTP1B), Src homology 2 domain-containing tyrosine phos-phatase 1 (Shp1) and Shp2 together with transmembrane enzymes such as CD45 and PTPα (Roskoski, 2005).

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The involvement of SFKs in cellular signaling

A lot of studies on SFKs have led to the realization that these enzymes regu-late many cellular events e.g. cell proliferation, cytoskeletal alterations, dif-ferentiation, survival, adhesion and migration. This broad spectrum of activi-ties is a consequence of the ability of these kinases to couple with many di-verse classes of targets (Thomas, et al., 1997), some of which will be briefly summarized in this chapter.

The interplay between SFKs and receptor tyrosine kinases

SFKs are involved in signaling from many receptor tyrosine kinases (RTKs), including platelet derived growth factor receptor (PDGFR) (Kypta, et al., 1990), epidermal growth factor receptor (EGFR) (Luttrell, et al., 1988), fibroblast growth factor receptor (FGFR) (Zhan, et al., 1994), insulin-like growth factor-1 receptor (IGF-1R) (Kozma, et al., 1990) and insulin receptor (IR) (Sun, et al., 1996). SFKs can promote mitogenic signaling from these receptors in a num-ber of ways, including initiation of signaling pathways required for DNA synthesis, control of receptor turnover (Ware, et al., 1997, Wilde, et al., 1999), actin cytoskeleton rearrangement and motility (Chang, et al., 1995, Weernink, et al., 1995) and survival (Karni, et al., 1999).

The biochemical connections between these different receptors and SFKs include phosphorylation of SFKs, association with the RTK, activation of SFKs and phosphorylation of RTKs (Thomas, et al., 1997).

G-protein-coupled receptors and SFKs

Unlike most other classes of cell surface receptors, heterotrimeric G-protein-coupled receptors (GPCRs) are classically thought to signal catalytically by regulating the activity of enzymatic effectors, such as adenylate cyclases, phospholipase Cβ isoforms, and ion channels leading to the generation of second messengers.

Several mechanisms of GPCR-mediated Src activation have been de-scribed, and often the suggested mechanism include forms of receptor crosstalk in which GPCR-derived signals activate SFKs associated with ei-ther RTKs or focal adhesions. It has been shown that GPCRs have the ability to stimulate Ras-dependent activation of mitogen-activated protein (MAP) kinases, and that these signals frequently require tyrosine protein phosphory-lation carried out by SFKs (Luttrell, et al., 1996, van Biesen, et al., 1995).

SFKs in synaptic transmission and plasticity

SFKs have been implicated in proliferation and differentiation during the development of the CNS (Stehelin, et al., 1976). However, Src, Fyn, Yes, Lck and Lyn are also expressed in fully differentiated neurons in the developed CNS (Cooke, et al., 1989, Cotton, et al., 1983, Sudol, et al., 1986, Zhao, et al., 1990), im-plying additional functions of these kinases. Over the past decade, a large body of evidence has accumulated showing that a main function of Src is to up-regulate the activity of N-methyl-D-aspartate (NMDA) receptors (Wang, et

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al., 1994) and other ion channels e.g. voltage-gated potassium channels (Fadool, et al., 1997), calcium channels (Cataldi, et al., 1996) as well as γ-aminobutyric acid type A (GABAA) receptors (Moss, et al., 1995) and nicotinic acetylcholine receptors (Wang, et al., 2004).

NMDA receptors (NMDARs) constitute a principal subtype of glutamate receptors, which mediate fast excitatory transmission at most central syn-apses. By up-regulating the function of NMDA receptors, Src gates the pro-duction of NMDA receptor-dependent synaptic potentiation and plasticity. Thus, Src may be critical for processes underlying physiological plasticity including learning and memory and pathological plasticity such as pain and epilepsy (Kalia, et al., 2004).

The interplay between integrins and other adhesion receptors

Integrins are heterodimeric receptors that mediate cell-matrix and cell-cell interactions. Following engagement by their adhesive ligands, integrins transduce signals within the cell that regulate cell adhesion and spreading, migration, proliferation, differentiation and other changes in cell behavior. SFKs are implicated in integrin-regulated events by the ability to phosphory-late β1 integrin as well as other proteins that are associated with integrin-nucleated focal adhesion complexes e.g. paxillin, p130cas and FAK (Burridge,

et al., 1996, Clark, et al., 1995, Petch, et al., 1995). SFK localization to focal adhesions requires myristoylation and the SH3

domain. The SH2 domain may participate in association with integrin com-plexes through interactions with tyrosine phosphorylated proteins that local-ize to focal adhesions. One such protein is FAK, a tyrosine kinase that is phosphorylated following engagement of many integrins (Schaller, et al., 1993, Schaller, et al., 1992) and several of the proteins that associate with FAK may be substrates of SFKs. FAK appears to serve as a scaffold to organize a network of signaling and cytoskeletal proteins (Thomas, et al., 1997).

SFKs in cytokine receptor signaling

Both class I (e.g. Epo-R, IL2-R) and class II (e.g. TNFR, p75 NGF-R) cyto-kine receptor superfamilies are capable of activating a signaling cascade that can regulate growth, differentiation, cell survival and multiple specialized cell functions (Briscoe, et al., 1994). As in other receptor systems, tyrosine phosphorylation is induced following receptor engagement. The receptor subunits do not contain any intrinsic tyrosine kinase activity, however, SFKs are one group of cytoplasmic PTKs that has been shown to be involved in cytokine superfamily signaling, especially for receptors mainly from class I (Ihle, et al., 1995). The evidence implicating SFKs in these receptor pathways includes cytokine-induced activation of SFKs, association of SFKs with the receptor subunits, cytokine-stimulated association of SFKs with other signal-ing components and SFK-mediated tyrosine phosphorylation of the receptor subunit(s) or downstream effectors (Thomas, et al., 1997).

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SFKs in neurodegenerative diseases

Together with our findings presented in this thesis (paper I and paper II), another study showed increases in the levels of activated SFKs associated with prion diseases both in vivo and in vitro (Nixon, 2005). In another work, PrP-(106-126)-mediated cytochrome c release and caspase 3 activation were observed in 1C11 bioaminergic neurons and that these events were fully blocked by the SFK inhibitor PP2 (Pietri, et al., 2006). This result indicates an involvement of SFKs in the apoptotic events in prion disease. Uncontrolled Src-kinase activity is mainly linked to cellular proliferation and/or transfor-mation (Thomas, et al., 1997). However, in post-mitotic cells like CNS neurons an inappropriately up-regulated Src kinase activity or mitogenic signaling may result in neuronal loss, as illustrated by neurodegeneration and cell death of SV40 T-antigen expressing Purkinje neurons (Feddersen, et al., 1992). In this respect, it is interesting that PrPSc is a well known trigger of microglia and astrocyte proliferation during prion disease. However, a paradox is that PrPSc is toxic to post-mitotic cells like neurons (DeArmond, et al., 1992).

Alzheimer’s disease (AD) is the most frequent neurodegenerative disorder and is characterized by numerous neuritic plaques and neurofibrillary tan-gles. The exact pathophysiologic mechanisms underlying this disease are still largely unknown, but the amyloid β fragment (Aβ), a proteolytic deriva-tive of the transmembrane protein amyloid precursor protein (APP), form aggregates that are thought to initiate a pathogenic cascade ultimately lead-ing to neuronal loss (Hardy, 1997). Several studies have shown an altered regu-lation of SFKs in AD, e.g. Fyn was shown to be up-regulated in a subset of neurons in AD brain that also contained hyperphosphorylated tau (Shirazi, et

al., 1993), that is the constituent of the neurofibrillary tangles in AD. An in-creased tyrosine phosphorylation of cytoskeletal and other neuronal proteins has also been shown in both primary human and rat brain cortical cultures after exposure to Aβ. This tyrosine phosphorylation was blocked by addition of the Src family tyrosine kinase inhibitor PP2. An increased association of Fyn with FAK as well as an increased tyrosine phosphorylation of FAK was also demonstrated (Williamson, et al., 2002).

It has also been demonstrated that Src deficiency or blockade of Src activ-ity in mice provides cerebral protection following stroke (Paul, et al., 2001). Further support for that SFKs are key players in neurodegeneration is pro-vided by an observation that the SFK-inhibitor PP2 reduces focal ischemic brain injury (Lennmyr, et al., 2004).

3.2 Receptor tyrosine kinases (RTKs)

RTKs activate numerous signaling pathways within the cell, leading to cell proliferation, differentiation, migration, metabolic changes and to keep the balance between cell survival and apoptosis in cells (Schlessinger, et al., 1992). The RTK family includes the receptor for insulin and for many growth fac-tors e.g. EGF, PDGF, FGF and IGF-1. Except for the insulin and the IGF-1

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receptors, the vast majority of RTKs exists as a single polypeptide chain and are monomeric in the absence of ligand. Most polypeptide ligands for RTKs are soluble, but some can either span the membrane or be anchored to the membrane via a GPI-linkage (Flanagan, et al., 1998, Holland, et al., 1998).

Signaling through RTKs begins with binding of ligand to the receptor, which causes dimerization of the receptor and activation of tyrosine kinase activity and autophosphorylation of specific C-terminal tyrosine residues (Heldin, 1996). These autophosphorylated residues in turn serve as docking sites for a variety of signaling protein-protein interacting molecules that con-tain SH2 domains. SH2 containing substrates include PI3-K, phospholipase Cγ (PLCγ), adapter proteins such as Grb2, Shc and as mentioned earlier, SFKs. Signals are subsequently transducted from these molecules to e.g. the nucleus via several pathways, including the MAP-kinase pathway and the PI3-K cascade resulting in proliferation and cell survival, respectively (Bonfini, et al., 1996, Denhardt, 1996). Another example of a RTK activated signal-ing pathway is the PLCγ/protein kinase C (PKC) pathway. Activated PLCγ hydrolyzes phosphoinositol (PI)4,5P2 in the plasma membrane generating the second messengers, diacylglycerol (DAG) and 1,4,5-inositol triphosphate (IP3) (Hunter, 2000). DAG is an activator of PKC and IP3 triggers Ca2+ release from membrane Ca2+ stores.

3.2.1 Insulin and insulin-like growth factor-1 receptors

IR and the IGF-1R both belong to the family of RTKs. The IR is widely dispersed throughout tissues and its main function is to stimulate GLUT-4 mediated glucose transport into cells. The IGF-1R has mainly been consid-ered mediating the actions of growth hormone by increased IGF-1 release in peripheral tissues. Both of these receptors are also implicated in the prolif-erative and cell survival actions of neuronal cells. The ligands, insulin and IGF-1 bind to IR and IGF-1R, respectively with high affinity for their own receptor, although they do bind the other receptor as well, albeit with lower affinity.

Insulin and IGF-1

Insulin has a molecular mass of 5.8 kDa and is synthesized as proinsulin in the β-cells of the islets of Langerhans in the pancreas. In its mature form insulin is a two-chain polypeptide hormone and it gains access to the CNS by crossing the blood-brain-barrier in an IR mediated process (Schwartz, et al., 1990). For binding IR, the affinity of insulin is in the nanomolar range (0.1-10 nM) and for binding to the IGF-1R, the affinity of insulin is 100-500 fold lower, i.e. in the micromolar range (Ota, et al., 1988, Van Schravendijk, et al., 1986).

Insulin-like growth factor-1 (IGF-1) is structurally homologous to proin-sulin. It is a single chain molecule with a molecular mass of 7.6 kDa. IGF-1 is synthesized in multiple tissues however, the liver is the major source, and it is also produced within the CNS. The affinity of IGF-1 for the IGF-1R lies

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within the low nanomolar range and the affinity of the IGF-1 to the IR is 100-500 fold lower (Navarro, et al., 1999).

Biosynthesis and structure of IR and IGF-1R

The IR gene is located on chromosome 19 in humans. The protein is synthe-sized as a proreceptor polypeptide and exists as two isoforms, IR-A and IR-B, containing 1343 and 1355 amino acids respectively. The two different isoforms are generated as a consequence of alternate splicing of exon 11 resulting in receptors with the presence (IR-B) or absence (IR-A) of 12 amino acids at the C-terminus of the α-subunit (Fig. 5) (Ebina, et al., 1985, Ull-rich, et al., 1985). Neuronal cells, including peripheral sensory and sympathetic neurons, express predominantly the A-isoform (Seino, et al., 1989). The IR proreceptor undergoes co-translational N-linked glycosylation and before exit from ER, two proreceptor monomers dimerize, forming two interchain disulfide bonds (Lu, et al., 1996), and the dimerized proreceptor is subsequently transferred to the Golgi apparatus for further modifications. After maturation of the N-linked oligosaccharides and proteolytic cleavage at a tetrabasic amino acid sequence (Arg-Lys-Arg-Arg) into α- and β-subunits, these sub-units are further connected by a disulfide bond (Cheatham, et al., 1992, Schaffer, et

al., 1992) and the receptor is transported to the plasma membrane as a hetero-tetramer (Bravo, et al., 1994, Olson, et al., 1988).

The ligand-binding IR α-subunit is entirely extracellular, containing a cysteine-rich domain as well as 14 potential N-linked glycosylation sites. The β-subunit contains four potential glycosylation sites, an extracellular portion, a transmembrane domain and an enzymatic intracellular part that includes a tyrosine kinase domain with at least six tyrosine phosphorylation sites (Ebina, et al., 1985, Ullrich, et al., 1985). N-linked glycosylation of the IR is necessary for the formation of its tertiary and quaternary structure, as well as for correct trafficking and insertion into the plasma membrane (Olson, et al., 1988, Ronnett, et al., 1984). The β-subunit, in addition to N-linked glycosylation, also undergoes O-linked glycosylation. However, this glycosylation is not a requirement for the structure and function of the receptor (Collier, et al., 1991).

The IGF-1R is very similar, both in structure and function to the IR. It is encoded on chromosome 15 in humans and the protein product is a glyco-protein with 1376 residues. Overall, the molecular topology and biosynthesis of IGF-1R are similar to that of the IR. Following the binding of ligand, the IR or IGF-1R is rapidly internalized into the cell and is either degraded or recycled back to the plasma membrane.

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Figure 5. Structure of IR, IGF-1R and IR/IGF-1R hybrid receptors. α-subunits are extracellular and coupled by a disulfide bridge. β-subunits are transmembrane and contain an intracellular tyrosine kinase domain.

IR/IGF-1R hybrid receptors

Cells that express significant levels of IR and IGF-1R also produce IR/IGF-1R hybrid receptors (Seely, et al., 1995, Soos, et al., 1989). Hybrid receptors are formed when the IR and IGF-1R precursors encounter and become linked by disulfide bonds, rather than to a homologous precursor (Fig. 5) in the ER. Hybrid receptors have been characterized in a number of cell lines (Moxham,

et al., 1989, Soos, et al., 1989) as well as in different tissues (Bailyes, et al., 1997, Soos, et al., 1989), including brain (Bailyes, et al., 1997). In certain tissues, hybrid receptors are the most represented receptor subtype and in the brain 50% of total IGF-1R was present as hybrid receptors (Bailyes, et al., 1997). The biologi-cal role of these receptors is not yet known, however, functional studies in-dicate that hybrid receptors behave more like the IGF-1R than the IR be-cause they bind to and are activated by IGF-1 with much higher affinity than insulin (Federici, et al., 1997, Soos, et al., 1993). Depending on the IR isoform in-volved, the hybrid receptor will have different pharmacological and biologi-cal features. Thus, a hybrid receptor containing IR-A, the IR isoform pre-dominantly expressed in neurons, binds insulin with ~10-fold lower affinity than the IR. A hybrid receptor containing IR-B, the form found in peripheral tissue, hardly recognizes insulin at all, with an IC50 > 100 nM (Pandini, et al., 2002).

Subcellular localization of IR and IGF-1R

In neurons the IR is enriched in post synaptic densities of the synapse and is localized to lipid rafts together with its downstream signaling targets (Abbott, et al., 1999, Wu, et al., 1997). To date, there are no reports on IGF-1R localiza-tion to lipid rafts in neuronal cells. However, the IGF-1R was found to be localized to lipid rafts in a human breast carcinoma cell line, MCF-7 after stimulation with IGF-1. In untreated cells the IGF-1R was found in non-raft fractions after ultracentrifugation of cell lysates in a sucrose gradient (Manes,

25

et al., 1999). Another group has found that localization of IGF-1R to lipid rafts is required for 3T3-L1 adipocyte differentiation (Hong, et al., 2004). Although this has not been studied in neuronal cells, the putative involvement of lipid rafts in IGF-1R signaling is suggested since practically all of IGF-1R signal-ing partners i.e. Grb2, Ras, Fyn, PI3-K and ERK2 reside in lipid rafts in neurons (Abbott, et al., 1999, Wu, et al., 1997).

Signaling

The IR and the IGF-1R are very similar in signaling potential within a given cellular background (Adamo, et al., 1992, Lund, et al., 1994, Weiland, et al., 1991). The binding of a ligand to the α-subunit prevents α-subunit inhibition of the in-herent β-subunit tyrosine-kinase activity and within seconds/minutes, the β-subunits are autophosphorylated through a transphosphorylation mechanism.

IR and IGF-1R signaling is similar to that of the other RTKs, thus activat-ing three major signaling pathways; PI3-K/PKB, ras/raf/MAPK and PLCγ pathways. Autophosphorylated IR/IGF-1R recruits SH2 domain containing IRS1-4 or IRS p58/53 which become tyrosine phosphorylated by the recep-tor (Butler, et al., 1998, Sun, et al., 1995). The IRS-2 has been shown to be exclu-sively expressed in neuronal cells and IRS p58/53 has been found to co-localize with the IR in the synapse (Abbott, et al., 1999). The autophosphory-lated receptors recruit Shc proteins (p46/p52/p66), also phosphorylated on tyrosine residues (Pelicci, et al., 1992), enabling the receptors to activate the PI3-K and MAPK pathways, as described in chapter 3.2.

The IGF-1R mediated activation of the PKB/Akt pathway has been shown to result in proliferation and also to maintain an anti-apoptotic ma-chinery in neuronal cells (Dudek, et al., 1997, Morrione, et al., 2000), whereas the MAPK pathway mediates IR/IGF-1R activated proliferation and differentia-tion (Morrione, et al., 2000, Ye, et al., 2002).

Role of IR and IGF-1R in neurodegenerative diseases

Several studies have demonstrated a correlation between decreased IR activ-ity and cognitive deficits in AD. In one study it is shown that brain IR densi-ties are increased and that the tyrosine kinase activity is reduced in post-mortem brains from AD patients (Blum-Degen, et al., 1995). In addition, IR bind-ing and autophosphorylation studies have revealed that the Aβ-peptide re-duces insulin binding and IR autophosphorylation in human placental mem-branes (Xie, et al., 2002). These studies show that the expression, ligand bind-ing and tyrosine kinase activity of IRs are affected in AD.

Studies on rat hippocampal neurons have shown that IGF-1R protects neurons against cell death induced by Aβ (Dore, et al., 1997). Additionally, chronic subcutaneous infusion of IGF-1 reduced the levels of Aβ in the hip-pocampus on aged rats with increased levels of Aβ in the hippocampus (Carro, et al., 2002).

Parkinson’s disease (PD) is characterized by selective neuronal loss in the substantia nigra (Takahashi, et al., 1996). Regarding changes in IR expression or

26

function in PD patients a loss of IR immunoreactivity together with de-creased IR mRNA levels in substantia nigra neurons have been demonstrated in post-mortem brains from PD patients (Moroo, et al., 1994, Takahashi, et al., 1996). It has also been shown that treatment with IGF-1 is neuroprotective although no differences in the expression of IGF-1/IGF-1R have been re-ported (Offen, et al., 1996).

Amyotrophic lateral sclerosis (ALS) is characterized by degeneration of cortical, brainstem and spinal motor neurons. A small percentage of ALS is caused by a mutation in the gene encoding SOD, leading to an accumulation of superoxide radicals resulting in cellular damage. In ALS cases the IGF-1R has been found to be up-regulated in the spinal cords shown by 125I-IGF-1 binding and IGF-1R immunoreactivity (Adem, et al., 1994). Interestingly, the expression of IGF-1R has been found to be up-regulated by reactive oxygen species (Delafontaine, et al., 1997, Du, et al., 1999).

Previous studies in our group showed that scrapie-infection induces a 4-fold increase in the expression of the IGF-1R protein accompanied by ele-vated IGF-1R mRNA levels in the scrapie-infected neuroblastoma N2a cell line (ScN2a). However, no increase in binding sites or cell growth was ob-served in these cells, albeit the massive increase in IGF-1R expression. This discrepancy was further shown to be caused by a 7-fold decrease in receptor binding affinity in ScN2a cells (Östlund, et al., 2001a). Additionally, scrapie-infection also induced a 2-fold increase in IR protein level in both ScN2a and in the scrapie-infected neuroblastoma N1E-115 cell line (ScN1E). This up-regulation of IR protein was not due to an increased gene expression, suggesting a post-translational IR up-regulation. Proliferation studies showed that the increased IR level in ScN2a did not result in an increased insulin-mediated cell growth compared to normal N2a cells. Binding studies indicated that this apparent paradox was due to a 65% decrease in specific 125I-insulin binding sites in ScN2a when compared to the amount of immu-noreactive IR, although the binding affinity was unchanged. In addition, ScN2a showed an increased basal IR tyrosine phosphorylation together with an increased basal tyrosine phosphorylation of ERK-2, however, the insulin stimulated ERK2 phosphorylation was subsequently decreased in ScN2a (Östlund, et al., 2001b).

3.3 Protein tyrosine phosphatases

One of the processes downregulating the activity of PTKs is the action of protein tyrosine phosphatases (PTPs) (Tiganis, et al., 2007, Tonks, et al., 1996). The PTP family can be divided into receptor-like PTPs (RPTPs) and intracellular cytoplasmic PTPs. The structure of RPTPs suggests that they function as an interface between the extracellular environment of a cell and its intracellular signaling pathways. Their extracellular domains are highly variable, but all of them contain motifs that are implicated in cell adhesion such as immu-noglobulin like and fibronectin III like domains and contains two intracellu-

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lar phosphatase domains. Members of the intracellular PTPs lack a trans-membrane domain, possess a single phosphatase domain and have multiple protein:protein, protein:lipid domains either in the N- or C-terminus. These domains can target them to specific intracellular compartments in which the effective local concentration of a particular substrate is high, alternatively to keep them separated from their substrates (Frangioni, et al., 1992, Goldstein, 1993). PTPs play an important role in the control of RTK activity and the signaling pathways that they regulate. PTPs that dephosphorylate regulatory phospho-tyrosine residues will inhibit RTK activity and the biological responses me-diated by downstream effectors that depend on PTK activity (Tiganis, et al., 2007).

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4. LPS mediated production of nitric oxide

Bacterial lipopolysaccharide (LPS) is the major outer surface membrane component present in almost all Gram-negative bacteria and acts as an ex-tremely strong stimulator of the immune response in diverse eukaryotic spe-cies ranging from insects to humans. This chapter briefly describes the cellu-lar response to LPS exposure.

4.1 The bacterial endotoxin LPS

LPS consists predominantly of a lipophilic component anchored to the membrane and lipid A with a polysaccharide or oligosaccharide portion co-valently linked to it (Raetz, et al., 2002). This toxic agent is termed endotoxin, because it is a part of the bacterium in contrast to exotoxins that are released. LPS stimulation induces the expression of inflammatory mediators e.g. NO, mainly by immunocompetent cells e.g. macrophages and microglia. How-ever, also certain neuronal cells can respond to LPS with increased inducible nitric oxide synthase (iNOS) expression in the same manner as immuno-competent cells (Heneka, et al., 2001).

4.2 LPS receptor complex and LPS signaling

Probably the first host protein involved in LPS recognition is the LPS-binding protein (LBP) (Schumann, et al., 1990). LBP is an acute-phase protein, produced in the liver, which circulates in the bloodstream where it recog-nizes and forms a high affinity complex with the lipid A moiety of LPS. The role of LBP appears to be that of aiding LPS to dock at the LPS receptor complex by initially binding LPS and then forming a ternary complex with cluster of differentiation 14 (CD14), thus enabling LPS to be transferred to the LPS receptor complex composed of Toll-like receptor 4 (TLR4) and MD-2 (Hailman, et al., 1994, Tobias, et al., 1995). CD14 is found in two forms. The first is soluble CD14 (sCD14), the second and more extensively studied form of CD14, is the membrane bound (mCD14) attached to lipid rafts in the plasma membrane via a GPI-anchor (Haziot, et al., 1988, Wright, et al., 1990). The function of CD14 seems to be that of binding LPS and subsequently present-ing it to MD-2 and TLR4. Disruption of lipid raft formation by methyl-β-cyclodextrin has been demonstrated to inhibit LPS-induced TNF-α produc-tion in isolated human monocytes, suggesting a model in which LPS initially binds to CD14 whereby this ligation recruits different signaling molecules such as MyD88 and c-jun NH2-terminal kinase (JNK) to lipid rafts. LPS is then briefly released into the lipid bilayer where it interacts with TLR4 (Triantafilou, et al., 2002).

TLR4 is a member of a still rapidly growing family of innate immunity signaling proteins closely related by sequence similarity (Anderson, 2000). The

29

Toll-like protein shares a domain of about 200 amino acids present in the family of mammalian IL-1 receptor (IL-1R)-like proteins and has thus been termed the Toll/IL-1 receptor homology (TIR) domain.

When the LPS/LBP/CD14/TLR4/MD-2 complex is formed, the complex dimerizes and initiates the signaling machinery. TLR4 activates the ser-ine/threonine IL-1R-associated kinase (IRAK) via the MyD88 that possesses two domains, the death domain, interacting with the death domain of IRAK, and the TIR domain which, upon ligand binding to TLR4 is associated to the TIR domain of the receptor (Burns, et al., 1998, Wesche, et al., 1997). TNF receptor associated factor (TRAF) 6, another adaptor protein is subsequently phos-phorylated and recruited to IRAK (Cao, et al., 1996). TLR4 can use either Myd88 adaptor-like/TIR domain-containing adaptor molecule (Mal/TIRAP) and Myd88 or TRIF-related adaptor molecule (TRAM) and TIR domain-containing adaptor protein inducing interferon-β (TRIF) to signal to NFκB (Fig. 6) (Gay, et al., 2007).

In resting cells, the transcription factor NFκB associates with inhibitory IκB proteins. To release NFκB from the inhibitory IκB, the phosphorylation of IκB by IκB kinases (IKK) is required, resulting in the degradation of IκB by the proteasome (Rodriguez, et al., 1996). NFκB subunits are translocated to the nucleus where inducing expression of several pro-inflammatory cyto-kines and enzymes e.g. iNOS and cyclooxygenase-2 (COX-2).

Several molecules have been reported to transduce the signal to the IKK complex downstream of TRAF6 in the LPS signaling cascade. NFκB-inducing kinase (NIK) is thought to directly interact with IKKs to activate them in response to upstream stimuli, such as LPS. Transforming growth factor β-activated kinase 1 (TAK1) is also thought to be involved in LPS-mediated NFκB activation (Irie, et al., 2000), presumably acting upstream of NIK. In addition, an evolutionary conserved signaling intermediate in Toll pathways (ECSIT) has been shown to interact with TRAF6 leading to the activation of MAPK/ERK kinase kinase 1 (MEKK1). Activation of MEKK1 mediated by ECSIT leads to activation of NFκB (Hwang, 2001). Additionally, LPS has also been shown to activate the three members of the MAPK family i.e. ERK 1/2, JNK and p38 pathways (Anderson, 2000).

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Figure 6. LPS signaling. The LPS receptor complex activates several intracellular signaling pathways including NFκB leading to induction of inflammatory genes e.g. iNOS and COX-2 Figure adopted from (Gay, et al., 2007).

4.3 Nitric oxide and inducible nitric oxide synthase

Nitric oxide (NO) differs from classical neurotransmitters in that it is a gas and is thus able to penetrate into target cells to directly regulate many en-zyme systems e.g. guanylate cyclase. Consequently, NO has a range of in-fluence that extends well beyond the cell of origin. This property of NO makes it a useful signal for coordinating the activities of multiple cells in a localized region. NO diffuses only a few tens of micrometers from its site of production before it is degraded by spontaneous oxidation or scavenged by compounds e.g. superoxide anion or oxyhaemoglobin (Paakkari, et al., 1995). NO mediates e.g. synaptic transmission, neuronal excitability and immune-mediated cytoprotection and/or cytotoxicity. NO can be both beneficiary and detrimental, short impulses of small amounts of NO have beneficiary effects, for example during vasodilatation. Prolonged duration and high concentra-

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tions of NO are cytotoxic and the detrimental NO actions culminate at the formation of peroxynitrite (NO3

-) and its toxic radical products (OH·, NO2·)

mediating the excitotoxic cell death (Prast, et al., 2001). NO is produced by a family of three cellular nitric oxide synthases (NOS)

expressed in the brain, neuronal NOS (nNOS or NOS-1), inducible NOS (iNOS or NOS-2) and endothelial NOS (eNOS or NOS-3). eNOS and nNOS are mainly regulated by intracellular calcium and are constitutively ex-pressed. iNOS is under normal conditions expressed at very low levels and rapidly transcribed and expressed after stimulation with various inflamma-tory mediators e.g. proinflammatory cytokines and LPS and generates a sus-tained and elevated NO release (Dawson, et al., 1998, Xie, et al., 1994). All NOS isoforms catalyze the oxidation of the guanidine nitrogens of arginine to form NO and citrullin.

iNOS is expressed in numerous cell types as a consequence of the in-flammatory processes that follow infection, disease or tissue damage. In brain, iNOS expression has been well characterized in astrocytes, microglia cells and to a lesser extent in endothelial cells (Brosnan, et al., 1997, Murphy, et al., 1993). Additionally, recent reports have convincingly demonstrated iNOS expression and activity in response to inflammatory stimuli in neuronal cells. For example, exposure of neuroblastoma cells to proinflammatory cytokines resulted in expression of iNOS (Obregon, et al., 1997, Ogura, et al., 1996). More-over, combined stimulation with bacterial LPS and TNFα led to iNOS gene transcription and protein expression in neuronal differentiated PC12 cells (Heneka, et al., 1998).

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Methodology

Cell lines

All experiments in this thesis have been performed on persistently scrapie-infected murine cell lines (ScGT1-1 and ScN2a) and non-infected cells (GT1-1, N2a and BV-2).

GT1, N2a and BV-2

The GT1-1 (GT1) cell line is a subclone of a cell line originally generated by SV40 T-antigen immortalized gonadotropin-releasing hormone (GnRH) expressing neurons (Mellon, et al., 1990). The N2a cell line is a clone from the tumor line C1300, which was established from a spontaneous brain neuro-blastoma of an albino mouse (Amano, et al., 1972). The BV-2 cell line was es-tablished from murine primary microglia cell cultures, infected with a v-raf/v-myc recombinant retrovirus and exhibits phenotypical and functional properties of reactive microglia (Blasi, et al., 1990). Both N2a and GT1 cells were chosen as they are able to be persistently infected with scrapie. The BV-2 cell line was used in paper IV to serve as a control for studying an inflammatory response i.e. LPS-induced iNOS and COX-2 expression in N2a cells.

Scrapie-infected cell lines

ScGT1-1 is the only cell line shown to undergo scrapie-induced apoptosis (Schätzl, et al., 1997). However, our scrapie-infected GT1 cells (ScGT1a/b) showed neither caspase 3 activation nor DNA fragmentation as apoptotic markers. The ScN2a cells do not undergo PrPSc-related apoptotic cell death as readily as ScGT1 or neurons in vivo. Nevertheless, ScN2a cells demon-strate for example alterations in membrane fluidity and receptor-mediated signaling, as have been shown by us and others (Kristensson, et al., 1993, Östlund,

et al., 2001a, Östlund, et al., 2001b). The N2a cell line has been used by a large number of research groups to study the expression and metabolism of PrPC and PrPSc, the conversion from PrPC to PrPSc as well as the cytopathological changes during scrapie-infection, despite that these cells do not readily un-dergo apoptosis. The fact that not all neuronal cell lines undergo apoptosis upon scrapie-infection could be explained by that PrPSc is not neurotoxic per

se, but that PrPSc in the brain induces activation of microglia and astrocytes which release various neurotoxic substances, including NO and inflamma-

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tory cytokines. Time course studies of prion-infected brains suggest such a course of event including an inflammatory component in prion-induced neu-rodegeneration. Alternatively, the transformed phenotypes of immortalized neuronal cell lines may mask putatively neurotoxic events in finally differen-tiated neurons in the brain. However, chronically scrapie-infected cell lines provide an excellent model to study discrete molecular and cellular damage induced by the presence and propagation of PrPSc. These sublethal changes are important to characterize as they may lower the threshold to additional stress and provide alternative therapeutic targets in prion disease.

An alternative approach to study prion-induced pathology would be to treat the cells with a neurotoxic fragment of the prion protein and a large number of studies have been performed using PrP (106-126) (Brown, 2000, Forloni, et al., 1993). We have chosen not to work with PrP (106-126) since our interest has been to study the molecular changes in cells propagating PrPSc.

Generation of scrapie-infected cell lines; ScGT1a/b and ScN2a

ScN2a cells were a kind gift from Prof. S.B. Prusiner and the ScGT1a/b were generated at two different occasions in our laboratory. Having two independently scrapie-infected GT1 cell lines provides a good model for studying prion-induced molecular changes. In typical N2a cultures exposed to prions, <2% of the cells become infected and only low levels of prions are produced. In order to obtain persistently prion-infected cultures that produce sufficient quantities of PrPSc, the ScN2a was subcloned (Butler, et al., 1988). For studies aiming at elucidating potential neurotoxic changes induced by prion infection, these subclones are less suitable. The observed differences be-tween ScN2a and the uninfected N2a population might just represent cloning artifacts or be due to a selection artifact during the prion infection. The main difference between ScN2a and ScGT1a/b is that the latter cells were not subcloned before or after infection, thereby offering a reliable cell culture model of prion-induced changes, avoiding clonal differences (Bosque, et al., 2000). Using both ScN2a and ScGT1a/b provide several cellular models in which prion-induced changes can be reproduced and confirmed.

Detection of PrPSc

Prion infectivity has traditionally been linked to the presence of the protease-resistant core of PrPSc (PrP 27-30) (Bolton, et al., 1984), although in some ex-perimental setups, protease-resistant PrP could not be found in samples that contain prion infectivity (Lasmezas, et al., 1997, Manuelidis, et al., 1987). The other way around, not all PK-resistant PrP species are associated with prion infec-tivity (Shaked, et al., 1999). To demonstrate a correlation between PK-resistant PrP and prion infection, animals must be inoculated with the protease resis-tant PrP and monitored for clinical signs of scrapie, such as cerebellar ataxia, lethargy and rigidity. As RML-infected ScN2a and ScGT1 demonstrate both

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PK-resistance and prion infectivity in vivo (Butler, et al., 1988, Schätzl, et al., 1997) we have chosen to detect protease-resistant PrP to confirm infection of our cells.

The most straightforward approach would be to use a PrPSc-specific anti-body, but only a few have been described with moderate success in diagnos-tics (Korth, et al., 1997, Paramithiotis, et al., 2003). Instead, infection is commonly checked for by taking advantage of the partial PK-resistance of PrPSc ena-bling to distinguish between the two isoforms. Upon PK-digestion of PrPSc PrP 27-30 is formed, an N-terminal truncated prion protein which also is detected by the anti-PrP antibodies D13 and anti-PrP (SC-7694) upon im-munoblotting (Fig. 7). There are several alternate procedures to detect PrPSc in cell cultures e.g. cell blot, dot blot, western blot and ELISA. The most frequently used method here has been western blot, as western blot is both quantitative and qualitative i.e. individual glycosylation pattern and molecu-lar mass of PrPC and PrP 27-30 can be determined.

Figure 7. Western blot detection of the PK-resistant core, PrP 27-30 of PrPSc. PK-treated (+) and non-treated (-) GT1, ScGT1a/b, N2a and ScN2a cellular proteins were separated by 12% SDS-PAGE, transferred to nitrocellulose membranes and immunoblotted with anti-PrP (SC-7694) antibody.

Phenotypic differences between GT1 and ScGT1a/b

Experiments on growth rate of GT1 and ScGT1a/b were performed in our lab. Sub-cultivation procedures of the cells revealed that GT1 and ScGT1a cells reach confluence approximately 20-25% faster that ScGT1b cells. The underlying reason has not been established, but suggestions are either that cell division is slower for ScGT1b cells or that ScGT1b cells have a higher level of cell death. However, no pronounced apoptosis seems to occur among the ScGT1a and ScGT1b cells as detected by absence of DNA frag-mentation when analyzed on agarose gels as well as no cleavage of caspase-3 as shown by immunoblotting of whole cell lysates with anti-caspase-3

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antibody (unpublished results). This argues for a slower cell division of ScGT1b cells.

The levels of PrPC and PrPSc in GT1 and ScGT1a/b were analyzed as well as the effect of confluence and age on the levels of PrPC and PrPSc in GT1 and ScGT1a/b. This showed that lower PrPSc levels are detected in ScGT1a than in ScGT1b cultures. This is clearly visible after western immunoblot-ting analysis of PK treated samples (Fig. 7). At present we cannot distin-guish if a larger proportion of cells in the ScGT1b population are infected i.e. produce PrPSc, or if certain individual cells in the ScGT1b population produce an increased amount of PrPSc. A possible explanation for the higher level of PrPSc in ScGT1b compared to ScGT1a could be that ScGT1b cells have a lower growth rate than ScGT1a or that ScGT1a cells are less infected with prions.

Additionally, PrPC and PrPSc levels are independent of the confluence and age of the GT1, ScGT1a and ScGT1b cells, respectively. No clear trend of how the PrPC or PrPSc expression varies over time and cell confluence could be seen in GT1 and ScGT1a/b (personal communication with K. Löfgren and A. Wahl-ström).

Src and Fyn antibodies

Epitopes of the antibodies used for either purification by immunoprecipita-tion or detection by immunoblotting of Src and Fyn are shown in Fig. 8. The N16 and the FYN3 antibodies were used for the in vitro Src and Fyn kinase assays respectively. These antibodies were chosen because i) they are spe-cific for the respective kinase, ii) they are directed to a region which is not involved in the kinase regulation and iii) binding of antibodies to other epi-topes on the kinase may per se stimulate activity as reviewed in (Roskoski, 2004). The monoclonal antibody Clone 327 binds specifically to the SH3 domain of Src kinase (Lipsich, et al., 1983) and has been used for detection of Src kinase. The levels of active Src and Fyn kinase were determined using the monoclonal antibody Clone 28 (Kawakatsu, et al., 1996) that recognizes ac-tive Src and active Fyn. The Clone 28 antibody binds to the regulatory C-terminal tyrosine in its dephosphorylated state, indicating that autoinhibition of the SFK activity is prevented (see chapter 3.1.1 for Src regulation). Unfor-tunately, no antibodies worked in our hands for immunoblotting immuno-precipitated Fyn. Instead, we determined the protein levels by analyzing the intensity of the Fyn immunosignaling from whole cell lysates. However, to analyze the amount of active Fyn, immunoblotting of Clone 28 immunopre-cipitates with the specific Fyn antibody worked excellent.

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Figure 8. Antibodies used in this study directed to Src and/or Fyn. Clone 28 is the only antibody recognizing active forms of Src and Fyn by binding to the regulatory C-terminal Y527 in its unphosphorylated state.

Isolation of lipid rafts

The most widely used assay for studying lipid rafts is to isolate DRMs. When the plasma membrane is extracted with the non-ionic detergent Triton X-100 at 4°C, placed at the bottom of a density gradient and subjected to centrifugation, the DRMs float away from the soluble and non-membranous material. The nontoxic B subunit of cholera toxin (CTXB) binds the glyco-sphingolipid GM1, and has been widely used as a putative reporter for the distribution of lipid rafts (Harder, et al., 1998). GT1 and ScGT1 cells were incu-bated with CTXB for 30 minutes, before harvesting and subjected to density centrifugation, in order to work as a raft marker. CTXB was further visual-ized by immunoblotting with an anti-CTXB antibody.

As discussed in chapter 2.6 in this thesis, as isolation of DRMs requires detergent extraction at 4°C, the membrane is certainly affected by the de-creasing temperature and probably more of the membrane is in the Lo phase at lower temperatures than at 37°C. Additionally, an experimental artifact could be that the detergent per se may induce liquid ordered domains. Thus, studying lipid rafts in vitro may not exactly reflect the physiological state in the living cell. However, model membrane studies showed that Lo-Ld phase separation could occur at 37°C in lipid mixtures with physiological levels of each of these lipid classes, making phase separation in cell membrane plau-sible (Ahmed, et al., 1997, Silvius, 2003). As long as the assay is performed identi-cally for both cell types and as long as differences between non-infected and infected cells are analyzed this method is definitely appropriate.

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Protein deglycosylation

Deglycosylation of glycoproteins can be performed by either enzymatic or chemical methods. Enzymatic methods involve the incubation of the glyco-protein with exo- and/or endoglycosidases. However, extensive incubation with a number of enzymes is sometimes necessary for near-complete degly-cosylation. In paper III four different deglycosylating enzymes were used in an attempt to achieve an aglyco-IRβ. PNGase F cleaves the innermost N-acetylglucoseamine of most N-linked glycan groups, Endo H cleaves the chitobiose core of N-linked high mannose and some N-linked hybrid oligo-saccharides leaving the innermost GlnNAc intact and O-glycanase releases Ser- and Thr-linked N-acetylgalactosamine (GalNAc) from glycoproteins. While PNGase F, Endo H and O-glycanase are all endoglycosidases, neura-minidase is an exoglycosidase, removing the terminal sialic acid residues, which may inhibit the accessibility of PNGase F and inhibit the enzymatic activity of O-glycanase. Treatment with neuraminidase, PNGase F and O-glycanase would in theory result in a completely deglycosylated protein. Deglycosylation of glycoproteins can also be carried out using chemical reagents. Among the principal methods that have been employed for chemi-cal deglycosylation, anhydrous trifluoromethane-sulfonic acid (TFMSA) has been found to be the most successful reagent for cleaving N- and O-linked glycans non-selectively from glycoproteins, leaving the primary structure of the protein intact (Edge, et al., 1981).

Determination of nitrite production

The half-life of released NO is only a few seconds and it is rapidly degraded by spontaneous oxidation or scavenged by e.g. superoxide anion, and there-fore the direct measurement of NO requires specialized equipment (Sung, et

al., 1992). Nitrite (NO2-), together with nitrate (NO3

-) is the stable end prod-ucts of NO metabolism and accumulates in the cell culture medium. In order to measure the total amount of NO, nitrate is reduced to nitrite prior to de-termination of nitrite levels. However, as the nitrate concentration repre-sented only 10% of the total nitrite concentration, this step was omitted in our study. In addition, the aim was to detect a difference in the LPS-induced NO production between N2a and ScN2a cells, and the absolute amount of NO was not required, therefore omission of this additional step was appro-priate. NO production was measured as NO2

- production according to the protocol of Stuehr and Nathan (Stuehr, et al., 1989).

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Results and Discussion

Paper I and II

In paper I and II, studies of the expression, activity and localization of SFKs have been performed. Previous studies have shown that crosslinking of PrPC with specific antibodies leads to clustering of lipid rafts in Jurkat and human peripheral blood T cells. This study also showed that the crosslinking of PrPC induced signal transduction shown by recruitment and activation of Fyn kinase (Stuermer, et al., 2004). Other studies have shown that anti-GM1 medi-ated crosslinking of lipid rafts (Kasahara, et al., 2000), as well as that of GPI-anchored proteins, including PrPC (Hugel, et al., 2004, Mouillet-Richard, et al., 2000), induce a rapid activation of SFKs and a transient increase in the tyrosine phosphorylation of several substrates (Kasahara, et al., 2000). Thus, we hypothe-sized that a PrPSc-oligomer may bind and ligate several PrPC molecules and thereby cluster PrPC-containing lipid rafts and induce an aberrant activation of lipid raft-associated SFKs (Fig. 9). This in turn may be a contributing event leading to synaptic degradation and/or apoptosis of nerve cells in prion disease. Further, PrPC has been shown to co-immunoprecipitate with Src kinase (Mattei, et al., 2004, Morel, et al., 2004), indicating co-localization of PrPC and SFKs. In addition, crosslinking of PrPC with specific monoclonal anti-bodies triggers neuronal apoptosis in vivo (Solforosi, et al., 2004). These studies together, strengthen our hypothesis that PrPSc in lipid rafts may cluster PrPC and induce an activation of intracellular SFKs that may contribute to apop-tosis of neurons exposed to prions.

Figure 9. Model for how prions may induce an uncontrolled SFKs activity (figure by K. Bedecs).

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Increased level of active Src kinase in scrapie-infected cells (paper I)

Src kinase has been shown to be important in neurodegeneration including prion disease, see previous chapter “SFKs in neurodegenerative disease”, therefore we first wanted to further investigate the expression and activity of Src kinase and its downstream signaling targets.

The level of active Src kinase was determined using the monoclonal anti-body Clone 28. No increase in the specific activity of Src was found al-though a significant increase in the levels of active Src protein was observed in ScGT1a and ScGT1b as well as in ScN2a compared to their non-infected counterparts. The increased Src kinase activity in scrapie-infected cells was confirmed by an in vitro kinase assay. The kinase activity was efficiently inhibited by the Src specific inhibitor PP2, confirming that specific Src kinase activity was studied.

Despite the vast number of reports published regarding Src kinase activ-ity, virtually nothing is known about the transcriptional regulation of this kinase. This is remarkable considering that Src gene expression is develop-mentally regulated in the brain (Cartwright, et al., 1988) and transcriptionally upregulated in regenerating neurons (Le Beau, et al., 1991). However, besides a transcriptional and/or translational regulation, recent evidence suggest that Src is also subject to ubiquitin-dependent degradation. Specifically, activated forms of Src-kinases are turned over more rapidly than wild-type or kinase-inactive Src proteins (Harris, et al., 1999). Activated forms of Src and Fyn bind to and phosphorylate c-Cbl that acts as an E3 ubiquitin ligase (Andoniou, et al., 2000, Joazeiro, et al., 1999). As an E3 ubiquitin ligase, c-Cbl promotes poly-ubiquitination of associated Src-kinases, targeting them for proteasomal degradation. Ubiquitin-dependent proteolysis regulates the stability and function of many key regulatory proteins and is also responsible for the elimination of misfolded proteins. Evidence suggest that newly synthesized misfolded and/or mismodified PrPC is degraded by the proteasome (Yedidia, et

al., 2001) and increased ubiquitin-immunoreactivity has been found in both human and animal prion diseases. Additionally, in scrapie-infected mouse brains, the level of ubiquitinated proteins was significantly increased, con-current with the appearance of PrPSc and a decline in two endopeptidase activities associated with proteasome function (Kang, et al., 2004). This sug-gests that the accumulation of PrPSc in infected cells overwhelms the ubiq-uitin-proteasome system and therefore, if this system in general, or c-Cbl in particular, is deregulated in scrapie-infected cells, the result may well be elevated levels of activated Src.

To evaluate whether the elevated levels of active Src protein in ScGT1b and ScN2a cells are due to an increase in gene transcript number or altered post-transcriptional/translational processing of the protein, semiquantitative RT-PCR was performed. As shown in Fig. 10, the Src mRNA level was significantly increased in ScGT1b compared to GT1 as well as in ScN2a

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compared to N2a. No difference in GAPDH mRNA expression was ob-served between the cells. Determination of the Src/GAPDH ratio, showed a 29% increase in Src mRNA level in ScGT1b compared to GT1 cells. In ScN2a compared to N2a cells the increase in Src mRNA levels was deter-mined to 31%. The increase in Src mRNA levels corresponds to the increase in Src protein levels in ScGT1b cells compared to GT1 cells that was 32% indicating an elevated transcription of Src in ScGT1b cells. However, in ScN2a cells an increase in gene transcript number was shown to be 31%. This has to be compared to the increase in Src protein levels that was 75%. Thus, the increase in Src protein levels cannot only be explained by the in-crease in Src mRNA levels in ScN2a cells (unpublished results).

Figure 10. Protein and mRNA levels of Src in GT1/ScGT1b and N2a/ScN2a. Left, levels of active Src protein in respective cell type (paper I). Right, total RNA from the indicated cell types were extracted, reverse transcribed and amplified by PCR with specific primers for Src and GAPDH. Samples from 30 PCR cycles were sepa-rated on a 1% agarose gel and visualized by ethidium bromide staining. Determina-tion of the Src/GAPDH ratio, showed a 29 ± 1.5% (P = 0.0051, n = 3) increase in Src mRNA level in ScGT1b compared to GT1 cells. In ScN2a compared to N2a cells the increase in Src mRNA level was determined to 31 ± 3.9% (P = 0.048, n = 3). IP=immunoprecipitation, IB=immunoblotting.

To test the hypothesis that the accumulation of PrPSc in prion-infected cells overwhelms the ubiquitin-proteasome system and therefore, if this system is deregulated in scrapie-infected cells leads to an accumulation of active Src, treatment with the proteasome inhibitors MG 132 and Lactacystin was per-formed. Preliminary results are shown in Fig. 11 that show Src immunoreac-tivity of whole cell lysates from GT1, ScGT1b, N2a and ScN2a after treat-ment with MG 132 for 5 hours and Lactacystin for 24 hours. If the increase

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in Src protein level in scrapie-infected cells is due to an inhibited proteasome activity, treatment of non-infected cells with a proteasome inhibitor is ex-pected to yield a similar increase/accumulation of the Src protein level in non-infected cells as seen in scrapie-infected cells. N2a cells (Fig. 11B) but not GT1 cells (Fig. 11A) showed a dose-dependent increase in the accumula-tion of a possible degradation product of Src at ~55 kDa, recognized by the Src-specific antibody, Clone 327 after MG 132 and Lactacystin treatment. Densitometric measurements of total Src i.e. both the mature 60 kDa protein and the ~55 kDa degradation product revealed a dose-dependent increase up to 126% after treatment with 1 µM MG 132 compared to non-treated cells. The same result was shown after treatment with Lactacystin although the increase was not as obvious as after treatment with MG 132. The accumula-tion was dose-dependent and after treatment with 2 µM Lactacystin for 24 hours the increase in Src protein level compared to non-treated N2a cells was 90%. However, in ScN2a cells no similar changes in Src protein accu-mulation were detected after treatment with either MG 132 or Lactacystin. These results suggest that the accumulation of active Src protein in ScN2a is due to both a disturbed transcriptional process and an altered degradation via the ubiquitin/proteasome system (unpublished results).

Figure 11. Src protein accumulation after MG 132 and Lactacystin treatment. 2 x 106 GT1 or ScGT1b cells or 5 x 105 N2a or ScN2a cells were seeded in 6 cm culture dishes three days before treatments. Cells were treated for 24 hours in presence or absence of 0.1-2 µM Lactacystin for N2a/ScN2a and GT1/ScGT1b. MG 132 treat-ment was performed for 5 h in 0.05-1 µM for N2a/ScN2a and in 0.05-0.1 µM for GT1/ScGT1b. After treatment, whole cell lysates were separated on 10% SDS-PAGE and analyzed by western blot using the specific Src antibody Clone 327.

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At present, we have no clear explanation for the increased Src expression since we have not studied the transcriptional and translational regulation of Src kinase. In addition, few reports are published regarding the transcrip-tional regulation of Src kinase. One study however, shows that Src transcrip-tion is increased in regenerating Schwann cells (Le Beau, et al., 1991). Accord-ingly, scrapie-infected cells may upregulate Src protein in response to nerve cell loss in prion disease as an attempt to regenerate destroyed neurons through a mechanism that is still unknown.

A prion-induced dysregulation of the proteasome activity as indicated in ScN2a could not be confirmed in ScGT1b. However, the increase in Src expression in ScGT1b compared to GT1 can be explained by the increased transcription i.e. the increase in mRNA levels corresponds to the increase in protein expression. Further, the increase in Src protein expression in ScN2a compared to N2a seems to be due to an increase in transcription number in combination with inhibited proteasome activity. The different results ob-tained in ScGT1b and ScN2a may be due to their different neuronal origins i.e. CNS and peripheral nervous system (PNS) respectively. The cell lines may react with alternative response mechanisms to prion infection. Other scrapie-infected cell lines have to be treated with MG 132 and Lactacystin in order to evaluate if proteasome activity is affected during scrapie-infection.

Increased specific tyrosine kinase activity of Fyn in scrapie-infected cells (paper II)

The SFK member Fyn is dually acylated i.e. it is both myristoylated and palmitoylated. This allows Fyn to be attached to lipid rafts to a larger extent than mono-acylated kinases (e.g. Src) (Alland, et al., 1994). Palmitoylation has been shown to be required for localization of SFKs to rafts (Robbins, et al., 1995, Shenoy-Scaria, et al., 1994). Therefore, Fyn has a central role in our hy-pothesis that the presence of PrPSc in lipid rafts may induce cross-linking of PrPC and a subsequent raft-clustering and activation of raft-signaling by e.g. raft-resident SFK members (Fig. 9). Due to lack of suitable antibodies for immunoblotting immunoprecipitated Fyn, we determined the protein levels by analyzing the intensity of the Fyn immunosignaling from whole cell lys-ates. To analyze the amount of active Fyn, immunoblotting of Clone 28 im-munoprecipitates with the specific Fyn antibody was performed. This, to-gether with an in vitro kinase assay of Fyn activity showed a significant in-crease in the level of active Fyn in scrapie-infected cells even though the protein level of Fyn kinase was decreased. This indicates an increased spe-cific activity of Fyn in scrapie-infected cells. Since no differences were shown in Fyn mRNA levels between non-infected and infected cells we con-cluded that the decrease in protein levels of Fyn in scrapie-infected cells is due to a post-transcriptional/translational disturbance. As mentioned earlier, SFKs, including Fyn are subject to ubiquitin-dependent degradation involv-ing the SFK-substrate c-Cbl E3 ubiquitin ligase. Activated forms of Fyn are

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turned over more rapidly and if this system works satisfactory, one explana-tion for the decreased Fyn protein level in scrapie-infected cells is that in-creased proteasomal degradation would be a result from increased specific activity of Fyn kinase. This is however in contrast to one of the suggested mechanisms of the increased Src level in paper I.

One possible explanation for the increased specific Fyn kinase activity in scrapie-infected neurons is described by our original hypothesis (Fig. 9) that PrPSc oligomers in raft domains may induce ligation of PrPC (analogous to antibody mediated crosslinking) and thereby recruit and activate Fyn kinases. Due to the inherent metabolic stability of PrPSc, this could be a long lived association and result in a chronically elevated Fyn kinase activation.

Increased tyrosine phosphorylation in scrapie-infected cells (paper I and II)

An increased overall tyrosine phosphorylation was shown in scrapie-infected cells compared to non-infected cells, indicating a disturbed signaling. In ScGT1 this increase was shown to be due to the increased level of active Src kinase as well as the increased specific activity of Fyn kinase as shown by PP2 treatment (paper I, Fig. 4). However, in ScN2a cells the tyrosine phos-phorylation was not decreased to the basal levels as in N2a cells after treat-ment with the Src specific inhibitor PP2. Higher concentrations than 1 µM PP2 were lethal to both N2a and ScN2a cells, which could be one explana-tion for that the tyrosine phosphorylation in ScN2a could not be decreased completely with PP2. It can also be a result of that other tyrosine kinases might be upregulated or that one or several tyrosine phosphatases are down-regulated. It remains to be determined which one of these mechanisms is the correct one.

Both quantitative and qualitative changes in protein tyrosine phosphoryla-tion were observed. In ScGT1 cells, mainly usually tyrosine phosphorylated proteins showed an increased tyrosine phosphorylation, especially phospho-proteins of approximately 66, 120 and 180 kDa. In ScN2a cells phosphopro-teins of approximately 97, 120, 150 and 180 kDa, were heavily phosphory-lated in ScN2a whereas some of these were not at all phosphorylated in N2a.

Of note, the tyrosine kinase inhibitor STI571 has been shown to induce cellular clearance of PrPSc in prion-infected cells by activation of lysosomal degradation (Ertmer, et al., 2004). Thus, a possible scenario is that PrPSc induces an increased tyrosine kinase activity, which slows down cellular clearance of PrPSc and thereby facilitates PrPSc replication.

At present it is not known which proteins are affected by the increased ty-rosine phosphorylation. It is of great interest to identify these proteins to further understand the mechanisms underlying neuronal cell death in prion disease. Several approaches may be taken to identify them, some of which were initiated in our lab.

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To investigate if any of the affected proteins in ScN2a or ScGT1 cells is a membrane protein, WGA purification, recognizing N-linked glycosylated proteins, of whole cell extracts was performed. This study revealed that a fraction of the tyrosine phosphorylated proteins from ScN2a cells detected in anti-phosphotyrosine immunoprecipitates was also retained on the WGA-beads. This indicates that some, but not all, of the phosphoproteins were glycosylated (data not shown). In WGA-purified fractions from ScGT1a/b one predominantly affected protein was the phosphoprotein at 120 kDa (data not shown).

To further identify the phosphoproteins, in-gel trypsin digestion of the WGA-purified proteins or anti-phosphotyrosine immunoprecipitates fol-lowed by separation by SDS-PAGE may be performed. The resulting frag-ments can be identified by mass spectroscopy using matrix-assisted laser desorption ionization mass fingerprinting (MALDI-TOF) and on-line liquid chromatography tandem mass spectrometry (LC-MS/MS) peptide sequenc-ing. We performed this, in collaboration with Dr. Tomas Bergman at Karo-linska institute, on the 150 kDa tyrosine phosphorylated protein from ScN2a cells. Results revealed a yet unknown protein, corresponding to a full-length transcript, RIKEN cDNA 4932439K10 (unpublished results). In a first attempt to characterize this protein, antibodies directed against a peptide sequence of the full length protein may be generated.

Another approach to identify the affected phosphoproteins in prion-infected cells, anti-phosphotyrosine immunoprecipitates can be immunoblot-ted with a panel of antibodies directed against putative candidate proteins. Criteria for the proteins would be that they are SFK substrates, phosphopro-teins and have a molecular mass corresponding to those phosphoproteins that showed an increased tyrosine phosphorylation. Interestingly, c-Cbl has a molecular mass of 120 kDa. Other SFK substrates interesting to evaluate in this respect could be e.g. FAK and PYK2 that also have molecular masses of approximately 120 kDa.

ERK2 was previously shown to have an increased basal activity, i.e. was more phosphorylated in ScN2a compared to N2a (Östlund, et al., 2001b). Since ERK2 is a well known substrate of SFKs, this protein may be one of the proteins with an increased tyrosine phosphorylation visible on the blot in paper I, Fig. 3, at least in ScN2a cells a phosphoprotein at 42/44 kDa is visi-ble which could correspond to ERK1/2.

Both ScN2a and ScGT1 showed an increased tyrosine phosphorylation of phosphoproteins of ~120 kDa and ~180 kDa, compared to uninfected cells. Interestingly, proteins of the same molecular mass have been reported to be phosphorylated upon treatment with H2O2 (Monteiro, et al., 1996) or peroxyni-trite in human neuroblastoma cells (Li, et al., 1998). Therefore, an alternative explanation to the aberrant tyrosine phosphorylation in scrapie-infected cells could be an increased oxidative stress, previously described in both prion-infected brains and ScGT1 cells (Guentchev, et al., 2000, Milhavet, et al., 2000, Wong, et al., 2001). Both PTKs and PTPs are highly susceptible to oxidative

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agents (Griendling, et al., 2000, Nakashima, et al., 2002). To determine whether oxi-dative intracellular conditions may contribute to the increased tyrosine phos-phorylation in scrapie-infected cells, oxidizing conditions may be mimicked by treating uninfected cells with prooxidants. Correspondingly, scrapie-infected cells can be treated with antioxidants in an attempt to reverse the scrapie-induced changes.

Plasma membrane distribution of PrPC, PrPSc and active Fyn (paper II)

Since the conversion of PrPC to PrPSc has been shown to occur in lipid rafts (Kaneko, et al., 1997, Naslavsky, et al., 1997), studies of the distribution of PrPC/PrPSc and Fyn kinase in lipid rafts were performed. Interestingly, a shift in PrPC/PrPSc distribution in scrapie-infected cells to heavier floatation frac-tions compared to non-infected cells was shown.

One possible explanation for the distribution of PrPC/PrPSc/Fyn in heavier density fractions is as mentioned earlier the clustering of rafts as a result of PrPSc-induced crosslinking of PrPC. Clustering of rafts leads to an activation of lipid rafts as signaling platforms. Upon activation of rafts, several signal-ing molecules are recruited to rafts (Simons, et al., 2000) and this could increase the raft density and lead to an apparent redistribution of PrPC and PrPSc to heavier fractions due to their higher protein/lipid ratio.

An alternative explanation for the presence of PrPC/PrPSc in heavier floa-tation fractions in ScGT1 could be that these proteins move from lighter to heavier raft-populations in the presence of prions. In fact, different raft-populations with different densities have been suggested to exist in the mem-brane (Naslavsky, et al., 1997).

The first explanation goes well with our hypothesis that prion propagation may induce clustering of rafts via a PrPSc-mediated ligation of PrPC and a following recruitment and activation of Fyn kinase in rafts. This suits the finding that active Fyn follows the distribution of PrPC and PrPSc in the plasma membrane.

In addition, Fyn associated with rafts is catalytically active whereas those interacting with non-raft membranes are catalytically down-regulated (Ilangumaran, et al., 1999). This could be one of the reasons for the increased specific activity of Fyn kinase in scrapie-infected cells compared to non-infected cells.

Physiological implications (paper I and II)

Although mechanisms for the prion-induced increase in Src and Fyn kinase activities are suggested in previous chapters, the physiological consequences for an increased tyrosine kinase activity in prion disease are unknown. This part suggests some models for how an increased Src and Fyn tyrosine kinase activity may contribute to neuronal death in prion disease.

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One possible mechanism may be a sustained increased ERK activity as ERKs are SFK substrates. ERK2 was previously shown to have an increased basal activity, i.e. was more phosphorylated in scrapie-infected than in unin-fected neuroblastoma cells (Östlund, et al., 2001b). Additionally, clustering of PrPC with monoclonal antibodies in GT1-7 cells induced phosphorylation of ERK1/2 (Monnet, et al., 2004, Schneider, et al., 2003). Activated ERK initiates downstream events regulating cell growth, differentiation, survival and neu-ronal cell death (Seger, et al., 1995). Evidence indicates that neurotoxicity can be mediated through increased, prolonged or persistent ERK1/2 activation. Persistent activation of ERK has been shown to be responsible for gluta-mate- (Stanciu, et al., 2000) and ischemia- (Tsuji, et al., 2000) induced oxidative toxicity in neuronal cultures and brain. Increased ERK phosphorylation has also been shown to occur in neurons with an abnormal phosphorylation in brains of AD patients (Perry, et al., 1999). The increased basal ERK activity might be due to activation by an increased activity of SFKs in prion-infected cells and may reflect a neuro-stressing condition for differentiated neurons in the adult brain.

An alternative explanation for how increased Src and Fyn kinase activity may contribute to synaptic reorganization and neuronal death in prion dis-ease is through their regulation of potassium and/or calcium channels e.g. the NMDAR. An important function of SFKs in the adult CNS is to regulate the activity of K+ and Ca2+ channels. The NMDAR is an important receptor par-ticipating in the fast excitatory synaptic transmission throughout the CNS. The function of NMDARs is regulated by e.g. glutamate and glycine ex-tracellularly but also intracellularly by SFKs (Salter, et al., 2004). Several stud-ies have shown an involvement of NMDARs and excitotoxicity contributing to neuronal death in prion disease (Scallet, et al., 1997). Antagonists to NMDAR e.g. MK-801, have been shown to have a cytoprotective effect on PrPSc-induced toxicity in rat cortical cell cultures (Müller, et al., 1993). The toxicity of the synthetic fragment PrP(106-126) has been shown to involve Ca2+ uptake through voltage-sensitive Ca2+ channels as well as the activation of NMDA receptors (Brown, et al., 1997). However, the exact mechanism for the prion-induced excitotoxicity is not known. The results presented in this thesis i.e. the increased levels of active Src (paper I) and the increased specific Fyn kinase activity (paper II) in scrapie-infected cells may offer an explanation for the upregulated NMDAR activity and the following excitotoxicity found in prion infected brains. An interesting study showed that binding of the NMDAR-antagonist, MK-801 was decreased in hippocampal regions of prion-infected mice (Diez, et al., 2001) indicating a decreased expression of NMDARs during prion disease. This downregulation might be a defense mechanism to excitotoxicity caused by the increased SFK activity.

A third possibility is that a prolonged SFK activity is toxic to neuronal cells. In fact, in post-mitotic cells such as CNS neurons an inappropriately upregulated SFK activity and/or mitogenic signaling may lead to nerve cell death as illustrated by neurodegeneration and cell death of SV40 T-antigen

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expressing Purkinje neurons (Feddersen, et al., 1992). In addition, PrPSc is a well known trigger of microglia and astrocyte proliferation during prion disease. However, a paradox is that PrPSc is toxic to post-mitotic cells like neurons (DeArmond, et al., 1992).

It has also been demonstrated that Src deficiency or blockade of Src activ-ity in mice provides cerebral protection following stroke (Paul, et al., 2001). Further support for that SFKs are key players in neurodegeneration is pro-vided by an observation that the SFK-inhibitor PP2 reduces focal ischemic brain injury (Lennmyr, et al., 2004). However, these studies do not exclude the involvement of other cell types than neurons in the neuroprotective effect offered by SFK inhibitors.

Finally, SFKs are implicated in integrin-regulated events by the ability to phosphorylate β1 integrin as well as other proteins that are associated with integrin-nucleated focal adhesion complexes e.g. paxillin, p130cas and FAK (Burridge, et al., 1996, Clark, et al., 1995, Petch, et al., 1995). Recently, one study showed that integrins mediate Aβ induced cell cycle activation and neuronal death and that PP2 treatment rescued the Aβ-induced cell death. This indi-cates an involvement of SFKs and integrin activation in neuroprotective conditions involving an unscheduled neuronal cell cycle activation. Possibly, a similar mechanism may be responsible for neuronal cell death in prion infected brains (Frasca, et al., 2007).

Paper III

As summarized in chapter 3.2.1, previous results reported in our group showed that scrapie-infection may affect IR and IGF-1R expression and function in scrapie-infected neuroblastoma cell lines. Those studies showed that scrapie-infection induces a 2- and 4-fold increase in IR and IGF-1R protein levels respectively, without a corresponding increase in specific 125I-insulin and 125I-IGF-I binding sites or cell growth in insulin- or IGF-1-containing medium. In contrast to the elevated IGF-1R expression in ScN2a cells, receptor binding studies revealed an 80% decrease in specific 125I-IGF-1-binding sites. This decrease in IGF-1R-binding sites was shown to be caused by a 7-fold decrease in IGF-1R affinity. A similar decrease (65%) in specific 125I-insulin binding was found in ScN2a cells when the amount of 125I-insulin binding sites was related to the increase in immunoreactive IR; however, in the case of the IR, the receptor binding affinity was unchanged (Östlund, et al., 2001a, Östlund, et al., 2001b). The significant decrease in 125I-insulin-binding sites in ScN2a cells, despite the unchanged affinity, indicated the presence of two populations of IRs: one receptor population that is nor-mally expressed, processed and functional, as determined by an unchanged receptor binding affinity, and another receptor population that does not rec-ognize insulin, but is detected by the anti-IRβ antibody.

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In addition, the IRβ-chain in ScN2a cells exhibited an increased electro-phoretic mobility, with a molecular mass of 85 kDa compared to 95 kDa in N2a cells (Östlund, et al., 2001b). These findings indicate a scrapie-induced dys-regulation of the post-translational processing of the IR, including protein folding, glycosylation and trafficking. In paper III, the localization and al-tered post-translational processing of the atypical IR subunits in ScN2a have been studied.

Localization of IR in ScN2a cells and elevated expression of IR/IGF-1R hybrid receptors in ScN2a

The indication of two populations of IRs in ScN2a where one of them has no or very low affinity for insulin, but is detected by the anti-IR antibody was suggested as described above. At first, this was thought to be due to aberrant processing of the IR, leading to a population of receptors not reaching the plasma membrane but accumulating intracellularly. To examine whether the increased IR protein level in ScN2a could be detected on the cell surface i.e. if the IR is correctly processed and transported to the cell surface or if the increased IR level in ScN2a reflects an increased intracellular accumulation, surface-biotinylation followed by anti-IRβ immunoprecipitation was per-formed. Detection of biotinylated IR revealed no difference in the ratio of surface-localized IR versus total amount of IR between N2a and ScN2a, indicating a correct processing and trafficking of IR in ScN2a despite the decreased number of 125I-insulin binding sites.

However, this unresponsive population of IR, immunoreactive, present on the cell surface but not binding insulin, could be represented by IR/IGF-1R hybrids. An important comment here is the existence of the two IR isoforms. They are the result of alternative splicing of exon 11 in a tissue-specific manner (Seino, et al., 1989) and the IR-A (Ex11-) lacks, whereas the IR-B (Ex11+) contains the sequence coding for 12 amino acids in the C-terminal part of the α-subunit (Ebina, et al., 1985, Ullrich, et al., 1985). Brain and also pe-ripheral neurons show predominantly the IR-A type, which displays higher affinity to insulin compared to the peripheral IR-B type (Mosthaf, et al., 1990, Sugimoto, et al., 2000). Thus, a hybrid receptor containing IR-A binds insulin with ~10-20-fold lower affinity than the IR, whereas a hybrid receptor con-taining IR-B hardly recognizes insulin at all (Pandini, et al., 2002). Preliminary data show that N2a and ScN2a cells express predominantly the IR-A form as determined by RT-PCR, consequently, the hybrid receptors formed with the IR-A isoform will display a 10-20-fold lower affinity than homotypic IR and thus hardly bind 125I-insulin at the low tracer concentration used (40 pM). Hence, the unaltered surface localization of IR and the two-fold increase in the level of hybrid receptors in ScN2a, suggest that this “second” unrespon-sive population consists of hybrid receptors, since these have a much lower affinity (Kd~10 nM) and will not bind insulin at the low tracer concentration used.

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Altered molecular mass of the IRβ-subunit in ScN2a

The β-subunit of the IR in ScN2a was found to display a changed electro-phoretic mobility, with an apparent molecular mass of 85 kDa instead of 95 kDa in the non-infected cells, although the unprocessed IR precursor in ScN2a cells migrated similarly as in the non-infected cells. A similar, but not as significant shift of the IRβ-chain was also observed in ScNIE-115 (Östlund,

et al., 2001b). After excluding both a difference in the degree of phosphoryla-tion between the IRβ-subunit in N2a and ScN2a as well as differences in the proteolytic processing, the degree of glycosylation was examined. The IR was immunoprecipitated and enzymatically deglycosylated with PNGase F, Endo H, neuraminidase and O-glycanase, separately or in different combina-tions. All of these treatments, except treatment with O-glycanase, which had no significant effect, significantly increased the electrophoretic mobility of the IRβ from both N2a and ScN2a, however the shift remained. Following treatment with a combination of PNGase F, neuraminidase and O-glycanase, the difference in molecular mass between IRβ in ScN2a and N2a disap-peared. Moreover, it could be concluded from these results that IRβ in N2a and ScN2a contains high-mannose glycans (released by Endo H), hybrid- and complex types oligosaccharides (released by PNGase F) as well as ter-minal sialic acid (released by neuraminidase), but the relative amounts and possible differences of the amounts of the above mentioned carbohydrates between N2a and ScN2a remain to be established. The results from the en-zymatic deglycosylation were confirmed by chemical deglycosylation, using anhydrous TFMSA, which cleaves N- and O-linked glycans non-selectively from glycoproteins.

Interestingly, alterations in glycosylation have also been found between PrPC and PrPSc. Although they contain the same set of bi- tri- and tetraanten-nary N-linked oligosaccharides, the relative proportions of individual gly-cans differ. Compared to PrPC, PrPSc contains decreased levels of glycans with bisecting GlnNAc residues and increased levels of tri- and tetraanten-nary sugars. This change is consistent with a decrease in the activity of N-acetylglucosaminyltransferase II toward PrPC in cells where PrPSc is formed (Gambetti, et al., 2003, Hill, et al., 2003, Rudd, et al., 2001, Rudd, et al., 1999). Conse-quently, a plausible explanation for the altered glycosylation of IRβ in ScN2a could be a scrapie-induced perturbation of the glycosylation machin-ery.

Paper IV

At first this study aimed at investigating if ScN2a cells co-cultured with im-munocompetent cells such as microglia and astrocytes, could elicit an in-flammatory response i.e. if scrapie-infected cells could act as an inflamma-tory stimuli. Two microglia cell lines BV-2 and NTW8 and an astrocytic cell

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line Ast-1 were cultivated together with ScN2a cells. A readout for possible inflammatory responses, production of NO, was measured as accumulated nitrite in the medium of co-cultured cells. However, no response was ob-tained. In order to generate a synergistic response in the immunocompetent cells, the bacterial endotoxin LPS was added to the co-cultures, but unfortu-nately, no effect except from the LPS stimulation, was observed. However, in control wells containing only N2a or ScN2a, the N2a cells reacted to the LPS stimulation with a production of NO, whereas the ScN2a completely lacked this ability. LPS treatment in N2a induced a concentration- and time-dependent increase in NO production, from 0.01 to 10 µg/ml LPS between 24 and 96 hours with a 50-fold increase in NO production in presence of 1 µg/ml LPS after 96 hours. Consequently, this project changed its focus to investigate the mechanism underlying the scrapie-induced inhibition of LPS-stimulated NO production.

Lack of LPS-induced iNOS expression in ScN2a

To verify that the increased NO production in N2a was caused by increased expression of iNOS and not increased expression of nNOS, amplification of nNOS cDNA from N2a and a microglia cell line, BV-2 used as a reference for LPS-induced iNOS expression, was performed. The cells showed no increase in nNOS mRNA after LPS treatment for 4 hours, a time point at which the corresponding levels of iNOS mRNA were significantly elevated in both cell types. In addition, N2a and BV-2 were treated with LPS in the presence or absence of 2 mM aminoguanidine (AG), a selective iNOS in-hibitor. In the presence of 2 mM AG, the LPS-induced NO production was completely inhibited at all time points in both cell types, and this lack of NO production was due to inhibition of iNOS activity and not to its expression, as shown by unchanged iNOS expression in AG-treated N2a.

In contrast, LPS treatment of ScN2a induced no NO production at all. This absence of LPS-stimulated NO production was shown to be due to a complete lack of LPS-induced iNOS expression. Several mechanisms could explain the lack of iNOS expression in ScN2a. The LPS receptor complex is built up of several interacting proteins of which the presence is necessary to elicit proper signaling and the co-localization of prions in lipid rafts could putatively disrupt the receptor complex, or the necessary clustering to lipid rafts. Alternatively, molecules acting downstream of the LPS receptor com-plex could be disturbed by prion infection. As the first candidate, we studied the expression of CD14, since it is in the LPS-recognizing part of the recep-tor complex and after the preparation of paper IV, RT-PCR studies on CD14 revealed a complete lack of CD14 mRNA in ScN2a, even though the gene encoding CD14 was still present, as shown by PCR of genomic DNA (Fig. 12). This finding explains the complete lack of iNOS expression in ScN2a, as CD14 is a necessary component, although not signaling itself, in the re-

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ceptor complex mediating the LPS-induced response (Akashi, et al., 2000, Stelter, 2000).

Figure 12. Total lack of CD14 mRNA in ScN2a. Genomic DNA was isolated or cDNA was prepared from each cell type and amplified by PCR with CD14 or GAPDH specific primers for 28 cycles. Samples were run on 1% agarose gel and visualized by ethidium bromide staining.

This complete lack of iNOS expression, caused by the deficiency of CD14 mRNA, might appear paradoxical, as iNOS immunoreactivity is upregulated in scrapie-infected brains of mice and hamster. However, in scrapie-infected brains, the increased iNOS expression is generated by activated/reactive astrocytes (Ju, et al., 1998, Kim, et al., 1999), either by reacting on scrapie-infected neurons or extracellular PrPSc deposits or by their innate PrPSc pro-duction (Ye, et al., 1998).

Down-regulation of CD14 transcription as a defense towards invading bacteria or virus is not an unusual effect in immunocompetent cells. For example, mRNA levels of CD14 are down-regulated by cytomegalovirus infection of human alveolar macrophages (Hopkins, et al., 1996) and human resident intestinal macrophages, while TLR expression is intact (Smith, et al., 2001). Thus, the absence of CD14 mRNA in ScN2a could be an action to increase survival. An intracellular accumulation of CD14 has been observed in human polymorphonuclear neutrophils (Wagner, et al., 2003), acting as a pool of readymade proteins that can be recruited to the outside of the plasma membrane within minutes after stimulation by e.g. LPS.

Further studies are needed to characterize the mechanisms behind prion-induced CD14 down-regulation. Also the protein levels can be determined as intracellular accumulation of CD14 has been reported. In a first attempt, to restore LPS-responsiveness in ScN2a cells, exogenous administration of soluble CD14 together with LPS-treatment can be performed. Furthermore, in order to study if prion infection inhibits any downstream signaling, in addition to CD14, CD14 independent activation of the same signaling cas-cade can be performed. This can be done by treatment with different cyto-kine combinations, however, we found that non-infected N2a cells did not respond with iNOS expression or NO production to treatment with any cyto-kine mixtures and therefore, we had no appropriate control and further stud-ies need to be undertaken.

52

Sammanfattning på svenska – Hur reagerar nervceller på prioninfektion?

Prionsjukdomar är en grupp av sjukdomar som innebär att hjärnvävnad förstörs och som kan drabba nästan alla däggdjur. Sjukdomen har alltid dödlig utgång och då den påverkar hjärnan är de tidiga symptomen ofta demens, att det bildas ”hålrum” i hjärnan och att proteiner som vanligtvis kan brytas ner av kroppen klumpas ihop. Detta leder till att cellerna i hjärnan succesivt dör och därmed också till slut den drabbade individen.

Den mest uppmärksammade prionsjukdomen är BSE (bovine spongiform encephalopathy) mer känd som ”galna-ko-sjukan” och som namnet antyder drabbar den nötkreatur. En förödande upptäckt var att ”galna-ko-sjukan” kan överföras till människor efter intag av smittat nötkött. Hos människor kallas sjukdomen för Creutzfeldt-Jacobs sjukdom och förutom att smitta via intag av infekterat kött så har den också visats kunna smitta från patient till patient inom sjukvården till exempel via ej tillräckligt rengjorda hjärnkirurgi-instrument eller via hormonbehandlingar. Sjukdomen kan också vara ärftlig då olika mutationer i genen för prion-proteinet leder till Creutzfeldt-Jacobs sjukdom.

Prionsjukdomar orsakas av att ett protein som kallas prion-proteinet (PrPC) och som vanligtvis finns hos alla däggdjur, ändrar sin tre-dimensionella form. Mycket forskning pågår kring vilken funktion PrPC normalt har i cellerna och troligtvis har proteinet en rad olika uppgifter i hjärnan. Bland annat har PrPC visats hjälpa nervceller att ta upp metallen koppar vilken är viktig i cellens försvar mot fria syreradikaler. Man har också visat att PrPC kan vara involverad i kommunikationen mellan olika proteiner i cellen för att slutligen avgöra om just den cellen ska överleva och därmed föröka sig, eller om cellen ska begå självmord. Nya PrPC-proteiner bildas hela tiden i cellen och efter ca sex timmar bryts de ner av lysosomen, en av cellens egna ”sopstationer”. Under prionsjukdom ändrar PrPC sin tredimensionella struktur från att exempelvis se ut som ett klot till att se ut som en kub. Kuben kallas i detta fall för PrPSc eller prion och med denna förändring ändras också många av PrPCs egenskaper. Till exempel så är PrPSc motståndskraftig mot nedbrytning vilket leder till att kuberna ansamlas och bildar klumpar i hjärnan. Klumparna blir större och större eftersom nya PrPC-proteiner hela tiden bildas. Mekanismen för omvandlingen av PrPC till PrPSc är fortfarande ett mysterium och stor forskningskraft läggs världen över för att förstå hur detta går till. Hittills har man klargjort att

53

förvandlingen till PrPSc sker på cellens yta i det så kallade cellmembranet som omger hela cellen. Cellmembranet är uppbyggt av fetter och vissa delar av cellmembranet innehåller speciella fetter som har hög bindningsförmåga till bland annat PrPC samt till många andra proteiner som deltar i kommunikationen mellan cellen och dess omgivning. Dessa speciella delar i cellmembranet kallas ”lipid rafts” (=fettflottar) och det är bland annat dessa som jag studerat i denna avhandling. Eftersom förvandlingen från PrPC till PrPSc sker i lipid rafts och eftersom dessa också innehåller många andra proteiner som är viktiga för cellens överlevnad antog jag att ihop-klumpningen av PrPSc i lipid rafts kan påverka dessa livsnödvändiga proteiner negativt och bidra till att hjärncellerna dör.

I avhandlingen har både friska och prioninfekterade celler från mushjärnor använts för att jämföra hur olika proteiner påverkas under prionsjukdom.

De viktigaste upptäckterna som presenteras i det här arbetet är att två proteiner som heter Src och Fyn, som också är lokaliserade till lipid rafts, är mer aktiva i prioninfekterade celler och detta i sin tur leder till att många andra kommunicerande proteiner inne i cellen aktiveras och cellen får en signal att begå cellulärt självmord. Troligtvis beror detta på att när förvandlingen av PrPC till PrPSc sker i lipid rafts och när PrPSc klumpas ihop i lipid rafts, så klumpas även andra proteiner ihop och startar en okontrollerad aktivering av dessa Src- och Fyn-proteiner. Mitt arbete visar en av troligtvis många förklaringar till varför hjärncellerna dör under prionsjukdom. Att förstå vad som påverkas negativt i hjärnan under prionsjukdom är viktigt för att i framtiden veta hur och mot vad botemedel och/eller vaccin ska tillverkas.

54

Acknowledgements

I would like to thank ALL people at DBB who makes the department to such a great place to work at. There are some who deserve a special thank;

Coco, min handledare, tack för att du gav mig möjligheten att doktorera på DBB trots oviss framtid för priongruppen. Tack också för dina smarta ideér och din värdefulla feedback på allt jag gjort. Du har lärt mig massor!!

Astrid, för att du håller bra ordning och reda på DBB samt att du gjort det möjligt för mig att avsluta mina doktorandstudier i lugn och ro. Tack.

Tack Stefan för att du bryr dig om allt och alla som rör doktoranderna på DBB och att du samtidigt håller ordning på hela naturvetenskapliga fakulteten. Du ett praktexemplar till chef!

Elzbieta, tack för att du adopterat mig till din grupp under mitt sista år. Det har varit lärorikt samt bra för mig att få lite feedback från ”icke-prion-människor”.

Tack till all administrativ personal med Ann i spetsen. Ni är alltid otroligt hjälpsamma och ni är superbra på att ordna fester!

Stefan Nilsson, utan dig vet jag inte om jag hade fixat de där 15 obligatoriska poängen... Tack för grymt plugg-samarbete! Tack också för 4 års utomordentligt samarbete på BK10!

Ullis, du ska ha ett fett tack för att du förmodligen är en av världens bästa på att peppa! Din positivitet smittar av sig på hela DBB, keep on smiling!! Tack också för bra samarbete i doktorandrådet!

Thanks to the whole bunch of people at the 5th floor! You all contribute to such a great atmosphere in our corridor. The SN-group people and my for-mer office mates; Anders, Helen, Pedro, Tiago and Tomas (yes Tomas, I know you did not sit in our office but you spent more time there than in your own…). The ÅW-group for nice lunch discussions; Maria, Malin, Alex,

Tuulia, Amélie and Hanna E.

55

Onsdagsklubben, lovely onsdagsklubben. Anna N, Anna Karin och

Nyosha. Det är ni som har fått mig att gå till jobbet med ett leende även de dagar då själva jobbet inte riktigt har gått som man vill... Ni fattar inte (jo, det gör ni) hur värdefullt det är att ventilera jobbet och allt annat i livet med er varje onsdagkväll över lite billigt vin och BJ-glass. Yoshi loves you!

Tack hela priongruppen! Nina, du var den första att förklara för mig vad en prion är... Heléne och Pernilla, det är er förtjänst/ert fel (?) att jag började doktorera. Att jobba med er var superkul! Tack för allt ni lärt mig, all peppning och att ni nu har läst min avhandling i omgångar. Snart kan vi BARA prata om annat än prionforskning... Tack också till Daniel, Erik och Ana för bra samarbete den tid ni var i labbet. Tack Anna W för din korta men uppskattade tid i prionlabbet, du är ”Fröken korrekturläsning”, för att du gör det så bra. Kajsa, tack för fantastiskt samarbete den tid du varit i gruppen, utan dig hade det varit tungt. Du är en sann forskare med grymma ideér och ett glödande intresse, fortsätt så!! Tack också för alla roliga konferenser vi varit på!

Ett STORT tack till ALLA mina vänner utanför DBB och universitetet! Ni är många, ni vet vilka ni är och ni betyder alla väldigt mycket för mig!

John Gustaf, för att du är du och för att du är min! Tack också för att du har fått mig att tänka på annat än avhandlingsskrivande de senaste månaderna.

Och till slut, min familj;

Farmor, för ditt aldrig sinande intresse för vad jag gör och för att du alltid får mig på gott humör. Du är en förebild.

Mina småsyskon, Eva och Johan. Tack för er sjuka humor och för att ni alltid finns där. Ni är vansinnigt duktiga på allt ni gör och det inspirerar!

Mamma och Pappa, ni hade rätt igen... jag klarade det. Tack för att ni ALLTID stöttat mig, för all uppmuntran, omsorg och kärlek! Jag kunde inte valt bättre föräldrar.

56

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