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Oncostatin M and leukemia inhibitory factor in excitotoxicity Moidunny, Shamsudheen

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ONCOSTATIN MAND LEUKEMIA INHIBITORY FACTOR IN EXCITOTOXICITY

Shamsudheen Moidunny Stellingen behorende bij het proefschrift

Oncostatin Mand Leukemia Inhibitory Factor in Excitotoxicity

Shamsudheen Moidunny

1. Although LIF and OSM are known to have several overlapping properties, their neuroprotective effect is mediated by differentmechanisms (Chapters 2 and 6). 2. STAT-3 signal transduction is not required for LIF- and OSM- induced protection of mouse embryonic cortical neurons against glutamate-toxicity (Chapter 3). 3. In mouse astrocyte cultures, STAT-3 activation byOSM reduces cellular glutamate uptake and thereby promotes excitotoxic death of neurons (Chapter 5). 4. OSM plays "opposite" roles in excitotoxicity (promote or inhibit), depending on the target cell of the cytokine action (Chapters 2 and 5). 5. Adenosine and its receptors play an important role in mediating the effects of IL-6-type cytokines in the central nervous system. (This thesis) 6. Knowledge on the effects of IL-6-type cytokines on different cell types of the central nervous system is incomplete. It is therefore premature to state that elevated levels of these "inflammatory" cytokines as observed in injured or diseased brain, is good or bad. 7. The man who makes no mistakes does not usually make anything. (Edward Phelps) 8. Right data only pave half the way to a good research article, the other half depends on how you build and sell the story to your target audience. 9. Working in the west seems to be a good way for an Indian to improve his or her time management skills. 10. Lack of hierarchy while providing the freedom to learn, without imposing it ... these are the strongest.a!::;etsof Dutch culture. 11. Good researchers and photographers share a similar aptitude: the ability to en er le observable. •:.!ntralc ll Ml!dische M Bibliotheek C Groningen G The research described in this thesis was conducted at the Department of Neurosciences, Section Medical Physiology, University Medical Center Groningen (UMCG), University of Groningen (RUG), The Netherlands. This work was supported by the RUG, and the graduate school of Behavioral and Cognitive Neurosciences (BCN). Printing of this thesis was financiallysupported by: RUG, UMCG and BCN.

ISBN: 978-90-367-5382-1

Copyright© S. Moidunny 2012. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means without permission of the author and the publisher holding the copyright of the articles.

Cover design: Ming-San Ma Layout and Printing: OffPage, Amsterdam, www.offpage.nl RIJKSUNIVERSITEIT GRONINGEN

ONCOSTATIN M AND LEUKEMIA INHIBITORY FACTOR IN EXCITOTOXICITY

Proefschrih

ter verkrijging van het doctoraat in de Medische Wetenschappen aan de Rijksuniversiteit Groningen op gezag van de Rector Magnificus, dr. E. Sterken, Ccntrale -.., in het openbaar te verdedigen op Me V di�che M woensdag 14 maart 2012 1! ib1iotheek C om 16.15 uur Groningcn G

door L

Shamsudheen Moidunny geboren op 11 oktober 1982 te Kunnamkulam, Kerala, India Promotores: Prof. dr. K.P.H. Biber Prof. dr. H.W.G.M. Boddeke

Beoordelingscommissie: Prof. dr. U.L.M. Eisel Prof. dr. M. Schmidt Prof. dr. A.P. IJzerman CONTENTS Page

Abbreviations 6

Chapter 1 General Introduction and Thesis Outline 9

Chapter 2 lnterleukin-6-type cytokines in Neuroprotection and Neuromodulation: 47 Oncostatin M, but not Leukemia inhibitory factor, requires neuronal

Adenosine A1 Receptor function

Chapter 3 Neuroprotection induced by Oncostatin Mand Leukemia Inhibitory 71 Factor against excitotoxicity requires Akt, ERKl/2 and NF-KB, but not STAT3 signaling

Chapter 4 Adenosine A28 receptor mediated LIF release from astrocytes protects 91 cortical neurons against excitotoxicity

Chapter 5 Oncostatin M promotes excitotoxicity by inhibiting astrocytic 115 glutamate uptake

Chapter 6 Summary and Discussion 139 Nederlandse samenvatting (l)Ql(}Jd!f6IDnuotUJnOo Acknowledgements ABBREVIATIONS

adenosine A1 receptor A?aCSF artificialcerebrospinal fluid ALS amyotrophic lateral sclerosis AMPA a-amino-3-hydroxy-5-methyl-4-isoxazole propionate ATP adenosine-5' -triphosphate BBB blood-brain barrier BDNF brain derived neurotrophic factor BFA brefeldin A CADO 2-chloroadenosine cAMP cyclic adenosine monophosphate CBM cytokine binding module CLC cardiotrophin-like cytokine CNS central nervous system CNTF ciliary neurotrophic factor CNTFR ciliary neurotrophic factor receptor COX-2 cyclooxygenase-2 CPA N6-cyclopentyladenosine CT-1 cardiotrophin 1 DMEM Dulbecco's modifiedeagle medium DMSO dimethyl sulfoxide DPCPX 8-cyclopentyl-l, 3-dipropylxanthine DRG dorsal root ganglia EAAC l excitatory amino acid carrier 1 EAAT excitatory amino acid transporter EAE experimental autoimmune encephalomyelitis EDTA ethylenediaminetetraacetic acid ELISA enzyme-linked immunosorbent assay EPSC excitatory post-synaptic current ER endoplasmic reticulum ERKl/2 extracellular signal regulated kinase 1/2 FCS fetal calf serum GAPDH glyceraldehyde-3-phosphate dehydrogenase GFAP glial fibrillaryacidic protein GLAST glutamate-aspartate transporter GLT-1 glutamate transporter 1 GM-CSF granulocyte macrophage colony-stimulating factor gpl30 glycoprotein 130 HBSS Hank's Buffered Salt Solution HPRTl hypoxanthine phosphoribosyl transferase 1 IFN-y interferon y IL interleukin iNOS inducible nitric oxide synthase IP3 inositol triphosphate

6 JAK janus kinase JNK/SAPK c-jun N-terminal kinase/stress-activated protein kinase LDCV large dense-core vesicles LIF leukemia inhibitory factor LIFR leukemia inhibitory factor receptor LME L-leucine methyl ester LPS lipopolysaccharide MAP2 microtubule-associated protein 2 MAPK mitogen-activated protein kinase MCAO middle cerebral artery occlusion mGluR metabotropic glutamate receptor MMP matrix metalloproteinases MTT 3-(4, 5-dimethylthiazol-2-yl-) 2, 5-diphenyltetrazolium bromide NECA 5'-N-Ethylcarboxamide NF-KB nuclear transcription factor-kappa B NGF nerve growth factor NGS normal goat serum NMDA N-methyl-D-aspartic acid NMG N-methyl-D-glucamine NNT-1 novel neurotrophin 1 088 Odyssey blocking buffer OSM oncostatin M OSMR oncostatin M receptor PBS phosphate buffered saline PCR polymerase chain reaction

PGE2 prostaglandin E2 Pl propidium iodide Pl3K phosphatidylinositol-3-kinase PIAS protein inhibitor of activated STAT PKA protein kinase A PKC protein kinase C PLC phospholipase C PNS peripheral nervous system PVDF polyvinylidene fluoride rpl32A ribosomal protein L32-A socs suppressor of cytokine signaling STAT signal transducers and activators of transcription TBOA DL-threo-�-Benzyloxyaspartic acid TGF-� transforming growth factor � TNF-a tumor necrosis factor a VDCC voltage-dependent calcium channel XIAP X chromosome-linked inhibitor of apoptosis protein

7

GENERAL

CHAPTER

INTRODUCTION

AND

THESIS

OUTLINE CHAPTER 1

CONTENTS CENERAL INTRODUCTION

1. EXCITOTOXICITY 11 1.1. The neurotransmitter glutamate 11 1.2. Glutamate metabolism 11 1.3. Glutamate transporters 12 1.4. Neurons and astrocytes contribute to extracellular glutamate 12 1.5. Glutamate receptors 13 1.6. Excessive calcium influx leads to excitotoxicity 15 1.7. Glial cells, excitotoxicity and neuroinflammation 15 1.8. Glutamate transport and excitotoxicity in neurological disorders 17 1.9. Potential targets to prevent excitotoxic damage 17 1.9.1. NMDA receptor antagonism 17 1.9.2. Regulation of glutamate transporter expression and activity 18 1.9.3. Modulation of NMDA receptor activity and glutamate release 18

2. ADENOSINE IN THE CENTRAL NERVOUS SYSTEM 19 2.1. Adenosine receptors 20 2.2. Neuromodulation by adenosine: role in excitotoxicity 20 2.3. Immune modulation by adenosine 22

3. INTERLEUKIN (IL)-6-TYPE CYTOKINES IN THE CENTRAL NERVOUS SYSTEM 24 3.1. Receptors: types, activation and intracellular signaling pathways 24 3.2. Cytokine and receptor expression in the CNS 27 3.3. IL-6-type cytokines and CNS inflammation 28 3.4. Anti-excitotoxic and neuroprotective effectsof IL-6-type cytokines 30

4. THESIS OUTLINE 31

10 GENERAL INTRODUCTION

1. EXCITOTOXICITY 1.1. The neurotransmitter glutamate II Glutamate is recognized as the major excitatory neurotransmitter in the mammalian central nervous system (CNS). In addition to having pivotal role in cell metabolism as non-essential amino acid, excitatory input by glutamate is considered crucial for physiological functions of the brain including cognition, learning and memory. Moreover, glutamate contributes to the CNS development by induction and elimination of synapses as well as by its effects on migration, differentiation and death of cells. The brain contains very high amounts of glutamate (about 5 - 15 mM per kg wet weight, depending on the region) (Schousboe, 1981), which is mostly located intracellularly with the highest concentrations present inside nerve terminals (Ottersen et al, 1992; Storm-Mathisen et al, 1992). Only a small fraction of brain glutamate is normally present in the extracellular fluid (outside or between cells) and in the cerebrospinal fluid, 3 - 4 µMand about 10 µM, respectively (Hamberger et al, 1983; Lehmann et al, 1983). Maintenance of extracellular glutamate at low levels is of critical importance for the following reasons: 1) High extracellular glutamate might result in low signal-to-noise ratio in both synaptic and extrasynaptic transmission, and 2) Excessive activation of glutamate receptors can be toxic.

1.2. Glutamate metabolism The fact that glutamate concentrations in the extracellular fluid need strict regulation requires the existence of a highly efficientsystem forthe clearance of extracellular glutamate. Since there does not seem to be any enzyme that is capable of metabolizing glutamate extracellularly, the only way to regulate extracellular glutamate concentrations is by transporter-mediated uptake into glial cells and neurons (Figure 1), a highly efficient process for extracellular glutamate removal that has been widely reported (for review see, Danbolt, 2001). Glutamate that is taken

Figure 1: The glutamate-glutamine cycle. This pathway represents a way of recycling transmitter glutamate. Glutamate released from a nerve Nerve terminal terminal by exocytosis is taken up by glutamate transporters present presynaptically (1), postsynaptically (2) and extrasynaptically in astroglia (3). Astroglia detoxifies glutamate by converting it to glutamine in an ATP-dependent process. Glutamine is subsequently released from the glial cells by means of other glutamine transporter (4). Neurons take up extracellular glutamine through specialized transporters {S) and convert it back to glutamate. Synaptic vesicles are loaded with glutamate from cytosol by means of vesicular glutamate transporter (6). The glutamate released from into the Postsynapticneuron synaptic cleft activates its receptors at postsynaptic terminal (7). Adapted from reference: Danbolt, 2001.

ll CHAPTER 1

up by neurons is either reused as transmitter or used for metabolic purposes such as protein synthesis. In astrocytes, glutamate may be converted into glutamine by the glia-specific enzyme glutamine synthetase (Martinez-Hernandez et al, 1977). Glutamine is then released into the extracellular fluid, taken up by neurons through the glutamine transporters where it can be used for synthesis of new glutamate (by the mitochondrial enzyme glutaminase), which is subsequently packed into vesicles and released by exocytosis (Nicholls and Attwell, 1990; Sudhof, 1995). This process is generally referred to as the glutamate-glutamine cycle (Figure 1). The concentration of glutamine in the extracellular fluid under normal conditions ranges between 200 and 500 µM (Gjessing et al, 1972; Hamberger et al, 1983). Glutamine, in contrast to glutamate, is non-toxic, does not activate glutamate receptors and does not seem to compromise normal neurotransmission.

1.3. Glutamate transporters As mentioned before, glutamate transport across plasma membranes is mainly mediated through glutamate transporters. At least five different "high-affinity" excitatory amino + + acid transporters (EAATs), which catalyze Na - and K -coupled transport of glutamate and aspartate, have been described (Table 1): EAATl or glutamate-aspartate transporter (GLAST) (Storck et al, 1992; Tanaka, 1993), EAAT2 or glutamate transporter 1 (GLT-1) (Pines et al, 1992), EAAT3 or excitatory amino acid carrier 1 (EAACl) (Kanai and Hediger, 1992), EAAT4 (Fairman et al, 1995) and EAAT5 (Arriza et al, 1997). In addition, there is evidence for alternative spliced variants of GLT-1 (Meyer et al, 1998) and GLAST (Huggett et al, 2000). Moreover, transport of glutamate through sodium-independent, chloride-dependent high-affinity transporters (Zaczek et al, 1987) and glutamate-cystine exchangers (Warr et al, 1999) have been reported, further increasing heterogeneity of glutamate transporters and complexity of the transport process. Furthermore, the absence of any other "significant" mechanism for the clearance of extracellular glutamate makes these transporters highly important for long-term maintenance of non-toxic levels of glutamate in the extracellular fluid. Moreover, glutamate transporter activity might modulate duration and intensity of synaptic transmission and might contribute to the cellular supply of glutamate (for review see, Danbolt, 2001). In the CNS, expression of GLAST, GLT-1 and EAACl is widespread (Table 1), the former two are predominantly expressed in glial membranes and the latter (neuron-specific glutamate transporter) in post-synaptic neuron terminals. Expression of GLT-1 is absent during early developmental stages, whereas GLAST expression is consistent throughout development. However, in the adult brain GLT-1 is the most abundant glutamate transporter (especially in the hippocampus and the cerebral cortex, but not in cerebellum where GLAST expression is higher), and is responsible for the majority of the glutamate transport. On the other hand, expression of EAAT4 and EAAT5 seems to be restricted to the cerebellum and the retina, respectively.

1.4. Neurons and astrocytes contribute to extracellular glutamate In the extracellular fluid, release of glutamate and its transport into cells occur constantly and in a rapid manner. For many years, it was thought that the presence of glutamate in the extracellular fluid was merely the result of synaptic release from nerve terminals by exocytosis of synaptic vesicles. However, eventually it became clear that in fact most of the released glutamate is of non-exocytotic (non-vesicular) origin (Jabaudon et al, 1999).

12 GENERAL INTRODUCTION

Table 1. Nomenclature, distribution and Km values (for L-glutamate) of cloned glutamate transporters. Adapted from reference: Tilleux and Hermans, 2007. II Human Human proteins Rodent proteins Localization genes (Km) (Km) Cell types Tissue distribution SLClAl EAATI (48 µM) GLAST (77 µM) Astrocytes, oligodendrocytes Bergmann glia of cerebellum SLC1A2 EAAT2 (97 µM) GLT-1 (2 µM) Astrocytes, microglia, Cerebral cortex, oligodendrocytes, neurons hippocampus SLC1A3 EAAT3 (62 µM) EAACl (12 µM) Neurons, Astrocytes Brain (Postsynaptic terminals) SLC1A6 EAAT4 (2.5 µM) Purkinje cells, Astrocytes Cerebellum

SLC1A7 EAAT5 (64 µM) Photoreceptors Retina

Furthermore, growing evidence clearly suggests that astrocytes contribute largely to the released glutamate content in the extracellular fluid (for review see, Volterra and Meldolesi, 2005). During pathological conditions like ischemia, insufficientsupply of ATP causes excessive release of glutamate, both from the nerve terminals and astrocytes (Rossi et al, 2000; 2007). The excessive glutamate release from neurons has been proposed to occur largely by the reversal of uptake by glutamate transporters (Jensen et al, 2000; Rossi et al, 2000), and from astrocytes by calcium-dependent vesicular exocytosis (Araque et al, 2000; Innocenti et al, 2000; Jeremie et al, 2001). The calcium-dependent exocytosis of glutamate from astrocytes can be induced by the activation of ATP- (Jeremie et al, 2001) and metabotropic glutamate receptors (Chen et al, 2005) and might be regulated by calcium levels inside mitochondria (Reyes and Parpura, 2008). Despite this evidence, it is not yet clear whether astrocytes can release glutamate by exocytosis in vivo (Hamilton and Attwell, 2010), since most of the studies so far were performed in vitro. Furthermore, it is now clear that additional mechanisms may be involved in astrocytic glutamate release including release through swelling-induced anion channel opening, glutamate exchange via the cystine-glutamate antiporter, release through ionotropic purinergic receptors and functional unpaired connexons, "hemichannels", on the cell surface (for review see, Malarkey and Parpura, 2008). On the other hand, increased levels of extracellular glutamate might also arise from impaired uptake as a result of reduced glial glutamate transporter expression and/or activity, as commonly observed in neurological disorders (Gegelashvili et al, 2001; Maragakis and Rothstein, 2004). In line with this, Jabaudon and colleagues have shown that pharmacological inhibition of glutamate uptake using DL-threo-�-Benzyloxyaspartic acid (TBOA) resulted in rapid accumulation of extracellular glutamate (Jabaudon et al, 1999). In conclusion, excessive release from neurons and astrocytes as well as impaired uptake by astrocytes can all result in elevated extracellular concentrations of glutamate that might be detrimental to neurons and glial cells of the CNS.

1.5. Glutamate receptors Glutamate exerts its effectsby acting on receptors that are located at the plasma membrane of target cells (Figure 2). In the CNS, most neurons and even glial cells express receptors for glutamate (Bergles et al, 2000; Conti et al, 1999; Pocock and Kettenmann, 2007; Shelton and

13 CHAPTER 1

NMDA KAINATE

+ Na Ca2+

AMPA mGLuR

Glu -._

GDP-+GTP

Figure 2: The ionotropic (NMDA, Kainate, AMPA) and metabotropic (mGLuR) glutamate receptors.

McCarthy, 1999; Verkhratsky and Kirchoff, 2007), indicating the broad range of its targets of action. Glutamate receptors are broadly classified into ionotropic (ligand gated ion channels) and metabotropic (G-protein coupled transmembrane protein) receptors. Three ionotropic receptor types have been characterized based on specific binding of glutamate analogues, cx-amino-3-hydroxy-5-methyl-isoxazole-4-propionic acid (AMPA), kainate and N-methyl-0- aspartate (NMDA). AMPA receptors are subdivided into GluRl-4, and kainate receptors into GluRS, GluR6, GluR7, KAl and KA2. NMDA receptors consist of several subunits: NRl, NR2 (A, B, C and D) and NR3 (A and B). The ionotropic receptors conduct only sodium or both sodium and calcium ions. AMPA receptor channels, which display low affinityfor glutamate, open rapidly when exposed to glutamate, but desensitize quickly. In contrast, NMDA receptor channel complexes have much higher affinities and are slowly inactivated. In addition to glutamate, the NMDA receptor requires the binding of a co-agonist, glycine, to allow the receptor to function. A glycine-binding site is found on the NRl and NR3 subunits, whereas the glutamate binding site is found on NR2 subunits (Figure 2). Hence, expression of the NRl subunit alone does not produce a functional receptor, but co-expression of one or more NR2 subunits is required to form functional channels. At resting membrane potentials, NMDA receptors are largely inactive due to a voltage-dependent block of the channel pore by magnesium ions. Depolarization releases this channel block and permits passage of calcium as well as other ions. The metabotropic glutamate receptors consist of 8 types (mGluRl-8) which are subdivided

14 GENERAL INTRODUCTION into groups I (mGluRl and mGluRS), II (mGluR2 and mGluR3) and 111 (mGluR4, mGluR6, mGluR7 and mGluR8). Group I receptors are coupled to phospholipase C and thereby to inositol II triphosphate and diacylglycerol production, whereas groups II and Ill are negatively coupled to adenylate cyclase. The receptor groups I and II are activated by (±)l-amino-cyclopentane­ trans-1,3-dicarboxylic acid (trans-ACPD) while group Ill receptors are activated by L(+)-2- amino-4-phosphonobutyric acid (L-AP4). Activation of glutamate receptors induces calcium influx through the ionotropic glutamate receptor channels (NMDA and AMPA) and through the voltage-dependent calcium channels (VDCC), resulting in the increase of cytosolic calcium concentrations. Moreover, activation of metabotropic glutamate receptors induces production of inositol triphosphate (IP3) that results in calcium release from the endoplasmic reticulum (ER), through the IP3 receptor channels. Intracellular calcium regulates several calcium-dependent enzymes such as phospholipases, nucleases and proteases and thereby controls most physiological processes, cell injury and programmed cell death (apoptosis).

1.6. Excessive calcium influx leads to excitotoxicity As mentioned before, excessive amounts of extracellular glutamate might result in continuous activation of neuronal NMDA receptors in a tonic rather than a phasic manner, which ultimately leads to cell death by a process generally referred to as excitotoxicity (Olney and Sharpe, 1969). It is noteworthy that vulnerability to excitotoxicity might vary between neurons of different brain regions depending on the expression levels of glutamate receptors (Kovacs et al, 2001). The major intracellular mediator of the excitotoxic process is calcium (Choi, 1992), which therefore requires strict regulation. Cytosolic calcium loads may be bufferedby calcium-binding proteins (calbindins), which serve as endogenous anti-excitotoxic proteins (Mattson et al, 1991 ). In addition, mitochondria and ER play a crucial role in the regulation of intracellular calcium levels, and thereby in excitotoxicity (Duchen, 2000; Frandsen and Schousboe, 1991; Mattson et al, 2000). Activation of IP3 receptors on the ER induces calcium release into the cytosol, whereas transient elevation of cytosolic calcium may induce its uptake by mitochondria. There is evidence that both inhibition of calcium release from the ER as well as enhancement of mitochondrial calcium sequestration, can protect neurons against excitotoxicity (Duchen, 2000; Frandsen and Schousboe, 1991 ). Figure 3 illustrates how excess calcium influx leads to excitotoxic death of neurons.

1.7. Clial cells, excitotoxicity and neuroinflammation Apart from neurons, glutamate receptors are also expressed by astrocytes, oligodendrocytes and microglia, the three major glial cell types of the brain. It is therefore logical that these cells are affected, in addition to neurons, by glutamate-induced excitotoxicity (for review see, Mattson, 2003). Astrocytes are very resistant to glutamate toxicity, probably due to the presence of high affinity glutamate transporters. How astrocytes modify glutamate-induced synaptic transmission and influence excitotoxicity by regulating extracellular glutamate concentrations, has been discussed in section 1.2. In addition, several studies have shown that astrocytes can release neurotrophic factors and cytokines, and thereby influence survival of neurons under excitotoxic conditions (Gabriel et al, 2003; Mattson and Rychlik, 1990). Like astrocytes, oligodendrocytes also supply neurotrophic factors (Du and Dreyfus, 2002) and

15 CHAPTER 1

EXCESSIVE GLUTAMATE

Ca>+

Proteases (calpaln) + DNA NO Arachldonlc Cytoskeletal degradation synthase acid breakdown + + NO - Free radicals

Pro-caspases+ + caspases ------CELL DEATH

Figure 3: Calcium-mediated intracellular events during glutamate-induced excitotoxicity. High levels of extracellular glutamate in the synaptic cleft (extracellular space) persistently activate AMPA- and Kainate receptors (AMPA/KA-R) (1) at the post-synaptic terminal, which allows strong influx of Na•, thereby leading to membrane depolarization. The depolarized membrane activates voltage-sensitive calcium channels (VSCC) 2 (2) and results in the removal of Mg • from NMDA receptors (3), allowing glutamate-induced activation of the 2 NMDA receptor (NMDAR) (4). Both VSCC opening and NMDA receptor activation allows excessive influx of Ca •. In addition, activation of the metabotropic glutamate receptor (mGLUR) (S) might induce IP3 receptor (IP3R) 2 2 channel-mediated Ca • release from endoplasmic reticulum, further contributing tot he excessive intracellular Ca • levels. High levels of intracellular calcium may over-activate potentially harmful proteins such as endonucleases 2 (DNA degradation), proteases (breakdown of cytoskeleton / NCX (Na•/ca • exchanger)), phospholipases and calmodulin (free radical formation), protein kinases and others (7). Moreover, calcium overload inside mitochondria impairs ATP production and increases the mitochondrial membrane permeability. This causes release of cytochrome C and other factors including apoptosis inducing factor (AIF) and second mitochondrial activator of caspases (smac) (8), which activates caspase-mediated pathways leading to programmed cell death (apoptosis). Adapted from references: Choi, 1992; Duchen, 2000; Mattson, 2003).

therefore may modify glutamate-induced neurotoxicity. On the other hand , both in vitro and in vivo studies have shown that oligodendrocytes are highly sensitive to glutamate-induced cell death (Matute et al, 2001; Pak et al , 2003). Microglia , the resident immune cells of the brain, become rapidly activated upon any type of injury or insult (Hanisch and Kettenmann , 2007). During excitotoxicity and associated conditions microglia may play contrary roles , largely depending on the extent of its activation , which may be neurotoxic (Cardona et al , 2006; Rossum and Hanisch , 2004) or neuroprotective (Lalancette-Hebert et al, 2007; Streit , 2002). Microglial activation may lead to enhanced GLT-1 expression and thereby increased glutamate uptake , as has been reported following facial nerve axotomy (Lopez-Redondo et al , 2000) and LPS treatment (Persson et al , 2005). On the other hand , activated microglia produces inflammatory mediators such as nitric oxide that increases vulnerability of neurons

16 GENERAL INTRODUCTION

to excitotoxic degeneration (Liu et al, 2002; Zhao et al, 2004). Other inflammatory mediators produced by activated microglia include cytokines, reactive oxygen species and complement II factors. These factors influence excitotoxicity both by modifying vulnerability of neurons and by regulating glutamate transporter expression (for review see, Tilleux and Hermans, 2007). Moreover, neuroinflammatory events initiated by microglia might also activate astrocytes and oligodendrocytes (reactive gliosis), further potentiating excitotoxicity as a result of reduced glutamate uptake by gliotic astrocytes (Zhao et al, 2004). Thus, both oxidative stress induced by excessive calcium influx and neuroinflammation resulting from microglial activation can play major roles in excitotoxic conditions that may ultimately lead to neuronal and glial cell death.

1.8. Glutamate transport and excitotoxicity in neurological disorders Excitotoxicity, often resulting from impaired glial glutamate transport, is a major cause for neuronal death in several acute and chronic neurodegenerative conditions of the CNS. An earlier study by Tanaka and colleagues has shown that mice lacking GLT-1 develop both epilepsy and neurodegeneration (Tanaka et al, 1997). In amyotrophic lateral sclerosis (ALS), a motor neuron disease, both elevated glutamate concentrations as well as impaired glutamate transporter expression and activity have been reported along with the excitotoxic neuronal damage (Rothstein et al, 1995; Spreux-Varoquaux et al, 2002; Vanoni et al, 2004). Similar observations have been made under pathological conditions such as Alzheimer's disease (Li et al, 1997), ischemia (Rao et al, 2001) and traumatic brain injury (van Landeghem et al, 2006), supporting the hypothesis that excitotoxicity might be a key contributing factor to the nervous damage associated with these central degenerating conditions. Other neurological disorders where altered glutamate transporter expression and/or activity along with excitotoxic neurodegeneration have been found include Parkinson's disease, Huntington's disease, etc. (for review see, Mattson, 2003; Seifert et al, 2006; Tilleux and Hermans, 2007).

1.9. Potential targets to prevent excitotoxic damage Several events downstream of the glutamate receptor activation have been targeted in preclinical studies to prevent excitotoxic damage. This includes hyperpolarizing agents, calcium channel antagonists, antioxidants, DNA damage response inhibitors, mitochondrial stabilizers, protease inhibitors and cytoskeleton-modulating agents etc. (for review see, Mattson, 2003). Some of the other potenti.:::treatment strategies against excitotoxicity are discussed below.

1.9.1. NMDA receptor antagonism Since excitotoxicity mediated by overactivation of neuronal NMDA receptors is attaining wide acceptance in conditions like Alzheimer's disease, treatment strategies aiming at the blockade of NMDA receptor activation have been discussed (Mattson, 2003). Some of the frequently used NMDA receptor antagonists are dizocilpine (MK 801), memantine and kynurenate. Of particular importance is memantine, an NMDA receptor channel blocker with fast kinetics, moderate affinityand strong voltage-dependency. Use of memantine has been proven to restore memory and to largely reduce glutamate-mediated neuronal loss in Alzheimer's patients (for review see, Parsons et al, 2007) and is now also suggested in treatment of Parkinson's disease and vascular dementia (Olivares et al, 2011), especially since other treatment strategies such as use of acetylcholinesterase inhibitors failed due to side-effects like nausea, vomiting and diarrhoea

17 CHAPTER l

(for review see, Birks, 2006). However, it is to be noted that NMDA receptors play a crucial role in the development of neurons (Mattson et al, 1988) and in synaptic plasticity (Kullmann et al, 2000), and therefore its blockade using antagonists might impair normal learning processes (Collingridge and Bliss, 1995; Danysz et al, 1995; Morris et al, 1986). In line with this, a recent study has stated that neuroprotection by NMDA receptor blockade using memantine, which sometimes requires high doses of the drug, is impossible to achieve without having side-effects such as memory impairment (Creeley et al, 2006). Furthermore, studies in rats have shown that memantine metabolism and/or elimination in fe males is slower than males (Zajaczkowski et al, 2000). The relatively low (but significant) side effects caused by the use of memantine augments the need for development of alternative treatment strategies against NMDA­ receptor mediated excitotoxic conditions without compromising the synaptic plasticity.

1.9.2. Regulation of glutamate transporter expression and activity Regulation of glutamate transporters for short-term or long-term involves a wide variety of mechanisms including modulation of their phosphorylation, cellular trafficking, gene transcription etc. (Beart and O'Shea, 2006; Robinson, 2002). Several other factors have been identified that regulate glutamate transporter expression and/or activity. Among these factors, glutamate itself has been fo und to regulate glutamate transporters, either through receptor activation (Aronica et al, 2003; Vermeiren et al, 2005) or by direct interaction with its transporters (Duan et al, 1999; Munir et al, 2000). Others include neuronal soluble factors such as neurotransmitters, growth factors and neuropeptides (Figiel and Engele, 2000; Schlag et al, 1998; Swanson et al, 1997; Zelenaia et al, 2000). In addition, the regulation of glutamate transporter expression by several inflammatory mediators such as cytokines and free radicals is now being discussed (for review see, Tilleux and Hermans, 2007). Understanding the mechanisms underlying regulation of glutamate transporter expression and/or activity by the different factors mentioned above might help to suppress excitotoxic neurodegeneration. In line with this, a very recent study has shown enhancement of GLT-1 and GLAST transporter expression using ceftriaxone (a third -generation cephalosporin antibiotic) and also 2R,4R­ APDC (a selective group II metabotropic glutamate receptor agonist) induces protection against excitotoxic neurodegeneration in vitro (Beller et al, 2011).

1.9.3. Modulation ofNMDA receptor activity and glutamate release The NMDA receptor can be modulated by a number of endogenous and exogenous compounds including, sodium, potassium and calcium ions that can not only pass through the NMDA receptor channel but also modulate the activity of receptors. Zinc blocks the channel through NR2A- and NR2B-containing receptors in a noncompetitive and voltage-independent manner. Polyamines can also either potentiate or inhibit glutamate-mediated responses. In addition, modulation of NMDA receptor activity as well as glutamate release by adenosine (see section 2 of this chapter) and cannabinoid receptors (Gerdeman and Lovinger 2001; Hampson et al, 2011; Hoffman et al, 2010; Kawamura et al, 2006; Liu et al, 2009) are attaining wide interests due to their potent anti-excitotoxic properties.

18 GENERAL INTRODUCTION

2. ADENOSINE IN THE CENTRAL NERVOUS SYSTEM Adenosine is a purine nucleoside and a component of ATP, the universal "currency" of free a energy in the cell. Besides having an important role in energy transfer, adenosine is well recognized to have its role in intercellular signaling (for review see, Dunwiddie and Masino, 2001). In the CNS, adenosine modulates many functions including regulation of sleep, arousal, locomotion, anxiety, cognition, and memory (Dunwiddie and Masino, 2001; Ribeiro et al, 2002). There is no de nova synthesis pathway for adenosine. Its synthesis intracellularly may occur either by dephosphorylation of adenosine monophosphate (AMP) or by hydrolysis of 5-adenosyl-L-homocysteine (Zimmermann, 2000). There are two sources of extracellular adenosine: 1) release of adenosine from the intracellular space via specialized transporters and 2) extracellular conversion of released adenine nucleotides (ATP, ADP and AMP) by a cascade of ectonucleotidases that includes CD39 (also known as nucleoside triphosphate dephosphorylase) and CD73 (also known as 5' -ectonucleotidase) (Sperlagh and Vizi, 1996; Zimmermann, 2000; Linden, 2001) . Thus, extracellular levels of adenosine are determined by its generation, release, cellular uptake and metabolism. In the CNS, virtually all cell types add to the extracellular adenosine, astrocytes being the key player in adenosine homeostasis (Figure 4; Baison et al, 2010). Nevertheless, the extracellular adenosine level in the brain is generally low (about 30 - 300nM) under physiological conditions (Rudolphi and Schubert, 1997) . However, its concentration increases rapidly (up to 30 - 100 folds) following metabolic stress associated with ischemia, hypoxia, head injury, seizure and inflammation (van Lubitz, 1999). For instance, the concentration fo llowing a 15 min ischemia has been measured to be around 10 - 50 µM (Hagberg et al, 1987). During high-frequency neuronal activity and seizure, neurons release large quantities of ATP (Cunha et al, 1996; Mitchell et al, 1993), which can be converted to adenosine by CD39 and CD73. Glial cells, astrocytes in particular, contribute largely to the rapid increase of extracellular adenosine levels following

ADP / � / Astrocy�e \ Figure 4: The adenosine cycle. The regulation of extracellular levels of adenosine is largely dependent on the astrocyte-based adenosine ATP AMP cycle. A major source of synaptic adenosine r is vesicular release of ATP (in orange circle) followed by its extracellular degradation to adenosine (ADO) via ectonucleotidases. Nucleoside transporters (NT) equilibrate AD0 4 extra- and intracellular levels of adenosine. Intracellular metabolism of adenosine depends t on the activity of adenosine kinase (ADK), which, together with 5'-nucleotidase (5'-NT), forms a substrate cycle between AMP and adenosine. Thus, intracellular metabolism of adenosine via ADK drives the influx of adenosine into the cell. Adapted from reference: Baison, 2008. ADO

19 CHAPTER l

episodes of ischemia and hypoxia (Jennings et al, 2001; Lynch et al, 1998; Martin et al, 2007; Parkinson and Xiong, 2004; Parkinson et al, 2005).

2.1. Adenosine receptors Extracellular adenosine exerts its effects by acting on four types of trans membrane receptors

, that are evolutionarily conserved and pharmacologically well-characterized namely, A1 A2A, A28

and A3 adenosine receptors (Fredholm et al, 2001). These receptors are G-protein coupled.

The A1 and A3 receptors are coupled to inhibitory G proteins (see Table 2) and their activation + induces membrane hyperpolarization by activating K channels, inhibits voltage-dependent 2 Ca + channels and inhibs cAMP formation. Activation of A2A and A28 receptors, which are both coupled to facilitatory or stimulatory G proteins (see Table 2), on the contrary, increases cAMP

formation. In the CNS, expression of A1 andA2A adenosine receptorsubtypesare higher, compared

to A28 and A3 receptor subtypes (see Table 2). The distribution of adenosine receptors in the

CNS varies between regions. For example, the expression level of A1 receptors is particularly

high in cortex, hippocampus, cerebellum and spinal cord, whereas A2A receptors are highly expressed in olfactory bulb and striatum (Figure 5). Thus, both the differential distribution of the receptors as well as their coupling to different G proteins (inhibitory or facilitatory) causes a high degree of complexity in adenosine signaling. Moreover, the identification of receptor heteromerization, either with other adenosine receptors or with neurotransmitter receptors, has further increased the complexity of adenosine signaling in the CNS (Fuxe et al, 2007; Sebastiao and Ribeiro, 2009a).

2.2. Neuromodulation by adenosine: role in excitotoxicity Adenosine is regarded as the brain's endogenous neuromodulator and/or neuroprotectant, and has been widely discussed to reduce damage associated with several acute and chronic excitotoxic conditions of the brain including seizures, ischemia, hypoxia, Alzheimer's disease, Parkinson's disease and Huntington's disease (for review see, Boison, 2008).

Ta ble 2. Adenosine receptors: types, properties and distribution in the central nervous system. G (guanine nucleotide binding protein); AC (adenylate cyclase); GIRK (G-protein gated inward

rectifying potassium channels); PLA2 (phospholipase A2); PLD (phospholipase D); PLC (phospholipase C). Adapted from reference: Benarroch, 2009.

G protein Receptor coupling Transduction pathway Main distribution

� G;, Go Inhibition of AC Cerebral cortex N and P/Q channels inhibition Hippocampus GIRK activation Thalamus

Activation of PLA2 and PLD Cerebellum Brain stem Spinal cord

A2A Gs, Golf Activation of AC Striatum

A2B Gs Activation of AC, PLC activation Uniform at medium or low abundance

A3 G Activation of AC, PLC activation Hippocampus and cerebellum at medium or low abundance

20 GENERAL INTRODUCTION

cortex hippocampus II thalamus

olfactory bulb

amygdala

tractus solitarius Figure S: Distribution of adenosine A1 and A2A receptors in the brain. Bigger fonts indicate high levels of expression. spinal cord Adapted from reference: Ribeiro et al, 2002. (dorsal horn)

The inhibitory neuromodulation as well as neuroprotective effects of adenosine are mediated through A1 receptors. Pre-synaptic activation of A1 receptors inhibits release of excitatory neurotransmitters such as glutamate. A1 receptors also regulate release of other neurotransmitters such as dopamine, serotonin and acetylcholine. At the post-synaptic terminal, A1 receptor activation causes membrane hyperpolarization, thereby restrains NMDA receptor activation and intracellular calcium influx and thus prevents the generation and/ or propagation of excitatory action potentials. Thus, by regulating glutamate release and by modifying NMDA receptor activation, adenosine can influence excitatory neurotransmission as well as excitotoxicity by activating A1 receptors at glutamatergic synapses. The inhibitory actions of A1 receptors can be modifiedby the facilitatory A2A receptors. For example, activation of the A2A receptors has been shown to downregulate A1 receptor responses on synaptic transmission (Ciruela et al, 2006). Thus, extracellular adenosine may also aggravate excitotoxic tissue damage by activating the A2A receptors (Cunha, 2005; Picano and Abbracchio, 2000).

Consistently, inhibition of A2A receptors has neuroprotective effects (Silva et al, 2007). The protective effects of A2A receptor blockade are prominent in the striatum, since striatal neurons express high levels of A2A receptors (Svenningsson et al, 1999). A2A receptors are also expressed by microglia and astrocytes (Hasko et al, 2005), but usually in low levels under physiological conditions (Baison et al, 201 0). Their expression, however, is induced following brain insults (Cunha, 2005) and by pro-inflammatory cytokines such as interleukin (IL)-1 � (Baison et al, 201 0). Activation of glial A2A receptors can induce and/or exacerbate neurotoxicity. For example, activation of A2A receptors on astrocytes can increase extracellular glutamate levels,

21 CHAPTER 1

both by reducing glial glutamate uptake and by direct release (Li et al, 2001; Nishizaki et al, 2002), thereby potentiating excitotoxic neuronal damage. Consistently, injection of CGS 21680,

an adenosine A2A receptor agonist, enhanced extracellular glutamate levels in rat striatum

(Popoli et al, 1995) and blockade of striatal A2A receptors protected neurons against quinolic­ acid induced excitotoxicity (Popoli et al, 2002). In a middle cerebral artery occlusion (MCAO) model of transient focal cerebral lschemia, both striatal outflcw of glutamate and brain injury were strongly reduced in A2A receptor deficientmice compared to the wild-type littermates (Gui

et al, 2009). Moreover, adenosine A2A receptor activation in microglia may lead to neurotoxicity and this may be due to potentiation of nitric oxide release in activated microglia (Saura et al, 2005). In contrast to the above findings, there is accumulating evidence that tonic activation

of adenosine A2A receptors may regulate neuronal survival, plasticity and differentiation by infl uencing the actions of neurotrophic factors (for review see, Sebastiao and Ribeiro, 2009b). Ta ken together, the neuromodulatory effects of adenosine in the CNS are largely determined

by a balanced activation of inhibitory A, receptors and facilitatory A2A receptors (Cunha, 2005; Gomes et al, 2011). Excitotoxic conditions including ischemia, hypoxia, seizure and trauma, are associated with a rapid build up of extracellular ATP that undergoes degradation resulting in large amount of adenosine in the extracellular fluid. How adenosine modu lates excitotoxicity, by acting on A,

and A2A receptors located on neurons and glial cells, has already been discussed above. Several

studies suggest anti-excitotoxic properties of A, receptor activation and A2A receptor blockade. For example, agonists of A, receptors reduce epileptic seizures (Vianna et al, 2005) and have anticonvulsant (Hosseinmardi et al, 2007) and antidepressant effects (Vinade et al, 2003).

However, the use of A1 receptor agonists to treat CNS pathologies has been limited because

they developed severe peripheral side effects (Stone, 2002). On the other hand, use of A2A receptor antagonists showed only minor side effects, compared to A, receptor agonists (Stone,

2002). Thus, A2A receptor antagonists that show potent anticonvulsive effects (Hosseinmardi et al, 2007) have also been shown to improve symptoms in Parkinson's disease patients

(Chen et al, 2001; Schwarzschild et al, 2006). Furthermore, A2A receptor antagonists have been shown to reduce motor abnormalities in Huntington's disease (Popoli et al, 2000) and to protect motor neurons in ALS (Mojsilovic-Petrovic et al, 2006). Apart from these effects of adenosine receptors, a recent study has shown that adenosine receptor activation modifiesthe permea bility of blood-brain barrier (BBB) in vivo (Carman et al, 2011). In this study, the authors

showed that the activation of A, and A2A receptors on endothelial cells at the BBB fa cilitates better entry of intravenously applied compounds into mouse brain. Since, drug delivery across the BBB is a major challenge in treating CNS disorders, these findings suggest that adenosine receptor signaling can be exploited to assist the entry of therapeutic compounds into the CNS.

2.3. Immune modulation by adenosine In addition to the neuromodulatory effects,ther e is clear evidence for immune modulation by adenosine in the CNS (Boison et al, 2010; Hasko et al, 2005). Most receptors for adenosine, if not all, are expressed by astrocytes (Bjorklund et al, 2008) and immune cells in the CNS including microglia (Dare et al, 2007; Hammarberg et al, 2003; van Ca Iker and Biber, 2005) and infiltrating leukocytes (Hasko et al, 2008). Adenosine can induce distinct and sometimes contrasting effects on these cells by activating specific receptor subtypes. In astrocytes, activation of the

22 GENERAL INTRODUCTION

A, receptors reduces cell proliferation (Ciccarelli et al, 1994; Rathbone et al, 1991), activation of A2A receptors increases proliferation and can cause reactive astrogliosis (Brambilla et al, II 2003; Hindley et al, 1994) and activation of A28 receptors enhances TNF-a.induced astrogliosis

(Trincavelli et al, 2004). Adenosine can also affect survival of astrocytes. Activation of A1 receptors protects astrocytes from damage and/or promotes their survival (Bjorklund et al,

2008; Ciccarelli et al, 2007; D'Alimonte et al, 2007), whereas activation of A3 receptors may either promote their survival (Bjorklund et al, 2008) or induce apoptosis in these cells (Abbracchio et al, 1997; Appel et al, 2001; Di Iorio et al, 2002). In addition to regulating proliferation and

survival of astrocytes, adenosine has potent effectson the secretory functions of these cells: A1

receptor stimulation induces NGF release (Ciccarelli et al, 1999), A2A receptor stimulation inhibits inducible nitric oxide synthase (iNOS) expression and nitric oxide production (Brodie et al, 1998),

A28 receptor stimulation induces production of IL-6 (Fiebich et al, 1996a; Fiebich et al, 2005;

Schwaninger et al, 1997), and A3 receptor stimulation induces synthesis of the neuroprotective chemokine CCL2 (Wittendorp et al, 2004). Adenosine action on microglia can induce apoptosis

(Ogata and Schubert, 1996). On the other hand, specific activation of A1 and A2A receptor subtypes has been shown to enhance microglial proliferation (Gebicke-Haerter et al, 1996).

In addition, microglial A2A receptor activation has been implicated in regulating inflammatory and neurotrophic actions of this cell. This includes, upregulation of cyclooxygenase 2 (COX-2), induction of prostaglandin E2 (PGE2) release (Fiebich et al, 1996b), and synthesis and release of nerve growth factor (NGF) (Heese et al, 1997). Adenosine also regulates inflammatory actions

of infiltratingcells in the brain. For example, A2A receptor stimulation increases production of IL-1, IL-6 and IL-12 by infiltratingcells (Yu et al, 2004), whereas A1 receptor stimulation inhibits production of IL-lB and matrix metalloproteinase 12 by infiltrating macrophages (Tsutsui et al, 2004). In conclusion, by acting on specific receptor subtypes in astrocytes, microglia and infiltratingimmune cells, adenosine differentially modulates neuroinflammation. As described before, inflammation is now recognized to play an important role in excitotoxic conditions of the CNS. In recent years, cytokines of the IL-6 family have been shown to influence excitotoxicity associated with injury and disorders of the CNS (see section 3.4). By acting as important signaling molecules and by regulating development and survival of differentneural cell types, these cytokines mediate several neurotrophic, neuroprotective and neuroinflammatory effects in the CNS (see section 3). Although adenosine is clearly shown to have immuno­ modulatory effectsin the brain, it is not clear whether adenosine also regulates the function or expression of IL-6-type cytokines in the CNS. There is however evidence that adenosine induces synthesis and release of IL-6 in astrocytes (Fiebich et al, 1996a; Fiebich et al, 2005; Schwaninger et al, 1997). IL-6, in turn, can induce expression of adenosine A, receptors in astrocytes (Biber

et al, 2001) and in neurons (Biber et al, 2008). Furthermore, A1 receptor activity in neurons has

been shown to be crucial for IL-6-induced neuroprotection (Biber et al, 2008). How A1 receptor activity induces protective effectsin neurons and astrocytes has already been discussed. There is no evidence on whether adenosine and its receptors regulate expression and/or activity of other IL-6-type cytokines in the CNS. Since neuroprotective properties of both adenosine and IL-6- type cytokines are taken into account, understanding the "interplay" or "interaction" between the adenosinergic system and IL-6-type cytokine family is important for the development of potential treatment strategies against excitotoxic conditions of the CNS.

23 CHAPTER 1

3. INTERLEUKIN (IL)-6-TYPE CYTOKINES IN THE CENTRAL NERVOUS SYSTEM The family of IL-6-type cytokines consists ofseven pleiotropic molecules which are structurally and functionally related namely, IL-6, IL-11, leukemia inhibitory factor (LIF), ciliary neurotrophic factor (CNTF), oncostatin M (OSM), cardiotrophin-1 (CT-1) and novel neurotrophin-1 (NNT-1) or cardiotrophin-like cytokine (CLC) (for reviews see, Heinrich et al, 1998; Taga and Kishimoto, 1997). They all have a similar three dimensional structure comprising a four helix conformation, where two pairs of anti-parallel helices are connected by a polypeptide loop (Figure 6). Unlike other cytokines, all members of this family contain relatively long helices ranging from 15 to 22 residues (Bravo and Heath, 2000). Among the members of this family, the structural similarity is higher (about 27% homology) between OSM and LIF (Schrell et al, 1998). In addition, the genes encoding OSM and LIF are very closely located (only 20 kbp apart) on chromosome 22, and have been suggested to be the result of gene duplication (G6mez-Lech6n, 1999; Jeffery et al, 1993); the locations of other IL-6-type cytokine genes are scattered across multiple chromosomes (Table 3). Clearly these cytokines are important for normal development of the brain by regulating proliferation and maintenance of stem cells, neurogenesis, gliogenesis, cell survival etc. (Bauer et al, 2007; Hatta et al, 2002; Holtmann et al, 2005; Islam et al, 2009; Koblar et al , 1998; Lee et al, 2008; Nakashima et al, 1999; Pitman et al, 2004; Shimazaki et al, 2001 ). Apart from their physiological role, IL-6-type cytokines mediate several pro- and anti­ inflammatory effects and are widely implicated in pathology of the CNS. In this section, both trophic and harmful effects of IL-6-type cytokines (LIF and OSM in particular) as well as their role in excitotoxic conditions of the CNS, will be discussed.

3.1. Receptors: types, activation and intracellular signaling pathways IL-6-type cytokines exert their effects by acting on three types of receptor complexes at the plasma membrane of target cells: a homodimer of glycoprotein 130 (gp130/gp130; that binds

Figure 6: Structure of IL-6. The four long helices A, B, C, and Dare highlighted in different colours. Receptor-binding sites I, II and Ill are indicated by circles. Adapted N from reference: Heinrich et al, 2003.

24 GENERAL INTRODUCTION

Table 3. Biochemical properties of IL-6-type cytokines. Adapted from reference: Heinrich et al, 1998. Property IL-6 IL-11 LIF CNTF CT-1 OSM II Number of Am ino acids Precursor 212 199 202 200 201 252 Mature Protein 184 178 180 200 201 196 Molecular mass (kDa) Predicted 20.8 19.1 20 22.9 21.2 22.1 Observed 21-28 23 45 21-28 21.5 28 Glycosylation Potential N-glycosylation sites 2 0 6 0 0 2 N glycosylation demonstrated Yes.1 of 2 0 Yes 0 0 Yes Number of cysteine residues 4 0 6 1 2 5 Number of S-S bridges 2 0 3 0 ? 2 mRNA size (kb) 1.3 1.5, 2.5 1.8, 4 1.0 1.7 2 Number of exons 5 5 3 2 3 3 Chromosomal localization 7p21-p14 19q13.3-14.3 22q14 ll q12 16pll.1-11.2 22q12

IL-6 and IL-11), a heterodimer of gpl30 and the LIF receptor (gpl30/LIFR-p; that binds LIF, human OSM, CNTF, CT-1 and NNT-1) and a heterodimer of gpl30 and the OSM receptor (gp130/OSMR; that binds OSM) (Figure 7; Mosley et al, 1996; Turnley and Bartlett, 2000). IL-6, IL-11 and CNTF require an additional ligand recognition subunit: IL-6R, IL-llR and CNTFR respectively (Davis et al, 1993; Heinrich et al, 1998). The ligand recognition subunit of the receptor complex can either be a signaling (LIFR-P and OSMR) or non-signaling (IL-6R, IL-llR and CNTFR) component. Gp 130, the common receptor subunit shared by all the cytokines of this family, is responsible for most of the downstream intracellular signaling pathways (Heinrich et al, 1998). Several redundant properties have been reported among the members of this family due to the shared use of gpl30, hence they are also referred to as gpl30 cytokines. All lL-6-type cytokines, except OSM, bind firstto their ligand recognition subunit (IL-6R, CNTFR, IL-llR or LIFR-P) and then recruit one or two gpl30 subunits to activate downstream intracellular signaling pathways (Gearing et al, 1992; Mosley et al, 1996). The relative amount of different receptor subunits present on the cell strongly determines the specificityof cytokine binding as well as the quantitative response of the cell to the ligands (Wang et al, 2000). In order to facilitate their action on target cells, all IL-6-type cytokines contain three distinct binding sites that interact with their respective receptors, namely sites I, II and Ill (Figure 6). Site I interacts with the non-signaling (ligand recognition) subunit of the receptor complex; sites II and Ill bind to cytokine binding module (CBM)-domain and lmmunoglobulin (lg) like-domain of the gpl30 subunit, respectively (Figure 7). Sites I and II of the ligand cause high-affinitybinding to the receptors, whereas site Ill is responsible for recruiting the second receptor subunit (for review see, Heinrich et al, 2003). Upon receptor oligomerization, different signal transduction pathways are activated by IL-6-type cytokines (Figure 8). The downstream signaling is initiated by phosphorylation of gpl30-associated tyrosine kinases (JAK kinases: JAKl, JAK2, JAK3 and TYK2). These kinases subsequently phosphorylate gpl30, followed by recruitment, phosphorylation and activation

25 CHAPTER 1

N-terminus

= 7Cys residues - CBM II 1wsxws D1 [J lg-like Q 1g-like

residues u s D2 l7cys • CBM : � -; D3 lwsxws 11 wsxws FNIII I D4 1 osl FNIII I FNIII I D6TM 1 1--;;-X1 I 'BOX1 Dileucine oa.��f� .. TABQT3 motifsX2 gp1 30 LIFR OSMR

C-terminus

Figure 7: Receptor components required for LIF (gp130/LIFR) and OSM (gp130/OSMR) signaling.

of signal transducers and activators of transcription (STAT), mainly STAT3 and STATI (Heinrich et al, 1998). In addition to STAT3 and STATI, OSM has been reported to activate STATS (Figure 8) (Hintzen et al, 2008; Wang et al , 2000). Phosphorylation of gpl30 by JAKs also results in the activation of mitogen-activated protein kinases (MAPKs) , which are important serine/threonine protein kinases that mediate signal transduction from the cell surface to the nucleus (Thoma et al , 1994). The three MAPK cascades include: extracellular signal regulated kinase (ERKl/2), p38 and c-jun N-terminal kinases/stress-activated protein kinases (JNK/SAPK). The most important MAPK cascade resulting from gpl30 activation is recruitment of ERKl/2, which is mediated by the guanine nucleotide-binding protein Ras and the serine/threonine kinase Raf (Stancato et al , 1997; Thoma et al, 1994). In addition to JAK/STAT and MA PK cascades , IL-6-type cytokines are also known to activate the phosphatidylinositol-3-kinase (PI3K/Akt) signaling pathway (Figure 8; Kamimura et al, 2003). In order to control excessive activation of the receptors and the downstream signaling pathways by IL-6-type cytokines, the cells make use of certain classes of regulatory proteins such as suppressor of cytokine signaling (SOCS) and protein inhibitor of activated STAT (PIAS) (For review see, Croker et al , 2008). They regulate cytokine activity by acting at various levels ranging from inhibition of tyrosine phosphorylation at gpl30 receptor subunit to STAT binding to the DNA. Thus, the net effect of cytokine binding and activation of the receptors on a target cell is , at least in part, determined by the regulation of the intracellular signaling cascades by SOCS and PIAS.

26 GENERAL INTRODUCTION II

Figure 8: Intracellular signaling pathways activated by LIF and OSM. Both cytokines activate STAT3 (indicated by a fu ll shown pathway at LIFR and only by a STAT3 bound to OSMR). Both cytokines also activate the Pl3K/Akt and the MAPK pathways, but in different ways.

3.2. Cytokine and receptor expression in the CNS In the CNS, many members of the IL-6-type cytokine fa mily as well as their receptors are constitutively expressed, mainly in the hippocampus (Rosell et al, 2003) and the pituitary (Akita et al, 1995; Auernhammer and Melmed, 1999; Hanisch et al, 2000). Other brain regions where expression of the cytokines have been found include, visual cortex, superior colliculus, thalamus, hypothalamus and cerebellum for LIF (Yamakuni et al, 1996; Yamamori, 1991); forebrain and choroid plexus for CT-1 (Gard et al, 2004; Ochiai et al, 2001); cortex, olfactory bulb, cerebellum and hypothalamus for IL-6. In the CNS, astrocytes are likely to be the major source forIL -6 (Fujita et al, 2009; Schwaninger et al, 1997; Va n Wagoner and Benveniste, 1999), LIF (Yamakuni et al, 2002) and CNTF (Dal Iner et al, 2002). OSM is mainly produced by microglia (Glezer and Rivest, 2010) and by infi ltrating cells such as monocytes, macrophages and T-cells (Brown et al, 1987; Grenier et al, 1999; Kastl et al, 2008; Ta naka and Miyajima, 2003). Microglia, when activated, may also produce LIF (Nakanishi et al, 2007) and IL-6 (Lee et al, 1993; Nakanishi et al, 2007; Schmitt et al, 2006). In addition to glial cells, several neuronal subpopulations from different brain regions have been identified to express many of these cytokines including IL-6 (Gadient and Otten, 1994; Juttler et al, 2002; Schmitt et al, 2006; Sch6bitz et al, 1994), LIF (Lemke et al, 1996), OSM (Tamura et al, 2003a; Znoyko et al, 2005), CT-1 (Barnabe-Heider

27 CHAPTER 1

et al, 2005) and CNTF (Dutta et al, 2007). In short, most IL-6-type cytokines are constitutively expressed by cells of the CNS under physiological conditions. As mentioned before, many receptor subunits for IL-6-type cytokines are expressed in the hippocampus (Rosell et al, 2003), which therefore seems to be a major site for the cytokine action. In addition, olfactory bulb may be a target region for OSM action, since expression of OSMR-p is high in this region (Tamura et al, 2003b). The shared receptor subunit gp130 is ubiquitously expressed on neurons, glial cells and blood-vessel associated cells throughout the brain (Rosell et al, 2003; Watanabe et al, 1996), whereas other receptor subunits are expressed in a dispersed manner. For example, LIFR-P and CNTFR-a are specifically and highly expressed by oligodendrocytes (Butzkueven et al, 2002; Ishibashi et al, 2009; Marriott et al, 2008; Mayer et al, 1994; Stankoffet al, 2002), whereas OSMR- P and CNTFR-a are highly expressed in astrocytes (Dallner et al, 2002; Kordula et al, 1998; Ta mura et al, 2003b). OSMR-P is also highly expressed on endothelial cells (Modur et al, 1997; Ruprecht et al, 2001). On the other hand, neurons seem to be a common target for most IL-6-type cytokines, since they express IL-6R (Ohtaki et al, 2006), LIFR-P (Yamakuni et al, 1996), OSMR-P (Tamura et al, 2003a) and CNTFR-a (Lee et al, 1997; Watt et al, 2009). Although IL-6-type cytokines were primarily thought to modulate immune activity of glial cells, profound expression of their receptors on neurons has led to the suggestion that these cytokines can have direct neuromodulatory effects (neurotrophic or neuroprotective) in the CNS. Moreover, from the cell-type specific expression of the receptors, one might infer that the major targets of IL-6-type cytokines in the CNS are neurons and oligodendrocytes. OSM, however, may also have its effects on astrocytes and endothelial cells, in addition to certain neuronal subpopulations.

3.3. IL-6-type cytokines and CNS inflammation Microglia and astrocytes are the primary mediators of inflammation in the brain. In addition, disru ption of the BBB may allow the entry of peripheral blood-derived mononuclear cells (mainly monocytes, macrophages and T-lymphocytes) into the brain, which contributes significantly to the pathogenesis of CNS inflammation. A variety of pro- and anti-inflammatory cytokines as well as chemokines regulate inflammation in the brain, and the key players include IL-l p, TNF-a, interferon y, and IL-6. In addition, other soluble mediators such as nitric oxide and oxygen radicals also play a major role in the inflammatory processes of the CNS. Elevated levels of IL-6-type cytokines and/or their receptors have been reported in various chronic and acute pathological conditions of the CNS including injury, seizure, ischemia, hypoxia, HIV AIDS, tumour, multiple sclerosis, Alzheimer's disease, Pa rkinson's disease etc. (Banner et al, 1997; Chiaretti et al, 2008; Choi et al, 2003a; Choi et al, 2003b; Dutta et al, 2007; Getchell et al, 2002; Hergenroeder et al, 2010; Hori et al, 1996; Jankowsky and Patterson, 1999; Lee et al, 1998; Lin et al, 1998; Minami et al, 2002; Rosell et al, 2003; Slevin et al, 2008; Soilu­ Hanninen et al, 2010; Suzuki et al, 2000; Suzuki et al, 2005; Suzuki et al, 2009; Winter et al, 2004; Yokota et al, 2005). Elevated levels of IL-6-type cytokines can be due to endogenous production by the CNS cells or due to their transport from the periphery across the BBB (Pan et al, 2000; 2006; 2008). Presence of high levels of IL-6-type cytokines has been believed to cause damage to the brain. In line with this, evidence suggests that IL-6-type cytokines mediate various pro-inflammatory and neurotoxic effects in the CNS. For instance, IL-6 mediates neurotoxic effects either directly by inducing production of oxygen radicals in target neurons

28 GENERAL INTRODUCTION

(Conroy et al, 2004; Dugan et al, 2009) or indirectly by inducing expression and release of inflammatory mediators such as IL-1�, TNF-a and COX-2 in microglia (Krady et al, 2008) and by II promoting astrogliosis (Sriram et al, 2004), leading to severe neurodegeneration. In animals overexpressing IL-6 in the CNS, severe neurological dysfunction was observed, characterized by tremor, ataxia, seizures, astrocytosis and gliosis (Campbell et al, 1993). These animals also showed progressive decline in avoidance learning parallel to inflammatory neurodegeneration (Heyser et al, 1997). In another study, although IL-6 per se did not affect the survival of cortical neurons, IL-6 augmented neuronal death induced by beta-amyloid peptide and NMDA (Qiu and Gruol, 2003). This suggests that, IL-6 can cause neuronal damage in Alzheimer's disease, when acting in concert with other toxic factors. Other IL-6-type cytokines also contribute to inflammatory neurodegeneration, by direct or indirect mechanisms. For example, LIF induces expression of peripherins in hippocampal neurons (Djabali et al, 1993). Accumulation of peripherin in neurons has been implicated in neurodegenerative conditions such as ALS (Millecamps et al, 2006; Robertson et al, 2003). Consistently, exogenous administration of LIF or IL-6 exacerbated peripherin formation following stab lesion in adult mouse brain (Beaulieu et al, 2002). Furthermore, LIF signaling in the mouse retina causes cell death by inducing iNOS (Elliot et al, 2006). LIF also promotes migration of infiltrating cells into the brain, since the inflammatory cell infiltration into crushed nerve was significantlyslower in LIF deficientmice (Sugiura et al, 2000). OSM, which is primarily produced by microglia in the brain and also by infiltrating immune cells, can also contribute to the pro-inflammatory processes leading to neuronal demise. For instance, OSM produced by mononuclear cells of HIV-1 infected patients induces apoptosis of neurons (Ensoli et al, 1999). OSM can also induce and/or act in concert with other proinflammatory mediators. For example, OSM induces TNF-a and iNOS gene expression in microglia (Baker et al, 2010). In astrocytes, activation of STAT3 signaling by OSM and LIF induced reactive astrogliosis (Sriram et al, 2004). OSM also induces expression of PGE2 (an eicosanoid associated with inflammation) and COX-2 (an enzyme involved in PGE2 synthesis) in astrocytes (Repovic and Benveniste, 2003). More important, OSM in combination with the inflammatory mediators such as IL-1�, TNF-a , and LPS, exhibits a striking synergy, resulting in up to a SO-foldhigher PGE2 production by astrocytes, astroglioma and neuroblastoma cell lines (Repovic and Benveniste, 2003). Furthermore, in transgenic mice overexpressing OSM in neurons of the brain and the spinal cord, severe neuronal abnormalities leading to death of the animal have been observed (Malik et al, 1995). On the other hand, IL-6-type cytokines may also suppress CNS inflammation. For example, CNTF inhibits microglial activation by reducing COX-2 levels (Krady et al, 2008), inhibits production of TNF-a in the brain and periphery (Benigni et al, 1995) and protects from inflammatory damage (Kuhlmann et al, 2006). LIF inhibits production of oxygen radicals and TNF-a and mediates strong anti-inflammatory effects (Hendriks et al, 2008). Taken together, IL-6-type cytokines are pleiotropic in nature and their effects (pro- and anti-inflammatory) in the CNS are determined by multiple factors including target cells, intracellular signaling pathways, concentration of the cytokine, general environment, presence of other inflammatory mediators etc. Thus, predicting the relevance (trophic or toxic effects) of elevated levels of IL-6-type cytokines in the neuropathological conditions is difficult, considering the complex behaviour of these cytokines. Nevertheless, there is growing evidence that suggests that IL-6-type cytokines principally mediate trophic effects in the CNS, since susceptibility of animals to seizure is higher in IL-6 knock-out mice

29 CHAPTER 1

(De Sarro et al, 2004) and these animals also showed reduced hippocampal neurogenesis (Bowen et al, 2011). Hence, IL-6-type cytokines are also referred to as neuropoeitic cytokines or neurokines (Patterson, 1992).

3.4. Anti-excitotoxic and neuroprotective effectsof IL-6-type cytokines Apart from the neurotrophic effects, there is substantial evldence suggesting neuroprotective effects of IL-6-type cytokines in the CNS against various lnsults including excitotoxicity. For instance, IL-6 exhibits several anti-excitotoxic properties both in vitro and in vivo including protection of neurons against glutamate-induced cell death (Biber et al, 2008; Ya mada and Hatanaka, 1994), inhibition of neurotransmitter release and the spread of excitation in the cerebral cortex (D'Arcangelo et al, 2000) and protection of the brain from ischemic attacks (Ali et al, 2000; Loddick et al, 1998). Other members of the IL-6 family also exhibit potent neuroprotective properties against excitotoxicity. For example, OSM inhibits NMDA receptor­ mediated intracellular calcium influx in neurons and selectively reduces expression of the NR2C subunit of the neuronal NMDA receptor, both in vitro and in vivo (Weiss et al, 2006). The neuroprotective effect of LIF has been reported in neurons from different regions of the brain and spinal cord. In the retina, photoreceptors are protected by LIF synthesized by Muller glia (Joly et al, 2008). In another study, intravitreal injection of LIF preserved photoreceptor function and protected photoreceptor cell death against lig ht-induced oxidative damage, and this effect of LIF is mediated by STAT3 signaling (Ueki et al, 2008). CNTF and CT-1, two other cytokines that use LIFR for their signaling, are also potential factors that protect photoreceptors from degeneration (Song et al, 2003; Tao et al, 2002) and treatment of retinal degeneration using CNTF has attained a successful outcome in the phase 1 clinical trials (Sieving et al, 2006). Furthermore, CNTF promotes survival of injured ganglion cells after optic nerve transection and crush injuries (Leaver et al, 2006) and protects against retinal ischemic injury (Unoki and LaVail, 1994) . In the peripheral nervous system (PNS), CNTF enhances survival and neurite outg rowth of neurons from dorsal root ganglia (DRG) (Sango et al, 2008), and CT-1 induces protection of motor, sensory and sympathetic neurons (Bordet et al, 1999; Oberle et al, 2006; Pennica et al, 1996). In addition, CT-1 also protects cultured cortical neurons from oxidative stress-induced damage (Wen et al, 2005). In conclusion, IL-6-type cytokines are potent neuroprotectants in the CNS and in the PNS. Despite the existing evidence, the mechanism(s) underlying neuroprotective effects of IL-6-type cytokines against excitotoxicity is largely unknown. The neuromodulatory, neuroprotective and/or anti-excitotoxic effects of adenosine A 1 receptors in the CNS have already been described (see section 2.2). This includes pre-synaptic inhibition of excitatory neurotransmitter release, post-synaptic inhibition of calcium influx through voltage dependent calcium channels and activation of G-protein coupled inward ly rectifying potassium channels that mediate post-synaptic membrane hyperpolarization (de Mendonc;a et al, 2000; Dunwiddie and Masino, 2001). These effects also reduce metabolic demand, thereby preserving ATP stores, suppressing glutamatergic transmission, and thus protecting neurons from excitotoxicity (Schubert et al, 1997). We have recently shown that IL-6 exerts its neuroprotective properties by upregulating neuronal A receptors (Biber et al, 1 2008). In this study, when neuronal A receptors were pharmacolog ically blocked by a selective 1 antagonist, IL-6 failed to protect neurons against excitotoxicity. The dependence of IL-6 on A receptor activity for inducing its neuroprotective effects has been further confirmed in A 1 1

30 GENERAL INTRODUCTION

receptor deficientmice, where IL-6 failed to protect cortical and hippocampal neurons against excitotoxicity (Moidunny et al, unpublished observations). Furthermore, we have shown that IL-6 potentiates A receptor mediated inhibition of electrically evoked excitatory post-synaptic a 1 currents (EPSCs) in ex vivo hippocampal slice cultures (Biber et al, 2008). It would be highly interesting to know whether the anti-excitotoxic effects of other I L-6 family members is also dependent on A1 receptor activity, since several redundant properties have been reported for these cytokines. So far, survival of neurons and oligodendrocytes has been the focus of most studies performed to understand the effects of IL-6-type cytokines in excitotoxicity. However, excitotoxicity is a complex process, that may involve damage due to oxidative stress and inflammation (see section 1.7). In the CNS, microglia are the major cell type that mediate neuroinflammation, although their exact role in excitotoxicity is controversial. There is only limited evidence on how IL-6-type cytokines affect microglial activity such as cytokine secretion, phagocytosis and chemotaxis (Krady et al, 2008; Lee et al, 2009; Lin et al, 2009), whereas there is no existing evidence on how IL-6-type cytokines affect microglial actions during excitotoxicity. This is important in order to better understand the role of these cytokines in inflammatory neurodegenerative conditions of the CNS such as multiple sclerosis. Finally, a better understanding of the nature and regulation of toxic and trophic effects of IL-6-type cytokines will provide a foundation on which the molecular- and cytokine-based therapies can be developed to treat prevalent diseases of the CNS.

4. THESIS OUTLINE

Background In the previous sections we have discussed in detail excitotoxicity, neuromodulatory and immunomodulatory effects of adenosine in the CNS and the neuroprotective effects of IL- 6-type cytokines in excitotoxic conditions of the CNS. We have also discussed some of the existing evidence on "interplay" or "inter-regulatory effects" between the adenosinergic system and IL-6 in excitotoxicity, which includes: 1) IL-6-induced neuroprotection against glutamate is dependent on neuronal expression and function of A1 receptors (Biber et al, 2008),

2) A28 receptor activation induces expression and release of IL-6 in astrocytes (Fiebich et al,

1996a; Schwaninger et al, 1997) and 3) A1 receptor-mediated inhibitory actions on the EPSCs is potentiated by IL-6, most likely by increasing A1 receptor expression in neurons (Biber et al, 2008). Since most members of the IL-6-type cytokine family possess neuroprotective properties (see section 3.4), since their expression is enhanced in several pathological conditions of the CNS and since they exhibit several redundant properties due to shared signaling through the gpl30 receptor subunit, we hypothesized that a similar "interplay" exists between the adenosinergic system and other members of the IL-6 family during excitotoxicity. In this PhD thesis, we have just focussed on OSM and LIF to examine the existence of such "interplay" during excitotoxicity. In chapter 2, we started with investigating the neuroprotective properties of OSM and LIF against excitotoxicity in wild-type as well as in A1 receptor deficient (-/-) neurons isolated from the mouse cortex and hippocampus. In addition, we examined the modulatory effects of OSM and LIF on A1 receptor mediated EPSC inhibition in hippocampal slice cultures.

31 CHAPTER 1

All lL-6-type cytokines activate three major intracellular signaling pathways downstream of their receptor activation namely, JAK/STAT3, ERKl/2 and P13K/Akt (see section 3.1). In chapter 3, we studied the signaling mechanisms involved in the neuroprotective properties of OSM and LIF against glutamate-induced excitotoxicity using selective inhibitors of the three different signaling pathways. Astrocytes are the major source for IL-6 and LIF (see section 3.2) . Adenosine, which is highly released following excitotoxic conditions such as seizure and ischemia (see section 2), induces the synthesis and release of IL-6 in astrocytes. It is not known whether adenosine also induces the expression and/or release of LIF in astrocytes during excitotoxicity. Therefore, in chapter 4, we examined whether excitotoxicity in cortical neuron cultures induces LIF expression in cultured cortical astrocytes. The involvement of specific adenosine receptor activation in astrocytes required for LIF induction was controlled by using selective inhibitors of adenosine receptors. This study also included a detailed analysis of the intracellular signaling pathways downstream of adenosine receptor activation, in addition to analyses of the LIF carrying cytoplasmic compartments. Astrocytes play a major role in excitotoxicity by regulating extracellular glutamate concentrations. There is evidence that CNTF enhances both expression and activity of the glutamate transporter GLT-1 in astrocytes (Escartin et al, 2006), and thereby promotes survival of neurons against excitotoxicity (Beurrier et al, 201 0). Like for CNTF, the receptor subunits required for OSM binding are strongly expressed in astrocytes (Dallner et al, 2002; Kordula et al, 1998) . However, it is not understood how OSM affects astrocytic glutamate uptake. Therefore, in chapter 5,we studied the effect of OSM treatment on astrocytic glutamate uptake using radioactive measurement of glutamate transport by liquid scintillation spectroscopy. In addition, the effect of OSM treatment on the expression of the glutamate transporters, GLAST and GLT-1, was investigated both at mRNA and protein levels. Furthermore, the influence of OSM regulation of astrocytic glutamate transport on excitotoxicity was examined in cultured cortical neurons. Finally, the cell-specificroles of OSM and LIF in excitotoxicity are summarized and discussed in chapter 6. This chapter also contains a conceptual overview of the "interplay" between the adenosinergic system and IL-6 type cytokines during excitotoxicity.

32 GENERAL INTRODUCTION

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33 CHAPTER 1

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43 CHAPTER 1

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ABSTRACT Neuroprotection is one of the prominent functions of the interleukin (IL)-6-type cytokine family, for which the underlying mechanism(s) are not fully understood. We have previously shown that neuroprotection and neuromodulation mediated by IL-6 require neuronal adenosine A, receptor (A,R) function. We now have investigated whether two other IL-6-type cytokines (Oncostatin M (OSM) and Leukemia inhibitory factor (LIF)) use a similar mechanism. It is presented here that OSM, but not LIF, enhances the expression of A,Rs (both mRNA and protein levels) in cultured neurons. The neuroprotective effect of LIF was unchanged in A,R deficient neurons, whereas OSM failed to protect neurons in the absence of A1R. In addition, OSM pre-treatment for 4 hours potentiated the A,R-mediated inhibition of electrically-evoked excitatory post-synaptic currents (EPSCs) recorded from hippocampal slices either under normal or hypoxic conditions. No such effect was observed after LIF pre-treatment. Our findingsthus strongly suggest that, despite known structural and functional similarities, OSM and LIF use differentmechanisms to achieve neuroprotection and neuromodulation.

48 IL-6-TYPE CYTOKINES IN NEUROMODULATION AND NEUROPROTECTION

INTRODUCTION

Glutamate-induced excitotoxicity is a major cause for neuronal loss in many neurodegenerative disorders. This process is mediated by excessive activation of glutamate receptors and has been associated with damage observed after epileptic convulsions , stroke , spinal cord trauma , El head injury (Dirnagl et al, 1999; Faden et al , 1989; Vincent and Mulle , 2009) , and several neurodegenerative disorders such as Parkinson's disease , Alzheimer's disease , Huntington's disease , Amyotrophic lateral sclerosis and Multiple sclerosis (Dong et al , 2009). Excitotoxicity therefore remains a challenging problem in neuropathology ever since it was first described in the 1970's (Olney et al , 1972). Excitotoxicity causes excessive ATP hydrolysis , resulting in pronounced release of adenosine in affected brain tissue (Dunwiddie and Masino , 2001). Extracellular adenosine suppresses neuronal activity by at least three cellular mechanisms: 1) pre-synaptic inhibition of neurotransmitter release , 2) post-synaptic inhibition of calcium influx through voltage dependent calcium channels , and 3) activation of G-protein dependent inwardly rectifying K• channels (GIRKs) that mediate post-synaptic membrane hyperpolarization (de Mendonc;a et al , 2000; Dunwiddie and Masino , 2001). These effects also reduce metabolic demand , thereby preserving ATP stores , suppress glutamatergic transmission and thus protect neurons from excitotoxicity (Schubert et al , 1997). These effects in neurons are predominantly mediated by A,R.s. Although several adenosine receptor agonists like N6-cyclopentyladenosine (CPA) or 2-chloroadenosine (CADO) are neuroprotective , their therapeutic application in neurodegenerative diseases failed , as they provoke severe peripheral side effects and/or receptor desensitization (de Mendonca et al , 2000; Stone , 2002). In addition , attempts to rescue neurons using anti-excitotoxic drugs , seriously affected various aspects of synaptic plasticity rendering them useless for therapy (Mimica and Presecki , 2009; Parsons et al , 2007). This augments the urge for the development of new therapeutic approaches to suppress excitotoxicity. Other neuroprotective factors include the family of IL-6-type cytokines , which therefore are also called neurokines (Patterson , 1992). Members of this family like IL-6 , IL-11 , LIF , OSM, ciliary neurotrophic factor (CNTF) , novel neurotrophin 1 (NNTI) and cardiotropin-1 (CT-1) have been shown to have neuroprotective properties in various neuronal subpopulations (Gurfein et al , 2009; Holtmann et al , 2005; Suzuki et al , 2009; Weiss et al , 2006; Wen et al , 2005). Despite this large body of evidence it is currently not very well understood how IL-6-type cytokines mediate their neuroprotective effects. Recently , we demonstrated that IL-6 affords neuroprotection and neuromodulation by enhancing the expression and function of neuronal

A1 Rs (Biber et al , 2008). All IL-6-type cytokines require gpl30 receptor subunits for signaling leading to many shared redundant functions (Heinrich et al , 2003; Kamimura et al , 2003). We therefore hypothesized that IL-6-type cytokines in general exert their neuroprotective properties via a facilitation of neuronal Al function. IL-6 and IL-11 are the only IL-6-type cytokines that signal via gpl30 homodimers , while the remaining members signal via heterodimers of either gpl30 and LIFR (LIF , CNTF , CT-1 and NNT-1) or gpl30 and OSMR (OSM) (see for review: Bauer et al , 2007). Using LIF and OSM allows investigation of all major possible receptor signaling combinations that are known for the IL-6- type cytokine family. We thus here compared the neuroprotective properties of OSM and LIF in

49 CHAPTER2

cultured cortical and hippocampal neurons and investigated the electrophysiological effects of these cytokines in mouse hippocampal slices.

MATERIALS AND METHODS

Chemicals and Reagents Neurobasal media , BME media , HBSS , PBS , sodium pyruvate , L-glutamine , penicillin-streptomycin , HEPES , glutaMAX-1 and B27 supplement were from Gibco (Breda , The Netherlands). DMEM media and FCS were from PAA laboratories (Colbe , Germany). Mito serum extender was from Becton Dickinson Labware (Breda , The Netherlands). Trypsin was from Life technologies (Breda , The Netherlands) and papain from Worthington Biochemical Corporation (Lakewood , NJ , USA). Other chemicals were from Sigma-Aldrich (Zwijndrecht , The Netherlands). Cytokines , recombinant mouse LIF was from Millipore (Amsterdam , The Netherlands) and recombinant mouse OSM from Sigma-Aldrich. Mouse monoclonal anti-adenosine A, receptor antibody supernatant was kindly provided by Prof. Yuko Sekino and mouse monoclonal anti-�-actin was from Abeam (Cambridge , MA , USA). Fluorescence conjugated secondary antibody , donkey anti-mouse (IR Dey680) used separately to detect adenosine A1 receptor and �-actin bands was from LI-COR biosciences (Cambridge , UK).

Animal experiments Experiments were performed using wild-type C57BL/6J mice (Harlan , Horst , The Netherlands or Harlan lberica , Spain) and adenosine A1 receptor knockout (A1 R -/-) mice with the same genetic background (Universita di Roma "La sapienza" , Rome , Italy). All procedures were in accordance with the regulation of the Ethical Committee for the use of experimental animals of the University of Groningen , The Netherlands (License number DEC 4623) , as well as with the Portuguese law on Animal Care and European Union guidelines (86/609/EEC). Mice were housed in standard makrolon cages and maintained on a 12 hour light/dark cycle. They received food and water ad libitum.

Primary neuronal culture from mouse embryo and neonates

) Primary neuronal cultures from mouse embryo (~E15 were established as described previously (Biber et al , 2008). Primary neuronal cultures from newly born mouse neonates (P0 ) were established as described previously (Limatola et al , 2005). The neuronal purity , determined by MAP2-staining , was 90-95% (data not shown). Cortical and hippocampal neurons were used for the experiments after 5 and 9 days of in vitro culture respectively.

Induction of excitotoxicity Following the number of days in culture , cortical and hippocampal neurons were incubated without or with recombinant mouse OSM or LIF (0.1 , 1 or 10 ng/mL) for the indicated periods of time. Where indicated , neurons were incubated with a selective A1 R antagonist , 8-cyclopentyl-1 , 3-dipropylxanthine (DPCPX , 100 nM) or agonist , CPA (100 nM) for 15 min before they were subjected to excitotoxicity with glutamate (50 µM) for 1 hour. Following glutamate treatment , the media was refreshed and neuronal cultures were incubated for 24 hours before assessing the degree of cytotoxicity.

so IL-6-TYPE CYTOKINES IN NEUROMODULATION AND NEUROPROTECTION

Determination of neuronal viability

1) MTT assay - Survival of embryonic (~E15) cortical neurons in culture was measured by the colorimetric 3-(4 , 5-dimethylthiazol-2-yl-) 2, 5-diphenyltetrazolium bromide (MTT) assay , as described previously (Biber et al , 2008; Mosmann , 1983). El 2) Lysis buffer assay - 24 hours after glutamate treatment , cultures of cortical and hippocampal neurons isolated from neonatal pups (P0) were treated with detergent-containing lysing solution (0.5% ethyl hexadecyl dimethylammonium bromide , 0.28% acetic acid , 0.5% Triton

, X-100 , 3 mM NaCl , 2 mM MgCl2 in PBS; pH 7.4; diluted 1:10) for 5 min at room temperature and the viable cells were counted on a hemacytometer (Thoma chamber , Brand , Germany) , as described previously (Limatola et al , 2005; Volante et al , 1994).

Western blotting Equal amounts of protein (30 µg) were loaded on to 15% SDS-polyacrylamide gels and subsequently transferred on to PVDF membranes. The membranes were blocked using Odyssey blocking buffer (088; diluted 1:1 in PBS) for 1 hour and incubated overnight at 4°C on shaker with one of the following primary antibodies diluted in 1:1 088 and PBS + 0.1% Tween 20

(PBS-T): mouse monoclonal anti-A1 R antibody supernatant (1:50) and mouse monoclonal anti­ j3-actinantibody (1:8000). The membranes were then washed in PBS-T (4 x 5 min) , followed by incubation with fluorescent conjugated secondary antibody , donkey anti-mouse (IR Dey680 LI-COR; 1:8000 in PBS-T) , for 1 hour at room temperature on gentle shaking in the dark. Membranes were washed again in PBS-T (4 x 5 min) and fluorescent bands detected using LI­ COR's Odyssey infrared imaging system. The densitometry analysis of protein bands was done using Image J program.

Real-time Polymerase Chain Reaction (Q-PCR) Total RNA extracted from cultured cortical neurons were purified and then transcribed into cDNA as described previously (Biber et al , 2008). Adenosine A1 receptor mRNA expression of cultured embryonic cortical neurons treated with OSM (1 ng/ml) or LIF (10 ng/ml) for different time periods (1 , 2, 4, 8, 12 , and 24 hours) was analyzed by real-time PCR using the iCycler (Bio-Rad , Veenendaal , The Netherlands) and the iQ SYBR Green supermix (Bio-rad). Neurons that were not treated with OSM or LIF, served as control. Mouse ribosomal protein L32-A (rpl32A) and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) primers were used for normalization to housekeeping genes and they showed no variations in response to experimental treatments (see Table 1 for primer sequences). The comparative Ct method (amount of target amplicon X in Sample S, normalized to a reference R and related to a control sample C, calculated by

2-((C/ ,S-CtR ,S)-(C/ ,C-CtR,C)) was used to determine the relative expression levels (Livak

Table 1. Primers used forrea l-time polymerase chain reaction (Q-PCR).

Accession Product Gene number Forward primer (5'-3') Backward primer (5'-3') size (bp) GAPDH AF106860 ATGGCCTTCCGTGTTCCTAC GCCTGCTTCACCACCTTCTT 104 rpL32A NM-172086 GCTGGAGGTGCTGCTGATGT ACTCTGATGGCCAGCTGTGC 114 �R NM-001008533 CCTCTCCGGTACAAGACAGT GGTGTCAGGCCTACCACAAG 92

51 CHAPTER 2

and Schmittgen , 2001; Biber et al , 2008). Linear regression analysis of the data was performed to understand the effect of cytokine treatment over time on the neuronal A,R expression.

Patch-clamp recordings in hippocampal slices Acute hippocampal slices (300 µm-thick) were prepared as previously described (Diogenes et al , 2004). Whole-cell patch clamp recordings of excitatory postsynaptic currents (EPSCs) were obtained from CAl pyramidal cells upon stimulation of the Schaffer collateral fibers (0.2 ms rectangular pulses delivered each 30 s). EPSCs were acquired at room temperature with an Axopatch 2008 amplifier (Axon Instruments; Clampex 10 software) at 10 KHz and were low-pass filtered at 2 KHz. CAl pyramidal cells were identified by visualization with an upright microscope (Zeiss Axioskop 2FS) equipped with infrared video microscopy and differential interference contrast optics , and were functionally distinguished from interneurons by their slower firing frequencies , longer action potentials and for featuring spike-frequency adaptation (Madison and Nicoll , 1984). Patch pipettes had resistances of 4-7 MO when filled with an internal solution containing (in mM): K-gluconate 125 , KCI 11 , CaCl2 0.1 , MgCl2 2, EGTA 1, HEPES 10 , MgATP 2, NaGTP 0.3 and phosphocreatine 10 , pH 7.3 , 280-290 mOsm. Junction potentials were not corrected nor were access resistance compensated for. Whole-cell excitatory postsynaptic currents were recorded in voltage-clamp mode (V h = -70 mV) and averages of four consecutive individual responses were determined. Access resistance and holding current were continuously monitored throughout the experiment , and if any varied by more than 20% , the experiment was discarded. Slices were perfused (2-3 ml/min) with artificial cerebrospinal fluid (aCSF) , which contained (in mM): NaCl 125 , KCI 3, NaH/O4 1.25 , NaHCO3 25 , caq 2, Mg SO4 1 and glucose 10. Drugs were added to the perfusion solution. Hypoxia was induced by substituting the aCSF with an identical aCSF pre-equilibrated with

95% N2 and 5% CO2 for 4 min. This manipulation reduces bath oxygen tension in the recording chamber from -600 mm Hg to -250 mm Hg (Sebastiao et al , 2001). Each slice was subjected to a single period of hypoxia , since the effects of hypoxia may be modifiedby subsequent episodes in the same slice (Perez-Pinzon , 1999).

Statistical data analysis The absolute data values were converted to percentage of control in order to allow multiple comparisons and statistical analysis performed by one-way ANOVA followed by the Bonferroni correction , using the Statistical Package for the Social Sciences (SPSS , Chicago , IL , USA). Analysis of the electrophysiology data was done using paired t-test and that of the western blot data , by Non-parametric Mann-Whitney U test . In all cases , p values < 0.05 were considered statistically significant.

52 IL-6 TYPE CYTOKINES IN NEU ROMODULATION AND NEU ROPROTECTION

RESULTS

OSM, but not LIF-induced neuroprotection against excitotoxicity is dependent on

neuronal A1Rs Treatment with various concentrations of glutamate (20 up to 200 µM) for 1 hour caused a El concentration-dependent cell death in cultured embryonic cortical neurons that was abolished by the NMDA receptor antagonist dizocilpine (MK 801), as described earlier (Suppl. Figure 1; Biber et al, 2008) . Pre-treatment with OSM or LIF (0.1, 1 and 10 ng/ml for 24 hours) alone had no effect on neuronal viability but significantly protected neurons against toxicity induced by glutamate (50 µM for 1 hour) (Suppl. Figure 2A). Subsequently, the effect of OSM (1 ng/ml) and LIF (10 ng/ml) treatment on neuronal survival was tested over various time periods (1, 2, 4, 8, 12, and 24 hours) (Suppl. Figure 2B-C). In addition, pre-treatment with OSM (1 ng/ml) or LIF (10 ng/ ml) for 24 hours showed a reduced caspase-3 activation and reduced propidium iodide uptake in MAP2-positive neurons subjected to glutamate toxicity (50 µM for 1 hour) (Suppl. Figure 3A-B).

Inhibition of A1 R activity in embryonic cortical neurons by DPCPX abolishes the neuroprotective function of IL-6 (Biber et al, 2008) . Indeed, pre-incubation with DPCPX (100 nM for 15 min before glutamate) completely abolished the neuroprotective effects of OSM (1 ng/ml for 24 hours) (Figure lA), but left neuroprotection by LIF unaffected (10 ng/ml for 24 hours) (Figure lA). DPCPX alone, however, did not influence glutamate-induced neurotoxicity (Figure lA-B).

A B 120 120

100 100 g e * C 80 § 80 08 0 60 60 � � � n 0 �·s: 40 �2'. 40 � � a:; a:;u 20 u 20

0 0 Figure 1: Effect of A,R antagonist 11 + + LIF + OSM - - and agonist on OSM and LIF induced + - I+ II+ + DPCPX - DPCPX - neu roprotection against glutamate Glutamate + + + Glutamate + + + toxicity. Primary cortical neurons from wild-type (C57BL/6J) mice embryo ( ~E) C D 120 120 were treated without or with OSM (1 ng/ [7 ml) (A, C) and LIF (10 ng/ml) (B, D) for 100 100 24 hours after S days in culture and were e subsequently challenged with glutamate § 80 1 80 (50 µM) forl hour. Where indicated, cultures 0 �0 were treated with A,R antagonist, DPCPX 60 � � f.1• 60 (100 nM) or agonist, CPA (100 nM) for 15 "iii "iii� > > min prior to glutamate insult. Neuronal ·s: 40 -� 40 � � survival was assessed 24 hours following a:;u 20 a:;u 20 the glutamate insult by a colorimetric MTT-assay. The optical densities were measured at 570 nm with a 630 nm and - - OSM + - + + LIF - + - - blank correction. The bars represent Mean II+ II CPA - - - CPA ± S.E.M of 3 independent experiments; ** p Glutamate - + + + + Glutamate value < 0.001, * p value < 0.05.

53 CHAPTER 2

Neuroprotection against glutamate toxicity induced by OSM, but not LIF, is enhanced by the A,R agonist, CPA

We have previously shown that treatment of embryonic cortical neurons with the A1 R agonist, CPA alone did affectglutamate- toxicity, but that neuroprotection induced by IL-6 was further enhanced by CPA (Biber et al, 2008). Similarly, we now describe that CPA (100 nM for 15 min before glutamate) significantlyenhanced the neuroprotective effect of OSM against glutamate (50 µM for 1 hour), but it did not affect neuronal survival in the absence of OSM treatment (Figure lC). In contrast, LIF-induced neuroprotection was not affected by CPA (Figure lD). In accordance with the survival experiments, CPA (100 nM for 30 min) alone did not affect the glutamate mediated calcium transients in cultured cortical neurons (Suppl. Figure 4C). On the other hand, in neurons pre-treated with OSM (1 ng/ml for 24 hours), CPA not only delayed the influx of calcium, but also decreased the rate of calcium entry (Suppl. Figure 4D).

OSM, but not LIF, enhances A,R expression in cultured cortical neurons

IL-6 treatment enhances A1 R expression in neurons (Biber et al, 2008). We here investigated whether treatment with OSM (1 ng/ml) or LIF (10 ng/ml) for various time periods (1, 2, 4, 8, 12 and 24 hours) has a similar effect in cultured embryonic cortical neurons. Linear regression analysis of RT-PCR experiments shows a time dependent increase in the A,R mRNA expression, when normalized to GAPDH (Figure 2A; R2 = 0.402, p < 0.001, n =3) and to rpl32A (Suppl. Figure 5; R2 = 0.284, p = 0.005, n =3), in neurons treated with OSM, with an effect already apparent after 1 hour of incubation. In addition, western blot analysis confirms this since treatment with OSM (1 ng/ml for 24 hours) significantly(p < 0.05; n = 3) increased A,R protein expression compared to the non-treated control (Figure 2B). LIF treatment, however, did not influence A,R mRNA or protein expression in cortical neurons (Figure 2 and Suppl. Figure 5).

The protective effectof OSM, but not of LIF, against glutamate-induced excitotoxicity is lost in A,R -/- neurons

To further evaluate the involvement of A1 Rs in OSM-induced neuroprotection, neuronal cultures were prepared from A1 R -/- mice and data compared with that obtained from wild type mice. Neuronal cultures were prepared from new-born mice (P0 ) to allow genotype confirmation, as well as evaluation of neuroprotection in distinct brain areas (cortex and hippocampus). Pre­ incubation of wild-type cortical and hippocampal neurons with OSM or LIF (1 or 10 ng/ml for 24 hours) significantlyinc reased their survival against glutamate toxicity (Figure 3A-B). As shown for embryonic neurons, the protective effects of OSM in neonatal neurons were also abolished by DPCPX (100 nM, for 15 min before glutamate challenge) (Figure 3A). Pre-incubation with OSM did not protect against glutamate toxicity in cultured cortical and hippocampal neurons from A1 R-/­ mice (Figure 3C). In contrast to OSM, DPCPX treatment (100 nM, added 15 min prior to glutamate challenge), did not change the protective effect of LIF (Figure 3B), thus confirmingthe findingsin embryonic neurons. Consistently, cortical and hippocampal neurons from A1 R -/- mice were still protected by LIF pre-treatment (10 ng/ml, for 24 hours before glutamate) (Figure 3D).

Modulation of A,R-mediated actions on synaptic transmission by OSM but not LIF Since receptors for OSM and LIF are strongly and specificallyexpressed in hippocampal principal layers (Rosell et al, 2003), we investigated whether sustained exposure of hippocampal slices to

54 IL-6-TYPE CYTOKINES IN NEUROMODULATION AND NEUROPROTECTION

A - OSM 11 ng/ml) - LIF110 ng/ml) CHOA1F Corticalneurons 350 (+ve control) control OSM LIF c .::::::- 300 ·-a ...e A1R (37 kDa) El ...� § u 250 a. .... I � 0 200 • c:i:l � :r 150 EO 140 � � 100 ·- l!J C e .., 120 �QJ �� 2 5 100 a: 50 O u � a...... 0 80 c:i:��� Time QJ C Oh 1h 2h 4h 8h 12h 24h > 0 60 ·,.::; ·.:;; ra"' - QJ 40 QJ a. ... a: 20 0 control OSM LIF

Figure 2: OSM, but not LIF, enhances A R expression in wild-type (CS7BL/6J) cortical neurons in vitro. (A) 1 Real-time PCR analysis of A1R mRNA expression in cultured embryonic ( ~E15) cortical neurons. Graph shows the relative expression level of A1R mRNA (normalized to GAPDH) in cultures incubated with OSM (1 ng/ml) or LIF (10 ng/ml) for 1 to 24 hours, compared to the non-treated controls. Data presented as Mean ± S.E.M of 3 separate experiments, expressed as percentage of the control. * p value < 0.05 (compared to the control); independent samples t-test. (B) Western blot analysis of Al protein expression in cultured embryonic cortical neurons incubated with OSM (1 ng/ml) or LIF (10 ng/ml) for 24 hours, compared to the control. Relative optical densitometry values of A1R proteins are shown in the lower panel. Data is presented as percentage of each respective ratio between optical density value of A1R band intensity and optical density value of the matched = �-actin band intensity (n 3); CHOA1F - Chinese hamster ovary cells transfected with full-length A1R gene (positive control for A1R). * p value < 0.05.

OSM or LIF could modify the neuromodulatory actions of A1 Rs on synaptic transmission, using whole-cell recordings of electrically-evoked EPSCs from CAl pyramidal cells. To prevent pre­ conditioned responses, the effects of the selective A1 R agonist (CPA, 30 nM) were investigated in only one neuron per slice, from control and OSM or LIF (1 0 ng/ml, for 4 hours) treated slices, prepared from the same hippocampus. Activation of Als is well known to decrease synaptic transmission in the hippocampus (Sebastiao et al, 1990). Accordingly, CPA (30 nM) inhibited EPSC peak amplitude by 48 ± 7.7% (n=4), in control conditions (Figure 4). When recording from slices pre-incubated with OSM, EPSC inhibition by CPA was significantlyin creased to 73 ± 6.7% (n=4, p < 0.05, compared to the effect of CPA in the absence of OSM). In contrast to OSM, EPSC inhibition by CPA was not significantly affected (p > 0.05, n =4 for each group, comparisons within slices from the same hippocampus) by pre-exposure to LIF (Figure 5). Furthermore, the experiments with LIF incubation served as control for the possibility that prolonged pre­ incubation time (4 hours) per se would be responsible for the potentiation of the inhibition caused by CPA, observed in slices incubated with OSM but not in slices incubated with LIF for similar time periods.

55 CHAPTER 2

A B Cortical neurons Hippocampal neurons Cortical neurons Hippocampal neurons

120 120 100 100 I I * g 80 gC 80 0 i 0 * 0 .....0 60 n· 60 lij· iiil iii� > nJ > '> 40 40 -�� ai n· u 20 aiu 20 0 11 1 0 OSM + -- + + + II-- + + LIF - + II- - + + - + II- - + + DPCPX + - + -- - + - + DPCPX + - + + - + Glutamate + + + + -- + + + + Glutamate + + + + -- + + + +

C D Cortical neurons Hippocampal neurons Cortical neurons Hippocampal neurons

120 120

100 100

gC 80 g 0 0 80 u u 0 0 60 60 11 iii� � iii> .> 40 ·s: 40 n � � ai u 20 aiu 20

0 0 OSM + I + + II + LIF + I + + I + Glutamate + + + + Glutamate + + + +

Figure 3: OSM and LIF induced neuroprotection against glutamate toxicity in wild-type (CS7BL/6J) and

A1R -/- neurons. Primary cortical and hippocampal neurons from (A, B) C57BL/6J, and (C, D) A1R -/- mice (A, C) neonates (P0) were pre-incubated without or with OSM (10 ng/ml for 24 hours) or LIF (10 ng/ml for 24 hours) (B, D). Where indicated, the neurons were also pre-incubated with the Al antagonist, DPCPX (100 nM) for 15 min and were subsequently challenged with glutamate (1 00 µM) for 1 hour. Neuronal viability was measured 24 hours after the glutamate challenge by Lysis-buffer assay. The bars represent Mean ± S.E.M of 3 independent experiments; ** p value < 0.001, * p value < 0.05.

OSM potentiates A1R-mediated depression of synaptic transmission during hypoxia The consistent increase in the extracellular concentration of endogenous adenosine that follows hypoxia and the subsequent A,R-dependent inhibition of synaptic transmission (Sebastiao et al, 2001) are main neuroprotective mechanisms in ischemic brain damage and hypoxia (Rudolphi et al, 1992). We therefore investigated if the OSM-induced modulation of Al fu nction could have any functional impact in hypoxic conditions in pyramidal cells. In control slices, brief (4 min) hypoxic insult caused a 42 ± 4.8% maximum inhibition of EPSC amplitude (n=7). In hippocampal slices previously exposed to OSM (1 0 ng/ml, for 4 hours),

56 IL-6-TYPE CYTOKINES IN NEUROMODULATION AND NEUROPROTECTION

A A o Control <1J 175 <1J 175 o Control "C o OSM (10 nglml) "C .€ 150 .€ 150 o LIF (10 nglml) a. 125 � t 125 rtJ � 100 � 100 El 75 � 75 -c -� 50 -� 50

� 25 E 25 0 0

-10 0 10 20 30 40 50 60 70 -10 10 20 30 40 50 60 70 Time (min) Time (min)

B B

...... 1,,/...---··--4 - --- It / 1 �/_J 1�✓;�__J

C C 100 100

uct 80 uct 80 s. C n 60 0 60 ·.:: � ·.:: E E 40 .!:: 40 u·=

20 20 if.

CPA (30 nM) I+ CPA (30 nM) II+ OSM (10 ng/ml) + LIF (10 ng/ml)

Figure 4: OSM potentiates inhibition of Figure 5: LIF does not alter inhibition of synaptic synaptic transmission caused by A R activation. transmission caused by A R activation. (A) 1 1 {A) Averaged time-course of afferent evoked Averaged time-course of afferent evoked EPSC peak EPSC peak amplitude changes caused by a 20 min amplitude changes caused by a 20 min application application of the A1R agonist CPA (30 nM), in of the A1R agonist CPA (30 nM), in control versus control versus OSM (10 ng/ml for 4 hours) treated LIF (10 ng/ml for 4 hours) treated slices, from the slices, from the same hippocampus (n=4). Each same hippocampus (n=4). Each point represents point represents average amplitude of 4 EPSCs average amplitude of 4 EPSCs evoked once evoked once every 30s by electrical stimulation every 30s by electrical stimulation of the Schaffer of the Schaffer collaterals; 100% corresponds collaterals; 100% corresponds to the averaged to the averaged amplitude calculated for 5-10 amplitude calculated for 5-10 EPSCs recorded just EPSCs recorded just before adding CPA. Panel (B) before adding CPA. Panel (B) shows superimposed shows superimposed current tracings of EPSCs current tracings of EPSCs recorded before (1, 2) and recorded before (1, 2) and 20-30 min after (3, 4) 20-30 min after (3, 4) introduction of CPA in the introduction of CPA in the superfusion medium, superfusion medium, in representative control (1, in representative control (1, 3) and OSM-treated, 3) and LIF treated (2, 4) cells. Histogram (C) shows (2, 4) cells. Percent of EPSC inhibition corresponds percent inhibition of synaptic transmission (EPSC) to the average EPSC decrease 20-30 minutes after caused by activation of Als is not significantly starting CPA application. Histogram (C) shows different between control and LIF treated slices; percent inhibition of synaptic transmission (EPSC) n.s. (not significant) p value > 0.05, paired t-test. caused by activation of A1Rs was significantly higher Calibration: 30 ms, 50 pA. in OSM treated slices compared to the control; * p value < 0.05, paired t-test. Calibration: 30 ms, 50 pA.

57 CHAPTER2

the same hypoxic insult caused a 69 ± 3.4% EPSC inhibition (Figure 6A), which was significantly higher (p < 0.05) than that observed in control slices prepared from the same hippocampus. The hypoxia-induced depression of EPSC amplitude and its potentiation by OSM were abolished upon Al blockade by DPCPX (50 nM, applied 30 min before hypoxia induction) (Figure 7).

A o Control QJ 175 -c o OSM (10 ng/ml) .€ 150 C. N2 /CO2 125 roE

V1u 100 CL LU --��-----+ 3 tt-tilfQ ❖ 94i#t� -c 75 ;H -� 50 ? + E 25 T • 0 .----,----,�-.-----,----,-- ,---, -10 0 10 20 30 40 50 Time (min)

B

Figure 6: OSM potentiates hypoxia-induced inhibition of synaptic transmission. (A) Averaged time-course of afferent evoked EPSC peak amplitude changes caused by a 4 min hypoxic insult, recorded from control (aCSF-treated) and OSM (10 ng/ ml for 4 hours) treated slices prepared from the same hippocampus. Each point represents average amplitude of 4 EPSCs evoked once every 30s by C electrical stimulation of the Schaffercoll aterals; 100% corresponds to the averaged amplitude calculated for 100 5-10 EPSCs recorded just before hypoxia. The hypoxic

C insult consisted of replacing oxygenated aCSF (95% 0 ) 80 02 and 5% CO2 by aCSF saturated with 95% N2 and 5% :-E rt1 .o ·x CO2 for 4 min. Maximum inhibition was determined :g·- >. g_ 60 as the lowest EPSC amplitude recorded for each u .c V1 C experiment, at either 6 or 8 min afterhypoxia onset. CL ·- 11 LU E Panel (B) shows superimposed current tracings of Es;t- 40 EPSCs recorded before (1, 2) and 8 min afterhypoxia :::, ... ·E- 4=ai (3, 4) was induced, in representative control (1, 3) X ro 20 and OSM treated (2, 4) cells. Histogram (C) shows maximum inhibition of synaptic transmission induced a by hypoxia is significantly higher when recording from slices that were pre-treated with OSM (n=7); * p OSM (10 ng/ml) I + value < 0.05, paired t-test. Calibration: 30 ms, 50 pA.

58 IL-6-TYPE CYTOKINES IN NEUROMODULATION AND NEUROPROTECTION

A B

CIJ 175 o Control 100 "O • OSM (10 ng/rnl) C .E 150 :E0 ro 80 t125 El ro � "§ - a. U 100 i· >, 60 VI u .c VI C w 75 a. ·- w E 40 E'<:I" -� 50 :, ro .... ·E- .:I:'. (I) E 25 X ro ro 20 Z 0 DPCPX (50nM) � 0 -10 0 10 20 30 40 50 + -+ DPCPX (50 nM) Time (min) OSM (10 ng/ml) • +

Figure 7: Hypoxia-induced inhibition of synaptic transmission and its potentiation by OSM is dependent on A1R activation. Averaged time-course of afferent evoked EPSC peak amplitude changes caused by a 4 min hypoxic insult, delivered under conditions of Al blockade by the selective antagonist DPCPX (50 nM); EPSCs were recorded from control ( o) and OSM treated (•) slices from the same hippocampus (n=S); DPCPX was applied 30 min before induction of hypoxia. Further details as in Figure 6.

DISCUSSION

We have recently provided a possible explanation for IL-6 mediated neuroprotection. IL-6 treatment (thus gp130 homodimer activity) caused an up-regulation of neuronal A1 R expression and function in vitro and in vivo, which was mandatory for IL-6-dependent neuroprotection (Biber et al, 2008). This hypothesis is corroborated by our findings that IL-6 does not affect glutamate-induced excitotoxicity in A1 R deficient neurons(unpubl ished observations). Notably, the members of IL-6-type cytokine family oftenshow overlapping biological properties, due to the shared usage of gpl30 receptor subunits in their signaling cascade (Bauer et al, 2007; Taga and Kishimoto, 1997). We here investigated the possible role of A1 R function in neuroprotection and neuromodulation by OSM (OSMR/gp130 heterodimer) and LIF (LIFR/gp130 heterodimer) in hippocampal slices and in cultured neurons from wild-type and A1 R -/- mice. Here we describe that pre-treatment for 24 hours with OSM or LIF attenuates excitotoxicity in cultured neurons from cortex and hippocampus. These findings reinforce the idea of a principal neuroprotective effect of IL-6-type cytokines (see for review: Ransohoff et al, 2002). Since both cytokines display a comparable efficacy, similar mechanisms of action might be expected. However, blocking neuronal A1 R function with a selective antagonist (DPCPX, 100 nM) completely abolishes the neuroprotective effects of OSM, but leaves LIF-induced neuroprotection unaffected. In addition, activation of A1 R function with CPA (100 nM; 15 min before glutamate) is effective in OSM-treated, but not in LIF-treated neurons. The difference between OSM and LIF was also found in cultured cortical and hippocampal neurons from

A1 R -/- mice. While the neuroprotective effect of LIF is preserved in A1 R -/- neurons, OSM pre-treatment does not affect neuronal survival against glutamate toxicity in the absence of functional adenosine A1 receptors. The lack of effects of CPA and DPCPX in untreated neurons in vitro. most likely reflects an insufficientA 1 receptor expression level in these cells Whether or not this is due to the culture conditions is not clear at the moment. However, since in untreated

59 CHAPTER 2

acute slices both CPA and DPCPX showed effects it is indicated that in brain tissue basal A1 receptor expression level is sufficient for neuroprotection and can be increased by IL-6-type cytokines. When evaluating consequences for synaptic transmission , we found that a 4 hour pre­ treatment with IL-6 potentiates the A1 R-mediated inhibition of synaptic transmission in hippocampal slices , an effect that was of particular importance during short periods of hypoxia (Biber et al , 2008). Here we present that OSM , but not LIF, significantly increases Al mediated inhibition of EPSCs. Similarly to IL-6 , OSM pre-treatment also causes a significantlyhigher drop in EPSCs in response to a 4 min hypoxic period , which is dependent on Al function. Thus , OSM , but not LIF, sensitizes neuronal Al-mediated responses , equally to the effects described earlier for I L-6. It may therefore be concluded that OSM-treatment potentiates the functioning

of neuronal A1 R, thereby increasing adenosinergic effects required for inhibition of synaptic transmission and neuroprotection. Moreover , real-time PCR and western blot analysis strongly support this hypothesis as Al mRNA and protein expression levels are increased in OSM­ treated , but not in LIF-treated neurons. Taken together , our findingsstrongly suggest that two

members of the IL-6-type cytokine family (IL-6 and OSM) depend on A1R function for their neuroprotective properties , whereas LIF induces neuroprotection via a different , unknown , mechanism. However , since both cytokines are neuroprotective in vitro and may reduce ischemic damage in vivo (see for review: Suzuki et al , 2009) a further understanding of their mechanisms of action may provide new therapeutic possibilities in stroke patients. Although LIF and OSM are highly related members of the IL-6 type cytokine family (Jeffery et al , 1993; Nicola et al , 1993) sharing most properties , distinct effects upon activation of LIF and OSM receptor complexes have been reported. For example , only OSMR/gp130 heterodimer activation is able to promote osteoblast differentiation , whereas activation of both OSMR/ gp130 and LIFR/gpl30 heterodimers in these cells inhibited the expression of osteocalcin , a protein required for bone-building (Malaval et al, 2005). Furthermore , selective roles of OSM and LIF during haematopoiesis and their effects on the regulation of certain target genes have been reported (Tanaka et al , 2003; Weiss et al , 2005) , reinforcing the idea that different IL-6- type cytokine receptor complexes may activate specificsignaling cascades. It is at the moment unclear which signaling pathways are important for the neuroprotective effects of LIF , OSM and IL-6. In our preliminary observations , using blockers for any of the possibly involved signaling pathways (ERKl/2 , STAT3 , Pl3K or NF-KB) , we were not able to identify the different signaling pathways for the neuroprotective effects of LIF, OSM and IL-6 (data not shown). In fact most of the blockers (ERKl/2 , PI3K or NF-KB) completely abolished neuroprotection of all cytokines , but did not discriminate between OSM and LIF (data not shown). The reasons for these unexpected results are not clear at the moment. More sophisticated experiments to interfere with different signaling cascades by RNAi interference might provide more clear results. It would be particularly relevant to unravel the cellular mechanism(s) by which IL-6 and

OSM differentially regulate A1 R expression. It has been shown that the basal as well as induced expressions of Als are regulated by the nuclear transcription factor - kappa B (NF-KB) (Jhaveri et al , 2007). Noteworthy , OSM regulates protein synthesis through NF-KB activation , in smooth muscle cells (Nishibe et al , 2001) . When subjected to oxidative stress , these cells show an increase in A1 R mRNA that is prevented by inhibitors of the NF-KB (Nie et al , 1998). Incidentally ,

60 IL-6-TYPE CYTOKINES IN NEUROMODULATION AND NEUROPROTECTION

IL-6 has also been shown to activate NF-KB in intestinal cells (Wang et al, 2003). Therefore, it

is likely that increased expression of A1 Rs by OSM and IL-6 is also, at least partially, regulated by NF-KB, an assumption that is corroborated by our unpublished findings. Nevertheless, it is irrefutable that adenosine A receptors are key players in neuroprotection against excitotoxicity 1 El induced by OSM and IL-6.

It has long been known that neuronal A1 R suppress neuronal activity and glutamatergic transmission, reduce oxidative stress, minimize metabolic demand thereby preserving ATP stores, and protect neurons from excitotoxicity (Schubert et al, 1997). The concept now emerges that cytokines from various families may utilize A1 R to exert their neuroprotective functions. In

the absence of functional adenosine A1 receptors, numerous cytokines, such as IL-6 (Biber et al, 2008), OSM (present work), chemokine (C-X3-C motif) ligand 1 (CX3CL1) (Lauro et al, 2008), fail to protect neurons under excitotoxic conditions. Given the urgent need for new therapies in neurodegenerative diseases, it is concluded that a detailed analysis of the intracellular signaling cascade activated by IL-6 and OSM is not only important for our molecular understanding of cytokine biology, but might furthermore provide ideas of cellular mechanisms by which the expression and function of neuronal Als can be therapeutically increased.

61 CHAPTER2

REFERENCES 1. Bauer S., Kerr B.J., Patterson P. H. (2007) The neuropoietic cytokine family in development, plasticity, disease and injury. Nat. Rev. Neurosci. 8(3), 221-32. 2. Biber K., Pinto-Duarte A., Wittendorp M. C. eta I (2008) lnterleukin-6 Upregulates Neuronal Adenosine A1 Receptors: Implications for Neuromodulation and Neuroprotection. Neuropsychopharmacology33(9}, 2237 SO. 3. De Mendonca A., Sebastiao A. M., Ribeiro J. A. (2000) Adenosine: does it have a neuroprotective role after all? Brain Res. Brain Res. Rev. 33, 258-74. 4. Diogenes M. J., Fernandes C. C., Sebastiao A. M., Ribeiro J. A. (2004) Activation of adenosine A2A receptor facilitates brain-derived neurotrophic factor modulation of synaptic transmission in hippocampal slices. J. Neurosci. 24(12), 2905-13. 5. Dirnagl U., ladecola C., Moskowitz M. A. (1999) Pathobiology of ischemic stroke: an integrated review. Trends Neurosci. 22, 391-397. 6. Dong X. X., Wang Y., Qin Z. H. (2009) Molecular mechanisms of excitotoxicity and their relevance to pathogenesis of neurodegenerative diseases. Acta. Pharmacol. Sin. 30(4), 379-87. 7. Dunwiddie T. V., Masino S. A. (2001) The role and regulation of adenosine in the central nervous system. Annu. Rev. Neurosci. 24, 31-55. 8. Faden A. I., Demediuk P., Panter S. S., Vink R. (1989) The role of excitatory amino acids and NMDA receptors in traumatic brain injury. Science 244(4906), 798-800. 9. Gurfein B. T., Zhang Y., LopezC. B., ArgawA. T., Zameer A., Moran T. M., John G. R. (2009) IL-11 regulates autoimmune demyelination. J. lmmunol. 183(7), 4229-40. 10. Heinrich P. C., Behrmann I., Haan S., Hermanns H. M., Mueller-Newen G., Schaper F. (2003) Principles of interleukin (IL)-6-type cytokines signalling and its regulation. Biochem. J. 374, 1-20. 11. Holtmann B., Wiese S., Samsam M., Grohmann K., Pennica D., Martini R., Sendtner M. (2005) Triple knock-out of CNTF, LIF, and CT-1 definescooperative and distinct roles of these neurotrophic factors for motoneuron maintenance and function. J. Neurosci. 25(7), 1778-87. 12. Jeffery E., Price V., Gearing D. P. (1993) Close proximity of the genes for leukemia inhibitory factor and oncostatin M. Cytokine. 5(2), 107-11. 13. Jhaveri K. A., Reichensperger J., Toth L. A., Sekino Y., Ra mkumar V. (2007) Reduced basal and lipopolysaccharide­ Neuroscience stimulated adenosine A1 receptor expression in the brain of nuclear factor-kB p50 mice. 146, 415- 426. 1 14. Kamimura D., lhsihara K., Hirano T. (2003) IL-6 signal transduction and its physiological roles: the signal orchestration model. Rev. Physiol. Biochem. Pharmacol. 149, 1-38. 15. Lauro C., Di Angelantonio S., Cipriani R., Sobrero F., Antonilli L., Brusadin V., Ragazzino D., Limatola C. (2008) Activity of adenosine receptors type 1 is required for CX3CL1-mediated neuroprotection and neuromodulation in hippocampal neurons. J. lmmunol. 180(11), 7590-6. 16. Limatola C., Lauro C., Catalano M., Ciotti M. T., Bertollini C., Di Angelantonio S., Ragozzino D., Eusebi F. (2005) Chemokine CX3CL1 protects rat hippocampal neuronsagainstglutamate-mediated excitotoxicity. J. Neuroimmunol. 166(1-2), 19-28. 17. Livak K. J., Schmittgen T. D. (2001) Analysis of relative gene expression data using real time quantitative PCR and the 2(-delta delta C(T)) method. Methods 25, 402-408. 18. Madison D. V., Nicoll R. A. (1984) Control of the repetitive discharge of rat CAl pyramidal neurons in vitro. J. Physiol. 354, 319-31. 19. Maiava! L., Liu F., Vernallis A. B., Aubin J. E. (200S) GP130/OSMR is the only LIF/IL-6 family receptor complex to promote osteoblast differentiation of calvaria progenitors. J. Cell Physiol. 204(2), 585-93. 20. Mimica N., Presecki P. (2009) Side effects of approved antidementives. Psychiatr. Danub. 21(1), 108-13. 21. Mosmann T. (1983) Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays. J. lmmunol. Methods 65(1-2), 55-63. 22. Nicola N. A., Cross B., Simpson R. J. (1993) The disulfidebond arrangement of leukemia inhibitory factor: homology to oncostatin Mand structural implications. Biochem. Biophys. Res. Commun. 190(1), 20-6.

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62 IL-6-TYPE CYTOKINES IN NEUROMODULATION AND NEUROPROTECTION

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63 CHAPTER 2

SUPPLEMENTARY FIGURES

Glut. Glut. Glut. Glut. (20µM) (SOµM) (l00�1M) (200�1M) 120

100

0 80 0 � 60 a

·;;: 40

a:; u 20

0 MK801 - + - + - + -+ -+ Glutamate + + + + + + + +

Supplementary Figure 1: Glutamate-induced excitotoxicity in neurons is mediated through NMDA receptors. Primary cortical neurons from wild-type (C57BL/6J) mice embryo (~E15) were treated for i hour with differentconcentrations of glutamate (20, 50, 100 and 200 µM) after 5 days in culture. Where indicated, cultures were pre-incubated with the specific NMDA receptor antagonist, MK 801 (30 µM; 30 min prior to glutamate). Neuronal survival was assessed 24 hours following the glutamate insult by a colorimetric MTT-assay. The optical densities were measured at 570 nm with a 630 nm and blank correction. The bars represent Mean ±. S.E.M of 4 independent experiments; HH p value < 0.001 (compared to the control); ** p value < 0.001 (compared to the respective glutamate treated condition).

64 IL-6-TYPE CYTOKINES IN NEUROMODULATION AND NEUROPROTECTION

·l· ? �? OJ OJ �? ? ? A � � OJ � (: (: � �OJ B r::,": -..::...._(;;' -..::.� � r::,": -..::...._(;;' -..::.� "'� � � 120 120 o 6' 6' �� � � 100 I 100 II e ** gC: E 80 80 u u .....0 0 0 60 �0 60 � i6 > > ·s: 40 -� 40 � C] � Qi Qi u 20 u 20

0 0 24h - 1h 2h 4h 8h 12h 2◄h Glutamate - + + + + + + + OSM (1 ng/ml) - I I Glutamate - - + + + + + + +

C 120 Supplementary Figure 2: LIF and OSM protect cortical neurons from glutamate-induced excitotoxicity. 100 Primary cortical neurons from wild-type (C57BL/6J) mice g embryo (-E15) were maintained in culture for 5 days and 80 treated with (A) OSM or LIF (0.1, 1, or 10 ng/ml) for 0 24 hours. Subsequently, the cultures were challenged with glutamate (SO µM for 1 hour) and neuronal survival � 60 i6 CJ assessed after 24 hours by a colorimetric MTT-assay (n�S). (B) Effect of 1 ng/ml OSM treatment (n=4) and ·s: 40 = (C) effect ofl0 ng/ml LIF treatment (n 3) over different � time period (1, 2, 4, 8, 12 and 24 hours) on neuronal Qi 20 u survival against glutamate (SO µM forl hour). The optical densities were measured at 570 nm with a 630 nm and 0 blank correction. Data normalized to percent of control LIF (10 ng/ml) - 2◄h - 1h 2h 4h 8h 12h 24h and presented as Mean :t. S.E.M; ** p value < 0.001, * p - I Glutamate - + + + + + + + value < 0.05.

65 CHAPTER 2

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Supplementary Figure 3: LIF and OSM reduce caspase-3 activation and protect cultured cortical neurons against glutamate toxicity. Primary cortical neurons from C57BL/6J (wild-type) mouse embryos (-E15) were cultured for 5 days and were incubated without or with LIF (10 ng/ml) or OSM (1 ng/ml) for 24 hours, followed by glutamate challenge (50 µM for 1 hour). (A) The cells were collected 4 hours after glutamate treatment and the protein lysates analyzed by Western blot. Activation of caspase-3 by cleavage was monitored using an anti­ cleaved caspase-3 antibody, that determines cleaved ( -17 kDa) form of the protein. Total cellular j3-actin served as loading control. (B) The cells were fixed in 4% paraformaldehyde, washed and co-stained with Hoechst 33342 (in purple, showing cell nuclei), anti-MAP2 antibody (in green, showing morphology of neurons) and propidium iodide (in red, showing cell death). White arrows indicate nuclei of dead cells in each conditions. Scale bars correspond to 50 µm.

66 IL-6-TYPE CYTOKINES IN NEUROMODULATION AND NEUROPROTECTION

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Supplementary Figure 4: Calcium transients in response to glutamate in cultured cortical neurons.

Embryonic ( ~E15) cortical neurons from wild-type mice were maintained in culture for 5 days. The cells were loaded with 5 µM Fura-2(AM) (Teflabs.com) together with 0.4% pluronic© F-127 (Molecular Probes) in 1 ml standard ° loading buffer for 30 - 45 min at 37 C. Digital images were taken at an emission wavelength of 510 nm using paired exposure at 340 nm and 380 nm excitation wavelengths at a frequency of 1 Hz. Changes in intracellular calcium levels were expressed as the ratio of the 340 and 380 nm excitation wavelengths (ti340/380) in time (seconds). During recording experiments, compounds were administered directly to the loading buffer using a pipette. Buffer was applied as pipetting control prior to glutamate stimulation. Each curve on the graph corresponds to calcium influx into separate neurons from (A) untreated controls, (B) pre-treated with OSM (1 ng/ml for 24 hours), (C) pre-treated with CPA (100 nM for 30 min) and (D) cells pre-treated with OSM (1 ng/ml for 24 hours) followed by CPA (100 nM for 30 min). Note that pre-treatment with OSM or CPA alone had no obvious effect on the rate and/or intensity of calcium influx induced by glutamate (B, C). On the other hand, the neurons pre­ treated with both OSM and CPA showed delayed influx of calcium with a decreased rate (D).

67 CHAPTER2

- OSM 11 ng/ml] LIF (10 ng/ml] 350 c: :::::::- 300 o e Supplementary Figure 5: OSM, but not -�w t:o 250 ,_ u LIF, enhances A R expression in wild-type Cl.� 1 (C57BL/6J) cortical neurons in vitro. ri) � 200

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ABSTRACT Neuroprotection is one of the prominent properties of interleukin (IL)-6-type cytokines, but the underlying intracellular signaling pathway(s) are not completely understood. We here investigated the downstream signaling mechanism(s) involved in OSM- and LIF-induced neuroprotection against glutamate-toxicity, using selective inhibitors of JAK/STAT3 (AG 490 and JSl-124), ERKl/2 (U 0126) and Pl3K/Akt (LY 294002) in cultured mouse embryonic cortical neurons. Western blot and immunocytochemistry experiments show that treatment of cultured cortical neurons with OSM (1 ng/mL) or LIF (1 0 ng/mL) for 1 hour induced phosphorylation at Tyr705, Thr202/Tyr204 and Ser473 residues of STAT3, ERKl/2 and Akt proteins, respectively. Pre-treatment for 2 hours with U 0126 (5 µM) and LY 294002 (25 µM), but not AG 490 (50 µM) or JSl-124 (5 µM), abolished LIF- and OSM-induced neuroprotection against glutamate. We have shown previously that OSM mediates neuroprotection against glutamate by upregulation of neuronal adenosine A, receptors (A,Rs). Here we show that, OSM-induced A,R mRNA expression in cortical and hippocampal neurons was abolished by a selective inhibitor of nuclear transcription factor (NF)-KB (BAY 11-7082). Furthermore, both OSM- and LIF-induced neuroprotection were abolished by pre-treatment with BAY 11-7082 (5 µM). In conclusion, our findings suggest that neuroprotection against glutamate mediated by LIF and OSM is independent of STAT3 signaling, but requires activation of Akt, ERKl/2 and NF-KB pathways.

72 NEU ROPROTECTIVE SIGNALING PATHWAYS OF OSM AND LIF

INTRODUCTION

Leukemia inhibitory factor (LIF) and oncostatin M (OSM) belong to the fa mily of interleukin (IL)-6 type cytokines that fu rther includes IL-6, IL-11, ciliary neurotrophic factor (CNTF), cardiotrophin 1 (CT-1) and novel neurotrophin 1 (NNT-1) (Heinrich et al, 2003). In the central nervous system (CNS), many members of this cytokine fa mily as well as their receptors are II constitutively expressed already at the developing embryonic states, albeit in low levels. Elevated expression of these cytokines in the CNS have been reported following injury, infection (bacterial and viral), ischemia, epilepsy, seizure, multiple sclerosis (MS), amyotropic lateral sclerosis (ALS), Alzheimer's and Pa rkinson's disease (Chiaretti et al, 2008; Jankowsky and Patterson, 1999; Li et al, 2011; Loddick et al, 1998; Soilu-Hanninen et al, 2010). The relevance of these elevated expression levels remains controversial since these cytokines exhibit both pro- and anti-inflammatory properties (Baker et al, 2001; Suzuki et al, 2009). Nevertheless, it is clear from the studies on knock-out animals, that IL-6-type cytokines play an important role in the normal development ofthe CNS by regulating proliferation and/or maintenance of stem cells, neurogenesis, cell survival etc. (Bauer et al, 2007; Cohen and Fields, 2008; DeChiara et al, 1995; Hatta et al, 2002; Holtmann et al, 2005; Horton et al, 1996; Lee et al, 2008; Li et al, 1995; Morikawa et al, 2004; Oberle et al, 2006; Turnley and Bartlett, 2000). In addition, several studies demonstrate that these cytokines possess neuroprotective properties against various insults and injury of the CNS (Gurfein et al, 2009; Holtmann et al, 2005; Ohtaki et al, 2006; Suzuki et al, 2009; Weiss et al, 2006; Wen et al, 2005). We have previously shown that IL-6, OSM and LIF protect mouse cortical and hippocampal neurons against glutamate-induced excitotoxicity (Biber et al, 2008; Moidunny et al, 2010). Intriguingly, we provided evidence that the neuroprotective properties of IL-6 and OSM, but not LIF, are mediated by enhanced neuronal expression of neuromodulatory adenosine A receptors (A Rs) (Moidunny et al, 2010), 1 1 which might suggest that OSM and LIF activate different intracellular signaling pathways in these cells. It has been shown recently that IL-6, which uses the gpl30/gpl30 homodimer in the receptor complex, requires JAK/STAT3, Pl3K/Akt and MAPK pathways to induce protection of primary rat cerebellar granule neurons against excitotoxicity (Liu et al, 2011; Wang et al, 2009). In mice, OSM and LIF act on target cells through a receptor complex consisting of a ligand recognition subunit (OSMR-� and LIFR, respectively) and a signal transducing subunit (gpl30) (Lindberg et al, 1998; Mosley et al, 1996). The activity of this receptor complex is also known to activate the three signaling pathways JAK/STAT3, PI3K/Akt and MAPK pathways (Heinrich et al, 1998; Hintzen et al, 2008; Kamimura et al, 2003; Thoma et al, 1994; Wang et al, 2000). Which of these pathways is activated in neurons upon OSM or LIF stimulation, however, is not yet known. Since a detailed understanding of the signaling pathway(s) involved in neuroprotection induced by IL-6-type cytokines is important for their putative use in treatment of neurodegenerative disorders we here analyzed the intracellular signaling pathway that is mediating the neuroprotective properties of LIF and OSM.

73 CHAPTER 3

MATERIALS AND METHODS

Chemicals and Reagents Neurobasal media, HBSS, PBS, sodium pyruvate, L-glutamine, penicillin-streptomycin, HEPES, glutaMAX-1 and 827 supplement were from Gibco (Breda, The Netherlands). Trypsin was obtained from Life technologies (Breda, The Netherlands). All other cell medium components, recombinant mouse OSM and the dye used to stain cell nuclei (Hoechst 33342) were purchased from Sigma-Aldrich (Zwijndrecht, The Netherlands). Recombinant mouse LIF was obtained from Millipore (Amsterdam, The Netherlands). MEKl/2 inhibitor (U 0126) and JAK2/STAT3 inhibitor (Cucurbitacin 1 or JSl-124) were obtained from Tocris bioscience (Bristol, UK). JAK 2/3 inhibitor (Tyrphostin or AG 490), Pl3K inhibitor (LY 294002) and NF-KB inhibitor (BAY 11-7082) were obtained from Calbiochem (Darmstadt, Germany). Reagents used in immunoblotting experiments were purchased from Bio-Rad Laboratories; except polyvinylidene fluoride (PVDF) membranes that were obtained from Millipore (Bedford, MA). Primary antibodies, mouse monoclonal anti-�-actin was obtained from Abeam (Cambridge, UK), mouse monoclonal anti­ MAP2 was obtained from Sigma-Aldrich (Zwijndrecht, The Netherlands). Primary antibodies against total and phosphorylated STAT3, ERKl/2, Akt and NF-KB p65 were obtained from Cell Signaling Technology (Leiden, The Netherlands). The fluorescence conjugated secondary antibodies used for western blot, donkey anti-mouse IR Dye 680 and donkey anti-rabbit IR Dye 800CW were obtained from LI-COR biosciences (Cambridge, UK). The fluorescence conjugated secondary antibodies used for immunocytochemistry, goat anti-rabbit CY3 was obtained from Jackson lmmunoResearch laboraties (Uden, The Netherlands) and donkey anti-mouse Alexa Fluor 488 was obtained from Molecular probes (Breda, The Netherlands).

Animals Wild-type C5 7BL/6J mice were obtained from Harlan (Horst, The Netherlands). All procedures were in accordance with the regulation of the Ethical Committee forthe use of experimental animals of the University of Groningen, The Netherlands (License number DEC 4623A). Animals were housed in standard makrolon cages and maintained on a 12 hour light/dark cycle. They received food and water ad libitum.

Primary neuronal culture

Primary culture of cortical neurons from mouse embryo (-E15) was established as described previously (Moidunny et al, 2010). Briefly, cortices from embryonic brains were dissected in ice-cold HBSS supplemented with 30% glucose. Meninges were removed, and the tissues were treated with trypsin before they were gently dissociated by trituration in neuronal culture media (neurobasal medium supplemented with 2% B27, 1 mM sodium pyruvate, 2 mM L-glutamine and 50 U/mL penicillin-streptomycin). Cell suspension was filtered using cell strainer (70 µm; Falcon, Franklin Lakes, NJ) before centrifugation (800 rpm for 10 min). Cells were then seeded on poly-D-lysine-coated 96 well plates (1 x 105 cells/well) or 6 well plates (1.5 x 106 cells/well) ° and maintained in neuronal culture media in a humidified atmosphere with 5% CO2 at 37C. Media was refreshed the next da y to get rid of debris. The neuronal purity as determined by MAP2-staining was around 98% (data not shown; Moidunny et al, 2010). Cultures were used after 5 days in vitro.

74 NEUROPROTECTIVE SIGNALING PATHWAYS OF OSM AND LIF

lmmunocytochemistry Activation of different intracellular signaling pathways (STAT3, Akt and ERKl/2) in cortical neurons upon treatment with OSM and LIF (1 and 10 ng/ml, respectively for 1 hour) were analyzed by immunocytochemistry. Briefly, neurons cultured on glass coverslips were fixed for 15 min in 4% Paraformaldehyde. After several washes in PBS, the cells were blocked for 45 min with 5% normal goat serum (Vector, Burlingame, CA, USA) in PBS containing 0.1% TritonX II (Sigma). The coverslips were then incubated overnight at 4°C with mouse anti-MAP2 (1:600) in combination with one of the following primary antibodies: rabbit anti-phospho-p44/42 MAPK (Thr202/Tyr204) (1:1000), rabbit anti-phospho STAT3 (Tyr705) (1:25), rabbit anti-phospho-Akt (Ser473) (1:200). The following day, cells were rinsed 3 times with PBS and incubated for 1 hour with the appropriate secondary antibodies: donkey anti-mouse Alexa Fluor 488 (1:400) and goat anti-rabbit CY3 (1:400). The coverslips were then rinsed with PBS, counter stained with Hoechst 33342 (1:1000) and mounted on microscopic slide with Mowiol (Sigma). The fluorescent signals were analyzed by confocal imaging with a Leica SP2 AOBS system (Leica Microsystems, Rijswijk, The Netherlands) . Pictures were deconvoluted using the software Huygens Pro (SVI, Hilversum, The Netherlands). The specificity for the primary antibodies for all signaling molecules were controlled by western blot experiments (see below). Primary antibodies omission served as control for the secondary antibodies used.

RNA isolation and Reverse Transcription-Polymerase Chain Reaction {RT-PCR) Primary cortical neurons were lysed in guanidinium isothiocyanate / mono-thioglycerol (GTC) buffer and total RNA was extracted using a phenol-chloroform/iso-amyl alcohol step, followed by DNAse I treatment. Purified RNA was then transcribed into cDNA as described previously (Moidunny et al, 2010). Quality of the cDNA was examined using the following housekeeping gene glyceraldehyde-3-phosphate dehydrogenase (GAPDH) primer pairs: Fw 5'-CATCCTGCACCACCAACTGCTTAG-3' and Rev 5'-GCCTGCTTCACCACCTTCTTGATG-3'. Potential contamination of RNA samples by genomic DNA was checked for by running reactions without reverse transcriptase, using GAPDH primers for the subsequent PCR amplification.

Real-Time Polymerase Chain Reaction (Q-PCR)

Effect of BAY 11-7082 and OSM treatments on expression of A1 R mRNA in cultured cortical and hippocampal neurons was analyzed by real-time PCR using the iCycler (Bio-Rad, Veenendaal, The Netherlands) and iQ SYBR Green supermix (Bio-Rad). Mouse ribosomal protein L32-A (rpl32A), Mouse hypoxanthine phosphoribosyltransferase 1 (HPRn) and GAPDH primers were used for normalization to housekeeping genes (data normalized to GAPDH are not shown) and they showed no variations in response to the experimental treatment (see table 1 for primer details). The comparative Ct method (amount of target amplicon X in Sample S, normalized to a reference Rand related to a control sample C, calculated by 2-((C/,S-CtR,S)-(C/,C-CtR,C)) was used to determine the relative expression levels (Livak and Schmittgen, 2001).

Western blotting Western blotting oncultured cortical neurons was performed as previously described (Moidunny et al, 2010). Equal amounts of protein (30 µg) were loaded on to 12.5% sodium dodecyl sulfate - polyacrylamide gels and subsequently transferred on to PVDF membranes. The membranes

75 CHAPTER 3

Table 1. Primers used for real-time polymerase chain reaction (qPCR).

Accession Product Gene number Forward primer (5'-3') Backward primer (5'-3') size (bp) GAPDH NM-008084 CATCAAGAAGGTGGTGAAGC ACCACCCTGTTGCTGTAG 209 HPRTI NM-013556 GACTTGCTCGAGATGTCA TGTAATCCAGCAGGTCAG 102 rpl32A NM-172086 GCTGGAGGTGCTGCTGATGT ACTCTGATGGCCAGCTGTGC 114 AR NM-001008533 CCTCTCCGGTACAAGACAGT GGTGTCAGGCCTACCACAAG 92

were blocked using Odyssey blocking buffer(O BB; diluted 1:1 in PBS) for 1 hour and incubated overnight at 4°C (on shaker) with diffe rent combinations of the primary antibodies, diluted in 1:1 OBB and PBS-T (PBS + 0.1% Tween 20): mouse anti-�-actin (1:8000), mouse monoclonal anti­ STAT3 (1 :1000), ra bbit monoclonal anti-phospho STAT3 (Tyr705) (1 :1000), mouse monoclonal anti-p44/42 MAPK (1 :2000), rabbit polyclonal anti-phospho-p44/42 MAPK (Thr202/Tyr204) (1 :1000), mouse monoclonal anti-Akt (Pan) (1:2000), ra bbit monoclonal anti-phospho-Akt (Ser473) (1 :2000), mouse monoclonal anti-NF-KB p65 (1:1000), rabbit monoclonal anti­ phospho-NF-KB p65 (Ser536) (1 :1000). Following primary antibody incubation, the membranes were washed in PBS-T (4 x 5 min) and incubated for 1 hour at room temperature (on gentle shaking in the dark) with appropriate fluorescence conjugated secondary antibodies (diluted in PBS-T): donkey anti-mouse IR Dye 680 (1 :10000) and donkey anti-rabbit IR Dye 800CW (1:10000). Membranes were washed again in PBS-T (4 x 5 min) and fl uorescent bands were detected using LI-COR's Odyssey infrared imaging system.

Induction of excitotoxicity Followi ng the number of days in culture, cortical neurons were treated without or with specific inhibitors of JAK/STAT3 (AG 490, 50 µM; or JSl-124, 5 µM), P13K (LY 294002, 25 µM), MEKl/2 (U0 126, 5 µM) or NF-KB (BAY 11-7082, 5 µM) for 2 hours before incubation with either recombinant mouse OSM (1 ng/ml) or recombinant mouse LIF (1 0 ng/ml) for 24 hours. Subsequently, neurons were subjected to an excitotoxic insult with glutamate (50 µM for 1 hour). Following glutamate treatment, the media was refreshed and neuronal cultures were incubated for 4 and 24 hours before they were assessed for the degree of cytotoxicity, by MTT assay.

Determination of neuronal viability: MTT assay Survival of cultured embryonic cortical neurons was measured 24 hours after glutamate treatment by the colorimetric MTT (3-(4,5-dimethylthiazol-2-yl-) 2,5-diphenyltetrazolium bromide) assay, as described previously (Moidunny et al, 2010; Mosmann, 1983). MTT solution (fi nal concentration: 0.5 mg/m l) was added to cultured neurons and incubated for 4 hours. Following incubation, the cells were lysed and MTT-formazan solubilized in dimethyl sulfoxide (DMSO) on a orbital shaker for 15 min. Optical density measure of each sample was determined using an automated ELISA reader (Varioskan Flash spectral scanning multi mode reader; Thermo scientific, USA) at 570 nm, with a background correction of 630 nm.

76 NEUROPROTECTIVE SIGNALING PATHWAYS OF OSM AND LIF

Statistical data analysis The absolute data values were normalized to control in order to allow multiple comparisons. Statistical analyses were performed by one-way analysis of variance (ANOVA) followed by Bonferroni post-hoc test, using the Statistical Package for the Social Sciences (SPSS, Chicago, IL, USA). In all cases, p values< 0.05 were considered statistically significant. II

RESULTS

OSM and LIF induces activation of STAT3, Akt and ERKl/2 signaling pathways in cultured cortical neurons We firstdetermined the effects of OSM and LIF on different intracellular signaling pathways in mouse embryonic cortical neurons, by immunocytochemistry. As shown in figure1, treatment of cultured neurons for 1 hour with OSM (1 ng/mL) or LIF (10 ng/mL) induced phosphorylation at Tyr705, Ser473 and Thr202/ Tyr204 residues of STAT3, Akt and ERKl/2 proteins, respectively. While phosphorylated Akt and ERKl/2 immunoreactivity was detected throughout the cytoplasm (as it co-localized with the neuronal marker MAP2), phosphorylated STAT3 immunoreactivity was also detected in the cell nuclei (shown with white arrows), indicating nuclear translocation of activated STAT3 (Figure 1). Activation of differentsignaling proteins by OSM and LIF further indicates that the cultured cortical neurons indeed expressed all receptor components required for the respective cytokine binding and signaling.

OSM· and LIF-induced neuroprotection against glutamate requires Akt and ERKl/2 signaling We next determined whether OSM- and/or LIF-induced neuroprotection against glutamate is mediated through the Akt pathway, using a selective inhibitor of the PI3K, LY 294002 (Vlahos et al, 1994). Pre-treatment for 2 hours with LY 294002 (25 µM) reduced both basal as well as OSM­ (1 ng/mL, 1 hour) and LIF- (10 ng/mL, 1 hour) induced levels of phosphorylated Akt proteins in cultured cortical neurons (Figure 2A). Glutamate treatment (50 µM, for 1 hour) reduced survival of cortical neurons by approximately 40%, whereas pre-treatment for 24 hours with OSM (1 ng/mL) or LIF (10 ng/mL) increased their survival against glutamate (Figure 2B), as previously described (Moidunny et al, 2010). Treatment with LY 294002 (25 µM) alone or in combination with OSM or LIF did not affect neuronal survival, when compared to the untreated control (Figure 2B). In addition, LY 294002 treatment did not change the effect of glutamate (50 µM) on neuronal survival. On the other hand, enhanced neuronal survival against glutamate-toxicity induced by both OSM and LIF were abolished by LY 294002 (Figure 2B), suggesting that OSM and LIF requires activation of Pl3K/Akt signaling in order to induce their neuroprotective effects against glutamate. The importance of ERKl/2 on OSM- and LIF-induced neuroprotection was investigated using the selective inhibitor of ERKl/2 activation (U 0126; Favata et al, 1998). As shown in figure 3A, pre-treatment for 2 hours with U 0126 (5 µM) reduced both basal as well as OSM- (1 ng/mL, for1 hour) and LIF- (10 ng/mL, forl hour) induced levels of phosphorylated ERKl/2 proteins in cultured cortical neurons. Treatment with U 0126 (5 µM) alone or in combination with OSM or LIF did not affect neuronal survival, when compared to the untreated control (Figure 3B). In addition, U 0126 treatment did not change the effect of glutamate on neuronal survival (3B), similar to LY 294002.

77 CHAPTER3

Figure 1: OSM and LIF treatment induces activation of STATI, Akt and ERK1/2 signaling pathways in cultured cortical neurons. Primary cortical neurons from wild-type (C57BL/6J) mice embryo (NE15) were maintained in culture for 5 days. Cells were fixed in 4% paraformaldehyde fo llowing treatment without or with OSM (1 ng/ml) or LIF (10 ng/ml) forl hour. Activation of different intracellular signaling pathways were monitored by immunocytochemistry (see methods) using primary antibodies that detected phosphorylation at Tyr705, Ser473 or Thr202/Tyr204 residues of STAT3, Akt and ERKl/2 proteins, respectively (in red), which co-localized with the neuronal marker MAP2 (in green). White arrows indicate translocation of phosphorylated STAT3 proteins into the nucleus, which was confirmed by counter staining with the nuclear marker Hoechst 33342 (not shown). Scale bars correspond to 25 µm.

On the other hand, enhanced neuronal survival against glutamate-toxicity induced by both OSM and LIF was abolished by U 0126, suggesting that OSM and LIF require ERKl/2 signaling in addition to PI3K/Akt, to induce neuroprotection against glutamate.

OSM- and LIF-induced neuroprotection against glutamate is independent of STAT3 signaling We next determined whether OSM- and/or LIF-induced neuroprotection requires STAT3 signaling, in addition to ERKl/2 and Akt pathways. For this purpose we applied two commonly used JAK/STAT3 inhibitors, AG 490 and JSl-124 (Blaskovich et al, 2003; Meydan et al, 1996).

78 NEUROPROTECTIVE SIGNALING PATHWAYS OF OSM AND LIF

A

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Figure 2: OSM- and LIF-induced neuroprotection against glutamate is dependent on Akt activation. (A) Shows treatment of cultured cortical neurons for 1 hour with OSM (1 ng/mL) or LIF (1 0 ng/mL) induces phosphorylation at Ser473 of Akt proteins, when analyzed by Western blot. Pre-treatment for 2 hours with Pl3K inhibitor LY 294002 (25 µM) reduced both basal as well as OSM- and LIF-induced Akt phosphorylation. Total Akt proteins (pan) in the samples were monitored using an appropriate antibody (see methods); !3-actin served as loading control. (B) Shows survival of cultured cortical neurons measured 24 hours following treatment without or with glutamate (Glut; 50 µM, for 1 hour). Where indicated, neurons were pre-treated for 24 hours with OSM (1 ng/mL) or LIF (10 ng/mL) in presence or absence of LY 294002 (25 µM), prior to glutamate insult. Neuronal survival was assessed using a colorimetric MTT-assay. The optical densities were measured at 570 nm with a 630 nm and blank correction. Data normalized to percent of control and presented as Mean ± S.E.M; *p < 0.05, n = 4.

Pre-treatment for2 hours with AG 490 (50 µM) or JSl-124 (5 µM) abolished OSM- and LIF­ induced phosphorylation of STAT3 proteins (Figure 4A). AG 490 treatment (50 µM, for 26 hours) alone did not affect survival of cortical neurons, whereas JSl-124 treatment (5 µM, for 26 hours) reduced neuronal survival, when compared to untreated control (Figure 48). On the other hand, both AG 490 and JSl-124 did not change the effect of glutamate on neuronal survival (Figure 48). Furthermore, neuroprotection induced by OSM and LIF was unaffected by pre-treatment with AG 490 or JSl-124 (Figure 48), strongly suggesting that OSM or LIF induces neuronal survival against glutamate by STAT3 independent mechanisms.

79 CHAPTER 3

A

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Figure 3: OSM- and LIF-induced neuroprotection against glutamate is dependent on ERK1/2 activation. {A) Shows treatment of cultured cortical neurons for 1 hour with OSM (1 ng/ml) or LIF (10 ng/ml) induces phosphorylation at Thr202/Tyr204 of ERKl/2 proteins, when analyzed by Western blot. Pre-treatment for 2 hours with MEKl/2 inhibitor U 0126 (5 µM) reduced both basal as well as OSM- and LIF-induced ERKl/2 phosphorylation. Total ERKl/2 proteins in the samples were monitored using an appropriate antibody (see methods); !3-actin served as loading control. {B) Shows survival of cultured cortical neurons measured 24 hours following treatment without or with glutamate (Glut; 50 µM, for 1 hour). Where indicated, neurons were pre-treated for 24 hours with OSM (1 ng/ml) or LIF (10 ng/ml) in presence or absence of U 0126 (5 µM), prior to glutamate insult. Neuronal survival was assessed using a colorimetric MTT-assay. The optical densities were measured at 570 nm with a 630 nm and blank correction. Data normalized to percent of control and presented as Mean ±.S.E.M; *p < 0.05, n = 4.

Both basal and OSM- and LIF-induced neuronal survival is regulated by the nuclear transcription factor (NF)-KB Since OSM treatment upregulated expression of neuromodulatory A,Rs in cultured cortical neurons (Moidunny et al , 2010) , and since A,Rs have been shown to be regulated by NF-KB

80 NEUROPROTECTIVE SIGNALING PATHWAYS OF OSM AND LIF

(Jhaveri et al, 2007), we next investigated whether a selective inhibitor of NF-KB activation (BAY 11-7082) blocked OSM-induced Al expression in cultured neurons. Treatment with OSM (1 ng/ml, for 24 hours) increased Al mRNA expression in cultured cortical and hippocampal neurons (Figure SA-B). This effectof OSM was abolished when neurons were pre-treated with BAY 11-7082 (5 or 10 µM; added 2 hours prior to OSM stimulation) (Figure SA-B). On the other

hand, the basal A1 R gene expression in neurons was unaffected by BAYll-7082 treatment (Figure II SA-B), when compared to the untreated control.

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Figure 4: OSM- and LIF-induced neuroprotection against glutamate is not dependent on STAT3 activation. (A) Shows treatment of cultured cortical neurons for 1 hour with OSM (1 ng/ml) or LIF (10 ng/ ml) induces phosphorylation at Tyr705 of STAT3 proteins, when analyzed by Western blot. Pre-treatment for 2 hours with either of the two JAK inhibitors (AG 490, 50 µM; or JSl-124, 5 µM) inhibited OSM- and LIF-induced STAT3 phosphorylation. Total STAT3 proteins in the samples were monitored using an appropriate antibody (see methods); �-actin served as loading control. (B) Shows survival of cultured cortical neurons measured 24 hours following treatment without or with glutamate (Glut; 50 µM, for 1 hour). Where indicated, neurons were pre­ treated for 24 hours with OSM (1 ng/ml) or LIF (10 ng/ml) in presence or absence of AG 490 (SO µM) or JSl-124 (5 µM), prior to glutamate insult. Neuronal survival was assessed using a colorimetric MTT-assay. The optical densities were measured at 570 nm with a 630 nm and blank correction. Data normalized"topercent of control and presented as Mean± S.E.M; *p < 0.05, n = 4.

81 CHAPTER3

We further investigated the effect of BAY 11-7082 on OSM- and LIF-induced neuroprotection against glutamate. As shown in Figure 6, treatment with BAY 11-7082 (5 µM, for 26 hours) alone reduced survival of cultured cortical neurons, but did not change the effect of glutamate on neuronal survival. In addition, both OSM- and LIF-induced neuroprotection against glutamate was abolished when neurons were pretreated with BAY 11-7082 (5 µM, added 2 hours prior to OSM and LIF) (Figure 6), indicating that NF-KB is a key transcription factor that regulates survival of cultured cortical neurons, both under basal conditions as well as during OSM- and LIF-induced neuroprotection against glutamate-toxicity.

A

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Figure S: OSM-induced expression of adenosine A receptors (A R) in cultured neurons is dependent on 1 1 NF-KB activation. Primary cortical (A) and hippocampal neurons (B) from wild-type (C57BL/6J) mice embryo (~E15) were maintained in culture for 5 days. Where indicated, specific inhibitor of NF-KB (BAY 11-7082, 5 or 10 µM) was added to the cultures 2 hours prior to treatment without or with OSM (1 ng/ml) for 24 hours. Graphs show relative measure of A1R mRNA expression (normalized to HPRTI or rpl32A) analyzed by real-time PCR. Data normalized to untreated controls and presented as Mean ±.S.E .M; *p < 0.05, n = 3.

82 NEUROPROTECTIVE SIGNALING PATHWAYS OF OSM AND LIF

120

e 100 c 8 80 0 El e:, 60

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Figure 6: OSM- and LIF-induced neuroprotection against glutamate is dependent on NF-KB activation. Graph shows survival of cultured cortical neurons measured 24 hours following treatment without or with glutamate (Glut; 50 µM, for 1 hour). Where indicated, neurons were pre-treated for 24 hours with OSM (1 ng/ml) or LIF (10 ng/ml) in presence or absence of BAY 11-7082 (5 µM), prior to glutamate insult. Neuronal survival was assessed using a colorimetric MTT-assay. The optical densities were measured at 570 nm with a 630 nm and blank correction. Data normalized to percent of control and presented as Mean .±S.E .M; *p < 0.05, n = 4.

DISCUSSION

Neuroprotection is one ofthe prominent properties of IL-6-type cytokines (Gurfeinet al, 2009; Holtmann et al, 2005; Ohta ki et al, 2006; Suzuki et al, 2009; Weiss et al, 2006; Wen et al, 2005) and it is of importance to understand how IL-6-type cytokines exert their neuroprotective fu nction. We have presented evidence that of the IL-6-type cytokine fa mily, IL-6 and OSM require an up-regulation and fu nction of adenosine A1 receptors for neuroprotection (Biber et al, 2008; Moidunny et al, 2010). The neuroprotective fu nction of LIF, however, seems independent of A receptor expression levels suggesting diffe rent mechanisms of IL-6-type 1 cytokines in neuroprotection. Among the members of IL-6 fa mily, OSM and LIF are highly related, both structurally and fu nctionally (Jeffery et al, 1993; Nicola et al, 1993) and indeed several overlapping physiological properties have been reported for the two cytokines (Thoma et al, 1994). However, few reports described different effects of these two cytokines (Maiava I et al, 2005; Ta naka et al, 2003; Weiss et al, 2005). All lL-6-type cytokines act through a receptor complex consisting of at least one common signal transducing subunit, gp130. Activation of the receptor complex by the ligands induces several intracellular signaling pathways, mainly ERKl/2, Pl3K/Akt and STAT3 (Heinrich et al, 1998; 2003). It is yet not known whether LIF and OSM activate different signaling pathways in order to exert their distinct functions. We therefore analyzed which neuronal signaling pathways are activated by LIF and OSM and compared by various selective inhibitors the potential influence of those signaling pathways in neuroprotection. Treatment of embryonic mouse cortical neurons with OSM or LIF induces phosphorylation of STAT3, ERKl/2 and Akt proteins. As shown earlier, pre-incubation for 24 hours with OSM (1

83 CHAPTER 3 ------ng/ml) or LIF (10 ng/ml) leads to enhanced survival of cultured neurons against glutamate­ induced excitotoxicity (Moidunny et al , 2010). We now show that OSM- and LIF-induced neuroprotection is completely abolished by selective inhibitors of PI3K and MEKl/2 suggesting the importance of these pathways in IL-6-type cytokine-dependent neuroprotection. In contrast , inhibition of JAK/STAT3 by two selective inhibitors did not affect the neuroprotective effects of OSM and LIF. Moreover , we provide evidence that inhibition of the NF-KB pathway prevents neuroprotection of LIF and OSM and inhibits OSM-induced up-regulation of Al expression in cultured neurons. Based on the shared signal transducing subunit gpl30 by all IL-6-type cytokines and their general activation of intracellular signaling pathways , being ERKl/2 , PI3K/Akt and STAT3 (Heinrich et al , 1998; 2003) various reports have addressed the role of these pathways in neuroprotection. It has been shown recently that neuroprotection against excitotoxicity induced by IL-6 requires all of the above mentioned signaling pathways (Liu et al , 2011; Wang et al , 2009). Moreover , it has been shown previously that STAT3 activation is necessary for IL-6 to promote neuroprotection against cerebral ischemia (Yamashita et al , 2005) and NMDA­ induced excitotoxicity (Liu et al , 2011). Several other lines of evidence strongly indicate a prominent role of STAT3 in IL-6-dependent protection of neurons against excitotoxicity. In primary hippocampal neurons it has been observed that activation of gpl30 by a fusion protein of IL-6 receptor and IL-6 (hyper IL-6) prevents glutamate-induced excitotoxicity. In these cultures gpl30 activation induced the phosphorylation of STAT3 and p42/p44 MAPK pathways (Sun et al , 2002). A gpl30-dependent activation of STAT and MAPK pathways has also been observed in pheochromocytoma (PC12) cells , a rat neuronal cell line (Marz et al , 2002). IL-6 treatment of organotypic hippocampal slice cultures protected neurons and oligodendrocytes from NMDA-induced excitotoxicity (Pizzi et al , 2004). Although the activation of STATl and STAT3 was associated with the protective effects of IL-6 in this study, it is not clear whether IL-6 induced activation of STATs is instrumental for the observed protection of neurons and oligodendrocytes (Pizzi et al , 2004). Moreover , involvement of STAT3 in IL-6 dependent neuroprotection in a stroke model (MCAO) has been shown (Yamashita et al , 2005). In this study , blocking of IL-6 signaling in vivo by antibody treatment significantly aggravated infarct volume and increased the number of apoptotic , caspase-3 positive neurons. The injection of IL-6 antibodies also prevented activation of STAT3 but did not influence phosphorylation of Akt and ERKl/2. Since apoptotic neurons did not express STAT3, it was concluded that the survival of neurons is correlated with a critical expression level of STAT3 (Yamashita et al , 2005). Taken together these results suggest that expression or activation of STAT3 is instrumental in the neuroprotective effect of IL-6. Less is currently known about the signaling routes that are involved in LIF-dependent neuroprotection. It has been observed that injection of LIF into the brain reduces brain damage in response to ischemic stroke (Suzuki et al , 2005). In this study, the authors observed significant phosphorylation of STAT3 after injectionof LIF and showed that STAT3 was only present in non­ apoptotic cells (Suzuki et al , 2005). No significant effects of LIF injection on Akt and ERKl/2 phosphorylation in the brain have been found. From the above mentioned findingsthe authors concluded STAT3 activation is at least associated with the neuroprotective effects LIF, although a direct involvement was not shown.

84 NEUROPROTECTIVE SIGNALING PATHWAYS OF OSM AND LIF

We show here that both Akt and ERKl/2 signaling in neurons are important for the LIF­ and OSM-induced neuroprotection against excitotoxicity. In line with this, there is evidence suggesting that activation of the Pl3K pathway promotes neuronal survival (Dudek et al, 1997; Franke et al, 1997), and its inhibition severely affects neuronalsurvival (Dudek et al, 1997). Our data add up to the complexity of IL-6-type cytokine signaling in neurons. Despite the conclusive evidence that STAT3 activity is vital for the neuroprotective properties of IL-6, we show that II STAT3 signaling is not crucial for OSM- and LIF-induced neuroprotection and that rather Akt and ERKl/2 are required. Thus from our previously published data (Biber et al, 2008; Moidunny et al, 2010) one might speculate that each of the so far investigated IL-6-type cytokines uses its own specific pathwayto protect neurons. IL-6 leads to STAT3 activation and A, receptor up-regulation, whereas OSM up-regulates A, receptors in a STAT3 independent manner. LIF requires neither STAT3 nor adenosine A, receptors to exert its neuroprotective properties. Further investigation that may involve knockout or knockdown approach, however, should be performed to investigate whether this holds true and what lies behind the specificity of IL-6- type cytokine signaling. The data presented here furthermore suggest potential influence of NF-KB in OSM- and LIF-induced neuroprotection. NF-KB is a dimeric transcription factor that is controlled by I-KBa. Degradation of I-KBa induces activation of NF-KB and its subsequent translocation from the cytoplasm to the nucleus (Gilmore, 2006). NF-KB has been shown to influence neuronal survival and plasticity (Chiarugi, 2002; Gutierrez et al, 2005; Mattson and Meffert, 2006). In line with this, neuroprotection against glutamate induced by cytokines including TNFa and IL-113 has been reported to be dependent on NF-KB signaling (Dolga et al, 2008; Pizzi et al, 2002). Anti-apoptotic function of NF-KB includes regulation of the activity of X chromosome­ linked inhibitor of apoptosis protein (XIAP) gene, and thereby enhancing neuronal survival (Kairisalo et al, 2009). In this study, overexpression of XIAP gene in hippocampal neurons caused NF-KB dependent increase in expression and activity of BDNF and IL-6, two factors that have been widely implicated for neurotrophic effects. Moreoverit is known that neuronal A,Rs are regulated by NF-KB (Nie et al, 1998), and A,R expression is substantially reduced in different brain regions of NF-KB pS0 -/- mice (Jhaveri et al, 2007). Thus, the data presented here, showing that OSM-dependent up-regulation of A? is abolished by an inhibition of the

NF-KB signaling pathway may be in favor of a general importance for NF-KB in A1R regulation. There is however, so far no study that would suggest direct signaling of LIF or OSM receptor to the NF-KB pathway. There is evidence that ERKl/2 and Akt may activate NF-KB (Duronio, 2008; Heck et al, 1999; Lee et al, 1997), whether or not however this also holds true in LIF- and OSM­ induced neuroprotection remains to be established. Taken together, neuronal signaling pathways that are specificallyactivated by OSM and LIF have not been found here. Our data suggest a so far unknown role of IL-6-type cytokines for the activity of NF-KB pathway, at least in neurons.

85 CHAPTER 3

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87 CHAPTER 3

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ABSTRACT Neuroprotective and neurotrophic properties of leukemia inhibitory factor (LIF) have been widely reported. In the central nervous system (CNS), astrocytes are the major source for LIF, expression of which is enhanced following disturbances leading to neuronal damage. It is however not understood, how neuronal damage mediates enhanced LIF expression in astrocytes. Since glutamate induced-excitotoxicity is common in CNS disorders and is associated with excessive production of extracellular adenosine, we investigated whether expression of LIF in mouse cortical astrocytes was mediated through adenosine receptor signaling. We show here that glutamate-stressed cortical neurons induce LIF expression through activation of adenosine receptors in cultured astrocytes. Furthermore, using the adenosine analogue, 5'-N-Ethylcarboxamide (NECA), we demonstrate that adenosine-induced

LIF expression in astrocytes is mediated through the adenosine A28 receptor subtype and requires signaling of protein kinase C (PKC) and mitogen-activated protein kinases (MAPKs: p38 and ERKl/2). Moreover, we demonstrate that LIF expression (both basal and NECA-induced) in astrocytes is regulated by the nuclear transcription factor (NF)-KB. We also show that NECA does not increase LIF secretion when Endoplasmic reticulum to golgi transport is blocked using Brefeldin A, suggesting that LIF secretion occurs by a constitutive release pathway. lmmunocytochemistry experiments show that LIF-containing vesicles co-localize with clathrin and Rabll, but not with pHogrin and Chromogranin A+B, suggesting that LIF might be secreted through recycling endosomes. We further show that pre-treatment with supernatants from NECA-treated astrocytes increased survival of cu ltured cortical neurons against glutamate, which was abolished when the supernatants were pre-treated with an anti-LIF neutralizing antibody. In conclusion, adenosine A28 receptor activation in cortical astrocytes stimulates PKC, MAPKs (p38 and ERKl/2) and NF-KB dependent expression and secretion of LIF that promotes survival of cortical neurons against excitotoxicity.

92 LIF RELEASE FROM ASTROCYTES

INTRODUCTION

Leukemia inhibitory factor (LIF) is a soluble glycoprotein that belongs to the family of interleukin (IL)-6-type cytokines. Other members of this family include IL-6, IL-11, ciliary neurotrophic factor (CNTF), oncostatin M (OSM), cardiotrophin-1 (CT-1) and novel neurotrophin-1 (NNT-1) (Heinrich et al, 2003), which display pronounced trophic as well as protective properties during pathophysiology of the central nervous system (CNS) and are hence referred to as neuropoeitic cytokines or neurokines (Patterson, 1992). Specificfunctions of LIF in the nervous system include a induction of cholinergic differentiation of sympathetic neurons, induction of neuropeptide and choline acetyltransferase (ChAT) gene expression (Patterson, 1994), regulation of polyneuronal innervation of neuromuscular junction (Kwon et al, 1995; Kurek et al, 1997) and regulation of the HPA axis (Akita et al, 1995; Chesnokova et al, 1998). Furthermore, LIF signaling is crucial for development of the nervous system including development of sensory and motor neurons (Murphy et al, 1991; Li et al, 1995) and glial cells (Mayer et al, 1994). Consistently, reduced numbers of astrocytes and oligodendrocytes are found in LIF knock-out mice (Bugga et al, 1998). During inflammation, LIF has been suggested to be both pro- and anti-inflammatory and appears to play a key role in neural injury and regeneration. We and others have previously demonstrated neuroprotective properties of LIF against damages caused by excitotoxicity, light etc. (Ueki et al, 2008; Leibinger et al, 2009; Moidunny et al, 2010). Moreover, promotion of axonal regeneration and oligodendrocyte growth and survival by LIF has led to its potential application in reducing damage associated with central inflammatory demyelinating disease like multiple sclerosis (Barres et al, 1993; Butzkueven et al, 2002; Slaets et al, 2010). In the CNS, astrocytes are considered to be the major source for LIF (Murphy et al, 1995; Ishibashi et al, 2006), and its expression in the brain is significantduring pathological conditions including ischemia (Suzuki et al, 2000; Slevin et al, 2008), multiple sclerosis (Mashayekhi and Salehi, 2011), Alzheimer's and Parkinson's disease (Soilu-Hanninen et al, 2010) and brain injury (Banner et al, 1997). The factors responsible for elevated LIF induction during CNS pathology are largely unknown. One of the candidates identified recently to induce LIF expression in astrocytes is ATP (Yamakuni et al, 2002; Ishibashi et al, 2006), levels of which also rise during conditions like high-frequency neuronal activity, seizure, ischemia and hypoxia (Mitchell et al, 1993; Cunha et al, 1996) . However, extracellular ATP is rapidly hydrolyzed by a cascade of ectonucleotidases resulting in an enhanced level of adenosine (Cunha et al, 1996; Zimmermann, 2000). Correspondingly, excitotoxic conditions such as ischemia, hypoxia, seizure and head injury are known to induce a rapid increase in extracellular adenosine concentrations, up to 100 times to that of the resting concentration (Lynch et al, 1998; van Lubitz, 1999; Berman et al, 2000; Parkinson and Xiong, 2004; Parkinson et al, 2005). There is abundant evidence for immune regulation by adenosine (Hasko et al, 2005) including expression and release of growth factors and cytokines such as nerve growth factor (NGF), Sl00beta, IL-6 and CCL2 in glial cells (Fiebich et al, 1996; Heese et al, 1997; Schwaninger et al, 1997; Ciccarelli et al, 1999; Wittendorp et al, 2004). However, it is not known whether adenosine can induce LIF expression in astrocytes. In the present study, we investigated the potential influence of adenosine receptors activity on LIF release from cultured astrocytes.

93 CHAPTER 4

MATERDA LS AND METHODS

Chemicals and Reagents Neurobasal media, HBSS, PBS, sodium pyruvate, L-glutamine, penicillin-streptomycin, HEPES, glutaMAX-1 and B27 supplement were obtained from Gibco (Breda, The Netherlands). DMEM media and FCS were obtained from PAA laboratories (Colbe, Germany) . Trypsin was obtained from Life technologies (Breda, The Netherlands). L-leucine methyl ester (LME) and the remaining cell medium components were purchased from Sigma-Aldrich (Zwijndrecht, The Netherlands). Recombinant mouse LIF was obtained from Millipore (Amsterdam, The Netherlands). Brefeldin

A (BFA), Caffeine, L-glutamate, adenosine A28 receptor antagonist (MRS 1754), PKA inhibitor (KT 5720), PKC inhibitor (Ro 31-8220), p38 MA PK inhibitor (SB 203580), and adenosine analogue (5'-N-Ethylcarboxamide or NECA) were obtained from Sigma-Aldrich. Non-hydrolysable ATP

(2MeSATP), adenosine A2A receptor antagonist (ZM 241385), adenosine A2A receptor agonist (CGS 21680) and MEKl/2 inhibitor (U 0126) were obtained from Tocris bioscience (Bristol, UK). NF-KB inhibitor (BAY 11-7082) and JNK inhibitor (SP 600125) were obtained from Calbiochem (Darmstadt, Germany). Reagents used in immunoblotting experiments were purchased from Bio-Rad Laboratories; except polyvinylidene fluoride (PVDF) membranes that were obtained from Millipore (Bedford, MA) .

Animals Wild-type C57BL/6J (1-2 days postnatal) mice were obtained from Central laboratory animal facility of Groningen. Wild-type C57BL/6J (14-15 days embryonic) mice were obtained from Harlan (Horst, The Netherlands). All procedures were in accordance with the regulation of the Ethical Committee for the use of experimental animals of the University of Groningen, The Netherlands (License number DEC 4623A and DEC 5913A). Animals were housed in standard makrolon cages and maintained on a 12 hour light/dark cycle. They received food and water ad libitum.

Primary neuronal culture

) Primary culture of cortical neurons from mouse embryo (~E15 was established as described previously (Moidunny et al, 2010) . Briefly, cortices from embryonic brains were dissected in ice-cold HBSS supplemented with 30% glucose. Meninges were removed, and the tissues were treated with trypsin before they were gently dissociated by trituration in neuronal culture media (neurobasal medium supplemented with 2% B27, 1 mM sodium pyruvate, 2 mM L-glutamine and 50 U/mL penicillin-streptomycin). The cell suspension was filtered using cell strainer (70 µm; Falcon, Franklin Lakes, NJ) before centrifugation (800 rpm for 10 min). Cells were then seeded on poly-D-lysine-coated 6 well plates (1 .5 x 106 cells/well) and maintained in neuronal culture ° media in a humidified atmospherewith 5% CO2 at 37 C. The culture medium was refreshed the next day to get rid of debris. The neuronal purity as determined by MAP2-staining was around 98% (data not shown; Moidunny et al, 2010). Cultures were used after 5 days in vitro.

Induction of excitotoxicity Cortical neuron cultures were subjected to an excitotoxic insult with glutamate (50 µM, for 1 hour), after which cultures were refreshed with fresh media and were incubated at 37 °C. Supernatants from neuron cultures (untreated and glutamate-treated) were collected 18 hours after glutamate challenge and applied to the primary astrocyte cultures.

94 LIF RELEASE FROM ASTROCYTES

Primary astrocyte cultures Primary astrocyte cultures were established from cerebral cortices of postnatal (1-2 days) C57BL/6J mice according to a previously described procedure (Matos et al, 2008), modified to reduce microglial contamination (Saura, 2007). Microglial cells were separated from the astrocytic monolayer by 1 hour shake-off at 150 rpm. This procedure was repeated 2 times with an interval of 4 days in vitro between each shake off, followed by an overnight (O/N) shake­ off at 240 rpm to remove oligodend rocyte precursor cells. Purified astrocytes were washed with HBSS buffer containing 1 mM EDTA and fu rther detached using HBSS with 0.1% trypsin. a Cells were reseeded with fresh astrocyte culture medium (DMEM supplemented with 5% FCS, 2 mM L-glutamine, 1 mM sodium pyruvate and 50 U/ml penicillin-streptomycin) in multiwell 4 plates (5 x 10 cells/cm2) and maintained in culture to confluency. To further reduce microglial contamination, confluent astrocyte cultures were treated with 5 mM LME, a lysosomotropic agent (Hamby et al, 2006), for 4-5 hours . Astrocytes were ready for experiments after 1-2 days. Our cell preparations had a high percentage of astrocytes (.!: 95%), which was confirmed by immunostaining against GFAP (astrocyte specific marker) and CDllb (microglial specific marker) (data not shown).

Real-Time Polymerase Chain Reaction (Q-PCR) Total RNA of primary astrocytes was extracted, purifiedand transcribed into cDNA as described previously (Moidunny et al, 2010). Quality of the cDNA was examined using the following housekeeping gene glyceraldehyde-3-phosphate dehydrogenase (GAPDH) primer pairs: Fw 5'-CATCCTGCACCACCAACTGCTTAG-3' and Rev 5'-GCCTGCTTCACCACCTTCTTGATG-3'. Effect of neuronal supernatants and NECA on LIF mRNA expression in cultured astrocytes was analyzed by Q-PCR using the iCycler (Bio-Rad, Veenendaal, The Netherlands) and iQ SYBR Green supermix (Bio-rad). Mouse hypoxanthine phosphoribosyltransferase 1 (HPRTI) and GAPDH primers were used for normalization to housekeeping genes (data normalized to HPRTI are not shown) and they showed no variations in response to the experimental treatments. The primer pairs used for Q-PCR were: LIF (Fw 5'-ATGTGCGCCTAACATGACAG-3' and Rev 5'-TATGCGACCATCCGATACAG-3'); GAPDH (Fw 5'-ATGGCCTTCCGTGTTCCTAC-3' and Rev 5'-GCCTGCTTCACCACCTTCTT-3') and HPRTl (Fw 5'-GACTTGCTCGAGATGTCA-3' and Rev 5'-TGTAATCCAGCAGGTCAG-3'). The comparative Ct method (amount of target amplicon X in Sample S, normalized to a reference R and related to a control sample C, calculated by 2-((C/,S-C R,S)-(C/,C-C R,C)) was used to determine the relative gene expression levels t t (Livak and Schmittgen, 2001).

Western blot Western blotting on cultured cortical astrocytes was performed as previously described (Moidunny et al, 2010). Equal amounts of protein (30 µg) were loaded on to 12.5 or 15% sodium dodecyl sulfate-polyacrylamide gels and subsequently transferred on to PVDF membranes. The membranes were blocked using Odyssey blocking buffer (OBB; LI-COR biosciences, Cambridge, UK; diluted 1:1 in PBS) forl hour and incubated O/N at4°Cwith diffe rent combinations of primary antibodies (diluted in 1:1 OBB and PBS-T (PBS + 0.1% Tween 20)): mouse monoclonal anti-!3-actin (1:8000, Abeam, Cambridge UK), rabbit monoclonal anti-phospho-NF-KB p65 (Ser536) (1 :1000, Cell Signaling Technology, Leiden, The Netherlands), rat monoclonal anti-LIF (MAB449; 1 µg/

95 CHAPTER 4

ml, R&D systems, Oxford, UK). The next day, membranes were washed in PBS-T (4 x 5 min) and incubated for 1 hour at room temperature (RT) with appropriate fluorescence conjugated secondary antibodies (diluted in PBS-T): donkey anti-mouse IR Dye 680 (1:10000, LI-COR), goat anti-rat IR Dye 680 (1:10000, LI-COR) and donkey anti-rabbit IR Dye 800CW (1:10000, LI-COR). Membranes were washed again in PBS-T (4 x 5 min) and the fluorescent bands were detected using LI-COR's Odyssey infrared imaging S'/Stem.

LIF ELISA 1 ml of supernatants from each well of 6 well plates of primary mouse astrocyte cultures was collected and stored at -20°C. 96-well ELISA plates (Costar, The Netherlands) were coated O/N at RT with 100 µI/well of primary antibody goat anti-LI F (AF449; 0.5 µg/ml, R&D systems, Oxford, UK) diluted in 0.01 M PBS (pH 7.4). The following day, the plates were washed 6 times with wash buffer (0.25 M Tris-He! pH 8, 0.15 M NaCl, 0.05% Tween-20) using an automated microplate washer (Titertek-Berthold, Germany) and air dried (this step is repeated after each incubation steps). Plates were subsequently incubated for 1 hour at RT with 200 µI/well of blocking buffer (0.01 M PBS, 2% BSA). Afterblocking, the plates were incubated with supernatants from astrocyte cultures (100 µI/well) for 2 hours at RT. 2 dilutions (1: 2 and 1:4) of each samples, diluted in incubation buffer (0.01 M PBS, 0.2% gelatin, 0.05% Tween-20), were used in triplicates. The plates were then incubated for 1 hour at RT with 100 µI/well of the detection antibody, biotinylated goat anti-LIF (BAF449; 0.05 µg/ml; R&D systems) diluted in incubation buffer, followed by an incubation for 30 min at RT with 100 µI/well of Streptavidin-HRP conjugate (1:8000, Sanquin Reagents, Amsterdam, The Netherlands). The plates were then incubated for 15 to 20 min at RT with 100 µI/well of TMB detection buffer (0.1 M acetate buffer, 0.1 M sodium-acetate, pH adjusted with 1 M citric acid (0.21 gms/ml; dissolve 2 tablets of 3, 3', 5, 5'-tetramethyl benzidine dihydrochloride in 11 ml of TMB buffer and add 2 µI of 30% Hp). Upon stable color formation

• the reactions were stopped by adding 100 µI/well of 1 M H 2SO 4 Absorbance of the samples was measured using a spectrophotometric ELISA plate reader (VersaMax microplate reader with SoftMax Pro software, Molecular Devices, CA, USA) at 450 nm, with a background correction at 575 nm. Recombinant mouse LIF (15 - 2000 pg/ml) was used to plot the standard curve.

MTT assay Survival of cultured embryonic cortical neurons was measured 24 hours after glutamate treatment by the colorimetric MTT (3-(4,5-dimethylthiazol-2-yl-) 2,5-diphenyltetrazolium bromide) assay, as described previously (Mosmann, 1983). MTT solution (0.5 mg/ml final concentration) was added to cultured neurons and incubated for 4 hours afterwhich, cells were lysed and MTT-formazan solubilized in DMSO on an orbital shaker for 15 min. Optical density measure of each sample was determined using an automated ELISA reader (Varioskan Flash spectral scanning multimode reader; Thermo scientific, U.S.A) at 570 nm, with a background correction at 630 nm. lmmunocytochemistry and confocal microscopy Astrocytes cultured on glass coverslips were fixed for 15 min in 4% Paraformaldehyde. After several washes in PBS, the cells were blocked for 45 min with 5% normal goat serum (Vector, Burlingame, CA, USA) in PBS containing 0.1% TritonX (Sigma) . The coverslips were then

96 LIF RELEASE FROM ASTROCYTES

incubated O/N at 4°C with rat anti-LIF primary antibody (5 µg/ml, R&D systems) in combination with one of the following primary antibodies: rabbit anti-Rabll (1:400, Zymed, San Francisco, LA, USA), rabbit anti-chromogranin A+B (1:100, Novus Biologicals, Cambridge, UK), mouse anti-clathrin (1:1000, Abeam), rabbit anti-giantin (1:1000, Covance, Princeton, NJ, USA). The following day, cells were rinsed 3 times with PBS and incubated for 1 hour with the appropriate secondary antibodies: donkey anti-rat CY3 (1:500, Jackson lmmunoResearch laboratories, Uden, The Netherlands), donkey anti-rabbit Alexa Fluor 488 (1:500, Molecular Probes), donkey anti­ mouse Alexa Fluor 488 (1:500, Molecular Probes). The coverslips were then rinsed with PBS and a mounted on microscopic slide with Mowiol (Sigma) and analyzed with a Leica SP2 AOBS system (Leica Microsystems, Rijswijk, The Netherlands). Pictures were deconvoluted using the software Huygens Pro (SVI, Hilversum, The Netherlands). Primary antibodies omission served as control.

Statistical data analysis The absolute data values were normalized to control in order to allow multiple comparisons. Statistical analyses were performed by one-way analysis of variance (ANOVA) followed by Bonferroni post-hoc test, using the Statistical Package for the Social Sciences (SPSS, Chicago, IL, USA). p values < 0.05 were considered statistically significant.

RESULTS

Glutamate-challenged cortical neurons induce LIF expression in cultured astrocytes through adenosine receptor activation We have previously shown that treatment of cultured cortical neurons with glutamate (50 µM, for 1 hour) reduces cell survival by 60%, when compared to untreated controls (Moidunny et al, 2010). In order to investigate whether neuronal death induces LIF synthesis in astrocytes, supernatants from cultured cortical neurons were collected 18 hours afterglutamate treatment (50 µM for 1 hour) and applied to cultured cortical astrocytes. It is shown here that treatment of cultured astrocytes for 2 hours with supernatant from untreated neurons did not change LIF expression (Figure 1). On the other hand, supernatant from glutamate-challenged neurons induced an approximately 3 times higher expression of astrocytic LIF mRNA (Figure 1). The induction of LIF mRNA expression by glutamate-challenged neuronal supernatants was abolished in the presence of the non-specific adenosinereceptor antagonist (caffeine, 50 µM)

(Figure 1) and by cocktail of the specific adenosine A2 receptors antagonists (A2A antagonist:

ZM 241385, 250 nM; A28 antagonist: MRS 1754, 250 nM) (Figure 1), suggesting that enhanced LIF expression in astrocytes induced by glutamate challenged-neuronal supernatants is mediated

through adenosine A2A and/or A28 receptor subtypes.

NECA-induced LIF expression and secretion levels in cultured mouse astrocytes is concentration and time dependent In order to further investigate adenosine receptor mediated LIF expression in astrocytes, a non-selective adenosine receptor agonist (NECA) was used. As shown in Figure 2A, NECA­ induced LIF mRNA expression in cultured astrocytes was concentration- and time-dependent, with maximum induction after 2 hours of incubation with 1 and 10 µM NECA. Subsequently, the effect of NECA (1 µM) on LIF protein expression was analyzed by Western blot. Elevated

97 CHAPTER4

4.0 *

-� 3.0 Ill Q) C.I <(� a:<( 2.0 �Q E� � ::::I.. iij 1.0

0.0 Control NS NS 11NS NS I NS INS (untr.) (glut.) (untr.) (glut.) (untr.) (glut ) Caffeine ZM241385 + MRS1754

Figure 1: Glutamate-stressed cortical neurons induce LIF gene expression in primary cortical astrocytes, by a mechanism dependent on astrocytic adenosine receptor activation. Supernatants of primary mouse cortical neurons (NS) collected 18 hours following treatment without (untr.) or with glutamate (glut.; 50 µM, for 1 hour) were applied (1:1 dilution by volume) to the primary cultured cortical astrocytes for 2 hours. Where indicated, astrocytes were pre-treated with Caffeine (50 µM) or a cocktail of adenosine A2A and A28 receptor antagonists (A2A antagonist: ZM 241385, 250 nM; A28 antagonist: MRS 1754, 250 nM), for 30 min before incubation with neuronal supernatants and were analyzed for LI F mRNA expression (relative to GAPDH) using real-time PCR. Data is normalized to the control and presented as Mean ± SEM of 3 independent experiments. *p < 0.05.

LIF protein expression was detected already after 1 hour of NECA treatment, with a maximum induction after2 to 4 hours (Figure 2B). Consistently, ELISA analysis revealed LIF protein content in supernatants from untreated astrocyte cultures, which increased in time upon treatment with NECA (1 µM) (Figure 2C).

NEC -induced LIF expression and secretion levels is dependent on adenosine A A28 receptor activation

In subsequent experiments, specifican tagonists of adenosine A2A and A28 receptors (ZM 241385 and MRS 1754 respectively) were used to identify the receptor subtype involved in NECA­ induced LIF expression and release in cultured astrocytes. Pre-treatment of astrocytes with ZM 241385 (250 nM, added 30 min before NECA) did not affect NECA-induced LIF mRNA and

protein expression (Figure 3A and 3D). In addition, specificA 2A receptor agonist, CGS 21680 (1 µM) fa iled to induce LIF mRNA or protein expression (Figure 3B and 3D). On the other hand, NECA-induced LIF mRNA expression was abolished by MRS 1754 pre-incubation (1 00 and 250 nM, added 30 min before NECA) (Figure 3C), and this effect of MRS 1754 was also observed on LIF protein expression (Figure 3D). Moreover, ELISA analysis of the supernatants from cultured astrocytes showed complete inhibition of NECA-induced LIF release by MRS 1754, but not ZM 241385 (Figure 3E). Furthermore, CGS 21680 treatment (1 µM for 4 hours) did not have an effect on LIF concentration in the culture supernatants (Figure 3E). Ta ken together, these results provide pharmacological evidence that NECA-induced LIF expression and release from

cultured mouse astrocytes is dependent on the activation of adenosine A28 receptors and not

A2A receptors.

98 LIF RELEASE FROM ASTROCYTES

A

2.5

II

0.0 NECA (time} 0.5hr 1hr 2hrs 4hrs 8hrs 12hrs 24hrs Figure 2: NECA increases LIF expression and secretion levels in primary mouse astrocytes. (A) primary cortical astrocytes were B treated without or with NECA (1, 10 or 20 µM) for 0.5, 1, 2, 4, 8, 12 and 24 hours. Cells were then analyzed for LIF LIF (20 kDa) j ...,__...... _., mRNA expression (relative to GAPDH) P-actin (42 kDa} by real-time PCR. Data is normalized to the control and presented as Mean NECA (time) 1hr 2hrs 4hrs 6hrs Bhrs 12hrs ±SEM of 3 independent experiments. *p < 0.05. {B) shows Western blot analysis of cultured astrocytes treated without or with NECA (1 µM) for 1, 2, 4, 6, 8 and 12 hours to C determine LIF protein levels. !3-actin served as loading control. {C) shows 400 a representation of sandwich ELISA ·-c�....J experiment performed to detect LIF C E o- 300 * * content in supernatants of cultured ���- astrocytes that were treated without cO+-Q)c cu 200 or with NECA (1 µM) for 1, 2, 4, 6, 8 CCU 0 C and 12 hours. Each bar corresponds 0 ai LL Q. to the mean concentration of LIF in - ::J 100 ....J

NECA-induced LIF expression and secretion levels in primary astrocytes is mediated through the C -PLC-PKC and MAPKs, but not through C ·CAMP-PKA pathway q111 5 Adenosine A28 receptors are coupled to two types of G-proteins: Gs and Gq/n (Feoktistov and Biaggioni, 1997). Activation of Gs proteins stimulates cyclic AMP (cAMP) leading to protein kinase A (PKA) activation or EPAC signaling pathways (Fredholm et al, 2001), whereas Gq111 proteins stimulate protein kinase C (PKC) via phospholipase C (PLC). In order to determine which pathway downstream of the A28 receptor is responsible for NECA-induced LIF expression and release, we used specific inhibitorsof PKA and PKC (KT 5720 and Ro 31-8220, respectively) (Feng et al, 2010). Pre-incubation of astrocytes with KT 5720 (250 nM, added 30 min before NECA) did not affect NECA-induced LIF expression (both mRNA and proteins) (Figure 4A and 4B). However, pre-treatment with Ro 31-8220 (250 nM, added 30 min before NECA) reduced

99 CHAPTER 4

A B C 30 25

20 g 20 C ·;;; � :;J 15 1.5 !� i� � � 10 i�E !!: 1 o i d E � � 0 r 5 I 00 - : Control NECA - NECA Control NECA NECA - : I ControlI NECA II- NECA IINECA ZM241385 CGS21680 MRS1754 MRS1754 I I (100nM) (250nM) D E 1500

- LIF (20kDa) 1....., -1 .S: ::J C E -act n (42 kDa) IL...-______,I � i ,g � 1000 Control NECA NECA jg:::-c C MRS1754 g� 8 � u. c. 500 ::J ii'l ':.�------..-..-----.. -..-..-. -.. - ..-...-.. - .. -. -.. �. I LIF (20 kDa) • I ,l -actin (42 kDa) Control NECA- - NECA - NECA Control NECA NECA NECA NECA ZM241385 CGS21680 MRS1754 ZM241385 CGS21680

Figure 3: NECA-induced LIF expression and secretion levels in primary astrocytes is dependent on adenosine A receptor activation. 28 Primary cortical astrocytes were treated with the adenosine analogue,

NECA (l µM) or a selective adenosine A2A receptor agonist (CGS 21680, l µM) for 2 hours (for real-time PCR) or 4 hours (for Western blot and ELISA). Where indicated, astrocytes were pre-treated for 30 min with selective

adenosine A2A receptor antagonist (ZM 247385, 250 nM) or adenosine A28 receptor antagonist (MRS 1754, 700 or 250 nM), prior to NECA stimulation. (A-C) show real-time PCR analyses of LIF gene expression (relative to GAPDH). Data is normalized to the control and presented as Mean ± SEM of 3 independent experiments. *p < 0.05. (D) shows Western blot analyses to detect LIF protein levels. 13-actin served as loading control. (E) shows a representation of sandwich ELISA experiment performed to detect LIF content in supernatants of cultured astrocytes. Each bar corresponds to the mean concentration of LIF in triplicate samples; error bars indicate SEM. Observations were confirmedby repeating the experiments 2 additional times. *p < 0.05.

NECA-induced LIF mRNA (Figure 4A) as well as protein levels (Figure 48). Consistently, Ro 31- 8220, but not KT 5720, significantlyabolished LIF content in the supernatant from NECA-treated

astrocyte cultures (Figure 4C) , indicating that adenosine A28 receptor mediated LIF expression and release from astrocytes requires PKC, but not PKA, activation. Activation of the PKC pathway has been associated with effects mediated through mitogen­ activated protein kinase (MAPK) signaling (Nagamoto-Combs et al, 1999; Fiebich et al, 2005). In order to determine the involvement of MAPKs in NECA-induced LIF expression and release , specific inhibitors of the three MAPK cascades: p38, extracellular signal-regulated kinase (ERK) 1/2 and c-Jun N-terminal kinase (JNK) (SB 203580, U 0126 and SP 600125 respectively), were used. It is shown here that pre-treatment (2 hours before NECA) of cultured astrocytes with SB 203580 (10 µM) and U 0126 (5 µM) significantlyreduced both basal as well as NECA-induced LIF mRNA expression (Suppl. Figure lA) and protein release (Suppl. Figure 18). On the other hand ,

100 LIF RELEASE FROM ASTROCYTES

A 3.0

* ��C.J: 2.0 l,j�

E!!:: 10 !�a,-' >- 0.0 II ControlI NECA NECA NECA I - - Figure 4: NECA-induced LIF expression and secretion levels in primary astrocytes 11KT5720 11Ro31-8220 are blocked by protein kinase (PK) C B inhibitor, but not by PKA inhibitor. Primary cortical astrocytes were treated __....- ____-______-�J LIF (20 kDa) without or with NECA (1 µM) for 2 hours �J .-... (for real-time PCR) or 4 hours (for Western r-· · ______] �-actin (42 kDa) blot and ELISA). Where indicated, astrocytes .__ ....., Control NECA NECA NECA were pre-treated for 30 min with specific inhibitors of PKA (KT 5720, 250 nM) or PKC (Ro KT5720 Ro31-8220 31-8220, 250 nM), prior to NECA stimulation. {A) shows real-time PCR analysis of LIF gene expression (relative to GAPDH). Data is normalized to the control and presented as C Mean ±.SEM of 3 independent experiments. 1500 * *p < 0.05. (B) shows Western blot analysis to

.5c: ::J' E detect LIF protein levels. 13-actin served as ,g � 1000 (C) g:::;- loading control. shows a representation C: C: * of sandwich ELISA experiment performed 8 Q) 500 to detect LIF content in supernatants of gt cultured astrocytes. Each bar corresponds ...J"'!:: g- 0 I to the mean concentration of LIF in Control NECA I NECA I NECA triplicate samples; error bars indicate SEM. I - - I I Observations were confirmed by repeating KT5720 Ro31-8220 the experiments 2 additional times. *p < 0.05.

pre-treatment with SP 600125 (10 µM, added 2 hours before NECA) neither affected basal, nor NECA-induced LIF mRNA expression or protein release (Suppl. Figure lA-B), suggesting that both p38 and ERKl/2, but not JNK, are important for basal as well as NECA-induced LIF expression and release in cultured astrocytes.

Both basal and NECA-induced LIF expression and secretion levels in primary astrocytes is dependent on NF-KB activation IL-6geneexpression in cultured astrocytes is enhanced by NECA (Fiebich et a I, 1996; Schwaninger et al, 1997), similar to our present findings with LIF. Since NF-KB is a key transcription factor that regulates IL-6 gene expression (Libermann and Baltimore, 1990; Spooren et al, 2010), we wondered whether NECA-induced LIF gene expression is similarly regulated by NF-KB. Analysis of the mouse LIF promoter region (using genomatix-Matlnspector) identified multiple consensus binding sequences for NF-KB at the first500 bps upstream to the transcription start site (data not shown). Treatment of cultured astrocytes with NECA (1 µM) for 0.5, 1, 2 and 4 hours, induced activation of NF-KB, detected by phosphorylation of NF-KB p65 subunit by Western blot (Figure SA) . In addition, the specific inhibitor of NF-KB activation, BAYll-7082 (10 µM; added 2 hours before NECA) reduced NECA-induced phosphorylation of NF-KB p65

101 CHAPTER4

A B 2.5 C 0 ':li 2.0 _...... ,,------· � - . -- -- phospho-NF-KB p65 (65 k a) � :co 1;_,-;-:_ • □ Cl.1.5 J < • · l 13-actin (42 kDa) � � I· . ·I E - 1 o Control 0.5 hr 1 hr 2 hrs 4 hrs - NECA g,!::::!.. NECA BAY11-7082 &! 0.5 0.0 Iii ControlI NECA I NECA BAY11-7082

C D

300

LIF (20 kDa)

p-actin (42 k□a)

Control NECA NECA BAY11-7082 11 Control NECA NECA ••BAY11-7082 Figure 5: Both basal and NECA-induced LIF expression and secretion levels in primary astrocytes are dependent on NF-KB activation. (A) shows Western blot analysis of primary cortical astrocytes treated without or with NECA (1 µM; for 0.5, 1, 2 and 4 hours) to detect phosphorylation at Ser536 of NF-KB p65 (RelA) proteins. Where indicated, cells were pre-treated with selective inhibitor of NF-KB (BAY 11-7082, 10 µM) for 2 hours prior to NECA stimulation. �-actin served as loading control. Subsequently, astrocytes were treated without or with NECA (1 µM) for 2 hours (for real-time PCR) and 4 hours (for Western blot and ELISA), in presence or absence of BAY 11-7082 (10 µM, added 2 hours prior to NECA stimulation). (B} shows real-time PCR analysis of LIF gene expression (relative to GAPDH). Data is normalized to the control and presented as Mean .± SEM of 3 independent experiments. *p < 0.05. (C) shows Western blot analysis to detect LIF protein levels. �-actin served as loading control. (D} shows a representation of sandwich ELISA experiment performed to detect LIF content in supernatants of cultured astrocytes. Each bar corresponds to the mean concentration of LI Fin triplicate samples; error bars indicate SEM. Observations were confirmed by repeating the experiments 2 additional times. *p < 0.05.

(Figure SA) and significantlyinhibited basal as well as NECA-induced LIF expression (both mRNA and protein) and release (Figure 58 , SC and SD), strongly indicating that LIF gene expression is regulated by NF-KB.

LIF secretion in primary astrocytes is constitutive and independent of NECA stimulation Supernatants from untreated astrocytes contained basal levels of LIF, suggesting that it could be constitutively released from these cells. Since we observed increased LIF concentrations in astrocyte supernatants after NECA stimulation, we investigated whether or not NECA played a direct role in LIF secretion. Thus , we blocked the early secretory pathway with brefeldin A (BFA), a fungal metabolite that causes Golgi-derived proteins to accumulate in the endoplasmic reticulum (Klausner et al , 1992). Cultured astrocytes were pre-treated for 1 hour with BFA (5 µg/ml) , followed by treatment without or with NECA (1 µM) for 1 or

102 LIF RELEASE FROM ASTROCYTES

II

C 900 *

i: ·- _J-

600 �_gIll� cc:...- �si: Ill 8 � 300 * u..:, Q. * -I- Ul

0 Control NECA ■BFA BFA+NECA Figure 6: LIF secretion in primary astrocytes is constitutive• and independent of NECA stimulation. (A) illustrates LIF immunocytochemistry in cultured cortical astrocyte, where LIF is found in vesicle-like structures through out the cytoplasm. Scale bar corresponds to 10 µm. (B) shows complete co-localization of LIF with Giantin, a marker of Golgi apparatus, when cultured astrocyte were treated with Brefeldin A (BFA, 5 µg/ml for 1 hour). Scale bar corresponds to 10 µm. (C) shows a representation of sandwich ELISA experiment performed to detect LIF content in supernatants of cultured astrocytes that were treated without or with NECA (1 µM) for 4 hours. Where indicated, cells were pre-treated with BFA (5 µg/ml) forl hour prior to NECA stimulation. Each bar corresponds to the mean concentration of LIF in triplicate samples; error bars indicate SEM. Observations were confirmedby repeating the experiments 2 additional times. *p < 0.05.

103 CHAPTER 4

4 hours (for immunocytochemistry and LIF-ELISA, respectively). In BFA-treated cells, all LIF­ immunoreactivity co-localized with the Golgi marker Giantin (Figure 68), compared to control conditions, where punctate LIF stainings could be observed all over the cytoplasm (Figure 6A). LIF levels in the supernatants from BFA treated (5 µg/ml, for 5 hours) astrocyte cultures were significantlylower than that of the control (Figure 6C), implying that LIF is indeed constitutively secreted. Moreover, NECA treatment (1 µM, for 4 hours) did not stimulate LIF secretion in BFA­ treated astrocytes (Figure 6C), further indicating that increased LIF levels in astrocytic culture supernatants after NECA stimulation requires synthesis of new proteins and does not involve a ready-releasable post-Golgi reservoir of LIF.

Figure 7: LIF co-localizes with clathrin and Rab11 but not with large dense-core vesicles markers. lmmunostaining performed in cultured astrocytes revealed that LIF do not co-localize with pHogrin (A) and Chromogranin A+B (B), which are both markers for large dense-core vesicles. However, we observed a massive co-localization of LIF with clathrin (C) and a partial co-localization with Rabll (D). Sclae bars correspond to 10 µm.

104 LIF RELEASE FROM ASTROCYTES

LIF secretion in primary astrocytes is mediated through recycling endosomes We further investigated the type of organelles responsible for LIF release. Several reports have shown that cytokines such as IL-6 and transforming growth factor p (TGFP) are secreted through specialized secretory granules called large dense-core vesicles (LDCV) (Specht et al, 2003; Moller et al, 2006). However, when astrocytes were co-stained for LDCV markers such as Chromogranin A+B or pHogrin and LIF no co-localization was observed (Figure 7A and 7B). In contrast, co-localization between LIF and clathrin was observed (Figure 7C). Clathrin is a marker for endosomal vesicles and is sometimes used as a marker for constitutive release (Deborde a et al, 2008; McPherson, 2010). Furthermore, LIF partially co-localized with Rabll (Figure 7D), which is a marker for recycling endosomes (van IJzendoorn et al, 2003; van IJzendoorn, 2006), suggesting that recycling endosomes rather than LDCV mediate secretion of LIF.

NECA-treated astrocytes induced LIF-mediated protection of cultured cortical neurons against excitotoxicity We have previously shown that recombinant mouse LIF (rmLIF) protein protects mouse cortical neurons against excitotoxicity (Moidunny et al, 2010). In order to understand whether NECA stimulation of astrocytes specifically would induce accumulation of neuroprotective LIF, astrocyte cultures were refreshed with new media shortly before NECA stimulation (1 µM for 4 hours) and the supernatant was collected. As shown in figure 8, pre-treatment (1:1 dilution, for 24 hours) with supernatant from NECA-treated astrocytes significantly reduced the glutamate-induced cell death of cultured cortical neurons. A similar protective effect was observed by pre-treatment with rmLIF (0.1 ng/ml, for 24 hours) (Figure 8). Pre-treatment of

120 * e 100

80 0 � 60 iii

-� 40

20

0 Control Glut Glut Glut AF449 Glut. Glut. AF449 I +Glut.I +Glut. NECA rm LIF Astro. Sup. Astro Sup. (untreated) (NECA-treated)

Figure 8: Effect of untreated and NECA-treated astrocytes supernatants on survival of cultured cortical neurons against excitotoxicity. Primary cortical neurons were treated without or with recombinant mouse LIF (rmLIF, 0.1 ng/ml), NECA (1 µM) or astrocyte supernatants (untreated or treated with 1 µM NECA for 4 hours; diluted 1:1 in neuronal culture medium) for 24 hours. Where indicated, rmLIF and NECA-treated astrocyte supernatants were treated with an anti-LIF neutralizing antibody (AF449; 100 ng/ml for 1.5 hours) before applying them to cultured neurons. Subsequently, the neurons were treated without or with glutamate (Glut.; 50 µM for 1 hour) and cell survival assessed 24 hours after glutamate treatment by a colorimetric MTT-assay. The optical densities were measured at 570 nm, with a 630 nm and blank correction. Data is normalized to percent of control and presented as Mean ± 5.E.M of 4 independent experiments. *p < 0.05, when compared to glutamate-treated condition.

105 CHAPTER4

the neurons (for 24 hours before application of glutamate) with NECA (1 µM) or supernatant from untreated astrocytes did not affect glutamate induced neuronal cell death (Figure 8) . We further investigated whether neuroprotection induced by NECA-treated astrocyte supernatant was mediated by LIF, by incubating the supernatants for 1.5 hours at 37°C with a LIF neutralizing antibody (goat polyclonal anti-LIF, AF449, 100 ng/ml, R&D systems) before applying to the neuronal cultures. The optimum concentration of LIF-neutralizing antibody was standardized by an efficiency test, performed according to manufacturer's recommendations (data not shown) . In addition, the effect of rmLIF protein treated with LIF-neutralizing antibody, on neuronal survival against glutamate, served as a control (Figure 8) . Interestingly, LIF-neutralized supernatant from NECA-treated astrocyte cultures failed to protect neurons against glutamate (Figure 8), suggesting a direct neuroprotective mechanism of the endogenous LIF produced by astrocytes in response to NECA stimulation.

DISCUSSION

We have previously shown that recombinant LIF protects neurons against glutamate-induced excitotoxicity (Moidunny et al, 2010). In this study, we investigated the mechanism by which astrocytes produce and release LIF. Here we show that glutamate-induced neuronal excitotoxicity leads to adenosine receptor-mediated increase in LIF mRNA expression in cultured cortical astrocytes. We demonstrate that the upregulation of LIF mRNA and protein is adenosine A receptor-dependent, and is mediated through G 1-PLC-PKC-NF-KB signaling 28 q11 pathways. We furthermore show that LIF is transiting through the Golgi and is found in recycling endosomes rather than large dense core vesicles. Finally, LIF produced by astrocytes can protect neurons against excitotoxicity. It is known for more than a decade that astrocytes are the major source for LIF in the CNS (Aloisi et al, 1994; Murphy et al, 1995; Dallner et al, 2002; Ishibashi et al, 2006) . However, the factors responsible for the regulation of LIF expression in these cells are still largely unknown. It is well known that stressed neurons release nucleotides such as ATP and adenosine (Lynch et al, 1998; Elliott et al, 2009) . Recently, it was demonstrated that astrocytes increase LIF production and release in response to ATP receptor stimulation (Ishibashi et al, 2006). In this study, the authors demonstrate that neurons can secrete ATP during action potentials, which triggers LIF production in astrocytes. This ATP-dependent upregulation of LIF by astrocytes is responsible for the promotion of oligodendrocyte-mediated myelination around neuronal axons. ATP is also known to be secreted by neurons during stressful conditions such as seizure, ischemia and hypoxia (Mitchell et al, 1993; Cunha et al, 1996) . However, when we blocked adenosine receptors with the non-selective antagonist caffeine or with specific A2/A28 receptor antagonists, the effect of glutamate-stressed neuronal supernatants on LIF expression in astrocytes was absent, suggesting that adenosine but not ATP is responsible for astrocytic LIF production during glutamate-induced neuronal stress. Thus, it might be hypothesized that depending on the CNS status astrocytic LIF expression and secretion is differentially regulated: during normal neuronal activity and development ATP is involved whereas during neuronal insults, adenosine might enhance LIF secretion by astrocytes.

Several studies have demonstrated the involvement of adenosine A28 receptors in the regulation of IL-6 expression in various cell types in vitro (Schwaninger et al, 1997; Rees et al,

106 LIF RELEASE FROM ASTROCYTES

2003; Fiebich et al, 2005; Ryzhov et al, 2008; Feng et al, 2010) as well as in vivo (Vazquez et al,

2008), suggesting that A28 receptors might also be essential in the regulation of other IL-6-type cytokines. Our results indicate that adenosine-dependent LIF regulation is mediated through

the A28 receptor. We furthermore demonstrated that increased LIF expression is dependent on the PKC, but not the PKA pathway. These data are in line with the study of Aloisi and colleagues, which demonstrated that LIF modulation by pro-inflammatory cytokines in human astrocytes was mediated through PKC activation (Aloisi et al, 1994). Moreover, PKC has also been shown to be essential in IL-6 regulation (Rees et al, 2003; Fiebich et al, 2005; Kim et al, 2009; Feng II et al, 2010), revealing a prominent role for PKC in the signaling pathway controlling LIF gene expression.

MAPKs have been reported to be involved in adenosine A28 receptor mediated regulation of IL-6 gene expression in astrocytoma cells (Fiebich et al, 2005). In our experiments, both basal as well as NECA-induced LIF gene expression and release in cultured astrocytes were inhibited by specificinhibitors of p38 and ERKl/2, but not JNK-MAPKs (Suppl. Figure 1). In line with this, it has been shown that LIF expression in Schwann cells is mediated through PKC pathway-induced ERKl/2 activation (Nagamoto-Combs et al, 1999). Furthermore, we show here that adenosine­ dependent LIF expression in astrocytes is regulated through the NF-KB transcription factor. This observation is in line with several studies showing an NF-KB-dependent regulation of IL-6 gene by this transcription factor in several cell types (Libermann and Baltimore, 1990; Matsusaka et al, 1993; Schwaninger et al, 1997; Kim et al, 2009; Spooren et al, 2010). It has been shown that NECA-induced NF-KB activation and the resultant IL-6 gene expression was abolished by inhibitors of MAPK pathways (Kim et al, 2009). In our study, preliminary observations indicate that NECA-induced activation of the NF-KB pathway is reduced by selective inhibitors of p38 and ERKl/2 pathways (data not shown), suggesting that these pathways might play as upstream mediators in NF-KB-dependent LIF expression in astrocytes. Recent evidence indicates that, depending on the cell type, differentsecretory pathways are employed for cytokine release (Stow et al, 2009). For example, T cells use two different release mechanisms: IL-2 and IFN-y are secreted at the immunological synapse whereas CCL3 and TNF-a are secreted multidirectionally, suggesting different secretory pathways (Huse et al, 2006). In neurons or neuron-like cells, secretory granules called LDCVs are the organelles used for the selective secretion of IL-6, TGF-�2 and CCL21 (Specht et al, 2003; Moller et al, 2006; de Jong et al, 2008). The same organelles are also used in immune cells such as mast cells and neutrophils (Stow et al, 2009). Here we show that LIF protein is transported through Golgi but its secretion by astrocytes is not mediated by secretory granules. Instead, LIF co­ localizes with Rabll, a known marker of recycling endosomes (van IJzendoorn et al, 2003; van IJzendoorn, 2006). Moreover, we observed a partial co-localization of LIF with clathrin, which also associates with recycling endosomes where it is implicated in protein sorting (Deborde et al, 2008). Recycling endosomes have now been shown to be responsible for cytokine secretion in several cell types. For example, IFN-y and TNF-a secretion from NK cells require Rabll (Reefman et al, 2010). Recycling endosomes are also responsible for the constitutive secretion of IL-6 and TNF-a in macrophages (Manderson et al, 2007). Further studies will be needed to better understand LIF sorting, traffickingand release by these vesicles. Interestingly, our data indicate that LIF is constitutively released from astrocytes. Indeed constant levels of LIF were present in the supernatants of untreated astrocytes when measured

107 CHAPTER4

by ELISA. Similar data were observed in human astrocyte cultures (Aloisi et al , 1994). Whether this observation is representative of the physiological behaviour of astrocytes in vivo or is due to the culture conditions , remains to be determined. We further show that by blocking the early secretory pathway with BFA , the LIF concentration in the culture supernatant was not increased upon NECA stimulation. The inhibitory effect of BFA indicates that LIF passes through the Golgi prior to its secretion , and thus does not follow non-coventional secretory pathways that by-pass the Golgi and is typically insensitive to BFA , which has recently been reported to be used by other cytokines (Nickel and Rabouille , 2009). Importantly, the inhibitory effect of BFA suggests that NECA-stimulated release of LIF by astrocytes requires de novo LIF synthesis , and does not involve a ready-releasable post-Golgi pool of LIF. It is now clear that one of the major roles of LIF is directed toward cell protection. Indeed , it has been shown that LIF is upregulated in astrocytes and neurons after cerebral ischemia (Suzuki et al , 2000) as well as in astrocytes after cortical brain injury (Banner et al , 1997) , suggesting a role of LIF in neuronal repair or protection. In line with these data , treatment of rat with LIF prevented loss of motoneurons after peripheral nerve injury (Cheema et al , 1994; Tham et al , 1997) and protection of retinal ganglia cells was compromised in LIF KO mice after lens injury (Leibinger et al , 2009). Finally, LIF was shown to limit demyelination in a EAE mouse model (Butzkueven et al , 2002) and has become a prominent therapeutic candidate for MS (Slaets et al , 2010). We have previously shown that LIF can protect cortical as well as hippocampal neurons against glutamate-induced excitotoxicity (Moidunny et al , 201 0) . Here we show that LIF coming from the supernatant of NECA-treated astrocytes has the same protective effect. Indeed , astrocytes produce several other cytokines and neurotrophic factors including IL-6 , nerve growth factor (NGF) , brain derived neurotrophic factor (BDNF) , neurotrophin-3 (NT-3) , S-100� protein and TGF� (Nakagawa and Schwartz , 2003) , that might help neurons to cope with excitotoxic stress. Accordingly , conditioned media from astrocyte cultures protected cortical neurons against glutamate (data not shown). In order to confine the neuroprotective effect of astrocytic factors that are released in response to NECA treatment , we had to refresh astrocyte culture medium prior to NECA treatment and testing supernatant on glutamate-stressed neurons. This together with LIF neutralization indicates that LIF produced by astrocytes after adenosine receptor stimulation is necessary to witness neuronal protection. Our results provide further evidence for a role of adenosine in neuronal protection. Indeed , it has been shown that adenosine can protect neurons during hypoxia (Gribkoff and Bauman , 1992; Fowler , 1993) , ischemia (Lloyd et al , 1993; Latini et al , 1999) and excessive neuronal activity (Arvin et al , 1989; Lloyd et al , 1989). This adenosine protection is often mediated through the A, receptor subtype (Dunwiddie and Masino , 2001; Ramkumar et al , 2001; Pingle et al , 2007) , but here we show that an indirect protection of adenosine through the stimulation of A26 receptor on astrocytes leading to LIF upregulation exists. This A26 receptor activation might be related to an anti-inflammatory process as observed previously by others (Rosi et al , 2003; Yang et al , 2006; Kuno et al , 2007). In conclusion , we demonstrate a protective role of LIF against glutamate neurotoxicity and we provide clear evidence that adenosine is required for an increased production of LIF by astrocytes. These data further confirm a neuroprotective role of adenosine in the brain.

108 LIF RELEASE FROM ASTROCYTES

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56. Nagamoto-Combs K., Vaccariello S. A. and Zigmond R. E. (1999) The levels of leukemia inhibitory factor mRNA in a Schwann cell line are regulated by multiple second messenger pathways. J. Neurochem. 72, 1871-1881. 57. Nakagawa T. and Schwartz J. P. (2003) Expression of neurotrophic factors and cytokines and their receptors on astrocytes in vivo, in Advances in Molecular and Cell Biology: Non-Neuronal Cells of the Nervous System: Function and Dysfunction Vol. 31, pp 561-573. 58. Nickel W. and Rabouille C. (2009) Mechanisms of regulated unconventional protein secretion. Nature reviews 10, 148-155. 59. Parkinson F. E. and Xiong W. (2004) Stimulus- and cell-type-specificrelease of purines in cultured rat forebrain astrocytes and neurons. J. Neurochem. 88, 1305-1312. 60. Parkinson F. E., Xiong W. and Zamzow C. R. (2005) Astrocytes and neurons: different roles in regulating adenosine Neurol. Res. II levels. 27, 153-160. 61 . Patterson P. H. (1992) The emerging neuropoietic cytokine family: first CDF/LIF, CNTF and IL-6; next ONC, MGF, GCSF? Curr. Opin. Neurobio/. 2, 94-97. 62. Patterson P. H. (1994) Leukemia inhibitory factor, a cytokine at the interface between neurobiology and immunology. Proc. Natl. Acad. Sci. U. S. A. 91, 7833-7835. 63. Pingle S. C., Jajoo S., Mukherjea D., Sniderhan L. F., Jhaveri K. A., Marcuzzi A., Rybak L. P., Maggirwar S. B. and

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82. Vazquez J. F., Clement H. W., Sommer 0., Schulz E. and van Ca Iker D. (2008) Local stimulation of the adenosine A28 receptors induces an increased release of IL-6 in mouse striatum: an in vivo microdialysis study. J. Neurochem. 105, 904-909.

111 CHAPTER 4

83. von Lubitz D. K. (1999) Adenosine and cerebral ischemia: therapeutic future or death of a brave concept? Eur. J. Pharmacol. 371, 85-102.

84. Wittendorp M. C., Boddeke H. W. and Biber K. (2004) Adenosine A1 receptor-induced CCL2 synthesis in cultured mouse astrocytes. Glia 46, 410-418. 85. Yamakuni H., Kawaguchi N., Ohtani Y., Nakamura J., Katayama T., Nakagawa T., Minami M. and Satoh M. (2002) ATP induces leukemia inhibitory factor mRNA in cultured rat astrocytes. J. Neuroimmunol. 129, 43-50. 86. Yang D., Zhang Y., Nguyen H. G., Koupenova M., Chauhan A. K., Makitalo M., Jones M. R., St Hilaire C., Seldin D.

C., Toselli P., Lamperti E., Schreiber 8. M., Gavras H., Wagner D. D. and Ravid K. (2006) The A28 adenosine receptor protects against inflammation and excessive vascular adhesion. J. Clin. Invest. 116, 1913-1923. 87. Zimmermann H. (2000) Extracellular metabolism of ATP and other nucleotides. Naunyn Schmiedebergs Arch. Pharmacol. 362, 299-309.

112 LIF RELEASE FROM ASTROCYTES

SUPPLEMENTARY FIGURE

A

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·c:-- ....J 2000 c: E 0 -- .:;�!!!- 15 00 c c (I) r:c * C:u- r:c 1000 * 0 C: * (.J ii; LL a. ::, ....J- Ul 500 0 I I I I ControlI NECA NECA NECA I NECA S8203580 U0126 SP600125

Supplementary Figure 1: Both basal and NECA-induced LIF expression and secretion levels in primary astrocytes is dependent on ERK1/2- and p38-, but not JNK-MAPK activation. Primary cortical astrocytes were treated without or with NECA (1 µM) for 2 hours (for real-time PCR) or 4 hours (for ELISA). Where indicated, astrocytes were pre-treated for 2 hours with specific inhibitors of ERKl/2 (U 0126, S µM), p38 (SB 203580, 10 µM) or JNK (SP 600125, 10 µM) pathways, prior to NECA stimulation. (A) shows real-time PCR analysis of LIF gene expression (relative to GAPDH). Data is normalized to the control and presented as Mean .± SEM of 3 independent experiments. *p < 0.05. (B) shows a representation of sandwich ELISA experiment performed to detect LIF content in supernatants of cultured astrocytes. Each bar corresponds to the mean concentration of LIF in triplicate samples; error bars indicate SEM. Observations were confirmed by repeating the experiments 2 additional times. *p < 0.05

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ABSTRACT

Background: Inflammation of the central nervous system (CNS) and insufficient astrocytic glutamate uptake mediated through Na•-dependent transporters, GLAST/EAATl and GLT-1/ EAAT2, are instrumental in excitotoxic neurodegeneration associated with Alzheimer's disease and amyotrophic lateral sclerosis. There is previous evidence that cytokines such as interleukin (IL)-1� can regulate glutamate transporters. Since neurodegenerative conditions trigger an elevation of the CNS expression levels of the pro-inflammatory interleukin-6-type cytokines, we now investigated whether oncostatin M (OSM), a member of IL-6-type cytokine family, might also regulate astrocytic glutamate uptake thereby influence excitotoxicity. Methods: Primary cultures of purified cortical astrocytes and cortical neurons were used in this study. The effect of OSM treatment on glutamate transporter expression in astrocytes was studied using Q-PCR and Western blot analyses. The transporter activity was assessed by measurement of tritium radiolabelled D-aspartate (D-[3H] aspartate) uptake. Neurotoxicity was measured using a colorimetric MTT assay and immunocytochemistry for propidium iodide labeling. Statistical analyses of the data were performed using one way ANOVA to allow multiple comparisons. Results: OSM treatment time-dependently reduced mRNA expression of the glutamate transporters in cultured mouse astrocytes. Down-regulation of GLAST by OSM was also confirmed at the protein level. Consistently, OSM treatment concentration-dependently inhibited D-[3H] aspartate uptake, which was dependent on JAK/STAT3 activation. Finally, down-regulation of astrocytic glutamate transporter expression and activity by OSM caused increased glutamate-induced cell death of cultured cortical neurons. Conclusions: Taken together, these results demonstrate that STAT3 activation by OSM in cultured astrocytes causes reduction of glutamate transporter expression and activity that result in excitotoxic death of neurons.

116 ONCOSTATIN M INHIBITS ASTROCYTIC GLUTAMATE UPTAKE

INTRODUCTION

Astrocytes are the most abundant cell type of the brain (He and Sun, 2007), where one of their major functions is to support neurons. They ensure optimal conditions by maintaining local ion and pH homeostasis, regulating neurotransmitters and clearing metabolic waste (Nedergaard et al, 2003). In addition, they store glycogen and supply nutrients to neurons (Brown and Ransom, 2007). Dysfunction of astrocytes could therefore have serious consequences for neurons. One major cause of neuronal death is excessive release of glutamate and subsequent activation of neuronal N-methyl D-aspartate (NMDA) receptors (Dong and Benveniste, 2001; Olney et al, 1972). Astrocytes prevent this excitotoxicity in neurons by sequestration of extracellular Bl + glutamate, which is transported mainly by two Na -dependent transporters: Glutamate aspartate transporter (GLAST/EAAn ) and glutamate transporter-1 (GLT-l/EAAT2) (Anderson and Swanson, 2000). Consistently, knock-down of GLAST or GLT-1 results in elevated levels of extracellular glutamate and excitotoxic neurodegeneration (Rothstein et al, 1996). In addition, loss of astrocytic glutamate uptake and metabolism is one of the key factors responsible for neurotoxicity in central nervous system (CNS) disorders like amyotrophic lateral sclerosis (ALS) (Bristol and Rothstein, 1996; Rothstein et al, 1992) and Alzheimer's disease (Mattson and Chan, 2003). In order to identify new mechanisms to protect neurons from excitotoxicity in the CNS disorders, it is important to understand essential factors that regulate astrocytic glutamate transporter expression and activity. Inflammation and the expression of pro-inflammatory cytokines is a hallmark in neurodegenerative diseases in general. Accordingly, increased levels of interleukin (IL)-6- type cytokines have been detected during CNS inflammation associated with injury, bacterial infection, ischemia, epilepsy and multiple sclerosis, ALS, Alzheimer's disease and Parkinson's disease (Chiaretti et al, 2008; Jankowsky and Patterson 1999; Li et al, 2011; Loddick et al, 1998; Soilu-Hanninen et al, 2010). Oncostatin M (OSM), a pleiotropic cytokine belonging to the IL-6 family, has been shown to cause reactive astrogliosis (Sriram et al, 2004). In addition, OSM induces expression of various pro-inflammatory factors in astrocytes including IL-6 (Van Wagoner et al, 2000), prostaglandin (PG) E2 and cyclooxygenase-2 (COX-2; an enzyme involved in PGE2 synthesis) (Repovic et al, 2003). Moreover, OSM regulates expression of differentmatrix metalloproteinases (MMPs) in human astrocytes such as MMP-1 (interstitial collagenase) and MMP-3 (stromelysin 1) (Korzus et al, 1997), and thereby promotes degradation of extracellular matrix components (Stetler-Stevenson et al, 1996). In cultured human astrocytes, OSM stimulates expression of an acute phase protein, al-antichymotrypsin (Kordula et al, 1998), which is a serine protease inhibitor of chymotrypsin (Nelson et al, 1993) and has been shown to be associated with the formation of amyloid-B deposits found in brain tissue of Alzheimer's disease patients (Abraham et al, 1988; Selkoe, 1991). These findings suggest that OSM might directly regulate the inflammatory activity of astrocytes in a variety of brain diseases. Inflammation of the CNS generally causes a reduction of GLAST transporter expression and consequently suppresses glutamate uptake capacity of astrocytes (Chao et al, 1995; Korn et al, 2005). The molecular mechanisms that link inflammation and decreased astrocytic glutamate uptake are, however, not well understood. Limited evidence links the cytokines tumor necrosis factor (TNF)-a and IL-lB differentially with glutamate transporter expression and glutamate uptake capacity in astrocytes (Chao et al, 1995; Hu et al, 2000; Korn et al, 2005; Zou and Crews, 2005). Possible involvement of OSM in the regulation of glutamate uptake in astrocytes, has not

117 CHAPTERS

been addressed yet. Since OSM is important in neuroinflammation, we investigated whether it regulates expression of GLAST and GLT-1, and modulates glutamate uptake in cultured mouse cortical astrocytes. We provide here the firstevidence that OSM downregulates expression and activity of GLAST and GLT-1 in cultured astrocytes. This effect is dependent on STAT3 signaling. Finally, we show that inhibition of glutamate uptake induced by OSM results in excitotoxic death of cortical neurons in vitro.

MATERIALS AND METHODS

Chemicals and Reagents D-[3H] aspartate (specific activity = 10-25Ci/mmol) was obtained from PerkinElmer (MA, USA) and Aquasafe 500 Plus liquid scintillation cocktail from Zinsser Analytic (Frankfurt, Germany). Neurobasal media, Hank's Buffered Salt Solution (HBSS), Phosphate Buffered Saline (PBS), sodium pyruvate, L-glutamine, penicillin-streptomycin, HEPES, glutaMAX-1 and B27 supplement were from Gibco (Breda, The Netherlands). DMEM media and fetal calf serum (FCS) were from PAA Laboratories (Colbe, Germany). Trypsin was obtained from Life Technologies (Breda, The Netherlands). All other cell medium components, recombinant mouse OSM, D-aspartate, L-leucine methyl ester (LME), N-methyl-D-glucamine (NMG) and the dyes used to stain cell nuclei (Hoechst 33342 and propidium iodide) were purchased from Sigma-Aldrich (Zwijndrecht, The Netherlands). DL-threo-j3-Benzyloxyaspartic acid (TBOA) and MEKl/2 inhibitor U 0126 were obtained from Tocris Bioscience (Bristol, UK). Janus kinase (JAK) inhibitor (AG 490) and phosphatidylinositol 3-kinase (PI3K) inhibitor (LY 294002) were obtained from Calbiochem (CA, USA). Reagents used in immunoblotting experiments were purchased from Bio-Rad Laboratories, except polyvinylidene fluoride (PVDF) membranes that were obtained from Millipore (Amsterdam, The Netherlands) . Primary antibodies, mouse monoclonal anti-j3-actin, rabbit polyclonal anti-GLT-1/EAAT2 and rabbit polyclonal anti-GLAST/ EAATl (-C-terminus) were obtained from Abeam (Cambridge, UK), mouse monoclonal anti­ GFAP was obtained from Millipore, mouse monoclonal anti-MAP2 and mouse monoclonal anti-a-tubulin were obtained from Sigma-Aldrich, primary antibodies against total and phosphorylated STAT3, ERKl/2 and Akt were obtained from Cell Signaling Technology (Bioke, Leiden, The Netherlands). The fluorescence conjugated secondary antibodies used for Western blot, donkey anti-mouse IR Dye 680 and donkey anti-rabbit IR Dye 800CW were obtained from LI-COR Biosciences (Cambridge, UK). The fluorescence conjugated secondary antibodies used for immunocytochemistry, goat anti-rabbit linked to CY3 was obtained from Jackson lmmunoResearch laboraties (Uden, The Netherlands) and donkey anti-mouse conjugated with Alexa Fluor 488 was obtained from Molecular Probes (Breda, The Netherlands).

Animals Wild-type C57BL/6J (1-2 days postnatal) mice were obtained from Charles River and Central laboratory animal facility of Groningen. Wild-type C57BL/6J (14-15 days embryonic) mice were obtained from Harlan (Horst, The Netherlands). All procedures were in accordance with the regulation of the Ethical Committee for the use of experimental animals of the University of Groningen, The Netherlands (License number DEC 4623A and DEC 5913A), as well as with the Portugese law on Animal Care and European Union guidelines (86/609/EEC). Animals were

118 ONCOSTATIN M INHIBITS ASTROCYTIC GLUTAMATE UPTA KE

housed in standard makrolon cages and maintained on a 12 hour light/dark cycle. They received food and water ad libitum.

Primary astrocyte cultures Primary astrocyte cultures were established from cerebral cortices of postnatal (1-2 days) C57BL/6J mice according to a previously described procedure (Matos et al , 2008) , modified to reduce microglial contamination (Saura , 2007). Microglial cells were separated from the astrocytic monolayer by 1 hour shake-off at 150 rpm. This procedure was repeated 2 times with an interval of 4 days in vitro between each shake off , followed by an overnight shake-off m at 240 rpm to remove oligodendrocyte precursor cells. Purifiedastrocytes were washed with

, , , HBSS buffer (in mM: 137 NaCl , 5.3 KCI , 0.3 Na2HPO4 0.4 KH/O4 4.2 NaHCO3 5.6 D-Glucose pH 7.4) containing 1 mM EDTA and further detached using 0.1% trypsin (diluted in HBSS). Cells were reseeded with fresh astrocyte culture medium (DMEM supplemented with 5% FCS , 2 mM L-glutamine , 1 mM sodium pyruvate and 50 U/ml penicillin-streptomycin) in multiwell 4 2 plates (5 x 10 cells/cm ) and maintained in culture to confluency. To further reduce microglial contamination , confluent astrocyte cultures were treated with 5 mM LME , a lysosomotropic agent (Hamby et al , 2006) , for 4-5 hours. Astrocytes were ready for experiments after 1-2 days. Our cell preparations had a high percentage of astrocytes (2: 95%) , which was confirmed by immunostaining against GFAP (astrocyte specific marker) and CDllb (microglial specific marker) (Suppl. Figure 1) .

RNA isolation and Reverse Transcription-Polymerase Chain Reaction (RT-PCR) Primary cortical astrocytes were lysed in guanidinium isothiocyanate / mono-thioglycerol (GTC) buffer and total RNA was extracted using a phenol-chloroform/iso-amyl alcohol step , followed by DNAse I treatment. Purified mRNA was then transcribed into cDNA as described previously (Biber et al , 2008). Quality of the cDNA was examined using primers for the housekeeping gene glyceraldehyde-3-phosphate dehydrogenase (GAPDH; see Table 1) . Potential contamination of mRNA samples by genomic DNA was checked by running reactions without reverse transcriptase , using GAPDH primers forthe subsequent PCR amplification.

Real-Time Polymerase Chain Reaction (Q-PCR) Effect of OSM treatment on expression of GLAST , GLT-1 and GFAP mRNA in cultured astrocytes was analyzed by real-time PCR using the iCycler (Bio-Rad , Veenendaal , The Netherlands) and iQ SYBR Green supermix (Bio-Rad). Mouse hypoxanthine phosphoribosyl transferase 1 (HPRn)

Table 1. Primers used forreverse transcription-polymerase chain reaction (RT-PCR). Accession Product Gene number Forward primer (5'-3') Backward primer (5'-3') size (bp) GAPDH NM-008084 CATCCTGCACCACCAACTGCTTAG GCCTGCTTCACCACCTTCTTGATG 346 OSMR NM-011019 ATTCTGGACACGAAGAGGTCAAG TTCCACTGCAAATCACAGCG 151 gpl30 NM-010560 TGGAAGGCACTGCCTCTTTC CTAGAGACGCGACATAGCGGT 151 GLAST NM-148938 CCTTCGTTCTGCTCACGGTC TTCACCTCCCGGTAGCTCAT 91 GLT-1 NM-001077514 GTGCAAGCCTGTTTCCAGC GCCTTGGTGGTATTGGCCT 86

119 CHAPTER S

and GAPDH primers were used for normalization to housekeeping genes (data normalized to GAPDH are not shown) and they showed no variations in response to the experimental treatment (see Table 2 for primer sequences). The comparative Ct method (amount of target amplicon X in Sample S, normalized to a reference R and related to a control sample C, calculated by 2-((C/,S-CtR,S)-(C/,C-Cl,C)) was used to determine the relative expression levels of all tested genes (Livak and Schmittgen, 2001). Linear regression analysis cf the data was performed to understand the effect of OSM treatment over time on the expression of GLAST and GLT-1 mRNA.

Western blotting Western blotting of cultured cortical astrocytic extracts was performed as previously described (Moidunny et al, 2010). Protein calibration controls were performed to ensure that the chosen working quantities were at non-saturating conditions (Suppl. Figure 28-C). Briefly, equal amounts of protein (30 µg) were loaded on to 12.5% sodium dodecyl sulfate-polyacrylamide gels and subsequently transferred on to PVDF membranes. The membranes were blocked using Odyssey blocking buffer (OBB; diluted 1:1 in PBS) for 1 hour and incubated overnight at 4°C with different combinations of primary antibodies (diluted in 1:1 088 and PBS-T (PBS + 0.1% Tween 20)): rabbit anti-GLAST, rabbit anti-GLT-1, mouse anti-�-actin, mouse anti-a.-tubulin, mouse anti-STAT3, mouse anti-p44/42 MAPK, mouse anti-Akt (Pan), rabbit anti-phospho STAT3 (Tyr705), rabbit anti-phospho-p44/42 MA PK (Thr202/Tyr204), rabbit anti-phospho-Akt (Ser473). The next day, membranes were washed in PBS-T (4 x 5 min) and incubated for 1 hour at room temperature (with gentle shaking in the dark) with appropriate fluorescence conjugated secondary antibodies (diluted in PBS-T): donkey anti-mouse IR Dye 680 and donkey anti-rabbit IR Dye 800CW. Membranes were washed again in PBS-T (4 x 5 min) and fluorescent bands were detected using LI-COR's Odyssey infrared imaging system. The densitometry analysis of protein bands was performed using Image J (Abramoff et al, 2004).

Determination of astrocytic glutamate uptake: D-[3H] aspartate uptake assay D-aspartate was used as index of glutamate uptake in this study as it is a substrate for the high-affinity glutamate transporters with slow intracellular metabolization, thus allowing a more precise measure of the activity of glutamate transporters (Bender et al, 1997; Danbolt, 2001; Furness et al, 2008). The analysis of D-[3H] aspartate uptake was evaluated as previously described (Matos et al, 2008). Briefly, cultured astrocytes were incubated with Krebs buffer (in 3 , , , mM: 132 NaCl, 4KCI, 1. 2 Na2HPO4 l.4MgCl2 6glucose, 10 HEPES, 1 CaCl2 pH 7.4) containing D-[ H] aspartate (0.1 µCi/ml) and 50 µM D-aspartate for lO min at 37 °C. Subsequently, the medium was

Table 2. Primers used forre al-time polymerase chain reaction (qPCR).

Accession Product Gene number Forward primer (5'-3') Backward primer (5'-3') size (bp) GAPDH NM-008084 CATCAAGAAGGTGGTGAAGC ACCACCCTGTTGCTGTAG 209 HPRTI NM-013556 GACTTGCTCGAGATGTCA TGTAATCCAGCAGGTCAG 102 GLAST NM-148938 CCTTCGTTCTGCTCACGGTC TTCACCTCCCGGTAGCTCAT 91 GLT-1 NM-001077514 GTGCAAGCCTGTTTCCAGC GCCTTGGTGGTATTGGCCT 86

120 ONCOSTATIN M INHIBITS ASTROCYTIC GLUTAMATE UPTAKE removed and the cultured cells were placed on ice, washed twice with cold NMG buffer (where NaCl is replaced by N-methylglucamine, NMG) to terminate the uptake process. The cells were lysed with 0.5 M NaOH and transferred to a scintillation vial to be mixed with liquid scintillation cocktail. The radioactivity content (disintegrations per minute) was determined using liquid scintillation counting on a TRICARB® 2900TR analyzer. The remaining cell suspension was used to determine the protein content with the BCA method (Pierce Technology) (Smith et al, 1985). The uptake rates for the cells in each well were expressed as the uptake per min per mg of protein. For saturation kinetics assays, the total D-aspartate concentrations ranged from 5 to 200 µM. The kinetic constants (i.e. maximum velocity, Vmax ' and Michaelis-Menten constant, KM) were determined by using nonlinear regression (curve fit) analysis followed by the application • of the one-site binding (hyperbola) equation to the data, using the Graph Pad Prism software (version 5.02, GraphPad Software Inc, La Jolla, CA, USA).

Primary neuronal culture

Primary culture of cortical neurons from mouse embryo ( ~E15) was established as described previously (Moidunny et al, 2010). Briefly, cortices from embryonic brains were dissected in ice-cold HBSS supplemented with 30% glucose. Meninges were removed, and the tissues were treated with trypsin before they were gently dissociated by trituration in neuronal culture media (neurobasal medium supplemented with 2% 827, 1 mM sodium pyruvate, 2 mM L-glutamine and 50 U/mL penicillin-streptomycin). Cell suspension was filtered using cell strainer (70 µm; Falcon, Franklin Lakes, NJ, USA) before centrifugation (800 rpm for 10 min). Cells were then seeded on poly-D-lysine (10 µg/mL)-coated 96 well plates (1 x 105 cells/well) and maintained in ° neuronal culture media in a humidified atmosphere with 5% CO2 at 37 C. Media was refreshed the next day to get rid of debris. The neuronal purity as determined by MAP2-staining was around 98% (data not shown) (Moidunny et al, 2010). Cultures were used after5 days in vitro.

Induction of excitotoxicity Confluent astrocyte cultures were treated without or with OSM (10 ng/mL for 24 hours) in presence or absence of STAT3 inhibitor AG 490 (25 µM, added 2 hours before OSM). The cultures were then washed with warm HBSS and incubated with glutamate (100 µM, diluted in neuronal culture media) at 37°C for 30 min. The supernatants from astrocytes incubated with glutamate (hereinafter referred to as buffered glutamate) were collected and applied (diluted 1:1 with neuronal media) on to 5 days old embryonic cortical neuron cultures and incubated for 1 hour at 37°C. Some neuron cultures were also treated with 50 µM glutamate for 1 hour (positive control for excitotoxicity). Where indicated, neurons were pre-incubated with NMDA receptor antagonist, dizocilpine (MK 801; 30 µM) for 30 min before the glutamate or the buffered glutamate treatments. Following glutamate treatment, the neuronal media was refreshed and cultures incubated for 4 and 24 hours before they were assessed for the degree of cytotoxicity by propidium iodide labeling and MTT assay, respectively.

Determination of neuronal viability 1) MTT assay - Survival of cultured embryonic cortical neurons was measured 24 hours after glutamate treatment by the colorimetric MTT (3-(4,5-dimethylthiazol-2-yl-) 2, 5-diphenyltetrazolium bromide) assay, as described previously (Moidunny et al, 201 0;

121 CHAPTERS

Mosmann, 1983). MTT solution (0.5 mg/ml finalconc entration) was added to cultured neurons and incubated for 4 hours. Following incubation, the cells were lysed and MTT-formazan solubilized in dimethyl sulfoxide (DMSO) on a orbital shaker for 15 min. Optical density measure of each sample was determined using an automated ELISA reader (Varioskan Flash spectral scanning multimode reader; Thermo scientific, USA) at 570 nm, with a background correction of 630 nm.

2) Propidium iodide labeling - Survival of cultured cortical neurons based on propidium iodide (Pl; DNA intercalating dye) labeling was analyzed by immu nocytochemistry, as shown previously (Moidunny et al, 2010). Briefly, neuronal cultures treated without or with glutamate (or buffered glutamate) were incubated with Pl (5 µg/ml; directly added to culture media) for 4 hours and fixed in 4% paraformaldehyde. After a few washes with PBS, cells were blocked using 5% normal goat serum (NGS) diluted in PBS+ (PBS containing 0.1% Triton-Xl00) for 1 hour at room temperature on shaker and subsequently stained with the primary antibody mouse anti-MAP2 (1 :600; diluted in PBS+ with 1% NGS) and incubated overnight at 4°C on shaker. Following primary antibody incubation, cells were washed with PBS (4 x 5 min) and incubated with the secondary antibody donkey anti-mouse Alexa Fluor 488 (1 :400; diluted in PBS+) for 1 hour at room temperature in the dark on an orbital shaker. A counter stain with Hoechst 33342 (1:1000; diluted in PBS) was performed to detect cell nuclei (not shown) and fluorescent signals analyzed by confocal imaging using a Leica SP2 AOBS system (Leica Microsystems, Heidelberg, Germany).

Statistical data analysis The absolute data values were normalized to control in order to allow multiple comparisons. Statistical analyses were performed by one-way analysis of variance (ANOVA) followed by Bonferroni and Dunnett post-hoc tests, using the Statistical Package for the Social Sciences (SPSS, Chicago, IL, USA) and GraphPad Prism software (version 5.02, GraphPad Software Inc, La Jolla, CA, USA). In all cases, p values < 0.05 were considered statistically significant.

RESULTS Primary mouse astrocytes express the receptor subunits for OSM and induce phosphorylation of STAT3, Akt and ERKl/2 upon receptor activation In mice, OSM acts on target cells through a receptor complex consisting of a ligand recognition subunit (OSMR-�) and a signal transducing subunit (gp130) (Lindberg et al, 1998; Mosley et al, 1996). It has been shown that all receptor components required for OSM signaling are expressed in human astrocytes (Kordula et al, 1998). We detected gp130 and OSMR-� mRNA in neonatal (P2) mouse cortical astrocytes (Figure lA). OSM binding to its receptor complex results in phosphorylation of tyrosine residues in the gp130 receptor. This results in the recruitment, phosphorylation and activation of signal transducers and activators of tra nscription (STAT) proteins, mainly STATl, STAT3 and STATS (Heinrich et al, 2003; Hintzen et al, 2008; Wang et al, 2000). In addition, OSM activates other signaling pathways such as phosphatidylinositol 3-kinase (Pl3K/Akt) (Kamimura et al, 2003) and mitogen-activated protein kinase (MAPK) namely, extracellular signal regulated kinase (ERKl/2), p38 and c-jun N-terminal kinases/stress-activated protein kinases (JN K/SAPK) (Thoma et al, 1994). We

1.72 ONCOSTATIN M INHIBITS ASTROCYTIC GLUTAMATE UPTAKE

A B

Control OSM �---_,,,,.,,.,,,--�'-+Phospho-ST AT-3 (~79-86 kDa) gp130 (151 bp) __. Phospho-Akt (60 kDa) ...... - �, ...... __. Phospho-ERK1 (44 kDa) , ..,.._, --- -+Phospho-ERK2 (42 kDa) • OSMR (151 bp) �-----•�I-+13-actin (42 kDa)

GAP DH (346 bp)

Figure 1: OSM treatment induces activation of STAT3, Akt and ERK1/2 proteins in primary mouse cortical astrocytes. (A) Total mRNA purified from wild-type (C57BL/6J) mouse neonatal (P2) brain cortex as well as from cultured cortical astrocytes established from P2 brains were analyzed for expression of OSM receptor components (gp130 and OSMR) by reverse transcriptase PCR; GAPDH primers were used as loading control (see Table 1 for primer details). (B) Wild-type cortical astrocyte cultures were treated without or with OSM (10 ng/ml) for 1 hour and analyzed for phosphorylation of STAT3, Akt and ERKl/2 proteins by Western blot. !3-actin served as the loading control.

next investigated whether OSM induces phosphorylation of the three principal downstream pathways: JAK/STAT3, PI3K/Akt and ERKl/2-MAPK (Van Wagoner et al, 2000). We report that OSM treatment (10 ng/ml for 1 hour) induced the phosphorylation at Tyr705, Ser473 and Thr202/Tyr204 residues of STAT3, Akt and ERKl/2 proteins, respectively, in primary mouse astrocytes culture (Figure 18).

OSM downregulates GLT-1 and GLAST expression in primary mouse astrocytes In order to determine whether OSM modulated astrocytic glutamate uptake, we next investigated its effect on GLAST and GLT-1 gene expression in primary cortical astrocyte cultures. Astrocytes treated with OSM (10 ng/ml) for different time periods (2, 4, 8, 12 and 24 hours) were analyzed for GLAST and GLT-1 mRNA expression by real-time PCR (Figure 2A). 2 Linear regression analysis shows a time-dependent reduction of the expression of GLAST (R = 2 0.525, p < 0.001, n = 3) and GLT-1 (R = 0.415, p = 0.001, n = 3) mRNA in OSM-treated astrocytes compared to the control, with an effect already apparent after4 and 8 hours of incubation, for GLAST and GLT-1, respectively. It is well known that pure astrocyte cultures, due to the absence of soluble neuronal factors, express GLT-1 protein at very low levels (Gegelashvili eta I, 2000; Swanson eta I, 1997). Consistently, in our study GLT-1 protein levels in cultured astrocytes were only faintly detected by Western blot technique (Suppl. Figure 2C), making it difficultto quantify. On the other hand, GLAST proteins

123 CHAPTER 5

A

1.2 ■GLT-1 EJGLAST 1.0 C: o� ·en T""" en 1- [8: 0.8 XI . z: "O8 0.6 �] Q) E o.4 Figure 2: OSM downregulates .!!l C: ioQ) � GLT-1 and GLAST expression in 0:: 0.2 primary cortical astrocytes. (A} Cortical astrocytic cultures were 0.0 treated without or with OSM (1 0 OSM (time) 2h 4h 8h 12h 24h ng/ml) for 2, 4, 8, 12 and 24 hours and analyzed for GLT-1 and GLAST mRNA levels (gene expression normalized to HPRTI) by real-time B PCR. Data is normalized to untreated controls and presented OSM (10 ng/ml) as mean ± SEM; n=3, ** p < 0.01, *** 3h 5h 24h p < 0.001; One way ANOVA using Bonferroni correction. Cortical GLAST (60 kDa) (B) astrocytic cultures were treated ,,...... _ :___. .... I a-tubulin (51 kDa) without or with OSM (10 ng/ml) for 3, S and 24 hours and were analyzed for GLAST proteins by Western blot. Relative optical densiometric values 120 •ffi of GLAST proteins are shown in the QCe 100 ... 0 lower panel. Data is presented as 0. u percentage of each respective ratio I-0 .... 80 ** Cl) between optical density value of ::it60 CJ C: GLAST band intensity and optical Q) ,Q > 1/) 40 density value of the matched a-tubulin (which served as the �Q) � 0. 20 0::: X Q) loading control) band intensity; 0 I n=3, ** p < 0.01, *** p < 0.001; one OSM (time) I3h I5h 24h way ANOVA.

were highly expressed in pure astrocytes culture (Suppl. Figure 28). Treatment with OSM (10 ng/ml) induced a time-dependent reduction of GLAST proteins (Figure 28), consistent with the reduction of GLAST mRNA levels (Figure 2A) . When compared to the untreated control, treatment with OSM during 5 hours led to a significant decrease of GLAST protein density (30.00 ± 7.00%, n=3, p < 0.01) and a treatment with OSM during 24 hours induced an even higher decrease of GLAST protein density (60.11 ± 2.40%, n= 3, p < 0.001) (Figure 28).

OSM impairs D-aspartate uptake in primary mouse astrocytes We next investigated whether the down-regulation of GLAST (mRNA and protein) and GLT-1 (mRNA) expression by OSM had consequences on the ability of astrocytes to clear extracellular glutamate. The rate of D-[3H] aspartate uptake, as determined by the saturation isotherm, was maximal when astrocytes were incubated with 50 µM of D- [3H] aspartate for 10 min at room temperature (Figure 3A) . As shown in figure 38, cultured astrocytes treated with OSM (1 or

124 ONCOSTATIN M INHIBITS ASTROCYTIC GLUTAMATE UPTAKE

A

0.5 • Total • Non-specific (NMG + TBOA) 0.4 • Specific{GLAST + GLT-1)

0.3

0.2 IJI

0.1

0.0 0 25 50 75 100 125 150 175 200 225 250

D-aspartate (µM)

B

120

Cl) 100 a.� :::, e 80 It: "6 (.) 60 !l"' 0- «I'#, 40 5:- c 20 0

OSM 1 ng/ml 10 ng/ml

Figure 3: OSM inhibits 0-aspartate uptake in primary corticalI astrocytes. (A) shows saturation isotherm 3 of Na•-dependent D-[ H] aspartate uptake by astrocytes. Primary astrocytes from wild-type (C57BL/6J) mouse 3 neonatal (P2) brains were incubated with D-[ H] aspartate (0 to 200 µM) dissolved either in a Na•-buffer or in a Na•-free buffer (NMG; N-methyl-D-glucamine chloride) containing TBOA (glutamate transporter inhibitor) for 10 min at room temperature. Each point represents the mean ± SEM of at least 3 separate experiments, measured in triplicate. Kinetic constants were determined by using nonlinear regression (curve flt) analysis followed by ± applying the One-site binding (hyperbola) equation, where V was 0.3 µM 0.08 and KM was 22.4 µM (95% confidence interval: 0.17 - 0.34 µM). (B) shows the effectof treatment (1 and 10 ng/ml; for 24 hours) on 3 D-[ H] aspartate uptake in cortical astrocytes culture. Data is noros·Mmaliz ed to untreated controls and presented as mean ± SEM; * P < 0.05, ** P < 0.01, n = 3 to 17; one way AN OVA followed by Dunnett's multiple comparison test.

10 ng/ml, for 24 hours) displayed a concentration-dependent decrease of D-[3H] aspartate uptake (OSM 1 ng/ml: 12.2 ± 2.2% decrease, n=3, p < 0.05; OSM 10 ng/ml: 24 ± 4% decrease, n=3, p < 0.01).

OSM-induced reduction of D-aspartate uptake in cortical astrocytes is dependent on JAK/STAT3 activation We next investigated the downstream signaling pathway(s) involved in OSM-induced reduction of glutamate transporter activity in cultured astrocytes. As shown in figure 4A, basal

125 CHAPTERS activation of ERKl/2 and Akt proteins, but not STAT3, was observed in untreated astrocytes culture. Activation of all three signaling proteins was induced after treatment with OSM (10 ng/ml for 1 hour) (Figure 4A). We then confirmed that the pre-treatment of astrocytes with selective inhibitors of either Pl3K (LY 294002, 25 µM), MEKl/2 (U 0126, 5 µM) or JAK (AG 490, 25

A

1------·--I - Phospho-Akl (60 kDa) t------1 - To tal Akt (pan) (60 kDa) Phospho-ERK1 (44 kDa) I -=------__ _ --- --@ • 1- - Phospho-ERK2 (42 kDa) 1:::.� -;. ..-= :71 =: i� :�: ���rn��g�l ·-1 - Phospho-STAT-3 (79-86 kDa) To tal STAT-3 (79-86 kDa) I�======:::::::: F5t-- __. aliiiil .._�-,..-. I -

B 120 110

Q) 100 � 90 C. � BO :::J Q)e ..... 70 .....ro C: o �ro 0 - 60 C. 0 50 gj :,!! 40 I !:!- 30 0 20 10 Ii

Figure 4: Inhibition of JAK/STAT3, but not Pl3K/Akt or MEK/ERK1/2 signaling pathways, abolishes 3 OSM-induced reduction of D-[ H]-aspartate uptake in primary cortical astrocytes. (A) shows the effect of 2 hours pre-treatment with selective inhibitors of Pl3K (LY294002, 25 µM), MEKl/2 (U0126, 5 µM) and JAK (AG490, 25 µM) on activation of Akt, ERKl/2 or STAT3 proteins respectively in OSM-treated (10 ng/ml for 1 hour) astrocytes culture. Activation of different signaling proteins were evaluated by Western blot using antibodies that selectively detect the phosphorylated form at Ser473, Thr202/Tyr204 and Tyr705 residues of Akt, Erkl/2 and STAT3 respectively. Total amounts of each protein were detected using appropriate antibodies (see methods); (3-actinserved as loading control. Note that high total protein loaded in lane 8 (see protein bands for (3-actin and total STAT3) accounts for high density band for the respective phosphorylated STAT3. (B) shows the effect of 3 pre-treatment with LY294002 (25 µM), U0126 (5 µM) and AG490 (25 µM) on D-[ H] aspartate uptake in untreated and OSM-treated (10 ng/ml, for24 hours) astrocytic cultures. Data is normalized to untreated controls and ## presented as mean ± SEM; ** p < 0.001 (compared to untreated control); p < 0.001 (compared to OSM); n=8; One-way AN OVA.

126 ONCOSTATIN M INHIBITS ASTROCYTIC GLUTAMATE UPTAKE

µM) abolished OSM-induced phosphorylation of Akt, ERKl/2 and STAT3 proteins, respectively (Figure 4A). Subsequently, these signaling pathway selective inhibitors were added to astrocyte cultures 2 hours prior to OSM treatment (10 ng/ml for 24 hours) and analyzed for their ability to prevent OSM-induced decrease of D-aspartate uptake. Blockade of Akt and ERKl/2 signaling pathways did not affectOSM- induced inhibition of D-[3H] aspartate uptake, whereas blockade of JAK/STAT3 pathway significantly (p < 0.001) prevented the OSM-induced decrease of astrocytic glutamate transporter activity (Figure 4B). Hence, these results indicate that STAT3 activation induced by OSM leads to the impairment of glutamate transporter activity in mouse cortical astrocytes. II

Impaired astrocytic glutamate uptake induced by OSM promotes excitotoxic cell death of cortical neurons in vitro Since OSM down regulated glutamate uptake in astrocytes (Figure 3B), we wondered whether this effect of OSM could be detrimental to neurons. In order to investigate this, cultured astrocytes treated without or with OSM (10 ng/ml for 24 hours) were incubated with glutamate (1 00 µM for30 min). The buffered glutamate (corresponding to the medium oftheseglutamate­ treated astrocytic cultures; see methods) was then applied to neuronal cultures. After 1 hour of incubation at 37°C, the medium was removed and neuronal viability was assessed after 4 and 24 hours by immuocytochemistry and an MTT assay, respectively. As a positive control, we used the direct addition of glutamate (SO µM) to neuronal cultures, which decreased neuronal viability by approximately 4S% (Figure SA). Furthermore, this glutamate-induced neuronal toxicity was NMDA receptor-mediated since the NMDA receptor antagonist MK 801 (30 µM) abolished the toxic effect of glutamate (F = 13.321; p < 0.001) (Figure SA). Finally, we observed that buffered glutamate from OSM-treated astrocytes, but not untreated astrocytes, also reduced the number of metabolically active (i.e. viable) neurons (OSM treated: 67.14% + 14.24% survival, n=4, p < 0.001; and untreated 98.39% + S.39% survival, n=4) (Figure SA); these results were confirmed by direct measure of neurotoxicity using Pl staining (Figure SB). On the other hand, neuronal survival was unaffected when the neuronal cultures were co-treated with MK 801 (Figure SA), confirmingth at the neurotoxicity induced by buffered glutamate from OSM-treated astrocytes was dependent on NMDA receptor activity. Since inhibition of JAK/ STAT3 pathway prevented the OSM-induced down regulation of glutamate transporter activity in cultured astrocytes (Figure 4B), buffered glutamate from astrocytic cultures pre-treated simultaneously with OSM and AG 490, should not cause neuronal death. Indeed, neuronal viability was not affectedwhen subjected to bufferedgl utamate from OSM-treated astrocytes where STAT3 activation was blocked (F = 13.321; p < 0.001) (Figure SA), reinforcing that OSM­ induced loss of astrocytic glutamate transporter activity is mediated by STAT3 signaling.

DISCUSSION We have shown that OSM, a member of IL-6-type cytokine family, reduced the activity of glutamate transporters in cultured mouse cortical astrocytes and thereby promote excitotoxic death of cortical neurons. This effectof OSM is mediated by the down-regulation of the Na+-dependent glutamate transporters, GLAST and GLT-1. We show that treatment of cultured astrocytes with OSM (10 ng/ml) time-dependently reduces expression of GLAST and GLT-1 mRNA (Figure 2A).

127 CHAPTER S

A

140

120

c 100 8

80

1 60 -�::, Ill 40

::, zQ) 20

I C, 0 Astrocyte supernatants

B

. _'--9 ' \

Control

Buffered glutam;ite Buffered glutcJmate (untrecJted astrocytes) (OS M-lrecJled,astrocytes)

Figure 5: Effect of reduced glutamate uptake by OSM-treated astrocytes on the viability of cultured cortical neurons. Primary cortical astrocytes culture from wild-type (C57BL/6J) mouse neonates (P2) were treated with OSM (10 ng/ml, for 24 hours) in absence or presence of JAK/STAT3 inhibitor (AG 490, 25 µM; added 2 hours prior to OSM). Following OSM treatment, the cultures were washed once using warm HBSS and incubated with glutamate (100 µM, diluted in neurobasal media) for 30 min. Thereafter, the resulting astrocytic culture media, named buffered glutamate (see methods), was applied (diluted 1:1 ratio in neuronal culture media) to 6 days old neuron cultures from embryonic ( �E15) mouse cortex. As a positive control, neuronal cultures were treated with 50 µM glutamate, in absence or presence of the NMDA receptor antagonist MK 801 (30 µM, added 30 min prior to glutamate treatment). Following glutamate treatment (for 1 hour), new medium was added to ►

728 ONCOSTATIN M INHIBITS ASTROCYTIC GLUTAMATE UPTAKE

Down-regulation of GLAST by OSM was also confirmedat the protein level (Figure 28); however , the low signals obtained with the GLT-1 antibody in the Western blot analysis (Suppl. Figure 2C) precluded any reliable analysis of the impact of OSM on GLT-1 protein levels. OSM treatment caused a concentration-dependent reduction of D-[3H] aspartate uptake by cultured astrocytes (Figure 38) , which involved the recruitment of the JAK/STAT3 pathway (Figure 48) rather than the PI3K or ERKl/2 pathways. This is in agreement with the previously reported ability of OSM to induce STAT3 phosphorylation in astrocytes (Schaefer et al , 2000) , which we now confirmed (Figure 1B). Several reports indicate that impaired glutamate uptake and metabolism by astrocytes is a major mechanism contributing to the cause of excitotoxic death of neurons in different Ill CNS disorders (Bristol and Rothstein , 1996; Mattson and Chan , 2003; Rothstein et al , 1992). In our study , buffered glutamate (see methods) from untreated astrocytes did not affect survival of cortical neurons , suggesting that the extracellular glutamate is rapidly taken up by the astrocytes. On the other hand , buffered glutamate from OSM-treated astrocytes caused excitotoxicity in cultured cortical neurons (Figure SA-8). It has been shown that gp130-mediated STAT3 activation precedes reactive gliosis in mouse astrocytes (Sri ram et al , 2004) , which might result in inflammatory death of neurons largely mediated by nitric-oxide (Bal-Price and Brown , 2001). In our current in vitro study , supernatants from OSM-treated astrocyte cultures did not affect neuronal survival (Figure SA). In addition , OSM-treatment did not induce the production of nitric-oxide in astrocyte cultures (Suppl. Figure 3) , thus excluding the possibility of indirect oxidative stress-induced neuronal damage. On the other hand , buffered glutamate from OSM­ treated astrocytes did not affect survival of neurons in the absence of neuronal NMDA receptor activity (Figure SA) , suggesting that the increased neurotoxicity directly co-relates with loss of glutamate uptake in OSM-treated astrocytes. The regulation of the extracellular glutamate by astrocytes is determined by the density and the activity of both glutamate transporters as well as glutamine synthetase , the enzyme that converts glutamate to glutamine (Danbolt , 2001). Whether or not OSM treatment regulates glutamine synthetase expression or activity has not been addressed in this study. Nevertheless , our findings provide direct evidence that the activation of the OSM receptor triggering STAT3 signaling in astrocytes has serious consequences for neuronal excitotoxicity. In line with our findings , gp130-mediated STAT3 activation in striatal astrocytes has been reported to be closely associated with neuronal damage in a MPTP model of neurodegeneration in vivo (Sriram et al , 2004). Thus , blockade of STAT3 signati:-,1:J in astrocytes might be beneficial to prevent excitotoxic neuronal death in models pertinent to many brain injuries with an inflammatory profile(Sriram et al , 2004) .

° ► the cortical neuronal cultures and they were incubated at 37 C in a CO2 incubator. (A) 24 hours after glutamate treatment, cell viability was measured using a colorimetric MTT assay. OD measurements were taken at 570nm, with a 630nm and blank correction. a: conditioned media from untreated astrocytes, b: buffered glutamate from untreated astrocytes, c: conditioned media from OSM-treated (10 ng/ml for 24 hours) astrocytes, d: buffered glutamate from OSM-treated astrocytes, e: buffered glutamate from AG 490 (25 µM) and OSM-treated astrocytes, f: buffered glutamate from OSM-treated astrocytes in presence of MK 801 (30 µM). Data are presented as percentage of untreated (control) neurons; Each bar represents the average of 4 independent experiments done in quadruplicates. ** p < 0.001 (compared to glutamate); ## p < 0.001 (compared to buffered glutamate: OSM-treated astrocytes); One way ANOVA using Bonferroni correction. (B) 4 hours after glutamate treatment, cells were fixedin 4% paraformaldehyde, washed and co-stained anti-MAP2 antibody (in green, neuronal marker) and propidium iodide (in red, showing cell death); scale bar corresponds to 25 µm.

129 CHAPTER S

Comparison of the findings reported in the present study with previous studies shows that OSM has a complex profile of action in the control of neurodegeneration. In fact, previous studies demonstrated anti-inflammatory as well as neuroprotective properties of OSM, both in vitro and in vivo. For example, OSM inhibits production of pro-inflammatory mediators such as TNF-a, granulocyte macrophage colony-stimulating factor (GM-CSF) and IL-8 (Richards et al, 1996; Wallace et al, 1999) and has been shown to suppress inflammatory processes associated with the murine experimental allergic encephalomyelitis model of multiple sclerosis (Wallace et al, 1999). In another study, direct activation of neuronal OSM receptors caused down regulation of the NR2C subunit of the NMDA receptor and thereby prevented NMDA-induced toxicity (Weiss et al, 2006). Moreover, we previously provided evidence for a neuroprotective effect of

OSM against glutamate by upregulating neuromodulatory adenosine A1 receptors (Moidunny et al, 2010). Taken together these findingsand our present study, it may be concluded that the pro- and anti-excitotoxic effects of OSM in the CNS are largely determined by the target cell addressed by this cytokine. In the mammalian brain, astrocytes are the predominant players in regulating the glutamate diffusion and spill over from perisynaptic areas, a pre-requisite for the high signal­ to-noise ratio of synaptic communication (for reviews see: Anderson and Swanson, 2000; Danbolt, 2001). Therefore, a compromised astrocytic glutamate uptake function, caused by cytokine overproduction such as OSM, concomitant or a resultant consequence of brain injury, might synergistically exacerbate accumulation of extracellular glutamate at excitotoxic concentrations causing neuronal damage (Aloisi, 2005). In spite of their various suggested roles in astrocytic metabolism, IL-6 family members such as OSM have been scantily explored in their effects on glutamate uptake. There is recent evidence that CNTF, in contrast to our present findings with OSM, enhances both expression and activity of GLT-1 in astrocytes (Escartin et al, 2006), and thereby promotes survival of neurons against excitotoxicity (Beurrier et al, 2010). On the other hand, IL-6 was shown to have no effect on glutamate uptake on cultured murine astrocytes (Okada et al, 2005; Piani et al, 1993) though it suppressed the increase of glutamate uptake induced by PGE2 (Okada et al, 2005). In this study we showed that OSM, through STAT3 activation, impairs the capacity of astrocytes to remove glutamate from extracellular space, which may contribute to excitotoxic neuronal damage. Thus, a closer understanding of the OSM signaling mechanisms, that regulate glutamate transporter level and activity may have important implications for developing novel strategies to limit excitotoxic brain damage in acute and neurodegenerative pathologies.

130 ONCOSTATIN M INHIBITS ASTROCYTIC GLUTAMATE UPTAKE

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Res. 22(6), 721-726. m 7. Beurrier C., Faideau M., Bennouar K. E., Escartin C., Kerkerian-Le Goff L., Bonvento G., Gubellini P. (2010) Ciliary neurotrophic factor protects striatal neurons against excitotoxicity by enhancing glial glutamate uptake. PLoS One 5(1), e8550. 8. Biber K., Pinto-Duarte A., Wittendorp M. C., Doig a A. M., Fernandes C. C., Von Frijtag Drabbe Kunzel J., Keijser J. N., de Vries R., ljzerman A. P., RibeiroJ.A., Eisel U., SebastiaoA. M., Boddeke H. W. (2008) lnterleukin-6upregulates neuronal Neuropsychopharmacology33(9), adenosine A1 receptors: implications for neuromodulation and neuroprotection. 2237-50. 9. Bristol L. A., Rothstein J. D. (1996) Glutamate transporter gene expression in amyotrophic lateral sclerosis motor cortex. Ann. Neural. 39(5), 676-9. 10. Brown A. M., Ransom B. R. (2007) Astrocyte glycogen and brain energy metabolism. G/ia 55(12), 1263-71. 11. Chao C. C., Hu S., Ehrlich L., Peterson P. K. (1995) lnterleukin-1 and tumor necrosis factor-alpha synergistically mediate neurotoxicity: involvement of nitric oxide and of N-methyl-D-aspartate receptors. Brain. Behav. lmmun. 9(4), 355-65. 12. Chiaretti A., Antonelli A., Mastrangelo A., Pezzotti P., Tortorolo L., Tosi F., Genovese 0. (2008) lnterleukin-6 and nerve growth factor upregulation correlates with improved outcome in children with severe traumatic brain injury. J. Neurotrauma 25(3), 225-34. 13. Danbolt N. C. (2001) Glutamate uptake. Prog. Neurobiol. 65(1), 1-105. 14. Dong Y., Benveniste E. N. (2001) Immune function of astrocytes. Glia 36(2), 180-190. 15. Escartin C., Brouillet E., Gubellini P., Trioulier Y., Jacquard C., Smadja C., Knott G. W., Kerkerian-Le Goff L., Deglon N., Hantraye P., Bonvento G. (2006) Ciliary neurotrophic factor activates astrocytes, redistributes their glutamate transporters GLAST and GLT-1 to raft microdomains, and improves glutamate handling in vivo. J. Neurosci. 26(22), 5978-89. 16. Furness D. N., Dehnes Y., Akhtar A. Q., Rossi D. J., Hamann M., Grutle N. J., Gundersen V., Holmseth S., Lehre K. P., Ullensvang K., Wojewodzic M., Zhou Y., Attwell D., Danbolt N. C. (2008) A quantitative assessment of glutamate uptake into hippocampal synaptic terminals and astrocytes: new insights into a neuronal role for excitatory amino acid transporter 2 (EAAT2). Neuroscience 157(1), 80-94. 17. Gegelashvili G., Dehnes Y., Danbolt N. C., Schousboe A. (2000) The high-affinity glutamate transporters GLTI, GLAST, and EAAT4 are regulated via different signalling mechanisms. Neurochem. Int. 37(2-3), 163-70. 18. Hamby M. E., UliaszT. F., Hewett 5. J., HewettJ. A. (2006) Characterization ofan improved procedure for the removal of microglia from confluent monolayers of primary astrocytes. J. Neurosci. Methods 150(1), 128-37. 19. He F., Sun Y. E. (2007) Glial cells more than support cells? Int. J. Biochem. Cell Biol. 39(4), 661-5. 20. Heinrich P. C., Behrmann I., Haan S., Hermanns H. M., Muller-Newen G., Schaper F. (2003) Principles of interleukin (IL)-6-type cytokine signalling and its regulation. Biochem. J. 374(Pt 1), 1-20. 21 . Hintzen C., Evers C., Lippok B. E., Volkmer R., Heinrich P. C., Radtke S., Hermanns H. M. (2008) Box 2 region ofthe oncostatin M receptor determines specificity for recruitment of Janus kinases and STATS activation. J. Biol. Chem. 283(28), 19465-77. 22. Hu S., Sheng W. S., Ehrlich L. C., Peterson P. K., Chao C. C. (2000) Cytokine effects on glutamate uptake by human astrocytes. Neuroimmunomodu/ation 7(3), 153-9. 23. Jankowsky J. L., Patterson P. H. (1999) Differential regulation of cytokine expression following pilocarpine-induced seizure. Exp. Neural. 159(2), 333-46. 24. Kamimura D., Ishihara K., Hirano T. (2003) IL-6 signal transduction and its physiological roles: the signal orchestration model. Rev. Physiol. Biochem. Pharmacol. 149, 1-38. 25. Kordula T., Rydel R. E., Brigham E. F., Horn F., Heinrich P. C., Travis J. (1998) Oncostatin Mand the interleukin-6 and soluble interleukin-6 receptor complex regulate alphal-antichymotrypsin expression in human cortical astrocytes. J. Biol. Chem. 273(7), 4112-8. 26. Korn T., Magnus T., Jung S. (2005) Autoantigen specificT cells inhibit glutamate uptake in astrocytes by decreasing expression of astrocytic glutamate transporter GLAST: a mechanism mediated by tumor necrosis factor-alpha. FASEBJ. 19(13), 1878-80. •

131 CHAPTER S

27. Korzus E., Nagase H., Rydell R., Travis J. (1997) The mitogen-activated protein kinase and JAK-STAT signaling pathways are required for an oncostatin AA-responsiveelement-mediated activation of matrix metalloproteinase 1 gene expression. J. Biol. Chem. 272(2), 1188-96. 28. Li G., Bauer S., Nowak M., Norwood B., Tackenberg B., Rosenow F., Knake S., Oertel W. H., Hamer H. M. (2011) Cytokines and epilepsy. Seizure 20(3), 249-S6. 29. Lindberg R. A., Juan T. S., Welcher A. A., Sun Y., Cupples R., Guthrie B., Fletcher F. A. (1998) Cloning and characterization of a specificreceptor for mouse oncostatin M. Mo/. Cell. Biol. 18(6), 3357-67. 30. Livak K. J., Schmittgen T. D. (2001) Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) Method. Methods. 25(4), 402-8. 31. Loddick S. A., Turnbull A. V., Rothwell N. J. (1998) Cerebral interleukin-6 is neuroprotective during permanent focal cerebral ischemia in the rat. J. Cereb. Blood Flow Metab. 18(2), 176-9. 32. Matos M., Augusto E., Oliveira C. R., Agostinho P. (2008) Amyloid-beta peptide decreases glutamate uptake in cultured astrocytes: Involvement of oxidative stress and mitogen-activated protein kinase cascades. Neuroscience 156(4), 898-910. 33. Mattson M. P., Chan 5. L. (2003) Neuronal and glial calcium signaling in Alzheimer's disease. Cell Calcium 34(4-5), 385-97. 34. Moidunny S., Dias R. B., Wesseling E., Seki no Y., Boddeke H. W., Sebastiao A. M., Biber K. (2010) lnterleukin-6-type cytokines in neuroprotection and neuromodulation: oncostatin M, but not leukemia inhibitory factor, requires neuronal adenosine A, receptor function. J. Neurochem. 114(6), 1667-77. 35. Mosley B., De Imus C., Friend D., Boiani N., Thoma B., Park L. S., Cosman D. (1996) Dual oncostatin M (OSM) receptors. Cloning and characterization of an alternative signaling subunit conferring OSM-specificreceptor activation. J. Biol. Chem. 271(51), 32635-43. 36. Mosmann T. 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(1993) Glutamate uptake by astrocytes is inhibited by reactive oxygen intermediates but not by other macrophage-derived molecules including cytokines, leukotrienes or platelet­ activating factor. J. Neuroimmunol. 48(1), 99-104. 42. Repovic P., Mi K., Benveniste E. N. (2003) Oncostatin M enhances the expression of prostaglandin E2 and cyclooxygenase-2 in astrocytes: synergy with interleukin-lbeta, tumor necrosis factor-alpha, and bacterial lipopolysaccharide. Glia 42(4), 433-46. 43. Richards C. D., Langdon C., Botelho F., Brown T. J., Agro A. (1996) Oncostatin M inhibits IL-1-induced expression of IL- 8 and granulocyte-macrophage colony-stimulating factor by synovial and lung fibroblasts.J. lmmunol. 156(1), 343-9. 44. Rothstein J. D., Dykes-Hoberg M., Pardo C. A., Bristol L. A., Jin L., Kunc! R. W., Kanai Y., Hediger M. A., Wang Y., Schielke J. P., Welty D. F. 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Neurosci. 17(3), 932-40.

132 ONCOSTATIN M INHIBITS ASTROCYTIC GLUTAMATE UPTAKE

54. Thoma B., Bird T. A., Friend D. J., Gearing D. P., Dowers. K. (1994) Oncostatin Mand leukemia inhibitory factor trigger overlapping and different signals through partially shared receptor complexes. J. Biol. Chem. 269(8), 6215-22. 55. Van Wagoner N. J., Choi C., Repovic P., Benveniste E. N. (2000) Oncostatin M regulation ofinterleukin-6 expression in astrocytes: biphasic regulation involving the mitogen-activated protein kinases ERKl/2 and p38. J. Neurochem. 75(2), 563-75. 56. Wallace P. M., MacMaster J. F., Rouleau K. A., Brown T. J., Loy J. K., Donaldson K. L., Wahl A. F. (1999) Regulation of inflammatory responses by oncostatin M. J. lmmunol. 162(9), 5547-55. 57. Wang Y., Robledo 0., Kinzie E., Blanchard F., Richards C., Miyajima A., Baumann H. (2000) Receptor subunit-specific action of oncostatin Min hepatic cells and its modulation by leukemia inhibitory factor.J. Biol. Chem. 275(33), 25273- 85. 58. Weiss T. W., Samson A. L., Ni ego B., Daniel P. B., Medcalf R. L. (2006) Oncostatin Mis a neuroprotective cytokine that inhibits excitotoxic injury in vitro and in vivo. FA SEBJ. 20(13), 2369-71. 59. Zou J. Y., Crews F. T. (2005) TNF alpha potentiates glutamate neurotoxicity by inhibiting glutamate uptake in Ill organotypic brain slice cultures: neuroprotection by NF kappa B inhibition. Brain. Res. 1034(1-2), 11-24.

133 CHAPTER 5

SUPPLEMENTARY FIGURES

95% 175 Supplementary Figure 1: Shows the 150 characteristic ratio of astrocytes/ microglia in the cortical astrocytes culture J!l. 125 a5 used in this study. The cells were fixed in 4% (.) paraformaldehyde, immunostained for GFAP 0 100 (astrocytes; in green) and CDllb (microglia; in red) and observed under a fluorescence .0Q) 75 E microscope with a total magnification of z:::, 50 lO00X. The graph shows the average astrocyte 5% cell number (-152) compared with the average 25 microglial cell number (~10) after counting 10 microscopic fields inseveral 16 mm coverslips 0 of astrocyte cultures in 4 independent = experiments. The astrocyte purity was Astrocytes Microglia determined to be between 94 - 96%.

134 ONCOSTATIN M INHIBITS ASTROCYTIC GLUTAMATE UPTAKE

A

GLAST GLT- 1

1- --, GLAST (60 kDa) ------______--L__

a-Tubulin (51 kDa)

Total protein (µg) 30 60 90 120

C -- - - L __ · - - - -- GLT-1 (70 kDa) a-Tubulin (51 kDa)

To tal protein (µg) 30 60 90 120

Supplementary Figure 2: Primary mouse astrocyte cultures express GLAST and GLT-1 transporters. Cortical astrocyte cultures were established from wild-type (C57BL/6J) mouse neonatal (P2) brains. (A) Total RNA was isolated and analyzed for the expression of GLAST (91 bp) and GLT-1 (86 bp) mRNA by reverse transcriptase PCR (see Ta ble 1 for primer details); GAPDH (not shown) served as the loading control. Protein lysates prepared from cultured astrocytes were analyzed for (B) GLAST and (C) GLT-1 protein expression by Western blot. a-Tubulin served as the loading control. Note: the fact that GLT-1 proteins were only faintly detected when high amounts of protein were loaded, is consistent with the previously published findingsthat showed GLT-1 protein in pure astrocyte cultures is expressed at almost undetectable levels (Gegelashvili et al, 2000; Swanson et al, 1997).

135 CHAPTERS

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Supplementary Figure 3: OSM treatment did not affectnitric oxide release in cultured cortical astrocytes. Cortical astrocyte cultures from wild-type (CS7BL/6J) mouse neonatal (P2) brains were treated without or with OSM (10 ng/ml) for 2, 4, 8, 12 and 24 hours. Following OSM incubation, the supernatants were directly analyzed for nitric oxide (NO) content using Griess reagent system as per manufacturer's protocol. Data represents absolute values of NO concentration and presented as mean .± SEM of 2 independent experiments performed in triplicates.

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SUMMARY AND DISCUSSION

SUMMARY AND DISCUSSION

For decades, the brain has been considered an "immune privileged organ", neither susceptible nor contributing to inflammation. This idea, however, has changed over the last few years and neuroinflammation is now recognized to principally mediate host defense to brain injury and infection (Lucas et al, 2006). It is noteworthy that although anti-inflammatory mediators may contribute to neuroprotection, pro-inflammatory mediators appear to actively participate in the neurodegeneration and this has been well documented in diverse neurological disorders (Block and Hong, 2005). Therefore, it is of interest to understand how the double-edged sword of cytokine function is regulated in the central nervous system (CNS). The role of interleukin (IL)-6-type cytokines in the CNS is controversial, since they exhibit DI both pro- and anti-inflammatory properties (Kamimura et al, 2003). This cytokine fa mily consists of IL-6, IL-11, leukemia inhibitory factor (LIF), oncostatin M (OSM), ciliary neurotrophic fa ctor (CNTF), cardiotrophin 1 (CT-1) and novel neurotrophin 1 (NNT-1). IL-6-type cytokines exert their effects on target cells by acting on three types of receptor complexes: a homodimer of glycoprotein 130 (g p130/g p130; that binds IL-6 and IL-11), a heterodimer of gp130 and the LIF receptor (gp130/LIFR-�; that binds LIF, humanOSM, CNTF, CT-1 and NNT-1) and a heterodimer of gp130 and the OSM receptor (gp130/OSMR; that binds OSM) (Heinrich et al, 1998; see chapter 1, section 3.1). Elevated levels of IL-6-type cytokines are found in the brain parenchyma during pathological conditions such as brain injury, ischemia, seizure, multiple sclerosis (MS) etc. (see chapter 1, section 3.2). This led to a general belief that the severe neuronal loss and brain damage associated with the above mentioned neurodegenerative conditions were, at least in part, a result of the elevated expression and the pro-inflammatory effects of IL-6-type cytokines in the CNS. This prompted several researchers to investigate the effects of various insults on the extent of neurodegeneration in knock-out animals that lack the genes coding forsp ecificcy tokines or their receptor subunits. Interestingly, the severity of damage is fu rther enhanced in these animals. For instance, IL-6 knock-out mice are highly susceptible to a variety of convulsant stimuli (De Sarro et al, 2004) and audiogenic seizures (De Luca et al, 2004). Furthermore, other studies have shown that IL-6-type cytokine knock-out mice show multiple developmental deficits in the CNS including reduced hippocampal neurogenesis in IL-6 knock­ out mice (Bowen et al, 2011), reduced astrogliogenesis in LIF knock-out mice (Cohen and Fields, 2008) and reduced noxious response due to impaired development of nociceptive neurons in OSM knock-out mice (Morikawa et al, 2004). The developmental abnormalities of the CNS are even more severe in the receptor knock-out mice (DeChiara et al, 1995; Hatta et al, 2002; Li et al, 1995). Taken together, this evidence strongly suggests that IL-6-type cytokines and their receptors are important for normal development and fu nction of the brain and their absence increases susceptibility to damage during CNS pathology. This evidence also suggests that the elevated levels during CNS pathology of IL-6-type cytokines might be associated with their anti-inflammatory effects and not pro-inflammatory effects. In line with this argument, substantial evidence suggests that most of these cytokines are locally produced by cells of the CNS (see chapter 1, section 3.2), where they exhibit anti-inflammatory and neurotrophic properties (hence, often referred to as neuropoeitic cytokines or neurokines). On the other hand, many studies still confirm their pro-inflammatory and neurotoxic effects in the brain (see chapter 1, section 3.3). Ta ken together, IL-6-type cytokines are pleiotropic in nature and their effects (pro- and anti-inflammatory) during CNS pathology are determined by multiple factors

141 CHAPTER 6

including target cells , intracellular signaling pathways , concentration of the cytokine , general environment , presence of other inflammatory mediators etc. Besides the pro- and anti-inflammatory effects , there is strong evidence that IL-6-type cytokines exhibit potent neuroprotective properties against various insults including glutamate­ induced toxicity (excitotoxicity) (see chapter 1, section 3.4). The underlying mechanisms , however , are not yet clearly understood. Recently , we have shown that IL-6-induced protection of cultured cortical neurons against excitotoxicity depends on the expression and activity of neuronal adenosine A1 receptors (A1 Rs) , a major neuromodulatory receptor in the brain (Biber et al , 2008). In this study , the authors showed that IL-6 treatment (thus , gp130/gp130 receptor complex activation) enhances the expression of neuronal A1 Rs , and blockade of A1 Rs in neurons using a selective antagonist (DPCPX) completely abolishes the IL-6-induced neuroprotection against glutamate. Since IL-6 family members show several redundant properties (due to the shared usage of atleast one common receptor subunit , gp130) , we hypothesized that neuroprotection induced by all lL-6-type cytokines is dependent on A1 Rexpression and activity. In order to investigate this , we chose two candidate cytokines of this family that activates either of the two remaining receptor complexes , gp130/LIFR and gp130/OSMR. Thus , we analyzed neuroprotective effects against glutamate and the underlying intracellular signaling pathways of LIF and OSM. We furthermore investigated how neurons during excitotoxicity may communicate with astrocytes and microglia (major source for LIF and OSM in the CNS , respectively) , in order to induce synthesis of the two neuroprotective cytokines. A part of this study also has focussed on the effect of OSM on glutamate uptake activity by astrocytes , the major regulator of extracellular glutamate levels and thereby excitotoxicity in the CNS. We used pure cell cultures of primary neurons , astrocytes and microglia , as well as ex vivo hippocampal slice preparations in our studies. The key findings of this research are summarized below.

Both OSM and LIF induce neuroprotection against glutamate, but by different mechanisms In chapter 2, we investigated whether OSM and LIF induced protection against glutamate in cultured mouse embryonic cortical and hippocampal neurons. The toxicity of glutamate was monitored using a colorimetric survival assay (MTT assay) , propidium iodide (Pl) staining (that specifically stains nuclei of dead cells) and cleaved-caspase 3 protein expression (a well known marker used to detect programmed cell death or apoptosis) by Western blot. The number of Pl staining positive cells and the expression of cleaved-caspase 3 proteins were higher in glutamate-treated neurons when compared to the untreated controls. Glutamate concentration-dependently reduced the survival of cortical neurons , and this toxicity was dependent on the activity of neuronal NMDA receptors. On the other hand , survival of neurons against glutamate was higher when they were pre-treated (for 24 hours before glutamate) with OSM or LIF (0.1 , 1 or 10 ng/ml). OSM and LIF pretreatment also reduced the number of Pl positive neurons and cleaved caspase 3 expression induced by glutamate , when compared to cells treated with glutamate alone. The neuroprotection induced by OSM, like IL-6 (Biber et al , 2008) , was abolished by DPCPX. These results were further confirmed in A,R knock-out mice, where OSM failed to induce neuroprotection in both cortical and hippocampal neurons. Moreover , by real-time PCR and Western blot analyses , we showed that OSM treatment time­ dependently enhanced A1 R mRNA and protein expression in cultured cortical neurons. On the

142 SUMMARY AND DISCUSSION

other hand, LIF-induced neuroprotection against excitotoxicity was neither dependent on A1 R

activity, nor did it induce A1 R expression in neurons.

A1 Rs modulate glutamate-toxicity by multiple mechanisms. Their activation at the

presynaptic nerve terminal inhibits glutamate release. At the postsynaptic terminal, A1 R activation hyperpolarizes the membrane, restrains NMDA receptor activation and inhibits intracellular calcium influx, which prevents the generation and propagation of excitatory action potentials and thereby prevents glutamate-induced excitotoxicity (de Mendonc;:aet al,

2000; Dunwiddie and Masino, 2001). In chapter 2, we further showed that OSM enhances A1 R­ mediated inhibition of calcium influx in response to glutamate, in cultured cortical neurons. Additionally, we showed in hippocampal slice cultures that OSM-, but not LIF-pretreatment enhances Al mediated inhibition of excitatory postsynaptic currents (EPSCs). This effect of

OSM was also observed in a hypoxia model, where inhibition of A1 Rs (using DPCPX) reduced both basal and OSM-induced EPSC inhibition.

Neuroprotection induced by OSM and LIF: common signaling pathways, but different "effectors" Despite the known structural and fu nctional similarites between OSM and LIF (Schrell et al, 1998; Jeffery et al, 1993), neuroprotection induced by these cytokines against glutamate is apparently mediated through different mechanisms. This made us hypothesize that the above mentioned differences are probably due to activation of different intracellular signaling pathways. Therefore in chapter 3, we investigated the effects of selective inhibitors of JAK/ STAT3, ERKl/2 and Pl3K/Akt, the three commonly activated pathways by OSM and LIF, on their neuroprotective effects against excitotoxicity. We found (by immunocytochemistry and Western blot) that activation of all the three signaling proteins was induced in cultured cortical neurons treated with OSM or LIF. Whereas, neuroprotection induced by both OSM and LIF were dependent on ERKl/2 and PI3K/Akt pathways, inhibition of JAK/STAT3 pathway (using two selective inhibitors AG 490 and JSl-124) did not abolish the neuroprotective effects of OSM or LIF. These results were in contradiction to our hypothesis, because we expected involvement of different signaling pathways. Furthermore, the fact that the STAT3 pathway was not involved in the neuroprotective effects against glutamate of OSM or LIF was surprising, since these cytokines are well known as STAT3 cytokines (since they often mediate most of their effectsthr ough this pathway). We fu rther showed in chapter 3, that inhibition of nuclear transcription factor (NF)-KB by the potent inhibitor (BAY 11-7082) abolished both OSM- and LIF-induced neuroprotection. Moreover, OSM-induced neuronal A R expression (chapter 2) 1 was abolished by BAY 11-7082 (chapter 3), in both cortical and hippocampal neurons. Since both OSM and LIF require common signaling pathways (ERKl/2, P13K/Akt and NF-KB) in our study to induce neuroprotection against glutamate, but only OSM-induced neuroprotection was dependent on A1 R expression and activity, we hypothesized that the downstream effect of the pathways might lead to expression and/or activation of different "effectors" (effectors in this context are A1 Rs, for example). This prompted us to look for possible involvement of other neuromodulatory "effectors" that might mediate neuroprotective effects of LIF. Neuromodulation by cannabinoid receptor 1 (CB R) is gaining wide attention. CB Rs, like 1 1

A1 Rs, are coupled to inhibitory G 1 proteins that negatively regulate adenylate cyclase activity. Recent evidence shows a neuroprotective role for CB Rs by modulating NMDA receptor 1

143 CHAPTER 6 mediated excitatory synaptic transmission including inhibition of glutamate release at the presynaptic terminal (Gerdeman and Lovinger 2001), inhibition of NMDA receptor mediated calcium influx (Liu et al, 2009) and regulation of calcium release from the intracellular stores

(Hampson et al, 2011). Interestingly, the expression level of CB1Rs is high at excitatory synapses of the hippocampus and cerebellum and is mainly localized at presynaptic nerve terminals (Kawamura et al, 2006), where they often co-exist and interact with A,Rs (l--1offman et al, 2010). We therefore investigated potential role for the neuromodulatory CB,Rs in LIF-induced neuroprotection. It is shown here that activation of neuronal CB,Rs using a specificagonist WIN 55,212-2 (100 nM, added 30 min before glutamate) enhanced survival of cultured hippocampal neurons against glutamate toxicity, and this effect of WIN 55,212-2 was abolished by a specific CB,R antagonist, AM 251 (Figure 1). These observations suggest that hippocampal neurons in our culture conditions do express functional CB,Rs. Interestingly, LIF-induced neuroprotection against glutamate in these neurons was also abolished by AM 251 (Figure 1), suggesting that LIF requires functional CB,Rs in order to induce its neuroprotective property. Furthermore, pre-treatment with LIF (10 ng/ml for 24 hours) enhanced the neuroprotective effect of WIN 55,212-2, suggesting that LIF might directly or indirectly modulate the expression and/or

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a5 20 (.) 0 c3-v� � c3-v � fl,.� c3-v� � c3-v� c3-v1'-. -s � �CJ' �" " &'V V� 0 0� X � X X X X X (j �� rp" �� v� �� �" �x x � �" {< X 'v {< 'v ��rr, Figure 1: LIF mediates neuroprotection against glutamate toxicity by modulating cannabinoid receptor 1 (CB R) effects. 1 Primary hippocampal neurons from wild-type (C57BL/6J) mice embryo (-E15) were treated without or with LIF (10 ng/ml) for 24 h after 5 days in culture and were subsequently challenged with glutamate (SO µM) for l h. Where indicated, cultures were treated with CB1R antagonist, AM 251 (1 00 nM) or agonist, WIN 55,212-2 (100 nM) for 30 min prior to glutamate insult. Neuronal survival was assessed 24 h following the glutamate insult by a colorimetric 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay. The optical densities were measured at 570 nm with a 630 nm and blank correction. The data is normalized to percent of control and represented as mean ± SEM of quadruplicate samples from one experiment; *p < 0.05.

144 SUMMARY AND DISCUSSION

activity of CB,Rs. In order to understand this , we next investigated whether sustained exposure of hippocampal slices to LIF could modify the neuromodulatory actions of CB,Rs on synaptic transmission , using the whole cell recordings of electrically evoked EPSCs from CAl pyramidal neurons. It is shown here that EPSC inhibition by the CB,R agonist WIN 55 ,212-2 was potentiated in slices pretreated with LIF (10 ng/ml for 4 hours) , compared with the effect of WIN 55 ,212- 2 in the absence of LIF (Figure 2). These results , however , are preliminary and need further investigation to establish whether LIF indeed modulates the neuroprotective effects of CB,Rs in hippocampal neurons. Nevertheless , these results collectively indicate that although OSM and LIF induce neuroprotection against glutamate by activating common intracellular signaling pathways , the "effectors" (A,R and CB,R , respectively) could be different.

Glutamate-stressed neurons release signals that induces LIF expression and release from astrocytes Astrocytes play a major supportive role to neurons in the CNS by supplying nutrients , oxygen and other blood-derived components. In addition , they regulate concentration of neurotransmitters and ions in the extracellular fluid as well as release several cytokines and neurotrophic factors that promote neuronal survival and development. One of the important neuroprotective factors that is produced by astrocytes during excitotoxic conditions is LIF, expression of which is elevated in pathological conditions of the CNS including injury (Banner et al , 1997) , ischemia (Slevin et al , 2008) , MS (Mashayekhi and Salehi , 2011) , Alzheimer's disease and Parkinson's disease (Soilu-Hanninen et al , 2010). It is not yet understood how neurons under excitotoxic stress induce LIF production in astrocytes. However , there is evidence for astrocytic IL-6 synthesis and release mediated by adenosine (Fiebich et al , 1996; Schwaninger et al , 1997). In chapter 4, we investigated whether glutamate-stressed cortical neurons regulate astrocytic LIF expression and release. We show that treatment with supernatants from glutamate-stressed cortical neurons induces LIF mRNA expression in cultured cortical astrocytes , and this was abolished when astrocytes were pretreated either with a non-selective

adenosine receptor antagonist (caffeine) or with a cocktail of antagonists for adenosine A2A and

Figure 2: LIF potentiates inhibition 150 WIN 55,212-2 • LIF of synaptic transmission caused (500 nM) A Control by cannabinoid receptor 1 (CB R} 1 Q) activation. "'O Averaged time-course .-e :'f of afferent evoked excitatory g- 100 --- H-�;��--A------post-synaptic current (EPSC) peak ro amplitude changes caused by a 35 (.) C/) min application of the CB1R agonist a.. ..-� -; ----E� � LU . WIN 55,212-2 (500 nM), in control - ... .�. versus LIF (10 ng/ml for 4 h) treated . slices, from the same hippocampus. .§ ...... 50 ...... -. .•.. .. . Each point represents amplitude of 0 z EPSCs evoked once every 30 s by electrical stimulation of the Schaffer collaterals (n=l); 100% corresponds 0 ------� to the averaged amplitude calculated for 5-10 EPSCs recorded just before -20 0 20 40 60 80 100 adding WIN 55,212-2. Calibration: 30 Time (min) ms, 50 pA.

145 CHAPTER 6

A28 receptors (ZM 241385 and MRS 1754, respectively). We further used the adenosine analogue 5'-N-Ethylcarboxamide (NECA) to demonstrate that adenosine-induced LIF expression and release in astrocytes is mediated through the adenosine A28 receptor subtype and requires protein kinase C (PKC) and mitogen-activated protein kinases (MAPKs: p38 and ERKl/2) and NF-KB signaling. We further showed that NECA-induced LIF release in astrocytes is constitutive and by a calcium-independent mechanism, and not through large dense core vesicle mediated (regulated) secretion. Furthermore, we speculated involvement of recycling ensodomes in the LIF release process based on the immunocytochemical evidence. Finally we showed that NECA­ treated astrocytic supernatants induced protection of cortical neurons against excitotoxicity, which was dependent on the LIF content in the supernatants. Thus for the first time, we provide evidence how neurons under glutamate-stress induce expression and release of LIF in astrocytes, that supports survival of neurons against excitotoxicity. This led us to investigate whether adenosine receptor activation induces OSM expression in microglia, the major source for OSM in the CNS. Preliminary observations indicate that treatment of primary shake-offmicroglia cultures with NECA (1 µM) time-dependently induces OSM mRNA expression, with maximum induction of NECA treatment after 2 hours (Figure 3). It is however not known whether glutamate-stressed neurons induce OSM expression and/or release in microglia. Nevertheless, our findings suggest that adenosine and its receptors play a key role in mediating the effects of OSM and LIF in the CNS during excitotoxicity.

Anti- and pro-excitotoxic effects of OSM in the CNS may depend on the target cell activated by the cytokine Astrocytes play a pivotal role in excitatory neurotransmission and excitotoxicity, mainly by regulating extracellular concentrations of glutamate (Seifert et al, 2006) . This is mediated through the high efficient sodium-dependent glutamate transporters located at the plasma

2.0

'in 1.5 Q) - '-I �oQ) a.. <( <( 1.0 z (9 o::- E� Q) 0 >- 0.5 0.0 I NECA (time) Oh 0.5h 1h 2h 4h 6h

Figure 3: NECA increases OSM expression in primary mouse microglia. Primary shake-off microglia were treated without or with NECA (1 µM) for 0.5, 1, 2, 4and 6 hours. Cells were then analyzed for OSM mRNA expression (relative to the housekeeping gene glyceraldehyde-3-phosphate dehydrogenase (GAPDH)) by real-time PCR, using the following primer pairs: OSM (Fw 5'-GTGGCTGCTCAACTCTTCC-3' and Rev 5'-AGAGTGATTCTGTGTTCCCCGT-3') GAPDH (Fw 5'-CATCAAGAAGGTGGTGAAGC-3' and Rev 5'-ACCACCCTGTTGCTGTAG-3'). Data is normalized to the control and presented as Mean ± SEM of two independent experiments.

146 SUMMARY AND DISCUSSION

membranes namely, glutamate aspartate transporter (GLAST) and glutamate transporter 1 (GLT-1) . There is growing evidence that dysregulation of glutamate transporters is associated with many chronic CNS disorders such as ALS (Rothstein et al, 1995; Vanoni et al, 2004). Furthermore, the transporter expression and activity can be highly influenced by inflammatory mediators such as TNFa, IL-lP, nitric oxide, etc. (for review see, Tilleux and Hermans, 2007). The receptors for CNTF and OSM are highly expressed by astrocytes (Kordula et al, 1998; Dallner et al, 2002). It has been shown that CNTF enhances both expression and activity of glutamate transporters in astrocytes (Escartin et al, 2006), and thereby promotes survival of neurons against excitotoxicity (Beurrier et al, 2010). However, there is no evidence so far how OSM affects astrocytic glutamate uptake. Therefore in chapter 5, we investigated how OSM receptor activation in astrocytes affects glutamate transporter expression and activity. DI Contrary to the direct neuroprotective effects of OSM on neurons against excitotoxicity (chapter 2) and the above mentioned effect of CNTF on astrocytic glutamate transport, we observed that OSM treatment downregulated GLT-1 (mRNA) and GLAST (mRNA and protein) expression and reduced glutamate uptake capacities in cultured cortical astrocytes, and thereby promoted excitotoxicity in cortical neurons. This effect of OSM on glutamate transporter function was dependent on JAK/STAT3 activation in astrocytes. Thus, OSM mediates a dual role in excitotoxicity by promoting neuronal survival (when acting on neurons) or promoting neurotoxicity (when acting on astrocytes). It is noteworthy that OSM can act through gpl30/ OSMR as well as gpl30/LIFR receptor complexes in humans, unlike in mouse where they act only through the gpl30/OSMR receptor complex (Mosley et al, 1996) . Hence, OSM might induce more complex effects in humans compared to mice. Thus, it would be interesting to know whether the here suggested dual role of OSM in excitotoxicity stays valid in humans.

CONCLUSIONS

Excitotoxic degeneration of the brain tissue has been a major challenge in neuroscience, ever since this process of severe cell death was first described (Olney and Sharpe, 1969) . There is substantial evidence suggesting that excitotoxic neurodegeneration is instrumental under most pathological conditions of the CNS including ischemia, seizure, injury, Alzheimer's disease, Parkinson's disease, ALS, huntington's disease and multiple sclerosis (Mattson, 2003; Seifert et al, 2006) . Indeed, many therapeutic measures have been employed against excitotoxicity, albeit with low or moderate success rates and/or with side effects. On the other hand, recent evidence suggests that IL-6-type cytokines are neuroprotective and therefore can be used against excitotoxic neurodegeneration. Our studies have confirmed that IL-6, OSM and LIF protect neurons against excitotoxicity. However, based on the findingsin chapter 5 of this thesis, IL-6-type cytokines can have dual roles (neuroprotective and neurotoxic) during excitotoxicity, depending on the target cell of activation. Thus, a detailed understanding of the pleiotropic properties of IL-6-type cytokines is important before considering these cytokines as drug targets against excitotoxicity. The work done in this thesis further suggests an "interplay" between IL-6-type cytokines and adenosinergic system (comprising of extracellular adenosine and its receptors) in the CNS (Figure 4) .

147 CHAPTER 6

Excitotoxicity (seizure/ischemia) r,;-,. A28R activation 9 \::.} LIFR 0 •• e � activation °cp e • � &Ii r:P LIF release • �\ ...... ______t 0,,. <:S'<§" 0 oO · IL-6 NMDAR V r�L __./'-____..-- L-� O oo (------... t LIF transcription I ���-s::_ c ; c:&;�- -,- � -:� ,,.i oo tra.. n. � :?/ ; ,,.!:;,�ptOa " � �J,i_,-,:<:� 0� ASTROCYTE � l- j� �r:A R act;,at;o �� a \ , 0 • � IL-6 release O .., NEURON '% ��og oi IL-6R i:g, IL-6R activation OSMR "g00 act1vat1on activation o o (';;'\ Li, , oO � e ----➔ • � • • • k/ �og • • • 4:::--- o --� •• Adenosme.• •• • ? OSM release • • • • 1' • release • •• • • • • • / Jf'1AR activation • • • tosM � transcription 0

MICROGLIA

Figure 4: Schematic overview of the "interplay" between adenosine and IL-6-type cytokines in excitotoxicity. Excessive activation of neuronal NMDA receptors leads to excessive calcium influx and subsequent excitotoxic death of neurons (1). Excitotoxic insults such as ischemia or seizure raises extracellular ATP to high levels which is degraded into adenosine (2). Extracellular adenosine mediates multiple neuromodulatory and immunomodulatory effects by acting on its receptors in neurons and glial cells. Thus, the activation of adenosine A28 receptor subtype (A2l) on astrocytes induces synthesis and release of IL-6 and LIF (3). Whereas, adenosine receptor (subtype yet not identified) activation in microglia induces OSM expression (4). Both IL-6 and OSM activate their receptors on neurons to up-regulate expression and function of adenosine A1 receptors (Als) (5). IL-6 has similar effects in astrocytes (6). The activity of A1Rs in neurons protects them against excitotoxicity (7), whereas, A1R activation in astrocytes inhibits apoptotic cell death (8). LIF activates its receptors on neurons and causes protection against excitotoxicity (9) by a mechanism that probably involves cannabinoid receptor 1 (CB1 R).

FUTURE DIRECTIONS

1) Excitotoxicity-induced apoptotic neuronal death is dependent on NF-KB signaling (Pizzi et al , 2002; Goffi et al , 2005). On the contrary , anti-apoptotic effects of LIF and OSM that lead to neuroprotection against glutamate also requires NF-KB signaling (Chapter 3). The precise role of NF-KB in neurodegeneration is still controversial since it regulates both pro- and anti­ apoptotic genes. The distinct effects of NF-KB on viability of neurons might depend on the selective activation of diverse NF-KB/Rel family members: NF-KBl , NF-KB2 , Rel A, Rel B and c-Rel (Pizzi et al , 2002). In line with this argument , previous studies have shown that p65 subunit is involved in the glutamate-mediated cell death , whereas the c-Rel subunit is implicated in promoting neuronal survival (Heck et al , 1999; Pizzi et al , 2002; Pizzi and Spano , 2006). Hence , a detailed analysis of the NF-KB/Rel family members that are activated by glutamate and the

148 SUMMARY AND DISCUSSION

cytokines is important to better understand the dual role of NF-KB in the regulation of cell survival during excitotoxicity and neuroprotection induced by OSM and LIF. 2) We suggested a possible mechanism for LIF-induced neuroprotection through the neuromodulatory CB Rs. A couple of questions remain to be answered: Does LIF induce CBl 1 gene expression or does it directly modulate CBl receptor fu nction, to induce neuroprotection?

If LIF induces CB1 R expression in neurons, how does selective inhibition of ERKl/2, Pl3K/Akt and NF-KB signaling pathways (those identifiedto be important for LIF-induced neuroprotection) affect CB,R expression? Does LIF induce neuroprotection against glutamate in CB R knock-out 1 mice? 3) Excitotoxicity in neurons induced LIF expression in cultured cortical astrocytes, which was dependent on adenosine A receptor activation (Chapter 4). It needs to be verifiedwhether 28 A receptor stimulation also accounts for the increased LIF expression in the CNS in vivo, 28 following injury or neurodegenerative conditions. Furthermore, it would be interesting to understand whether injury induced LIF expression is affected in A receptor knock-out mice, to 28 fu rther understand the importance of A receptor activation in elevated LIF expression during 28 neurodegenerative conditions. 4) In chapter 4, we showed by immunostaining that LIF proteins co-localize with Rabll, a recycling endosomal marker and therefore speculated LIF release in astrocytes might be mediated through recycling endosomes. In order to further understand this, it is important to study how LIF release is affected in Rabll deficientastrocytes. Rabll knock-out mice are not viable and therefore siRNA or sh RNA strategies may be undertaken to investigate this issue. 5) OSM treatment inhibited the glutamate uptake capacity of cultured cortical astrocytes and thereby promoted excitotoxicity (Chapter S). This effect of OSM is mediated through STAT3 signaling in astrocytes, blockade of which prevented the effects of OSM. Suppressors of cytokine signaling-3 (SOCS3), a potent regulator of STAT3 (Ta n and Rabkin, 2005; Yoshimura et al, 2005), are widely discussed for their therapeutic potential against STAT3 mediated effects in the CNS (Wang and Campbell, 2002). It is therefore of interest to understand the effects of OSM on glutamate uptake in astrocytes overexpressing SOCS3. In addition, it would be interesting to know whether inhibition of astrocytic glutamate uptake capacities by OSM is exacerbated under conditions of SOCS3 downregulation.

149 CHAPTER 6

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14. Fiebich B.L., Biber K., Gyufko K., Berger M., Bauer J., van Ca Iker D. (1996) Adenosine A28 receptors mediate an increase in interleukin (IL)-6 mRNA and IL-6 protein synthesis in human astroglioma cells. J. Neurochem. 66(4), 1426-31. 15. Gerdeman G., Lovinger D.M. (2001) CBl cannabinoid receptor inhibits synaptic releaseofglutamate in ratdorsolateral striatum. J. Neurophysiol. 85(1), 468-71. 16. Goffi F., Baroni F., Benarese M., Sarnico I., Benetti A., Spano P.F., Pizzi M. (2005) The inhibitor of I kappa B alpha phosphorylation BAY11-7082 prevents NMDA neurotoxicity in mouse hippocampal slices. Neurasci. Lett. 377(3), 147-51. 17. Hampson R.E., Miller F., Palchik G., Deadwyler S.A. (2011) Cannabinoid receptor activation modifiesNMDA receptor mediated release of intracellular calcium: implications for endocannabinoid control of hippocampal neural plasticity. Neuropharmacology. 60(6), 944-52. 18. Hatta T., Moriyama K., Nakashima K., Taga T., Otani H. (2002) The Role of gpl30 in cerebral cortical development: in vivo functional analysis in a mouse exo utero system. J. Neurosci. 22(13), 5516-24. 19. Heck 5., Lezoualc'h F., Engert 5., Behl C. (1999) Insulin-like growth factor-1-mediated neuroprotection against oxidative stress is associated with activation of nuclear factor kappa8. J. Biol. Chem. 274(14), 9828-35. 20. Heinrich P.C., Behrmann I., Mi.iller-Newen G., Schaper F., Graeve L. (1998) lnterleukin-6-type cytokine signalling through the gpl30/J ak/STAT pathway. Biochem. J. 334 ( Pt 2), 297-314. 21. Hoffman A.F., Laaris N., Kawamura M., Masino 5.A., Lu pica C.R. (2010) Control of cannabinoid CBl receptor function Neurosci. on glutamate axon terminals by endogenous adenosine acting at A1 receptors. J. 30(2), 545-55. 22. Jeffery E., Price V., Gearing D.P. (1993) Close proximity of the genes for leukemia inhibitory factor and oncostatin M. Cytokine. 5(2), 107-11. 23. Kamimura D., Ishihara K., Hirano T. (2003) IL-6 signal transduction and its physiological roles: the signal orchestration model. Rev. Physiol. Biochem. Pharmacol. 149, 1-38. 24. Kawamura Y., Fukaya M., Maejima T., Yoshida T., Miura E., Watanabe M., Ohno-Shosaku T., Kano M. (2006) The CBl cannabinoid receptor is the major cannabinoid receptor at excitatory presynaptic sites in the hippocampus and cerebellum. J. Neurosci. 26(11), 2991-3001. 25. Kordula T., Rydel R.E., Brigham E.F., Horn F., Heinrich P.C., Travis J. (1998) Oncostatin Mand the interleukin-6 and soluble interleukin-6 receptor complex regulate alphal-antichymotrypsin expression in human cortical astrocytes. J. Biol. Chem. 273(7), 4112-8.

150 SUMMARY AND DISCUSSION

26. Li M., Sendtner M., Smith A. (1995) Essential function of LIF receptor in motor neurons. Nature 378(6558), 724-7. 27. Liu Q., Bhat M., Bowen W.D., Cheng J. (2009) Signaling pathways from cannabinoid receptor-1 activation to inhibition of N-methyl-D-aspartic acid mediated calcium influx and neurotoxicity in dorsal root ganglion neurons. J. Pharmacol. Exp. Ther. 331(3), 1062-70. 28. Lucas S.M., Rothwell N.J., Gibson R.M. (2006) The role of inflammation in CNS injury and disease. Br. J. Pharmacol. 147, S232-40. 29. Mashayekhi F., Salehi Z. (2011) Expression of leukemia inhibitory factorin the cerebrospinal fluid of patients with multiple sclerosis. J. Clin. Neurosci. 18(7), 951-954. 30. Mattson M.P. (2003) Excitotoxic and excitoprotective mechanisms: abundant targets forthe prevention and treatment of neurodegenerative disorders. Neuromolecular Med. 3(2), 65-94. 31. Morikawa Y., Tamura S., Minehata K., Donovan P.J., Miyajima A., Senba E. (2004) Essential function ofoncostatin m in nociceptive neurons of dorsal root ganglia. J. Neurosci. 24(8), 1941-7. 32. Mosley B., De Imus C., Friend D., Boiani N ., Thoma B., Park L.S., Cosman D. (1996) Dual oncostatin M (OSM) receptors. Cloning and characterization of an alternative signaling subunit conferring OSM-specificreceptor activation. J. Biol. Chem. 271(51), 32635-43. 33. Olney J.W., Sharpe L.G. (1969) Brain lesions in an infantrhesus monkey treated with monsodium glutamate. Science 166(903), 386-8. 34. Pizzi M., Goffi F., Boroni F., Benarese M., Perkins S.E., Liou H.C., Spano P. (2002) Opposing roles for NF-kappa B/Rel factors p65 and c-Rel in the modulation of neuron survival elicited by glutamate and interleukin-lbeta. J. Biol. Chem. 277(23), 20717-23. 35. Pizzi M., Spano P. (2006) Distinct roles of diverse nuclear factor-kappaB complexes in neuropathological mechanisms. Eur. J. Pharmacol. 545(1), 22-8. 36. Rothstein J.D., Van Kam men M., Levey A.I., Martin L.J., Kuncl R.W. (1995) Selective loss ofglial glutamate transporter GLT-1 in amyotrophic lateral sclerosis. Ann. Neural. 38(1), 73-84. 37. Schrell U.M., Koch H.U., Marschalek R., Schrauzer T., Anders M., Adams E., Fahlbusch R. (1998) Formation of autocrine loops in human cerebral meningioma tissue by leukemia inhibitor factor, interleukin-6, and oncostatin M: inhibition of meningioma cell growth in vitro by recombinant oncr:�tatin M. J. Neurosurg. 88(3), 541-8. 38. Schwaninger M., Neher M., Viegas E., Schneider A., Spranger M. (1997) Stimulation of interleukin-6 secretion and gene transcription in primary astrocytes by adenosine. J. Neurochem. 69(3), 1145-1150. 39. Seifert G., Schilling K., Steinhauser C. (2006) Astrocyte dysfunction in neurological disorders: a molecular perspective. Nat. Rev. Neurosci. 7(3), 194-206. 40. Slevin M., Krupinski J., Mitsios N., Perikleous C., Cuadrado E., Montan er J., Sanfeliu C., Luque A., Kumar S., Kumar P., Gaffney J. (2008) Leukaemia inhibitory factor is over-expressed by ischaemic brain tissue concomitant with reduced plasma expression following acute stroke. fur. J. Neural. 15(1), 29-37. 41 . Soilu-Hanninen M., Broberg E., Roytta M., Mattila P., RinneJ., Hukkanen V. (2010) Expression ofLIF and LIF receptor beta in Alzheimer's and Parkinson's diseases. Acta. Neural. Scand. 121(1), 44-50. 42. Tan J.C., Rabkin R. (2005) Suppressors ofcytokine signaling in health and disease. Pediatr. Nephrol. 20(5), 567-75. 43. Tilleux S., Hermans E. (2007) Neuroinflammation and regulation ofglial glutamate uptake in neurological disorders. J. Neurosci. Res. 85(10), 2059-70. 44. Vanoni C., Massari S., Losa M., Carrega P., Perego C., Conforti L., Pietrini G. (2004) Increased internalisation and degradation of GLT-1 glial glutamate transporter in a cell model for familial amyotrophic lateral sclerosis (ALS). J. Cell Sci. 117(Pt 22), 5417-26. 45. Wang J., Campbell I.L. (2002) Cytokine signaling in the brain: putting a SOCS in it? J. Neurosci. Res. 67(4), 423-7. 46. Yoshimura A., Nishinakamura H., Matsumura Y., Hanada T. (2005) Negative regulation of cytokine signaling and immune responses by SOCS proteins. Arthritis. Res. Ther. 7(3), 100-10.

151 CHAPTER6

NEDERLANDSE SAMENVATTINC

Bij de processen die in het brein plaatsvinden zijn niet alleen de zenuwcellen (neuronen) van belang, maar spelen ook de ondersteunende cellen (gliacellen) een belangrijke rol. Deze gliacellen, waaronder de oligodendrocyten, astrocyten en microglia, verzorgen het 'welzijn' va n neuronen door verschaffing va n voedingsstoffen, verwijdering van afvalstoffen en door de productie van factoren die van belang zijn bij het voorkomen en bestrijden van infecties en hersenschade. Een voorbeeld va n deze laatstgenoemde factoren zijn de zogenaamde cytokines. Bij ziekte of beschadiging van de hersenen is de aanwezigheid van cytokines uit de 'interleukine-6 (IL-6) fa milie' in de hersenen verhoogd. Deze fa milie bestaat uit 7 leden, te weten IL-6, IL-11, leukemia inhibitie factor (LIF), oncostatine M (OSM), ciliaire neurotrofe factor (CNTF), cardiotronne 1 (CT-1) en novel neurotrophin 1 (NNT-1). Ze behoren tot een fa milie omdat ze overeenkomstige eigenschappen hebben en gebruik ma ken van gemeenschappelijke receptoren. In eerste instantie werd verondersteld dat deze cytokines onder patholog ische omstandigheden een schadelijk effect hebben op neuron en en verantwoordelijk waren voor het afsterven van deze cellen. Uit studies met knock out (KO) muizen bleek echter dat de cytokines uit de IL-6 fa milie en hun receptoren juist onmisbaar zijn voor de normale ontwikkeling en fu nctionering van de hersenen. Uit eerder onderzoek is gebleken dat cytokines een belangrijke 'd ubbelrol' spelen tijdens pathologische omstandigheden in de hersenen, omdat ze tegelijkertijd zowel negatieve als positieve effecten kunnen hebben op neuronen. De cytokines van de IL-6 familie zijn beschreven als 'ontstekingsremmend' (anti-inflammatoir) en als ontstekingsactiverend (pro­ inflammatoir). Daarnaast is IL-6 ook nog beschreven als neuroprotectieve factor, oftewel als een cytokine dat een beschermende werking kan hebben op zenuwcellen. Bij acute en chronische hersenziekten kan er sprake zijn van neuronale schade als gevolg va n gluta maat toxiciteit. Bij dit proces worden de zenuwcellen 'te sterk gestimuleerd' door een overmaat van de neurotransmitter glutamaat, wat uiteindelijk leidt tot celdood. Het is nog niet geheel duidelijk wat de rol van de IL-6 type cytokines is in dit proces. Veel basale processen die zich afspelen in de hersenen worden onderzocht m.b.v. celkweek technieken. Hierbij worden de verschillende hersencellen va n een proefdier, neuronen of gliacellen, buiten het lichaam gekweekt en kunnen fundamentele mechanismes worden bestudeerd, zoals bijvoorbeeld de rol van bovengenoemde cytokines in het proces van neurodegeneratie en excitotoxiciteit. In dit proefschrifthebben we twee led en van de IL-6 fa milie bestudeerd, namelijk OSM en LIF, en onderzocht welke rol ze spelen in neuroprotectie en neurodegeneratie. In eerste instantie hebben we aangetoond dat zowel OSM als LIF beschermend werken op neuronen die aan stress onderhevig zijn. Het proces van excitotoxiciteit werd nagebootst door embryonale neuronen van de muis in kweek te behandelen met glutamaat, wat resulteerde in massale celdood. Dit was echter niet het geval wanneer de neuronen v66r glutamaat toevoeging waren behandeld

met OSM of LIF. Uit voorgaand onderzoek was gebleken dat de adenosine A1 receptor (Al) een belangrijke rol speelt bij de beschermende neuronale werking van IL-6. Daarom herhaalden we onze experimenten met neuronale celkweken afkomstig van muizen die het gen voor de A R missen, zogenaamde A R knock out muizen. Het neuroprotectieve effect van OSM kon in 1 1 deze kweken niet meer worden aangetoond, terwijl het beschermende effect van LIF nog wel

152 NEDERLANDSE SAMENVATTI NG

tot uiting kwam. Hieruit konden we concluderen dat ondanks de structurele en functionele gelijkenissen tussen de beide cytokines, de werking via verschillende mechanismen verloopt

en dat de A1 R van belang is bij de neuroprotectieve werking van OSM en IL-6, maar geen rol speelt bij LIF. In hoofdstuk 3 hebben we onderzocht of het verschil in mechanisme tussen OSM en LIF gelegen is in het feit dat ze gebruik ma ken van verschillende intracellulaire signaal paden. De reeds in de literatuur beschreven paden die een belangrijke rol spelen bij neuroprotectie en excitotoxiciteit werden in onze embryonale neuronenkweken selectief ge"inhibeerd met specifiekebl okkers en daaruit bleek verrassend dat er geen verschil kon warden aangetoond tussen de gebruikte signaal cascades van OSM en LIF. Om het neuroprotectieve effect van LIF te verklaren, kwam na literatuurstudie de Ill cannabinoid receptor 1 (CB1 R) als een mogelijke factor naar voren en de neuronen in onze kweken bleken deze receptor inderdaad tot expressie te brengen. Door aan de gekweekte

neuronen stoffen toe te voegen die een rem mend of juist activerend effecthadden op de CB1 R konden we aantonen dat het neuroprotectieve effect van LIF via deze receptor verloopt. LIF en OSM maken dus gebruik van dezelfde intracellulaire signaal cascades, maar de uiteindelijke werking van de beide cytokines verloopt via verschillende receptoren. In geval van OSM (en IL-6) is dat de A,R, LIF maakt gebruik van de CB,R . Het is nog niet geheel duidelijk hoe de (gestresste) neuronen gliacellen aanzetten tot de productie van de beschermende cytokines, maar uit eerder onderzoek is gebleken dat de adenosine receptoren op de membraan van neuronen een belangrijke rol spelen bij de IL-6 productie van astrocyten. Uit onze experimenten bleek dat astrocyten, gekweekt in het medium van met glutamaat gestimuleerde neuronen, aangezet werden tot het produceren van LIF. Behandeling van gekweekte astrocyten en microglia met 5'-N-Ethylcarboxamide (NECA), een stof die veel overeenkomsten heeft met adenosine, resulteerde in de productie van cytokines door de gliacellen. Wanneer het celkweek medium van deze door NECA gestimuleerde glia cellen toegevoegd werd aan neuronenkweken, dan zagen we dat er na glutamaat behandeling minder celdood optrad. Uit deze experimenten, beschreven in hoofdstuk 4, concluderen we dat de factoradenosine een belangrijke rol spelen bijglutamaat -toxiciteit en er een soort van communicatie is tussen 'gestresste' neuronen en gliacellen. Astrocyten, de producenten van LIF, hebben in hun celmembraan eiwitten die zorgen voor de opname van glutamaat. Deze zogenaamde glutamaat transporters, spelen een belangrijke rol bij het voorkomen van glutamaat-toxiciteit. De glutamaat transporters GLAST en GLT-1 reguleren de concentratie extracellulaire glutamaat en verhoging van de expressie van deze beide transporters beschermt de neuronen tegen glutamaat-toxiciteit. Echter, na toevoeging van OSM aan gekweekte astrocyten zagen wij een verlaging van de expressie van GLAST en GLT-1, wat indirect een toxisch effect heeftop neuronen. Dit effect werkt het neuroprotectieve effect van OSM in hoofdstuk 2 tegen, waaruit wederom blijkt dat de cytokines uit de IL-6 familie een 'dubbelrol' spelen tijdens pathologische processen in de hersenen. Het voorkomen van degeneratie van neuronen ender pathologische omstandigheden blijft een uitdagend onderzoeksgebied en uit onze resultaten is gebleken dat de cytokines uit de IL-6 familie potentiele kandidaten zijn voor de behandeling van chronische en acute hersenschade.

153 CHAPTER 6

mc:rurm6ID O\JotlflnOo

moru.>"lacfbouo6ITT3un� [email protected] crunn acfbOCJ06ITT3§6o (supporting cells or glial cells) cme.ia.�,pol@og LnJruCOCllTlll m nJO!Prucrucm 6i00uBm"li000 @ru"lacfboCJ06ITT3@6@s cruomd:b316TDo @O'M! rum6CllTlll6cm6. @@cru@gocfb1o&cru ( cytokines) n{j)cm o1 CD.>@'Ms6cm @ruolw <:LnJo3"lm6cfbun @6TDfll06TD.

cme.ia�oo1m6 lcfb§},@S Ofllro'B !1.l"l

cm"ltrumow nJe.i mrrucml�cfb amo(J)6ITT3§1e.i6o moru>1cfb@6@s mouocmm1m6� 63m6 LnJWom cfb0«>6TDo glutamate n{j)CTn m,16@00-tsoo&rruml gol@ng am,ml ( am,cmow1cfb§6@s mouom�so) "n{j)cfh@@cruagoaso cfhcru1cru7g1'' ( excitotoxicity) n(j)CTn06TD @nJocm6@ru «rrao1CID@�s6cmlcfb@§ n{j)cfh@@cruagoasocfhcru1cru1g1 fll},e.Jfll],6TlsOcfb6cm cmce,moolarn m1cm6o cruom�1d36)omocfb6@mcm6o @6 moru>"lcruom�6TD LnJLcfb1CD.>CD.>6@s ne.ismCD.>60 LnJruCOCllTlllmru6o @1 mfll6d36)6 fllmq;\lle.JOd36)01m6cm1 'iM·

@'D nJomcmm1e.i6@s 6ml6ITT3LJn m6Tls6 LnJWom cruo1sorumcfb§06TD6 marnce, cmlcfb @},@S LnJClJCOlcfb@§ cruom�1d36)6cm LnJLdhlCIDCD.l},@S ne.is OSM mCD.>60 LnJClJCO6Tls}, cmmo @@cru@goce,7mcru6cfb@},@S (LIF @o @o) moru>"lcruom �6TD LnJLcfb1CD.>CID6@s ne.ismCD.>60 LnJClJCO1 am,amJn!l:!6mru1awCIDC2l0d36)},CTn},. 6ml6ITT3 @},@S LnJWom dh@Gm}, cruod:b311nJ6nJo cmo@\9 ru1 rum1i006cm6. OSM, LIF n{j)cm"l @@cru@goce,7o&cru6ce,uB lcfb@@ n{j)cfh@@cruagoasocfhcru1cru1g1w1arn m1cm6o cm"lm�wow;,,o cruom�1d36)6cm6. moru>[email protected]� "ad­ (A R) enosine A 1 receptor" n{j)cmo1w@'Ms6cm @fllo@@L6Tll"lcruo«>�6m LnJLdh1CD.>CID� ru@@«> «rracm,1oruuo,1 n (A R) l2lOCID 63©}, e.JScfbfll06TD n(j)"lcruo«>�6m LnJLdh1CD.>cw� @o C:LnJO§"lml@ng 1 �ruuo,1m1�. nJcfbmo, "cannabinoid receptor 1" (CB1 R) n(j)cmo1CD.>@�s6cm m@gom;,, @fllo@@L6Tll

(OlQJO�oolarn LIF @ng lnJUlom atcruocmcw astrocytes n(j)@�s;,,cm glial C:cfb0CJ06ITT3§06m moru>lcfbun

n{j)@CTID!fhle.i;,,o «[email protected]§6ITT3LJn (signals) ol e.i"lrru 6)ru(!Y},CTn},06TlsO n{j)CTn!o «r@6)(ll)ml} «m)5CD.l0@6ITT3§06)6TT)CTn},O 6ml6ITT3@6@s nJom1.e,uo as­ trocytes ae.id36)6 LIF @ro'BnJoB1'M7i00om6� ffi)OCBCJOl2lCD.>d36)},cfbCD.>6o «rracm6 ru\91 moM1cfb@@ �1d36)},cfbCD.>6o @!1.l1 6TID6ITT3§6@s nJOmo cfb06TD1d36)6cm6. m@gom6 cmmlro'B neuron - glia �uowru1m1fllCD.>o msd36)6cm6@6Tlsrro 6TID6ITT3@6@s nJomo ClJ_,\c6(ll)fllOd36)},

154 UJC:OJrm6ID CTUotUJnOo

tIDLDomanoamn1arn n(j)cfb@@cruc:goc:socfbcru1cru1g1w1ei1� OSM @OB LnJrumamnmamn1m «>6ml ruuo6GB@l�­ moM"lc:cfbouo6ID3@1arn Ln.Jrumam»1t66)lC:CllJOuil C:'2lm8n.J0611m»C:n.J06)eJ «ml«l'll moM"lcruom�Gmo n(j)CTn),\ B'D crumwo, astrocytes n(i)cmglial c:cfbouo6ID3�U@s c:r21arn LnJrumamn1t66)1c:C11Jouil OSM m1 ru1 ri!:!'2ltID'2lOtID <:BOril:ln.OeJ6ID3@l6mOruoo. c:mm@amn cru6!l.J1�-'2!t66)lcm«r>ow1 cfb6ml- @a»Extrace llular space n(i)cm '2lOW.,\'2lam»1arn glutamate @OB «>lnJ°1cfbmGmamn1mi cfbomGmr21oru1cfbwlo am,cmiruw1 moM1cfbuilc66> «r>cfbmooi6moruom1� cruofill.,\lO @!l.Jcg/lCTnl. c:mocnoruCT\.lllw1arn moM1cfb@l@S mouomn!Ms@amn «r>SWlcfb n(j)CTncm @C:?;:JOY>lo 63«>1 @ru�1ru1@1 «r>@cmtID06Tl1. @@cru@gocfb1oBcru1@0B 63©1 LnJWOffi Ln.JO«l'l.,\cfbc66> «mlOfficfbo nJO©UOJn.OeJ6ID3uil �� n(j)CTT>«l'l06Tl1. n(i)m,1ei10 1n!Mcfb

155 CHAPTER 6

ACKNOWLEDGEMENTS

....and it's finally time to end the story, but not before thanking all the people who have been a part of my PhD life. Remembering the past 5 years is indeed a challenging task, and so please fo rgive me if I have forgotten to mention your name here .... First and foremost, I offer my sincere gratitude to my supervisor Prof. Dr. K.P.H. Biber for giving me an opportunity to work with him. Knut, I appreciate the confidenceyou have shown in me despite the fact that I was inexperienced and unfocussed in the beginning. You have helped me improve my scientific skills (planning, presenting the data, writing articles, etc.) and to critically evaluate my work ... Your motivation and support was instrumental towards completion of this thesis. Most importantly, I thank you for giving me the freedom to implement my ideas. Prof. Dr. H.W.G.M. Boddeke (Erik) ... You have been a great mentor and I am glad to have gotten a chance to work with you!! You felt responsible, made sure that everything went alright and have supported me throughout my PhD. Moreover, scientific discussions with you have been very inspiring and fruitful. .. I am grateful to you for proof- reading the chapters in this thesis and giving valuable feedback ... As I promised, I owe you a nice holiday in Dubai. .. Thank you once again Mr. Bl! I express my sincere gratitude to the members of the reading committee: Prof. Dr. U.L.M. Eisel, Prof. Dr. M. Schmidt and Prof. Dr. A.P. IJzerman for their time and effort and for approving my thesis. Sincere thanks to all my collaborators: Marco, Raquel, Prof. Dr. Paola Bezzi, Prof. Dr. Sven van IJzendoorn, Johan Bijzet, Prof. Dr. Ana Sebastiao, Prof. Dr. Rodrigo Cunha, Prof. Dr. Paula Agostinho, Prof. Dr. Cristina Limatola, Guisy and Dr. Clotilde... for your scientific inputs into the research proj ects and sharing "shoulders" to reduce my work-load. Special thanks go to the staff at BCN graduate school ... Britta (who gave a nice city-tour in Groningen and helped me start living here by providing ample info rmation about the Dutch life-style) ... Diana and Janine (for all the favours, support, travel grants and for being very friendly). Gerry ... for the administrative work and for extending hands whenever I needed it. I like to extend my gratitude to Prof. Dr. Obaid Siddiqi and his lab members (Bilal, Hisham, Abu, Satya, Jawaid, Sunil) at the National centre for Biological sciences in Bangalore, India. Dear Obaid, thanks a lot for opening "the door" for me to scientific research, which would not have been possible without being a part of your research group. Yo ur dedication and strong passion for science has been highly inspirational, which in fact prompted me to take up a scientificcar eer. I cannot thank enough the people at Medical physiology (Med Fys) ... for accepting me as a part of them and for creating a friendly atmosphere ... I would specially like to thank all the individuals who has been associated with my stay here ... Amalia... for being so kind to teach how to isolate neurons from mice in the beginning of my PhD... and later on helping me with ELISAs ... thanks a lot!! Eiko ... dude, you have patiently helped me standardize the neuron culture protocol, and have cheered me up whenever the cultures failed. Afterall , "bad results are the source for new inspiration", right? ;) Marjolein... for the beginners Dutch lessons, for teaching how to properly hold a pipette ;)) and for the never ending "discussions" on MSN!! ;)) Evelyn.. . (my favourite working-partner) ... Eef, you have been such a great help for me in finishing up my proj ects ... Thanks for being very patient with me and

156 ACKNOWLEDGEMENTS

staying motivated even afterperforming hundreds of qPCRs ... a big THANK YOU and a tight hug for all the efforts you have put in ... Nieske ... (the multi-tasker)... thanks for being a wonderful person ... you totally understand the frustrations/problems of a PhD student irrespective of the cultural differences ... There were times when I failed to communicate/explain my difficultiesto others, and you helped me bridge the gap ... I respect you for what you are ... thanks a loooot!! (and heyy ...special thanks for organizing the bowling sessions, which in the end determined the "ultimate" winner ... Tadaann!!! ;)) Michel... was a pleasure working with you.. . enjoyed those long microscopy sessions and thanks for patiently trying to convince me with the "perfect" confocal images. D.J. Miguel ... thanks for bringing in the tech no feel to Med Fys, cheering up all the people ... many thanks for the cloning work buddy ... Trix ... thanks for kindly taking care of the paper works and of course ordering chemicals without disappointing me! ;) Sjef and Bart ... for proof-reading my chapters and giving valuable comments. Inge and Rob ... for help - with statistics, lessons on electrophysiology and ofcourse for the short n easy chat "in dutch" at the corridor. Tj alling ... for fixingcomputers, providing programs, cables (I often troubled you for USB cables) ;) and for hosting the Indian dinner party at your place!! :). Loes ... for keeping the cell-culture lab "in control" and re-organizing things ... letje ... for storing PFA stocks at the cold-room ... from which I stole once in a while!! ;) Reinhard ... (Mr. cool) well, NO .. NO ..NO .. NO .. ;))) bro, you have been one among my best buddies in Groningen ... we started off with PhD together ... and eventually got closer with time. You have been a highly tolerant office­ mate ;) and a trust-worthy friend, whom I felt extremely comfortable sharing personal things with. Really enjoyed "De Pintelier" sessions, bowling, swimming, barbecues, dinners (you name it!!!). Romy ... you have been a wonderful friend ... I will treasure those pictures from the Rome trip, when you and Reinhard visited me.. . I wish you both good luck and hopefully will bump onto each other in the years to come .. ;) Marcin ... (The Mr. iPS) .. thanks for the nice Polish evenings, dinners and barbecue in the rain ... ;) Bro, I would always remember my birthday in 2009 at "De Pintelier", where you cheered all of us with an "interesting" story on how to throw stones into the sea ... ;) I wish you and Kasia lots of luck and happiness in life, and good luck with the PhD!! Carlos and Ulrike ... thanks for those Wednesdays at "De Pintelier" ... wish you both good luck!! Vishnu ... it's going to be OKAY buddy.. . Please do not worry too much .. and take good care of your health. Thanks for helping me with the experiments, dinners at Kohinoor and coffee breaks in between the late night experiments ... Ming-san... (aka de shizzle) you have been my all-time-favourite buddy.. . probably someone to whom I connected the most in the department. Thanks for the unconditional care and for being sooo helpful throughout.. . what would have I done to translate those Dutch letters, without your help?!! ;) special thanks for designing the thesis cover ... and I really enjoyed our "dart" series at the office, "fish" hunting, chats on music, painting etc. ... Bro, long way to go!! Keep up the spirit and good luck with dentistry and PhD! :) Xin ... for being a quite yet cheerful office-mate while I wrote my thesis .. . ;) Loved those interesting chats over the Chinese and Indian cultural differences ... Ria... I hope you find the "white horse" and hey, good luck with the PhD!! Wandert ... you knew I was better in bowling, didn't u?!! ;)) thanks a lot for the fun times bro ... and good luck with the PhD. Fabrizio... (somebody whom I never dared to speak Italian to, after I saw him making fun of Jonathan). ;) It's a pity you are leaving half-way.. . but will stay in touch through FB... Good luck!! Divya ... I've really enjoyed sharing the officewith you and teasing u all the time.. . ;) thanks for tolerating my mean jokes .. and for the scientific discussions... good luck winding up the PhD!!

157 CHAPTER 6

Arun ... Couldn't get enough time to spend with you ... but I wish you good luck and will be in touch! Jonathan ... I really enjoyed the "chemistry" working with you... Bro, you have been very helpful and understanding ... I look forward to collaborate with you in the future. Do have fun in Italy! ! Kumar ... thanks for the advices, support and guidance when I needed them the most. I would like to thank all my students (Annika, Femke and Charlotte), ex-lab members (Falak, Marta, Hilmar, Sander) and new additions at the Med Fys (Suping, Laura, Silvia). I thank all people at the animal housing facility... especially Hester for being very friendly and ordering mice in time ... I thank all nice people at the 5th floor (Serena, Alexandra, Melania, Visnja, Balaji, Vaishali), 9th floor (Elena, Hans) and 10th floor (Zia, Hande., Herschel) ... where I constantly ended up asking for chemicals and technical help. Special thanks to all my good friends outside work, "Desis" in Groningen and NANMA (niranja nanmaites) for creating a home away from home ... Special thanks to Siva, Mariam, Jojo, Nibu, Sherin, Vinod, Toni, Sadia, Olah, Emmanuele, Gabriele, Saed, Saravanan, Arun, Sijeesh, Reeja, Prasanth, Pramod, Guru, Anil, Vinay, Reza ... and a lot more) for making my stay in NL memorable ... I like to extend my deepest gratitude to the best-ever college buddies: Shruthi and Shankar ... for supporting me at hard times ... "Professore" Jaykumar ... for the good times at the M.G. and Brigade.. . and for the 2009 New Year's Eve in Groningen .. . My dear Dutch counter parts: Pierre (pratlizzle), Egbert ("Lens"manizzle), Greg, Oscar, Bart and Djowie ... I am extremely thankful to you guys for introducing me to the "real" Dutch life ... ;)) cooking, partying and spending my weekends at the studio "turning it up a little" is all something I am gonna miss ... Keep rocking!! Special thanks to Ruth for being a great friend and for saying "yes" when I asked her to be my paranymph! ;)) Special thanks to Rajiv vijayaraghavan for proof-reading the malayalam summary of my thesis ... Special thanks to Karameh for supporting me throughout the final year of PhD and for the Farsi lessons and food ... ;)) A very special thanks to Azeez (Founder, Ponnappan school of drama), bahju, achoos and zia ... for considering me as one among them ... for being good hosts every weekend ;) and for being there at all times ... Az, thanks for the malayalam summary (nicely done!!) ;) I like to extend special thanks to my teachers at high school (Shiji, Nancy, Shaji, Ramla, Shyama) who shaped me up and encouraged me to do science ... Last but not the least ... I thank my family (Uppa, Umma, Rishad, Sahla, lshiqa, Seena, Shahbaz, lzzal, Sabeel, Shami and Afra) for the love, care and support ... for showing confidence in me ... and for being there for me at good and bad times ... love you!! Before I end, I like to remember my late Grandfather, who strongly inspired me to become a scientist but couldn't witness it happening ... I miss you vallippa ...

158