dissertations xavier ekolle ndode-ekane development of...

Publications of the University of Eastern Finland

Dissertations in Health Sciences

isbn 978-952-61-0990-9

Publications of the University of Eastern FinlandDissertations in Health Sciencesd

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Xavier Ekolle Ndode-EkaneDevelopment of Epileptogenic

Network Alterations in Rodent Models of Status Epilepticus:

Role of the urokinase-type plasminogen

activating system

Xavier Ekolle Ndode-Ekane

Development of Epileptogenic Network Alterations in Rodent Models of Status Epilepticus:Role of the urokinase-type plasminogen activating

system

Temporal lobe epilepsy is the

most common form of acquired

epilepsy. This thesis explored the

interaction between the cellular

reorganization processes occurring

during epileptogenesis, their

functional implications, and the role

of the urokinase-type plasminogen

activator (uPA) and its receptor

(uPAR) in epileptogenic network

alterations. It shows that components

of epileptogenic network alterations

may have different temporal

profiles, and the Plau and Plaur

genes encoding for uPA and uPAR

respectively are modifier genes for

acquired epileptogenesis.

XAVIER EKOLLE NDODE-EKANE

Development of Epileptogenic NetworkAlterations in Rodent Models of Status

Epilepticus:

Role of the urokinase-type plasminogen activating system

To be presented by permission of the Faculty of Health Sciences, University of EasternFinland for public examination in Tietoteknia Auditorium (TTA), Kuopio, on Saturday,

December 15th 2012, at 12 noon

Publications of the University of Eastern Finland Dissertations in Health Sciences

Number 144

Department of NeurobiologyA.I Virtanen Institute for Molecular Sciences

University of Eastern FinlandKuopio

2012

Kopijyvä OyKuopio 2012

Series Editors:Professor Veli-Matti Kosma, M.D., Ph.D.Institute of Clinical Medicine, Pathology

Faculty of Health Sciences

Professor Hannele Turunen, Ph.D.Department of Nursing Science


Professor Olli Gröhn, Ph.D.A.I. Virtanen Institute for Molecular Sciences


Distributor:University of Eastern Finland

Kuopio Campus LibraryP.O.Box 1627

FI-70211 Kuopio, Finlandhttp://www.uef.fi/kirjasto

ISBN (print): 978-952-61-0990-9ISBN (pdf): 978-952-61-0991-6

ISSN (print): 1798-5706ISSN (pdf): 1798-5714

ISSN-L: 1798-5706

III

Author’s address: Department of NeurobiologyA.I. Virtanen Institute for Molecular SciencesUniversity of Eastern FinlandKuopioFinland

Supervisors: Professor Asla Pitkänen, MD, PhDDepartment of NeurobiologyA.I. Virtanen Institute for Molecular SciencesUniversity of Eastern FinlandKuopioFinland

Professor Olli Gröhn, PhDBiomedical Imaging UniteA.I Virtanen Institute for Molecular SciencesUniversity of Eastern FinlandKuopioFinland

Reviewers: Docent Irma Holopainen, MD, PhDDepartment of Pharmacology, Drug development and TherapeuticsUniversity of TurkuTurku, Finlande-mail: [email protected]

Eleonora Aronica, MD, PhDDepartment of (Neuro) PathologyAcademic Medical CenterUniversity of AmsterdamAmsterdam, The Netherlandse-mail: [email protected]

Opponent: Professor Leszek Kacmarek, PhDDepartment of Molecular & Cellular NeurobiologyNencki InstituteWarsaw, Polande-mail: [email protected]

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Ndode-Ekane, Xavier EkolleDevelopment of epileptogenic network alterations in rodent models of status epilepticus: Role of theplasminogen activator system. 75 pUniversity of Eastern Finland, Faculty of Health Sciences, 2012Publications of the University of Eastern Finland. Dissertations in Health Sciences, Number 144, 2012.

ISBN (print): 978-952-61-0990-9ISBN (pdf): 978-952-61-0991-6ISSN (print): 1798-5706ISSN (pdf): 1798-5714ISSN-L: 1798-5706

ABSTRACTEpileptogenesis refers to a phenomenon whereby an initial brain damaging insult such as statusepilepticus (SE), stroke, infection, or traumatic brain injury triggers a cascade of molecular andcellular alterations that ultimately results in the occurrence of spontaneous seizures. Thesealterations which include neurodegeneration, gliosis, neurogenesis, granule cell layer dispersion,mossy fiber sprouting, inflammation, blood-brain barrier disruption and angiogenesis, are wellcharacterized; however, the molecular mechanisms that underlie them are poorly understood.

The aim of this thesis is to study the interaction(s) between the cellular reorganizationprocesses in the hippocampus during epileptogenesis, and in particular vascular and neuronalplastic events, since common trophic stimuli drive vascular and nerve network formation. Next,the molecular mechanisms that underlie these network alterations will be studied. The urokinase-type plasminogen activator (uPA) and its receptor (uPAR) were selected as a possible candidatemechanism, because; there is an increased expression of uPA-uPAR during epileptogenesis.Furthermore, uPA-uPAR has implications in various cellular reorganization processes, includingcell proliferation, migration and survival.

The pilocarpine-induced SE rat model and the intrahippocampal kainic acid-induced SEmouse model were employed in studying the interactions of the reorganization processes, and roleof uPA and uPAR in these processes, with the aid of immunohistochemistry methods. Thedevelopment of epilepsy was monitored using video-electroencephalography. Magnetic resonanceimaging (MRI) was used to study the functional outcome of vascular remodeling duringepileptogenesis.

The results demonstrate that the epilepsy phenotype following SE is severe in uPARdeficiency, where as it remains unchanged in uPA deficiency. SE induces vascular break down,followed by a progressive build-up of blood vessels. This process of vascular build-up is severelyimpaired in uPAR deficiency but not uPA deficiency. Nonetheless, the vessel length changescorrelate with cerebral blood volume increase in the hippocampus but lack association withneurodegeneration, neurogenesis or axonal sprouting. uPAR or uPA deficiency affects neuronalplastic events such as neurodegeneration and neurogenesis, but not mossy fiber sprouting.Furthermore, uPAR deficiency results in a delayed acute but chronic inflammatory response.

In conclusion, this study demonstrates that the components of epileptogenic networkalterations may have different temporal profiles during epileptogenesis. However, the uPA-uPARsystem is a part of the underlying mechanisms involved in the generation of these networkalterations. This suggests that the Plau and Plaur genes encoding for uPA and uPAR respectivelyare modifier genes for acquired epileptogenesis.

National Library of Medicine Classification: WL 385, WL 300, WL 302, WL 102.5, QU 142Medical Subject Headings: Status Epilepticus; Epilepsy; Blood Vessels; Brain; Cerebrovascular Circulation;Neurons; Nerve Net; Neural Pathways; Neuronal Plasticity; Nerve Degeneration; Neurogenesis; Urokinase-Type Plasminogen Activator; Receptors, Urokinase Plasminogen Activator; Hippocampus; Disease Models,Animal; Rats; Mice

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Ndode-Ekane, Xavier EkolleDevelopment of epileptogenic network alterations in rodent models of status epilepticus: Role of theplasminogen activator system. 75 pItä-Suomen yliopisto, terveystieteiden tiedekunta, 2012Publications of the University of Eastern Finland. Dissertations in Health Sciences Number 144,. 2012.

ISBN (print): 978-952-61-0990-9ISBN (pdf): 978-952-61-0991-6ISSN (print): 1798-5706ISSN (pdf): 1798-5714ISSN-L: 1798-5706

ABSTRAKTI

Epileptogeneesi tarkoittaa epilepsian kehittymistä, jonka voi käynnistää status epilepticus,aivoverenkiertohäiriö, infektio tai pään vamma (symptomaattinen epilepsia). Muutoksetmolekyyli- ja solutasolla kuten hermosoluvaurio, hermotukisolujen lisääntyminen,sammalsäikeiden versominen, veriaivoesteen vaurio ja verisuonten uudismuodostus voivatlopulta johtaa spontaaneihin toistuviin kohtauksiin. Vaikka tämä tapahtumaketju on hyvinselvillä, sen molekyylitason mekanismit tunnetaan vielä huonosti.

Tässä väitöskirjatyössä selvitettiin hippokampuksen alueella tapahtuvia neurobiologisiamuutoksia epileptogeneesin aikana. Erityisen mielenkiinnon kohteena olivat verisuonten jahermosolujen muovautuvuutta säätelevät molekyylitason mekanismit. Tutkimukseen valittiinurokinaasityyppinen plasminogeeniaktivaattori (uPA) ja sen reseptori (uPAR), sillä aikaisemmattutkimukset ovat osoittaneet, että uPA:n ja uPAR:in ilmentyminen on lisääntynyt epilepsiankehittymisen aikana. Tämä on myös yhteydessä solutason uudelleenjärjestäytymiseen (mm.solujen uudismuodostus, kulkeutuminen ja hengissäsäilyminen).

Kokeissa status epilepticus aiheutettiin rotalle pilokarpinnilla ja uPA/uPAR poistogeenisillehiirille hippokampukseen injisoidulla kainihapolla. uPA/uPAR:in osuutta kudostenuudelleenjärjestäytymisessä tutkittiin immunohistokemiallisin menetelmin. Epilepsiankehittymistä seurattiin video-EEG:llä ja verisuonimuutoksia magneettikuvausta käyttäen.

Tulokset osoittivat, että epilepsian ilmentyminen status epilepticuksen jälkeen onsidoksissa uPAR:in poistogeenisyyteen. Status epilepticus aiheutti myös häiriön veriaivoesteentoiminnassa, jota seurasi asteettainen lisääntynyt verisuonten uudismuodostus. Tämä prosessi olisidoksissa uPAR:in ilmentymiseen. Muutokset verisuonten pituudessa korreloivat verenkierronlisääntymiseen hippokampuksen alueella, mutta eivät hermosoluvaurioon, hermosolujenuudismuodostukseen tai hermosäikeiden versomiseen. uPA/uPAR:in poistogeenisyys vaikuttihermosolujen muovautuvuuteen (mm. hermosoluvaurio ja hermosolujen uudismuodostus), muttaei sammalsäikeiden versomiseen. Lisäksi status epilepticus aiheutti viivästyneen tulehdusreaktion,joka muuttui krooniseksi uPAR poistogeenisillä hiirillä.

Yhteenvetona voidaan todeta, että hermosolujen ja verisuonten uudismuodostuksella jatulehdusreaktiolla on erilainen ajallinen yhteys epileptogeneesiin. uPA:ta ja uPAR:ta koodittavillageeneillä, Plau ja Plaur, on tärkeä merkitys hermokudoksen muovautumisessa ja symptomaattisenepilepsian kehittymisessä

Luokitus: WL 385, WL 300, WL 302, WL 102.5, QU 142Yleinen suomalainen asiasanasto: epilepsia; aivot; hippokampus; verisuonet; hermosolut; plastisuus;verkostoituminen; koe-eläimet

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To my family

“The only true wisdom is knowing that you know nothing.”

Socrates

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AcknowledgementsThis study was carried out at the Epilepsy Research Laboratory and the Biomedical NMRlaboratory at the A.I Virtanen Institute for Molecular Sciences, University of EasternFinland during the years 2006-2012

I am sincerely and heartily grateful for my supervisor, Professor Asla Pitkänen MD,PhD, the head of the epilepsy research laboratory, for introducing me into the world ofepilepsy research and neuroscience in general. She has been a constant source of guidance,encouragement and support throughout my study and stay in Finland. Her constantsupply of fresh ideas and willingness to sit with us at the microscope, my favoritemoments in our interaction, made this work possible. Furthermore, I was constantlymotivated and inspired by her extensive knowledge, creative thinking and persistence.She has clearly demonstrated to me the level of professional standard and perfection Ishould be aiming at in my carrier and in other things that I do.

I would also like to extend my sincere appreciation to my second supervisor,Professor Olli Gröhn, PhD for his constant support and advice, on and of the pitch,especially during those moments where his expertise in NMR was solicited.

I owe sincere and earnest thankfulness to Professor Eleonora Aronica, MD, PhDand Docent Irma Holopainen, MD, PhD, the official reviewers for this thesis, for theirexpert comments and criticism, which helped to improve this thesis. I would like to thankProfessor Leszek Kacmarek for accepting our invitation to oppose this thesis.

I am obliged to my colleagues, former and present members of “Epiclub”: HeliMyöhänen, Tamuna Bolkvadze, Nino Kutchiashvili, Sofya Ziyatdinova, Noora Huusko,Olena Shatillo, Diana Miszczuk, Jukka Rantala, Laura Lahtinen, Nick Hayward, HeliKarhunen, Christine Einula, Cagri Yalgin and Annakaisa Haapasalo. Their warmfriendship, cooperation and understanding, took off some pressure from my work. I wantto extend special thanks to Jari Nissinen, Merja Lukkari and Jarmo Hartikainen, yourtechnical and loving support was a vital element in the success of this work.

I like to express my deepest gratitude to Jari Nissinen, Noora Huusko, and JukkaJolkonen, for helping to translate my abstract to Finnish. I extend a very big gratitude toMerja, for helping in the planning and organization of the “karonkka”.

I am truly indebted and thankful to my co-authors, Filip Barinka, Yumiko Akamine,Mohammed Hossein Esmaeli and the entire NMR group, especially those who helpedNick one way or the other during the NMR analysis.

I will like to thank the entire personnel of the A.I Virtanen Institute for MolecularScience for proving a pleasant and cooperative working environment. I extend specialthanks to Sari Koskelo and all the other past and present personnel of the office of A.IVirtanen Institute for the great help in practical issues over the years.

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I wish to thank Professor Hilkka Soininen, all the members of the BiND program,and the entire University of Eastern Finland for the BiND initiative. Special thanks to DrPauliina Korhonen, BiND coordinator, you were more than a coordinator, some kind ofguardian angel looking over us. Thanks to Mari Tikkanen for all the help with the BiNDrelated work. I am grateful to Dr Mikko Hiltunen, for the great advice he gave me alongthe way, especially at those moments when I needed them most. I can never say thank youenough to all the BiND fellows, Laku, Jaya, Gabriela, Alina and all the others who left usalong the way; you were all like a family.

I will like to thank all my friends here in Finland, especially those from Cameroon,Eta Valerie, Foncham Simon, Ako Collins, Ebote Daniel, and their families, who made mefeel like I was always at home. Special thanks to Karlhans Fru Che and Ngang ChiCelestine, whom, we started this journey together. The company you all provided hasbeen valuable to me.

I will always remain indebted to Sandra, your loving company, support and carewas a vital element in the successes of this thesis; especially during those moments when Ithought the dark tunnel was never ending, you were able to keep my focus on the light atthe end of it. Am also grateful to your family in Madrid for their kind, warm andwelcoming heart, especially during those times when I needed a break from my work.

Thank you is not enough to express my sincere and heartily gratefulness to DrHalle Gregory Ekane, without whom my entire stay and study abroad would not havebeen possible. Doc you have always been a constant source of inspiration, admiration andsupport to me. I can’t say how much your one-to-one guidance and support throughoutmy life have helped shaped who I am. You will forever remain a role model to me. ThanksDoc!

Finally, I will like to thank my entire family, Halle, Ekane, Diabe, Nzelle (of late),Sone, Akame, Ndode and most especially my dad, Mr Ekane Micheal Enongene and mom,Mrs. Elizabeth Ekane, not forgetting their grand children, for their constant unconditionallove and support throughout my life. You have all been a wonderful blessing to me.

This project was supported by the EU funded BiND program PhD position EC FP6,MEST-CT-2005-019217, The Nordic Center for Excellence, The Health Research Council ofthe Academy of Finland, The Sigrid Juselius Foundation, The Finnish Epilepsy FoundationThe Finnish Graduate School of Neuroscience, and The North Savo Regional Fund of theFinnish Cultural Foundation

Kuopio, November 2012


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List of the original publications

This dissertation is based on the following original publications:

I Ndode-Ekane XE, Hayward N, Grohn O, Pitkanen A (2010). Vascular changes in

epilepsy: Functional consequences and association with network plasticity in

pilocarpine-induced experimental epilepsy. Neuroscience. 166:312-332.

II *Lahtinen L, *Ndode-Ekane XE, Barinka F, Akamine Y, Esmaeili MH, Rantala J,

Pitkanen A (2010). Urokinase-type plasminogen activator regulates

neurodegeneration and neurogenesis but not vascular changes in the mouse

hippocampus after status epilepticus. Neurobiol. Dis. 37:692-703.

III Ndode-Ekane XE and Pitkänen A (2012). Urokinase-type plasminogen activator

receptor modulates epileptogenesis in mouse model of temporal lobe epilepsy.

Submitted for publication

* Shared 1st authorship. Neurodegeneration and neurogenesis are not included in this

thesis.

The publications were adapted with the permission of the copyright owners.

XV

Contents

1. INTRODUCTION 1

2. REVIEW OF THE LITERATURE 3

2.1 Epilepsy 32.1.1 Definition of epileptic seizure and epilepsy 32.1.2 Classification of seizures and epilepsies 32.1.3 Temporal lobe epilepsy (TLE) and the concept of epileptogenesis 32.1.4 Status epilepticus (SE) 4

2.2 Neuropathology of temporal lobe epilepsy 52.2.1 Hippocampal neurodegeneration 52.2.2 Granule cell dispersion (GCD) 62.2.3 Gliosis 62.2.4 Mossy fiber sprouting 72.2.5 Neurogenesis 82.2.6 Inflammation 82.2.7 Angiogenesis and blood-brain barrier (BBB) disruption 9

2.3 Experimental models of temporal lobe epilepsy 102.3.1 Post-status epilepticus models 10

2.3.1.1 Kainic acid (KA) Model 112.3.1.2 Pilocarpine Model 112.3.1.3 Electrical Stimulation-induced Model 12

2.4 The plasminogen activator system 122.4.1 Plasminogen/ Plasmin 122.4.2 Urokinase-type plasminogen activator (uPA) 142.4.3 Tissue type plasminogen activator (tPA) 152.4.4 Urokinase type plasminogen activator receptor (uPAR) 162.4.5 Plasminogen activator inhibitor (PAI) 172.4.6 The expression and role of elements of the plasminogen system in epilepsy 18

3. AIMS OF THE STUDY 23

4. MATERIALS AND METHODS 25

4.1 Animals 25

4.2 Anesthesia 25

4.3 Induction of status epilepticus (SE) 264.3.1 Pilocarpine model (I) 264.3.2 Intrahippocampal kainic acid model (II, III). 26

4.4 Electrode implantation 26

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4.4.1 Cortical electrodes (II, III) 264.4.2 Hippocampal electrodes (III) 26

4.5 Video-EEG monitoring (II, III) 274.5.1 Detection and analysis of severity of status epilepticus (II, III) 274.5.2 Detection and analysis of electrographic and behavioral seizures (II, III) 27

4.6 BrdU administration for study of endothelial cell proliferation (I) 28

4.7 Magnetic resonance imaging measurement of cerebral blood flow (CBF) andcerebral blood volume (CBV) (I) 28

4.8 Processing of brain for histology 294.8.1 Transcardiac perfusion - Timm fixation protocol (I, III) 294.8.2 Transcardiac perfusion - Para-formaldehyde fixation protocol (I-III) 294.8.3 Sectioning of brain 29

4.9 Histology 304.9.1 Nissl staining and analysis of neuronal damage (I - III) 30

4.9.1.1 Analysis of damage to principal cells (I, II, III) 304.9.1.2 Analysis of damage to hilar cells (II, III) 304.9.1.3 Analysis of granule cell layer volume (II, III) 31

4.9.2 Timm histochemistry and assessment of mossy fiber sprouting (I, III) 314.9.3 Doublecortin (DCX) immunohistochemistry and analysis of neurogenesis (I-III) 324.9.4 IgG immunohistochemistry and analysis of blood-brain barrier damage (I, III) 324.9.5 Thrombosis immunohistochemistry and assessment of thrombocyte (I) 334.9.6 CD68 immunohistochemistry and assessment of macrophage response (III) 334.9.7 CD3 immunohistochemistry and assessment of T cell response (III) 344.9.8 Blood vessel immunohistochemistry (I, II, III) 35

4.9.8.1 Rat endothelial cell antigen-1 (RECA-1) and endothelial barrier antigen(EBA) immunohistochemistry (I) 35

4.9.8.2 Lycopersicon esculentum (tomato) lectin staining (II, III) 354.9.8.3 Assessment of total blood vessel length (I, II, III) 354.9.8.4 Assessment of Blood vessel diameter (I) 36

4.9.9 BrDU-RECA-1 double staining and analysis of endothelial cell proliferation (I) 36

4.10 Statistical analysis (I, II, III) 37

5. RESULTS 39

5.1 Status epilepticus and seizure phenotype in uPA-/- and uPAR -/- mice (II, III) 395.1.2 Severity of status epilepticus (SE) (II, III) 395.1.2 Seizure phenotype characteristics (II, III) 395.1.3 Severity of behavioral seizures (III) 39

5.2 Neurodegeneration in the hippocampus (I, III) 405.2.1 Degeneration of principal cells 405.2.2 Degeneration of hilar neurons 405.2.3 Degeneration and dispersion of granule cells 41

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5.3 Neurogenesis in the dentate gyrus (I, III) 41

5.4 Mossy fiber sprouting in the dentate gyrus (I, III) 42

5.5 Blood-brain barrier impairment in the hippocampus (I, III) 42

5.6 Inflammatory response in the hippocampus (III) 43

5.7 Hippocampal vascular morphometry (I, II, III) 435.7.1 Endothelial cell proliferation, blood vessel length and diameter change (I, II, III) 435.7.2 Association of angiogenesis and hippocampal CBV and CBF (I) 445.7.3 Association of angiogenesis with neurodegeneration, neurogenesis and mossy

fiber sprouting (I) 445.7.4 Effect of uPA or uPAR deficiency on vascular morphometry (II, III) 44

6. DISCUSSION 47

6.1 Methodological considerations 47

6.2 Epilepsy phenotype in uPA and uPAR deficient mice 48

6.3 Neurodegeneration is affected by uPA and uPAR 49

6.4 uPA and uPAR are involved in neurogenesis 50

6.5 Angiogenesis is affected by uPAR but not by uPA 51

6.6 Association between angiogenesis and neuronal plastic events 53

6.7 Inflammation is modulated in uPAR deficient mice 54

7. CONCLUSIONS 57

8. REFERENCES 59

APPENDIX: Original publication I-III

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Abbreviations

Ach acetylcholineAD Alzheimer’s diseaseAMPA alpha-amino-3-hydroxyl-5-

methyl-4isoxazole-propionate

AP anterior posteriorasf area sampling fractionATF amino-terminal fragmentBBB blood-brain barrierBDNF Brain-derived neurotrphic

factorBrdU BromodeoxyuridneCA1 Cornu Ammonis 1 region of

the hippocampusCA3 Cornu Ammonis 1 region of

the hippocampusCBF cerebral blood flowCBV cerebral blood volumeCNS central nervous systemDCX doublecortineDV dorso-ventralEBA endothelia barrier antigenECM extracellular matrixEEG electroencephalographyEGF endothelial growth factorGCD granule cell layer dispersionGCL granule cell layerGPI glycosylphosphatidylinositolGPR G-protein coupled receptorHIV Human immunodeficiency

virusi.h. intrahippocampali.p. intraperitoneali.v. intraventricularIL1� interleukine 1 betaILAE International league Against

EpilepsyKA kainic acid

KPBS potassium sodiumphosphate buffer solution

LPR low-density lipoproteinreceptor

LPS lipopolysacharideMHC major histocompartability

complexML medial- lateralmRNA messenger ribonucleic acidMRI magnetic resonance imagingNF�B nuclear factor kappa BNHS normal horse serumNGS normal goat serumNIH national institutes of healthNMDA N-methyl-D-aspartatePA Plasminogen activatorPAI plasminogen activator

inhibitorPB sodium phosphate bufferPN1 protein nexin 1PTZ pentylenetetrazolRECA rat endothelial cell antigenROD relative optical densityROI region of interests.c. subcutaneousSE status epilepticusSERPIN serine protease inhibitorSGZ subgranular zoneSPD serine protease domainSSSE self-sustained status

epilepticusssf section sampling fractionSRPX2 sushi repeat protein x-linked

2SVZ subventricular zoneTBI traumatic brain injuryTGF-�1 transforming growth factor

beta-1

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TLE temporal lobe epilepsyTLR2 toll-like receptor 2TLR4 toll-like receptor 4TNF� tumour necrotic factor-alphatsf tissue sampling fractiontPA tissue-type plasminogen

activatoruPA urokinase-type plasminogen

activatoruPAR urokinase-type plasminogen

activator receptorVEGF vascular endothelial growth

factorVideo-EEG video-

electroencephalographyVN vitronectin

1. Introduction

Epilepsy is one of the most common neurological disorders, affecting approximately 1% ofthe world’s population. Temporal lobe epilepsy (TLE) is the most common form ofsymptomatic epilepsy (Engel, 1996b). Patients with TLE have usually had an initialprecipitating brain injury (initial insult) earlier on in life, which can be in the form of statusepilepticus (SE), head trauma, a stroke, tumor or infection. The initial precipitating insultis the first phase in the development of TLE. The next phase, the latency phase, is void ofsigns and symptoms of disease. However, during this phase there are cellular andmolecular reorganization processes going on in the brain including neurodegeneration,gliosis, neurogenesis, inflammation, gene expression, axonal plasticity, secretion of growthfactors and angiogenesis, which eventually make the brain prone to seizures. In the lastphase of the disease, the patient starts experiencing spontaneous seizures. During thisphase, there is still ongoing tissue remodeling (Pitkänen and Lukasiuk, 2009). Thesecellular reorganization events are well characterized, and their severity is associated withthe severity of the epilepsy (Pitkänen et al., 2007).

Epileptogenic cellular reorganization processes involve the cell-extracellular matrix(ECM), cell-cell interaction; and cell signaling. The urokinase-type plasminogen activator(uPA) and its receptor (uPAR), elements of the plasminogen activator (PA) system, haveimplications in these sorts of interactions and signaling events (Sidenius and Blasi, 2003).The uPA and uPAR play a role in physiological and pathological events that require tissueremodeling (Floridon C. et al., 1999; Smith and Marshall, 2010). Interestingly, there is anincreased expression of uPA and uPAR in the hippocampus during epileptogenesis(Lahtinen et al., 2006, 2009; Iyer et al., 2010; Liu et al., 2010). This suggests that uPA-uPARmay be involved in tissue reorganization during epileptogenesis. There is a growinginterest in understanding the molecular mechanisms underlying these epileptogenicnetwork alterations. Elucidating these mechanisms may provide new biomarkers andtreatment targets for epilepsy.

The aim of this thesis is to elucidate the interactions and functional implications ofthe cellular reorganization processes during epileptogenesis. The study focuses,particularly, on deciphering the relationship between angiogenesis and neuronal networkalterations such as neurodegeneration, neurogenesis and axonal sprouting in thehippocampus, since common trophic stimuli regulate vascular and nerve networkformation (Carmeliet and Tessier-Lavigne, 2005). In addition, the role of uPA and uPAR asa possible mechanism underlying epileptogenic network alterations was investigated. Thiswas studied effectively using uPA and uPAR deficiency mice in a mouse model oftemporal lobe epilepsy.

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2. Review of the literature

2.1 EPILEPSY

2.1.1 Definition of epileptic seizure and epilepsyAccording to the International League Against Epilepsy (ILAE) and the InternationalBureau for Epilepsy, an epileptic seizure is a transient occurrence of signs and/orsymptoms due to abnormal excessive or synchronous neuronal activity in the brain (ILAE,Fisher et al., 2005). Epilepsy is a disorder of the brain characterized by an enduringpredisposition to generate epileptic seizures and by neurobiologic, cognitive, physiologicaland social consequences of this condition. The definition of epilepsy should include ahistory of at least one seizure (ILAE, Fisher et al., 2005).

2.1.2 Classification of seizures and epilepsiesSeizures are classified as either partial or generalized seizures (ILAE, 1981). Partialseizures involve a network limited to one hemisphere and generalized seizures arewidespread, involving bilateral cortical networks (ILAE, 1981).

The classification of epilepsies and epilepsy syndromes is based on clinical featurestogether with appropriate EEG and brain imaging investigations. Epileptic syndromes areeither focal epilepsies (partial-onset seizures) or generalized epilepsies (generalizedseizures) (ILAE, 1989). However, as a result of fast technological and scientific advances,there is a growing concern for revising the concepts, terminology and approaches forclassifying seizures and epilepsies. The ILAE's Commission on Classification andTerminology has made proposals which will incorporate these advances (for details seeBerg et al., 2010).

2.1.3 Temporal lobe epilepsy (TLE) and the concept of epileptogenesisOften referred to as limbic epilepsy, TLE is the most common symptomatic epilepsy(Engel, 1996b). The seizures arise from the temporal lobe structures or limbic system,which includes the hippocampus, amygdala, subicular complex, entorhinal andparahippocampal cortices (Engel, 1996a). The habitual seizures begin with an aurafollowed by other symptoms referable to the limbic system, including epigastric rising,emotional change (typically fear) and occasionally olfactory or gustatory hallucinations(Williamson and Engel, 1997).

Patients with TLE usually have a history of febrile seizures or initial precipitatinginjuries such as a head trauma, infection, stroke, tumor or SE early on in life, suggestingthat the development of TLE begins with a brain insult (Mathern et al., 1995, 1997). The

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initial brain insult comprises the first phase in the development of TLE. The second phase(epileptogenesis or the latency phase), lasts from several months to years. It is void ofseizures but is characterized by tissue remodeling changes that make the brain prone todevelop seizures. In the final phase, epileptogenic network reorganization continues afterepilepsy diagnosis (Jutilia et al., 2002; Pitkänen and Lukasiuk, 2009). The process ofepileptogenesis is illustrated in figure 1.

Hippocampal sclerosis is a typical, pathological hallmark often associated withTLE. It has been described in 60-70% of patients with intractable TLE. Its characteristicsinclude loss of pyramidal neurons and gliosis in the CA1 and CA3 subfields of thehippocampus (Jutilia et al., 2001; Kälviäinen and Salmenperä, 2002). In addition, there isdentate hilar cell loss and extrahippocampal pathology in the amygdala and entorhinalcortex including the thalamus (Blumcke et al., 2002; Wieser et al., 2004).

Current treatment of epilepsy focuses on the prevention of seizures. Followingadequate medical treatment, patients can become seizure free. However, a handful ofthem, about 30%, become refractory (Hauser and Hesdorffer 1990). A subgroup of patientsmay obtain relief from epilepsy surgery by resecting the epileptic focus (Bertram, 2009).

Figure 1. The process of epileptogenesis in symptomatic epilepsy. The initial brain insult (headtrauma, infection, status epilepticus, or stroke) initiates epileptogenesis characterized byseveral cellular and molecular reorganization processes, leading ultimately to the appearanceof spontaneous recurrent seizure, i.e. epilepsy (modified from Pitkänen and Sutula, 2002).

2.1.4 Status epilepticus (SE)Status epilepticus is a single clinical seizure or repeated seizures lasting more than 30minutes without intervention or recovery of consciousness (ILAE, 1989; Waterhouse andDelorenzo, 2001). However, seizure activity persisting more than 5 minutes is consideredto be SE and is treated accordingly (Alldredge et al., 2001). According to epidemiologicalstudies in Switzerland, Germany, Italy, and among the white population of the USA, SE

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incidence is about 10-20/100,000 (see review Rosenow et al., 2007). The most common causeof SE in children is febrile seizures or infections, while in adults it is cerebrovascularaccidents, hypoxia, metabolic causes and low antiepileptic drug levels (DeLorenzo et al.,1996; Knake et al., 2001).

The risk of developing epilepsy after pediatric convulsive SE is within a wide rangeof 13-74% (Raspall-Chaure et al., 2006). The risk is 25-40% after 2 years following the firstunprovoked SE (Hesdorffer et al., 1998; Eriksson and Koivikko, 1997). In adults, the risk ofdeveloping epilepsy after SE is about 42%, 10 years later (Hesdorffer et al., 1998). Lowerrisk in children is due to the higher resistance of the juvenile brain against neuronaldamage (Sperber EF et al., 1991). In experimental settings, regardless of the model used(pilocarpine, kainic acid or electrical stimulation), about 50-100% of animals developspontaneous seizures (Hellier et al., 1999; Holtkamp et al., 2005; Goffin et al., 2007).

2.2 NEUROPATHOLOGY OF TEMPORAL LOBE EPILEPSY

2.2.1 Hippocampal neurodegenerationHippocampal neurodegeneration is the most common lesion in patients with temporallobe epilepsy. 50-70% of patients with medically intractable limbic epilepsy havehippocampal neurodegeneration (Honavar and Meldrum, 1997). In hippocampalneurodegeneration, there is loss of neurons in the hippocampal pyramidal cell layer (CA1,CA2 and CA3) and the dentate hilus. There is relative preservation of the CA2 neuronsand the dentate granule cells. Often neuronal loss is accompanied by dense gliosis(Blumke et al., 2002; Majores et al., 2007). In conjunction with hippocampalneurodegeneration are neuronal loss in the neighboring entorhinial cortex and theamygdala (Dawodu and Thom, 2005; Pitkänen and lukasiuk, 2009). Severe seizures and SEinduce neuronal cell death in the hippocampus within hours of seizure onset (Hauser,1983). The question still remains whether hippocampal neurodegeneration is theconsequence of repeated seizures or whether it plays a role in the development of epilepticfocus (Jefferys, 1999; Berkkovic and Jackson, 2000).

Animal models of human TLE also demonstrate hippocampal neurodegeneration.In particular, animal models of SE show that hippocampal excitatory principal cells of theCA1 and CA3 subfields and hilar neurons of the dentate gyrus are relatively sensitive toseizure-induced death. These models also demonstrate that dentate granule cells arerelatively resistant to damage (Franck, 1984; Sloviter, 1987; Morimoto et al., 2004). Like inhumans, neuronal cell death is seen in the amygdala, entorhinal, perirhinal andparahippocampal cortices. Dying neurons can be seen a few hours after SE andneurodegeneration may continue for up to 2 months (Pitkänen and Sutula, 2002).

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2.2.2 Granule cell dispersion (GCD)Granule cell dispersion (GCD) is a morphological hallmark observed in the sclerotichippocampus of TLE patients (Houser, 1990). In GCD, there is a loss of close appositionbetween the granule cell soma, enlargement of the stratum granulosum and the presenceof granule cells scattered in the molecular layer. About 40% of sclerotic hippocampalspecimens from TLE patients show GCD (Thom et al., 2002). The extent of granule celllayer dispersion correlates with the amount of neuronal cell death in the hippocampus andhilus of the dentate gyrus (Houser, 1990, Thom et al., 2002). Lurton and colleagues (1998)found that GCD was more likely to occure in TLE patients with early epileptic events,arising during the first 4 years of life, and is not associated with the duration or number ofseizures later on in life. However, animal studies have so far failed to show anyconnection between GCD and seizure activity. The mechanisms inducing thesemorphological changes and to what extent GCD contributes to seizure activity remainsunknown. However, current reports suggest reelin dysfunction causes GCD dispersion,since reelin is essential for the maintenance of layered structures in an adult brain andreelin deficiency is correlated with GCD in TLE patients (see review by Haas andFrotscher, 2010).

2.2.3 GliosisGliosis refers to the proliferation and activation of glial cells. It is a prominenthistopathological hallmark associated with TLE. It is only recently that its role in thedevelopment of epilepsy was fully appreciated (Yang et al., 2010). There are four majortypes of glia cells involved in gliosis: astrocytes, microglia, oligodendrocytes and NG2cells (Polydendrocytes). These cells play diverse roles in the normal and pathologicalbrain. However, astrocytes and microglia are the most widely studied and also the cellsmost implicated in TLE pathology.

Astrocytes constitute 20-50% of the volume in most brain areas (Bruzzone andGiaume, 1999). Astrocytes end-feet make contact with blood vessels and thus are anessential element of the blood-brain barrier (BBB) and as such control cerebral blood flow(Heneka et al., 2010). They also make contact with other cell types including neurons.Some of the most widely investigated roles of astrocytes include regulation of extracellularpotassium, glutamate uptake and synaptic glutamate concentrations, metabolism andsynthesis of various molecules. They also play a crucial role in synaptic transmission andseizure as well as in inflammation (Vezzani et al., 2007; Wetherington et al., 2008).

In a healthy brain, microglia are constantly surveying their environment(Nimmerjahn et al., 2005). In this way they are able to intervene in a host of events such assynaptic pruning during postnatal development (Katz and Shatz, 1996; Hua and smith,

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2004; Huberman et al., 2008). Other functions of microglia in a healthy brain includephagocytosis of apoptotic newborn neurons, regulation of neuronal activity by interactionwith synapses and astrocytes, and reorganization of neuronal circuits (see review byNapoli and Neumann, 2009). Most investigations on microglial function in the last decadeshow that they are a key player in brain injury and disease. In response to a brain injury,microglia become activated, proliferates, migrate or secrete compound into theextracellular matrix in a process generally termed microgliosis (Hailer, 2008). Studiesusing animal models show that microglial activation peaks at 3-5 days after a brain injuryand can remain elevated for several weeks (Jorgensen et al., 1993; Hailer et al., 1999). Someof the compounds secreted by microglia may be harmful to neurons while others may bebeneficial. Some of the harmful compounds include proinflammatory cytokines(interleukin-1, interleukin-6, tumor necrotic factor �), proteases and nitric oxide(Benveniste, 1992; Nathan, 1992; Kreutzberg, 1996). Secreted neuroprotective compoundsinclude transforming growth factor �, brain-derived neurotrophic factor (BDNF) and thenerve growth factor. Other beneficial actions of microglia include clearing the dyingneurons and glia. The net effect of the harmful and beneficial actions of microglia on thedevelopment and progression of TLE remains to be studied.

2.2.4 Mossy fiber sproutingSprouting of granule cell axons or mossy fiber is one of the most extensively studiedfeatures of human TLE (Mathern et al., 1996; Sutula and Dudek, 2007). Animal models ofepilepsy also demonstrate this feature, particularly models that utilize prolonged SE forthe induction of chronic epilepsy (Pitkänen et al., 2007). Mossy fiber sprouting refers tochanges in axonal projection of dentate granule cells. Normally, granule cell axonsinnervate hilar interneurons or the apical dendrites of the CA3 pyramidal neurons. Inhuman and animal models of TLE, mossy fiber forms synapses with granule cell dendritesin the inner molecular layer and the basal dendrites of CA3 pyramidal cells of thehippocampus (Represa et al., 1990). Experimental studies show that sprouting occursbefore spontaneous seizures and stays throughout the life-time of experimental animals(Nissinen et al., 2001).

The functional consequence of mossy fiber sprouting remains controversial. Earlierreports suggest that mossy fiber creates monosynaptic excitatory loops that contribute torecurrent excitation and thus enhances susceptibility to seizures (Sutula and Dudek, 2007).However, a recent study by Buckmaster and Lew (2011) shows that suppression of mossyfiber sprouting does not reduce seizure frequency, thus suggesting that mossy fibersprouting is neither pro- nor anticonvulsant. Their finding corroborates with earlier claimsby Harvey and Sloviter (2005) that granule cells become progressively less excitable rather

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than hyperexcitable as mossy fiber sprouting progresses, and granule cells do not initiatespontaneous behavioral seizures.

2.2.5 NeurogenesisErikson and colleagues (1998) provided the first evidence that neurogenesis occurs in theadult human brain. In adult human and rodent brains, neurogenesis occurs continuouslyin the subventricular zone (SVZ) and the subgranular zone (SGZ) (Eriksson et al., 1998;Benraiss et al., 2001; Bengzon et al., 1997). Other than normal physiological neurogenesis,several stimuli can increase neurogenesis in experimental animals. Examples includeexercise, enriched environment, adrenal steroids, and glutamatergic neurotransmission(Gage et al., 1998; Brown et al., 2003).

Despite the difficulty in labeling newly generated neurons in humans with TLE,evidence of neurogenesis was demonstrated in specimens from patients with TLE usingmarkers labeling neural progenitors and immature neurons (Blümcke et al., 2001;Hermann et al., 2006). Several experimental animal studies show compelling evidence ofseizure-promoting effects on neurogenesis. Increased neurogenesis occurs in a variety ofanimal models of epilepsy, including rapid animal kindling (Parent et al., 1998; Scott et al.,1998; Smith et al., 2005), electroconvulsive seizures (Segi-Nishida et al., 2008), the flurothylkindling model (Ferland et al., 2002) and pentylenetetrazol seizures (Jiang et al., 2003).

It is unclear how these newly generated neurons integrate in the local neuronalnetwork and whether they are anti- or pro-epileptogenic. A majority of the newly bornneurons migrate into the granule cell layer. Following prolonged seizure events somenewly generated neurons migrate aberrantly into the hilus and molecular layer anddifferentiate into ectopic dentate granular cells (Parent et al., 2006). Recently, Kron et al.(2010) showed that newly generated neurons are vulnerable to seizure-induced abnormalplasticity. They also showed that these neurons may contribute to epileptogenic networkdysfunction. Despite multiple experiments on seizure-induced neurogenesis in animalmodels, the role of this phenomenon in ictogenesis and epileptogenesis remains unclear.

2.2.6 InflammationChronic brain inflammation, which comprises the activation of microglia, astrocytes,endothelial cells of the BBB, peripheral immune cells and the production of inflammatorymediators, is a prominent feature in human TLE and animal models (Zimmer et al., 1997;Bien et al., 2007). Several inflammatory mediators, including NF�B, interferons,interleukins, chemokines, adhesion molecules, and toll-like receptor 4 (TLR4) are reportedin brain tissue from patients with TLE (see review by Choi and Koh, 2008). Recent

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evidence, especially from experimental animal models indicates that inflammation mightbe a consequence as well as a cause of epilepsy.

Experimental animal models of epilepsy are providing information on the role ofinflammation in seizures and the development of epilepsy. In these models, acute seizurescause activation of microglia and astrocytes, increase expressions of inflammatory relatedtranscription factors and cytokines such as IL1�, TNF-�, TGF-�1, and neuropeptidemRNAs (Zimmer et al., 1997; Palata-salaman et al., 2000). Turrin and Rivest (2004) showedthat following pilocarpine induced status epilepticus, the expression of toll-like receptor 2(TLR2) in microglia increases, leading to the activation of cytokines and chemokines, andMHC class I and II molecules. Evidence also suggests that activated glia and elevatedcytokines can in turn contribute to seizure-related hippocampal pathology such asneuronal death, neurogenesis, gliosis and mossy fiber sprouting (McNamara et al., 1994;Parent and Lowenstein, 1997; Koh et al., 1999; Jankowsky and Patterson, 2007).Experimental evidence suggests that inflammation can lead to neuronal excitability andneuronal injury either directly, by interacting with glutamatergic neurotransmission orindirectly, by activation of gene transcription (Viviani et al., 2003; Ravizza et al., 2008).

2.2.7 Angiogenesis and blood-brain barrier (BBB) disruptionAngiogenesis is the process that refers to the growth of new blood vessels from existingones by the proliferation of endothelial cells. Angiogenesis occurs during physiologicaland pathological conditions. In the brain, blood capillaries are composed of endothelialcells that line together forming a luminal structure and are connected by tight junctions.The abluminal surface of the endothelial cells is covered with a basement membrane,pericytes, and astrocytes’ end-feet, together forming the BBB. The BBB protects the brainfrom toxins in the blood while allowing essential metabolites to cross. Disruption of theBBB and subsequent angiogenesis occurs in several central nervous system (CNS) diseasepathologies, including traumatic brain injuri (TBI), strokes and human TLE (Morgan et al.,2007; Rigau et al., 2007; Tang et al., 2007). However, the question still remains whethervascular remodeling contributes to the pathogenesis of these diseases including epilepsy.

There is evidence suggesting an association between angiogenesis and seizuresusceptibility in experimental animal models of status epilepticus (Rigau et al., 2007). Also,electroconvulsive seizures can cause endothelial cell proliferation and increase vasculardensity in the rat hippocampus (Hellsten et al., 2005). These evidences indicate that bothepileptogenic brain injuries and brief seizures can trigger an increase in brain vascularity.Angiogenesis may be relevant for optimal functional recovery in an epileptogenic brain,providing oxygen and nutrients needed for tissue remodeling. However, questions remain

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concerning how this phenomenon is related to seizure frequency or patterns of neuronalplasticity that favor hyperexcitability.

There are many molecules that affect the development of both the vascular systemand the nervous system. Striking evidence stems from the recent recognition that classicalaxon-guidance cues, such as netrin, semaphorins, slits and ephrins also mediate thenavigation of blood vessels along predestined tracts during development (Carmeliet andTessier-Lavigne, 2005). Similarly, the angiogenic factor, the vascular endothelial growthfactor (VEGF), regulates the migration of various neuron types to their final destination(Schwarz et al., 2004). This evidence suggests that in the presence of such trophic stimulione would expect an association between angiogenesis and neuronal plastic events.However, it is yet to be established whether there is any interaction or association betweenthese processes during epileptogenesis.

BBB disruption is a prominent feature in experimental and human epilepsy (Sokrabet al., 1989; Seiffert et al., 2004). Leakage of serum-derived components into the brainparenchyma is associated with increased excitability and the occurrence of seizures(Seiffert et al., 2004). It was recently suggested that BBB impairment can lead toepileptogenesis and contribute to the progression of epilepsy (van Vliet et al., 2007). Eventhough BBB damage can lead to seizures and vice versa, it is yet to be demonstratedwhether this is a matter of causality or co-occurrence in epileptogenesis/ictogenesis

2.3 EXPERIMENTAL MODELS OF TEMPORAL LOBE EPILEPSY

The use of experimental animal models of TLE is with the sole aim of elucidating thefundamental mechanisms of epilepsy essential for devising new diagnostic, therapeuticand preventive approaches to human epilepsy. There are many epilepsy syndromescharacterized by the occurrence of one or more specific seizure type as well other clinicalfeatures. The current challenge is producing an animal model that reproduces any givenhuman epilepsy syndrome. So far the use of animal models in experimental epilepsy islimited to the study of particular aspect(s) of the epilepsy disorder.

2.3.1 Post-status epilepticus modelsAll post-SE animal models have in common at least three major stages: (1) the initialepisode of SE, (2) a seizure-free period of days to weeks and (3) the recurrent spontaneousseizure stage.

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2.3.1.1 Kainic acid (KA) ModelKA is a cyclic analogue of L-glutamate and an agonist of ionotropic, non-NMDAglutamate AMPA and KA receptors. It is commonly administered by intraperitoneal (i.p.),subcutaneous (s.c.), intravenous (i.v.) or intrahippocampal (i.h.) injection to causesustained neuronal depolarization, SE, lasting for several hours (Nadler et al., 1978; Ben-Ari, 1985; Longo and Mello, 1998). The receptors for KA are highest in density in thehippocampus, especially in the CA3 region, amygdala, perirhinal and entorhinal cortices(Miller et al., 1990). These areas are preferentially affected after KA administration andthus KA serves as a valuable model for partial seizures with secondary generalization(Ben-Ari, 1985; Engel, 2001).

Some of the electroencephalographic and behavioral alterations caused by KA aresimilar to those observed in human patients with TLE (Lothman and Collins, 1981;Krumholz et al., 1995). KA-induced SE models in rats and mice also mimic some of theneuropathological features observed in human TLE. These include loss of neurons in thehippocampal CA1 and CA3 subfields and mossy fiber sprouting (Lothman and Collins,1981; Okazaki et al, 1995; Sloviter, 1996; Chakravarty et al., 1997). However, neuronalnecrosis has also been reported in the piriform and olfactory cortices, amygdala, thalamusand neocortical layers III and VI (Ben-Ari et al., 1979).

2.3.1.2 Pilocarpine ModelPilocarpine is a muscarinic acetylcholine (Ach) receptor agonist. Systemic (i.p) injection ofpilocarpine in rats and mice produces seizures (Turski et al., 1984; Fujikawa, 2003). Itsseizure producing effect arises from the activation of muscarinic Ach receptors expressedespecially in the hippocampus, striatum and cortex (Kuhar and Yamamura, 1976). A dosewithin the range of 300 to 400 mg/Kg is needed to produce SE. This very high doseproduces significant peripheral effects; thus peripheral muscarinic antagonists such asscopolamine methylbromide (1 mg/kg, s.c.) is used to reduce these effects.

Behavioral alterations following pilocarpine injection include facial automatism,salivation, piloerection and behavioral automatism (Cavalheiro et al., 2006). Followingpilocarpine injection, the seizures are normally allowed to progress for at least 90 minutesbefore they are stopped using diazepam to reduce mortality. The first spontaneousepileptic seizures occur approximately 2 to 75 days after pilocarpine injection (for detailssee Nissinen, 2006). Some of the neuropathological alterations are similar to that producedby KA. However, unlike in the KA model, these lesions are more prominent in theneocortex than in the hippocampus (Schmidt-Kastner and Ingvar, 1996).

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2.3.1.3 Electrical Stimulation-induced ModelThis model is used less often as a model of TLE compared to KA and pilocarpine models.However, it produces clinical signs and lesions quite close to those produced bypilocarpine and KA models. Induction of self-sustained SE (SSSE) is achieved by electricalstimulation of a brain region with a 10-sec train of 1–msec square wave monophasicstimuli at 20 Hz delivered every minute (total of 30 trains) over a period of 30 minutes(Vicedomini and Nadler, 1987). SE is generated when the perforant path, hippocampus oramygdala is stimulated (Mazarati et al., 2004; Bastlund et al., 2005; Tilelli et al., 2005).

The behavioral features produced by this model depend on the stimulatedstructure. Some of the neuropathological findings as depicted by Nissinen and colleagues(2000) after stimulation of the amygdala included hippocampal damage in 67% of epilepticrats, characterized by cell loss in the hilus, CA1 and CA3 subfields of the hippocampus.They also reported that 87% of the animals develop epilepsy within 6 months after SE. Amajor advantage of this model is that it is free of chemical substances and the latencyperiod is long compared to the KA or pilocarpine model.

2.4 THE PLASMINOGEN ACTIVATOR SYSTEM

The plasminogen activation system is an enzymatic cascade which plays an important rolein various biological processes involving extracellular matrix proteolysis. It plays twoprincipal roles. First, it is the central pathway for the dissolution of fibrin clots. Second, itfacilitates cell proliferation and migration by proteolytic degradation of the ECM. As such,this system is involved in physiological processes such as development, including nervoussystem development and many pathological conditions (Sumi et al., 1992). It also plays arole in plastic events such as memory formation, neuronal death and reorganization (Meiriet al., 1994; Wu et al., 2000). The key players in this system include plasminogen activators:the urokinase-type plasminogen activator (uPA) and the tissue-type plasminogen activator(tPA); the uPA receptor (uPAR), and the plasminogen activator inhibitors (PAI-1, PAI-2).

2.4.1 Plasminogen/ PlasminThe human plasminogen is a 92-kDa zymogen composed of 791 amino acids. The aminoterminal of plasminogen includes a “finger” module and five “kringle” domains involvedin interactions with fibrin and matrix proteins. The carboxyterminal domain ofplasminogen is involved in serine protease activity (Mulichak et al., 1991). Plasminogen isconverted into a two-chain structure of plasmin by proteolytic cleavage of a single peptidebond (Arg560Val562). Over the years, researchers have demonstrated that this cleavage iscatalyzed by uPA or tPA, as well as by certain bacterial proteins (Robbins et al., 1967).

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Plasminogen is expressed in many tissues of the body; however, it is mainlyproduced in the liver and also in the testis and epidermal cells (Raum et al., 1980; Isseroff etal., 1983; Saksela et al., 1986). It binds principally to uPA and tPA; however, it can bind toother molecules including laminin, fibronectin, fibrin, thrombospondin, tetranectin andcytokeratin (Quigley, 1979; Salonen et al., 1984; Lucas et al., 1983). The two principalreceptors of plasminogen include �-enolase and anenexin (Miles and Plow, 1987; Hajjar etal., 1994) and are expressed on the cell surface of various cell types including monocytes,granulocytes, lymphocytes and endothelial cells (Hajjar et al., 1986; Silverstein et al., 1988;Miles et al., 1987). Upon interaction with its receptors, plasminogen is rapidly converted toplasmin which has enhanced enzymatic activity on the cell surface. However, thisproteolytic activity is regulated by the presence of uPA and tPA.

The multiple effects of plasmin activity emerge from the analysis of plasminogen-deficient mice. These mice are able to complete embryonic development, reach adulthoodand reproduce. However, they suffer from multiple spontaneous thrombotic lesions,organ damage, high early morbidity and impaired skin wound healing (Bugge et al., 1995;Romer et al., 1996). This thus indicates that plasmin proteolysis is indispensable for tissueremodeling and wound healing. In addition, the plasminogen knock-out mice model foraplasminogenaimia, the severe form of type I plasminogen deficiency, shows fewdifferences in behavioural development (reactivity and response to stress) when comparedto wild type mice (Hoover-Plow J et al., 2001).

Plasminogen deficiency has also been reported in humans. It is characterized by theformation of pseudo membranes on mucosal surfaces and often leads to organ damage ofaffected tissue. The most documented and most recognizable clinical syndrome is ligneousconjunctivitis (Schuster and Seregard, 2003). There are two types of plasminogendeficiency in humans. Type 1, hypoplaminogenaimea, is typically associated withpseudomembrane formation as a result of compromised fibrin clearance. Type IIplasminogen deficiency, dysplasminogenaemia, does not lead to specific clinicalmanifestations (Shuster et al., 2001). This indicates that plasminogen deficiency does notseverely affect brain function. However, there are speculations that plasmin deficiency inthe brain could lead to Alzheimer’s disease (AD) symptomatology, possibly due to theaggregation of amyloid, and trigger cell death signaling cascades (Dotti et al., 2004).

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Figure 2. Schematic representation of the interactions and role of uPA and uPAR as depictedin a tumor cell. Pro-uPA is converted to uPA when it binds to uPAR. uPA then convertsplanminogen (png) to plasmin, which there on degrades the ECM. uPAR can engage in non-proteolytic activities, independent of uPA, like signaling through GPR and integrin, and in celladhesion and spreading by interacting with vitronectin. Abbreviations: ECM, extracellularmatrix; GPR, G-protein coupled receptor; MMP, matrix metalloprotein; png, plasminogen;pngR, plasminogen receptor; suPAR; soluble urokinase-type plasminogen activator receptor;uPA, urokinase-type plasminogen activator; uPAR, urokinase-type plasminogen activatorreceptor; 1, 2, 3, homologous domains of uPAR.

2.4.2 Urokinase-type plasminogen activator (uPA)uPA is synthesized as a 53-kDA catalytically inactive single-chain polypeptide chain (pro-uPA) with 411 amino acids (Gunzler et al., 1982). When pro-uPA is secreted, it becomesconverted to an active two-chain form, uPA, by cleavage of the peptide bond K158-I159 byplasmin (Dano et al., 1985). Structurally, uPA is composed of two peptide chains linked bydisulfide bridges, forming three functional domains. The A chain corresponds to the non-catalytic amino-terminal fragment (ATF), and contains the kringle domain and theepidermal growth factor (EGF)-like domain that binds to uPAR. The B chain, whichcorresponds to the carboxyl terminal region, is a serine protease domain (SPD), which isresponsible for most of its proteolytic functions (Strassburger et al., 1983; Steffens et al.,1982).

Different cell types can synthesize and secrete uPA in normal conditions, such asduring development and pathological conditions. The first evidence of nonpathologicalsynthesis of uPA was demonstrated by Danø and colleagues (1985). The majority ofstudies demonstrating uPA expression under pathological conditions are in cancerresearch (see review by Mekkawy et al., 2009). However, increased uPA expression hasalso been reported in other diseases including human and animal models of TLE (Lahtinen

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et al., 2006; Iyer et al., 2010). Its expression is linked to a variety of physiological andpathological processes including development, wound healing, inflammation, cancerangiogenesis and invasion (Leonardsson et al., 1995; Gyetko et al., 1996; Andreasen et al.,1997). Expression of uPA in the brain is intense during development; however there islimited expression in an adult healthy brain (Del Bigio et al., 1999). Expression in the adultbrain is mainly in astrocytes and neurons (Kaldron et al., 1990; Masos and Miskin, 1996;Lahtinen et al., 2006).

uPA has proteolytic and non-proteolytic biological functions. The mostdocumented proteolytic action of uPA is its conversion of inactive plasminogen toplasmin. When pro-uPA binds to its receptor, uPAR, it is subsequently activated byplasmin to uPA, which then converts neighboring membrane-bound plasminogen toplasmin (fig. 2) (Stoppelli et al., 1986). uPAR localizes pro-uPA to a specific site on the cellsurface where proteolysis is required, and as such brings about regulated degradation ofthe ECM by plasmin.

The non-proteolytic functions of uPA include chemotaxis, cell adhesion andapoptosis. When uPA binds to uPAR, it may cause cleavage of the receptor betweendomain DI and DII. This exposes the uPAR domain DII/DIII (suPAR, SRSRY) that has astrong chemokin-like activity and sends signals through the FPRL/LXA4R receptor (fig. 2)(Sidenius and Blasi, 2003). Also, by binding to uPAR, uPA assists in the generation ofsignals via uPAR through the MEK/ERK and P13K/Akt pathways that favor theexpression of anti-apoptotic proteins of the Bcl family (fig. 2) (Alfano et al., 2006). Theactivity of uPA is controlled by PAIs together with its receptor uPAR (see below).

2.4.3 Tissue type plasminogen activator (tPA)tPA is a 70 kDa protein secreted as a precursor in the single-chain form (Pennica et al.,1983). It is converted to a two-chain active form by plasmin, with every single chainhaving significant activity (Neinaber et al., 1992). tPA is composed of four functionaldomains: (1) an amino-terminal domain or fibronectin-like domain, (2) an EGF-likedomain, (3) two kringle regions (K1 domain and K2 domain) and (4) a serine proteaseregion (Pennica et al., 1983).

The main biological role of tPA is fibrinolysis. Expression of tPA is observed inlocations with close contact to fibrin clots e.g. vascular endothelial cells (Kooistra et al.,1994). tPA is also induced in physiological situations prone to thrombosis, such asischemia (Schneiderman et al., 1991). However, there is interchangeability of activitybetween tPA and uPA. In studies where the tPA gene is deleted from mice, skin woundhealing proceeds normally. When both genes (tPA and uPA) are inactivated, skin woundhealing is severely impaired (Romer et al., 1996).

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In the CNS, the role of tPA is not well characterized, and its primary substrates areeven less known. In the normal nervous system, tPA mRNA is expressed in tissue derivedfrom neuronal ectoderm during development (Carroll et al., 1994). In the adult nervoussystem tPA was detected in the hippocampus, hypothalamus, cerebellum and amygdala(Sappino et al., 1993; Ware et al., 1995; Hastings et al., 1997). tPA genes are induced withneuronal activity, and their release is associated with morphologic differentiation(Krystosek and Seeds, 1981; Qian et al., 1993), thus indicating that tPA plays a role insynaptic plasticity. tPA has also been shown to be involved in learning and memory(Seeds et al., 2003) as well as in behavior and anxiety (Huang et al., 1996). Theseobservations suggest that tPA may be involved in CNS diseases.

2.4.4 Urokinase type plasminogen activator receptor (uPAR)uPAR is a glycosylphosphatidylinositol (GPI)-anchored protein belonging to thelymphocyte antigen (Ly-6) super family of proteins (Ploug and Ellis, 1994). The matureuPAR (283 residues) is highly glycosylated and composed of three homologous domains(about 90 residues each) designated as DI, DII and DIII (Ploug, 2003; Kjaergaard et al.,2008). The domains are connected by short linker regions, packed together forming aconcave-like structure that constitutes the binding site of its principal ligand, uPA (Huai etal., 2006).

Under normal conditions or in a healthy organism, uPAR is thought to beexpressed to a fair degree in various tissues including the kidneys, spleen, lungs, uterus,bladder, thymus, liver and heart (Solberg et al., 2001). However, there is a strongexpression of uPAR in human and animal tissue undergoing extensive tissue remodelingand in keratinocytes during wound healing (Floridon et al., 1999; Solberg et al., 2001;Uszynski et al., 2004). Also, uPAR expression is strongly induced in various pathologicalconditions including virtually all cancers (Bene et al., 2004; Jacobsen and Ploug 2008; Raschet al., 2008). For example, leukocyte expression of uPAR is increased during bacterial orhuman immunodeficiency virus-1 (HIV-1) infections (Speth et al., 1999; Coleman et al.,2001), in the kidneys during chronic proteinuric disease (Wei et al., 2008), in the CNSfollowing ischemia or trauma (Beschorner et al., 2000) and in human and animal models ofTLE (Lahtinen et al., 2009; Iyer et al., 2010; Liu et al., 2010). These findings are suggestive ofa role of uPAR in these diseases.

Generally, uPAR is known to mediate ECM proteolysis, cell- ECM adhesion andregulate intracellular signaling pathways that control cell proliferation, differentiation,migration and survival (fig. 2). uPAR executes its proteolytic function in conjunction withuPA, where it promotes cell surface activation of plasminogen to plasmin (fig. 2) (Ellis etal., 1992). uPAR is also involved in the proteolytic degradation of uPA in the lysosome, by

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the internalization of inactive complexes between uPA and PAI-1 in cooperation withmembers of the low-density lipoprotein receptor (LRP) family (Cubellis et al., 1990;Nykjaer et al., 1992). Many of the non-proteolytic functions of uPAR, most of which areindependent of the protease activity of uPA, are geared towards interactions between cellsand the ECM. By interacting functionally with matrix vitronectin (VN) (fig. 2), adhesionreceptors of the intergrin family and G protein-coupled receptors (GPR), uPAR mediatesin the migration of monocytes, melanoma, fibroblast and many other cells (Gyetko et al.,1994; Yebra et al., 1996; Degryse et al., 1999; Madsen et al., 2007. Another non-proteolyticfunction of uPAR is the chemoattractant property of its DIIDIII fragment as depicted inseveral cell lines (fig. 2) (Resnati et al., 1996). Finally, uPAR can non-proteolyticallyregulate cell proliferation by interaction with integrins, resulting in downstream signals,which activate the ERK/MAP1/2 pathways and suppress the growth inhibiting p38 MAPK(fig. 2) (Aguirre Ghiso et al., 1999). The activity of uPAR is regulated either by cleavagebetween its DI and DII domain, often mediated by uPA, or by forming a complex withuPA-PAI-1-LRP, which triggers internalization-recycling of uPAR (Cubellis et al., 1990).

uPAR deficient mice are used to study the functions of uPAR, especially inpathological conditions including epilepsy. Though uPAR knock-out mice developnormally, survive to adult hood and breed normally (Bugge et al., 1995), they showdefective leukocyte recruitment during infection (Gyetko MR et al., 2000), amelioration ofinflammatory proteinuria (Wei et al., 2008) and reduced brain damage in a model ofcerebral ischemia (Nagai et al., 2008). These mice also show increased susceptibility tochemically induced seizures, heightened anxiety and reduced social interactions (Powell etal., 2003). Recent reports suggest that mutation in one of the ligands for uPAR, Sushi-Repeat Protein X-linked 2 (SRPX2), can cause Rolandic epilepsy with speech impairment(RESDX syndrome) or developmental defects of the speech cortex (bilateral perisylvianpolymicrogyria) (Royer-Zemmour et al., 2008).

2.4.5 Plasminogen activator inhibitor (PAI)Inhibitors of plasminogen activators are members of the serine protease inhibitor superfamily (SERPIN). The inhibitors include PAI-1, PAI-2, and protease nexin 1 (PN1) andprotein C inactivators.

Plasminogen activator inhibitor type 1 (PAI-1)PAI-1 is the main inhibitor of uPA and tPA. It is a single-chain glycoprotein consisting of379 amino acids with an apparent molecular weight of 45 kDa. PAI-1 is secreted by manycells, usually in a dormant conformation (see review Gils and Declerck, 2004). This

18

conformation becomes physiologically relevant and stable by complexing with vitornectin(Ehrlich et al., 1991).

The principal function of PAI-1 is to inhibit ECM degradation by uPA. PAI-1 formsa complex with uPAR-bound uPA and LRP which is then internalized, engulfed bylysosomes and digested, while uPAR is recycled to the surface (Planus et al., 1997;Nykjayer et al., 1997). Also, when PAI-1 binds to vitronectin, it prevents vitronectin frombinding to vitronectin receptors and to ��3 integrin, as such interfering with virtronectinreceptor-dependent effects such as cell migration (Stefansson and Lawrence, 1996). In asimilar way, PAI-1 can prevent vitronectin binding to uPAR (Deng et al., 1996). In fact,PAI-1 mRNA is increased in mouse brains after a stroke (Ahn et al., 1999). PAI-1 mayprotect against neuronal cell injury caused by tPA (Docagne et al., 2002).

Plasminogen activator inhibitor type 2 (PAI-2)Human PAI-2 consists of a single chain protein with 415 amino acids and existspredominantly as a 47 kDa nonglycosylated intracellular form or as a 60 kDa glycosylatedsecreted form (Kruithof et al., 1995). Under normal conditions, PAI-2 has a limiteddistribution pattern with expression detected at high levels in keratinocytes, activatedmonocytes and the placenta (Sharon et al., 2001). Lower constitutive levels of PAI-2 can befound in other cells including cells of neuronal origin (Genton et al., 1987). PAI-2 mRNA isalso localized within the nucleus accumbens (Sharon et al., 2001). Furthermore, KAinduces intense expression of PAI-2 in several brain areas including the pyramidal layer ofthe hippocampus (Sharon et al., 2001).

The role of PAI-2 is not fully understood. PAI-2 blocks uPA and, less efficiently,tPA, although not as fast as PAI-1, raising the possibility that PAI-2 may have some yetunidentified roles. PAI-2 plays a role in the skin by regulating keratinocyte proliferationand differentiation (Hibino et al., 1999). Intracellular PAI-2 protects cells from tumornecrosis factor-� (TNF-�)-medicated apoptosis (Antali et al., 1998). Monocytes andmacrophages express high levels of PAI-2 upon stimulation with tumor necrosis factor(TNF) and lipopolysaccharide (LPS) (Suzuki T et al., 2000). As such, PAI-2 is involved inmonocyte differentiation (Schleuning et al., 1987). High levels of PAI-2 are present ineosinophilic leukocytes (Swartz et al., 2004).

2.4.6 The expression and role of elements of the plasminogen system in epilepsyRecently the role of the plasminogen activator system in neurological disorders has gainedmuch attention. This interest is driven by observations that this system plays a role in thenormal brain and in many brain pathologies including experimental models of seizuresand epilepsy (Gingrich and Traynelis, 2000; Yepes and Lawrence, 2004; Melchor and

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Strickland, 2005; Yepes et al., 2009). Different elements of the system are linked toneurodegeneration, axonal plasticity and seizure threshold, even though the mechanismsby which they alter these events remain unclear.

PlasminogenPlasmin, produced by proteolytic cleavage of plasminogen either by uPA or tPA, hasseveral substrates in the brain, including ECM proteins (Benarroch, 2007). Tsirka et al.(1997) showed plasminogen mRNA expression in most hippocampal neurons, withminimal expression of the protein in just a fraction of the cells. In addition, they showedthat intrahippocampal injection of kainate induces increased hippocampal expression ofplasminogen.

uPAuPA mRNA is one of the most upregulated genes in animal models of TLE includingangular bundle stimulation and amygdala stimulation models on rats (Lukasiuk et al.,2003; Lahtinen et al., 2006; Gorter et al., 2007). Lahtinen et al. (2006) showed using theelectrical stimulation SE model that increased mRNA expression is accompanied byincreased uPA enzymatic activity in the hippocampus and extrahippocampal temporallobe. They further reported that expression was prominent in small populations ofastrocytes, neurons and blood vessels. A similar finding was previously reportedfollowing KA-induced SE (Masos and Miskin, 1997). Increased uPA mRNA expression isobserved in neurons, astrocytes, microglia and blood vessels in humans with epilepsy(Iyer et al., 2010). The effects of increased uPA expression on the epileptogenic network arenot clearly understood. However, Morales and colleagues (2006) showed that uPAknockout mice developed significantly larger lesion volumes than wild-types followingTBI, indicating a possible role of uPA in neuroprotection.

tPAtPA was first shown to be linked to epilepsy by Qian et al. (1993). They showed increasedtPA expression in rat cortexes following pentylenetetrazol (PTZ) induced seizures. Also,there is an increase in tPA mRNA expression in the hippocampus following perforant pathstimulation and intrahippocampal injection of kainate (Qian et al., 1993; Salles andStrickland, 2002). In human epilepsy, there is increased tPA expression in tissues frompatients with gangliogliomas but not in those with TLE without hippocampal sclerosis orfocal cortical dysplasia (Iyer et al., 2010). Increased tPA mRNA expression is tied with anincrease in tPA enzymatic activity after intraamygdala or intraventricular KA injection(Endo et al., 1999; Yepes et al., 2002).

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The effect of tPA on seizure susceptibility was revealed using tPA knock-out mice.Tsirka et al. (1995) showed that tPA deficient mice require a higher dose of KA or PTZ toenter seizures compared to controls. Similarly, tPA deficient mice showed decreasedseizure progression and lacked generalized seizures following intra-amygdala KAinjection (Yepes et al., 2002). These effects were shown to be plasminogen-independent(Yepes et al., 2002). Various mechanisms have been proposed to be involved in thefacilitation of seizures by tPA, including the cleavage of the NR1 subunit of NMDAreceptor by tPA in cortical neurons (Benchenane et al., 2007).

uPARIt was recently demonstrated using the amygdala stimulation SE model that, uPARmRNA expression is increased at 1-4 days after SE (Lahtinen et al., 2009). Lahtinen andcolleagues showed that uPAR protein expression levels lasted 2 weeks after SE and weremostly concentrated in astrocytes, neurons and in parvalbumin positive interneurons inthe hippocampus and dentate gyrus, and in a subgroup of somatostatin and neuropeptideY positive hilar interneurons. However, in normal brains, expression was limited only toastrocytes and to a limited number of parvalbulmin-positive interneurons.

There is also uPAR mRNA expression in TLE patients with hippocampal sclerosisand focal cortical dysplasia, as well as in patients with frontal lobe epilepsy (Iyer et al.,2010; Liu et al., 2010). uPAR expression is mostly in neurons and very few astrocytes (Liuet al., 2010).

The role of uPAR in epilepsy was recently highlighted by observations made byPowell and colleagues (2003) using uPAR knock-out mice. They showed that uPARdeficient mice were susceptible to chemically induced seizures, and had poor socialinteractions. In addition, they linked these observations to poor interneuron development,particularly of the parvalbumin subtype.

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Table 1. Experimental studies describing behavioural and pathological changes associated withuPA and uPAR after brain damaging insults.

Model/ animal Finding Reference

FCI infarction inmice

No difference in infarct size 24h after injury betweenuPA-/- and Wt

Nagai et al.,1999

Severe malaria inmice

No difference in BBB impairment between uPAR-/- andWt. Absent of macrophage sequestration in uPAR-/-

Piguet et al.,2000

Brain stab woundin mice

Similar levels of BBB impairment in uPA-/- and Wt at3, 8, and 15 days after injury

Kataoka et al.,2000

Experimentalpneumococcalmeningitis in mice

Attenuated CSF pleocytosi in uPAR-/- at 24h post-injury. No difference in BBB impairment, intracranialpressure and chemokine

Paul et al.,2005

TBI (CCI model ) inmice

Increase size of lesion in uPA-/- Morales et al.,2006

TBI and focalcerebral infarctionin humans

Increase number of uPAR expressing granulocytes,microglia/ macrophages and endothelia cell, correlatedwith increase edema

Beschorner etal., 2000

Fluid percussioninjury in pigs

At 4 hr post-injury, uPA expression is associates withage dependent hyperaemia

Armstead et al.,2009

uPA overexpression in mice

Decrease food consumption, reduced body weight andsize

Miskin andMasos, 1997

uPA overexpression in mice

Impaired learning Meiri et al.,1994

uPAR-/- mice Disruption of GABAA receptor subunits; alfa 1, 2 and3, Beta 2 and 3, and gamma 2S and 2L in frontal andparietal cortices

Eagleson et al.,2010

PIFBD and MCAO inmice

Smaller lesion is uPAR-/- compared to Wt at 3 d post-MCO, whereas no difference with Wt in PIFBD injury

Nagai N et al.,2010

MCAO in mice Decreased brain damage in uPAR-/- mice at 3 d postinjury

Nagai N et al.,2008

EAE in mice uPAR-/- mice experienced delayed less acute diseasewhich subsequently become chronic

East et al.,2005

Corticaldevelopment inuPAR-/- mice

P21 uPAR-/- mice showed loss of cortical GAD67 andGABA positive cells. Only loss of cortical parvalbumincells in adults.

Eagleson et al.,2005

uPAR-/- mice(naïve animals)

Increase anxiety, spontaneous seizures and increasesusceptibility to PTZ-induced seizures. 50% loss ofGABAergic interneuron especially parvalbumin subtypein anterior cingulated and parietal cortices

Powel et al.,2003

Abbreviations: BBB, blood brain barrier; CCI, control cortical impact; d, day; EAE,experimental allergic encephalomyelitis; FCI, Focal cerebral ischemic; GAD67, glutamic aciddecarboxylase-67; hr, hour; i.h., intrahippocampal; KA, kainic acid; P, post-natal; PIFBD,photochemical induced thrombotic brain damage; PTZ, pentylenterazole; MCAO, medialcerebral artery occlusion; uPA, urokinase-type plasminogen activator; uPAR, urokinase-typeplasminogen activator; Wt, wild type.

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3. Aims of the study

The broad objective of this study is to increase our understanding of the interactions and

functional implications of the cellular and molecular reorganization processes known to

occur during epileptogenesis. Furthermore, it aims to study the underlying molecular

mechanisms involved in these cellular reorganization processes.

Therefore, the specific aims of this work are to:

1. Study the interaction between vascular remodeling and neuronal plasticity during

epileptogenesis

2. Study the functional implications of blood vessel length changes in the

hippocampus during epileptogenesis

3. Determine whether uPA or uPAR deficiency differentially affect the epilepsy

phenotype following an epileptogenic insult such as status epilepticus

4. Decipher the role of uPA and uPAR in the cellular reorganization processes during

epileptogenesis

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4. Materials and methods

4.1 ANIMALS

All animal experiments were approved by the Animal Care and Use Committee of theUniversity of Eastern Finland. All procedures were conducted in accordance withguidelines set by the European Community Council Directives 86/609/EEC. Animals werehoused in a control environment (temperature 22°C ± 1°C, humidity 50-60%, lights on07:00-19:00) with free access to water and food.

Rats (I)Adult male Sprague-Dawley rats (Harlan, the Netherlands) weighing 300-350g housed inindividual cages were used.

Mice (II, III)For study II, Adult male mice (12-14 weeks) lacking the uPA gene in C57BL/6 background(uPA-/-; B6.129S2-Plautm1Mlg/J from The Jackson Laboratory, Bar Harbor, USA; Carmeliet etal., 1994) and their wild type (Wt) littermates were used.For study III, adult male mice (12-14 weeks) lacking the uPAR gene (uPAR-/-, B6.129P2-Plaurtm1Jld, a generous gift from Dr. Van der Pol, University of Amsterdam, TheNetherlands) were used. uPAR-/- mice were backcrossed to C57BL/6 genotype (CharlesRiver, originally JAXC57BL/6J Stock 000664 from The Jackson Laboratory) for at least 8generations. The genotype of mice in all studies was determined by PCR.

4.2 ANESTHESIA

RatsFor MR imaging, animals were anesthetized with isoflurane in a carrier gas mixture of70% N2O, 30% O2. For transcardiac perfusion, animals were anesthetized with 6 ml/kg,(i.p.) of an anesthetic cocktail containing 58 mg/kg sodium pentobarbital and 60 mg/kgchloral hydrate.

MiceFor the induction of status epilepticus, implantation of cortical and/or hippocampalelectrodes and transcardiac perfusion, animals were anesthetized with a mixture of 18.75mg/ml ketamine and 0.25 mg/ml medetomidine (4 ml/kg, i.p.).

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4.3 INDUCTION OF STATUS EPILEPTICUS (SE)

4.3.1 Pilocarpine model (I)Animals were injected subcutaneously (s.c.) with scopolamine (1 mg/kg; #S-8502, SigmaChemical Co., St. Louis, MO) to reduce the peripheral adverse effects of pilocarpine. Thirtyminutes later, SE was triggered by intraperintoneal (i.p.) administration of pilocarpine(320 mg/kg, #P-6503, Sigma Chemical Co., St. Louis, MO). The development and severityof generalized SE was assessed by visual observation, and the behavioral severity ofseizures during SE was scored according to Racine (1972). Diazepam 20 mg/kg (i.p.,Stesolid Novum, Dumex-Alpharma) was administered 120 minutes following induction ofSE to reduce the mortality rate. Control animals were treated similarly except they wereinjected with 0.9% NaCl instead of pilocarpine

4.3.2 Intrahippocampal kainic acid model (II, III).Animals were anesthetized and placed into a stereotactic device. The skull was exposedand a small hole was drilled at the injection site. Using a glass pipette (inner diameter 0.2mm; outer diameter 0.75 mm; tip diameter 0.01-0.02 mm) connected to a pulse pressuredevice (Picospitzer, General Valve Corporation, Fairfield, USA), 60 nl of 20 mM (II) or 10mM (III) of KA or 0.9% of NaCl (controls) was injected into the right hippocampus withcoordinates AP-2.3; ML-1.5 (bregma as reference, mouse atlas of Franklin and Paxinos,2008) and DV 1.9 below dura. After injection, the capillary was kept in place for 5 minutesand then withdrawn slowly (100 μm at a time, 20-30 sec intervals). The aim was to get theinjection into the dentate gyrus. After the operation, the mice were injected withAntisedan� (0.5 mg/kg i.p; antipamezole hydrochloride; Orion Pharma, Espoo, Finland)and 0.9% saline (150μl/animal) to facilitate awakening from anesthesia.

4.4 ELECTRODE IMPLANTATION

4.4.1 Cortical electrodes (II, III)Following intrahippocampal kainate injection, four cortical screw electrodes wereimplanted. Two electrodes were placed bilaterally into the skull above the parietal cortex(AP -1.10; ML 2.5 with bregma as reference). The other two were placed bilaterally overthe cerebellum to serve as ground and reference electrodes.

4.4.2 Hippocampal electrodes (III)After kainate injection, a bipolar nylon-coated stainless steel wire electrode (� 0.127 μm;dorsoventral tip separation 400 μm, Franco Corradi, Milan, Italy) connected to a gold-

27

plated pin (Plastics One Inc., Roanoke, VA, USA) was implanted into the brain. The lowerelectrode tip was aimed to hit the suprapyramidal blade of the granule cell layer in thedentate gyrus (AP -2.3 ML -1.5, DV -1.9).

Electrodes were secured in place with dental acrylate (Selectaplus CN, DentsplyDeTrey GmbH, Dreieich, Germany). After the operation, animals were injected withAntisedan� (0.5 mg/kg i.p; antipamezole hydrochloride; Orion Pharma, Espoo, Finland)and 0.9% saline (150 μl/animal) to facilitate awakening from anesthesia.

4.5 VIDEO-EEG MONITORING (II, III)

After the implantation of cortical and/or hippocampal electrodes, animals were moved totheir home cages, connected to the video-EEG recording system, and continuouslymonitored for 30 d (24/7) as previously described by Nissinen et al. (2000). An infraredsensitive camera and infrared light were placed in front of the cages. Infrared lightpermitted video monitoring of animal behavior during the dark cycle. A wide-angle lenspermitted digital video-recording of up to 8 animals at the same time.

4.5.1 Detection and analysis of severity of status epilepticus (II, III)After the induction of SE, electrographic activity was recorded using a video-EEGrecording system from Nervus (Taugagreining, Iceland). The severity of status epilepticuswas assessed by counting the total number of spikes generated in EEG during the first 48hours after KA administration, using Clamfit� 9.0 software according to Pitkänen et al.(2005). A spike was defined as a high amplitude discharge (> 2x baseline) lasting less than70 ms.

4.5.2 Detection and analysis of electrographic and behavioral seizures (II, III)The development of spontaneous seizures was studied using data from the video-EEGrecording system. Seizures were analyzed by manually browsing through all EEG datafiles on a computer screen with the aid of the Nervus® EEG reader software. Anelectrographic seizure was defined as a high-frequency (> 5 Hz), high-amplitude (> 2xbaseline) discharge lasting for more than 10 s. When an electrographic seizure was found,its behavioral severity was analyzed from the matching video recording. Behavioralseizure activity was scored according to a slightly modified Racine's scale (Racine, 1972):score 0, electrographic seizure without any detectable motor manifestation; score 1, mouthand face clonus, head nodding; score 2, clonic jerks of a forelimb; score 3, bilateral forelimbclonus; score 4, forelimb clonus and rearing; score 5, forelimb clonus with rearing andfalling.

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4.6 BrdU ADMINISTRATION FOR STUDY OF ENDOTHELIAL CELLPROLIFERATION (I)

To investigate the proliferation of endothelial and neuronal cells after SE, animals wereinjected with 5-bromo-2'-deoxyuridine (BrdU) dissolved in 0.9% NaCl (BrdU, 50mg/kg,#280879, Roche Diagnostics Corp., Indianapolis, IN) twice a day (i.p.) at an interval of 12 hfor 5 d. The last injection was administered 24 h before sacrifice.

4.7 MAGNETIC RESONANCE IMAGING MEASUREMENT OF CEREBRALBLOOD FLOW (CBF) AND CEREBRAL BLOOD VOLUME (CBV) (I)

To assess the functional consequences of observed vascular changes, MRI was employedto measure absolute CBF and relative CBV in the rat hippocampus. Rats were anesthetizedand cannulae were inserted into the femoral vein and femoral artery, for contrast agentdelivery and arterial blood sampling to monitor blood gases, respectively. The rat wassecured in a stereotactic holder using ear bars and a bite bar. Breath rate was continuouslymeasured throughout imaging via a pressure probe between the rat and the holder. Thebreathing rate measurement helped us to control the anesthesia concentration, which wasmaintained at 0.7-1.7%. The holder was electronically heated to sustain a temperature of37°C throughout.

All MRI was performed using a 4.7 T scanner (Magnex Scientific Ltd, Abington,UK) interfaced to a Varian Inova console (Varian, Palo Alto, CA) with an activelydecoupled linear transmission volume coil and quadrature surface receiver coil pair(Rapid Biomedical, Germany). CBF was quantified using continuous arterial spin labeling(Williams et al., 1992) (duration of labeling pulse = 3 s, B1 = 0.018 mT, 800 ms delay afterlabeling pulse) with a fast spin echo read out (field of view = 4 x 4 cm, 128 x 128 points, 2mm slice thickness, repetition time = 6 s, 16 echoes/excitation, echo spacing = 7 ms, 6 pairsof label and control images) from one coronal hippocampal slice. Absolute CBF wascalculated in the hippocampus through region of interest (ROI) analysis of CBF mapsgenerated by the arterial spin labeling approach. CBF maps were corrected for T1 variationacross the brain by employing a T1 inversion recovery, fast spin echo sequence to provideT1 maps that were incorporated into CBF map calculations (Barbier et al., 2001). Thesequence parameters were as follows: field of view = 4 x 4 cm , 64 x 64 points, 2 mm slicethickness, repetition time = 5 s, echo spacing = 6 ms, inversion times = 5, 300, 600, 1000 and1500 ms.

�R2 was mapped by acquiring T2 weighted images (field of view = 4 x 4 cm, 256 x128 points, echo time = 70 ms, repetition time = 2500 ms, 15 coronal slices, 1 mm slicethickness) taken before and 3 min after iron oxide contrast agent injection (Sinerem,

29

Geurbet, France) at 6 mg/kg rat body mass. �R2 caused by the intravascular contrast agentwas assumed to be directly proportional to CBV (Dunn et al., 2004), which was quantifiedin the hippocampus and dorsal cortex from a single hippocampal �R2 map. The CBF mapand chosen CBV map were from an identical slice. Hippocampal CBV was normalized tothat of the healthy neck muscle in order to account for contrast agent concentrationvariations between animals, hence relative CBV was studied. Arterial blood pH, pCO2 andpO2 were monitored by blood sampling before contrast agent injection, around 60 minafter cannulae surgery, and again soon after imaging was completed. The total imagingtime was around 100 min per animal. After imaging, rats received recovery surgery toremove the cannulae.

4.8 PROCESSING OF BRAIN FOR HISTOLOGY

4.8.1 Transcardiac perfusion - Timm fixation protocol (I, III)Timm fixation was carried out according to Sloviter (1982). Briefly, animals were deeplyanesthetized and perfused transcardially with 0.37% sulfide solution for 10 min usingperistaltic pump followed by cold (4°C) paraformaldehyde (PFA) in 0.1 M sodiumphosphate buffer (PB, pH 7.4) for 10 min (flow rate, Rats: 30 ml/min; Mice: 5 ml/min).After perfusion the brains were removed from the skull, postfixed in 4%paraformaldehyde in 0.1 M PB at 4°C for 4 h, cryoprotected in 20% glycerol in 0.02 Mpotassium phosphate-buffered saline (KPBS) at 4°C for 24 h, frozen on dry ice, and storedat -70oC.

4.8.2 Transcardiac perfusion - Para-formaldehyde fixation protocol (I-III)Animals were deeply anesthetized and were transcardially perfused with 0.9% NaCl (2min) followed by 4% paraformaldehyde in 0.1 M in PB (Rats, 30 min and mice, 20 min).After perfusion, the brains were further processed as described above in the Timmperfusion protocol.

4.8.3 Sectioning of brainThe brains were cut serially in the coronal plane using a sliding microtome. For rats,sections were cut 30 μm thick in a 1-in-5 series. For mice, sections were cut 25 μm thick ina 1-in-6 series. The first series of sections was stored in 10% formalin and the other seriesof sections were stored in a cryoprotectant tissue collecting solution (30% ethylene glycoland 25% glycerol in 0.05 M PB) until further processed.

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4.9 HISTOLOGY

4.9.1 Nissl staining and analysis of neuronal damage (I - III)In order to identify the distribution and severity of neuronal damage, the first series ofsections (stored in 10% formalin) were stained using thionin, cleared in xylene, and cover-slipped using DePeX� (BDH Chemical, Poole, United Kingdom) as a mounting medium.

4.9.1.1 Analysis of damage to principal cells (I, II, III)Neuronal damage in the CA1 and CA3 subfield of the hippocampus was estimated fromnissl stained sections, using a semiquantitative scoring method. Severity of degenerationwas scored according to Freund et al. (1992) as follows: 0 = no cell loss, 1 = less than 20% ofneurons lost, 2 = 20 to 50% of neurons lost, 3 = over 50% of neurons lost. Eight consecutivesections from the level at which suprapyramidal and infrapyramidal blades of the granulecell layer become fused together were analyzed, blind to the experimental groups. Theseverity of neurodegeneration in each subfield in each section was scored, and the meanscore was used for statistical analysis.

4.9.1.2 Analysis of damage to hilar cells (II, III)Damage to dentate hilar cells was estimated bilaterally using unbiased stereology.Systematic sampling of nissl-stained sections with a random start, covering the entireseptotemporal axis of the hippocampus, was employed. Sampling was started at the levelwhere the supra and infra-pyramidal blades of the hippocampus fuse together. For mice, asection sampling fraction of 1:12 was used. Sections were analyzed usingStereoInvestigator� software (Version 2006, MicroBrightField Inc., Colchester VT,Germany) under an Olympus BX50 microscope equipped with a motorized stageinterfaced with a Hitachi HV-C20A camera. The total number of hilar cells was estimatedusing the optical fractionator method (West et al., 1991). The region of interest wasoutlined at low magnification and a sampling grid of 40 μm (x-axis) and 40 μm (y-axis)was placed on the section. For each x–y step, cell counts were derived from a knownfraction of 28 μm (x-axis) by 28 μm (y-axis) (an "optical dissector"). Counting wasperformed throughout the section and the total number of hilar cells was estimated usingthe following equation: Ntot = �Q · 1/ssf · 1/asf · 1/tsf, where ssf is section sampling fraction,asf is area sampling fraction (the area of the counting frame divided by area of samplinggrid), and tsf is tissue sampling fraction (the height of the mounted section thicknessdivided by the dissector height).

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4.9.1.3 Analysis of granule cell layer volume (II, III)To determining the extent of granule cell layer dispersion post-SE in mice following KA-injection, granule cell layer volume was measure bilaterally from nissl stained sectionsusing unbiased stereology. The volume was estimated using Cavalieri principle. A similarsampling scheme as described for hilar cell count in mice was employed. After delineatingthe region of interest (ROI) at low magnification, a grid of 100 μm (x-axis) by 100 μm (y-axis) was laid on the section. The total number of hit points within the selected area ofinterest was counted as described by Gundersen and Jensen (1987). The volume of thegranule cell layer was calculated using the formula: V = �Pi · a(p) · T, where P� is the totalnumber of hit points counted in all sections, a(p) is the area associated per point (0.01mm2), and T is the section thickness (300μm, ssf multiplied by section thickness).

4.9.2 Timm histochemistry and assessment of mossy fiber sprouting (I, III)

To be able to assess post-injury synaptic reorganization in the dentate gyrus, one series ofsections (Timm-fixed) were stained using Timm sulfide/silver method as previouslydescribed by Sloviter, 1982.

Assessment of mossy fiber sproutingDensity of mossy fibers was assessed in the supragranular region and inner molecularlayer of the dentate gyrus. Analysis was performed along the septotemporal axis of thehippocampus, starting at the level were the suprapyramidal and infrapyramidal blade ofthe granule cell layer form a continuous band of cells. In rats, analysis was performedseparately from the "tip", "mid" and "crest" portions of the granular cell layer as describedbefore (Nissinen et al., 2001) using a semiquantitative scoring scale according to Cavazos etal. (1991) (for details see Nissinen et al., 2001). In mice, only the contralateral dentate gyruswas analyzed since sprouting was absent in the severely damaged ipsilateral sidefollowing kainate injection. The analysis in mice was performed using National Institutefor Health (NIH) ImageJ® software version 1.45b. First, a digital photomicrograph wascaptured from each section using a Leica DMRD microscope at 5x objective. The areaincluding the supragranular region and inner molecular layer of the dentate gyrus wasdelineated. The image was then converted to grayscale and threshold to match with thestaining (Buckmaster and Lew, 2011). Thereafter, the area (Ai) covered by the mossy fiberswithin the delineated area was automatically calculated. The total volume (V) wascalculated using the formula: V = �Ai · T · n, where �Ai is the sum of areas from n sections,T is the section thickness and n is the number of sections sampled.

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4.9.3 Doublecortin (DCX) immunohistochemistry and analysis of neurogenesis (I-III)

In order to be able to assess post-injury neurogenesis in the dentate gyrus, sections werestained for doublecortin (DCX). Free floating sections were washed (0.02 M KPBS) andincubated for 15 min in 1% H2O2 in KPBS to remove endogenous peroxidase activity. Thenwere rinsed 6 times with 0.02 M KPBS and non-specific binding was blocked in a solutioncontaining 10% normal horse serum (NHS), 0.4 % Triton X-100, and KPBS at RT for 2 h.This was followed by incubation in goat-derived anti-doublecortin antibody (1:800; SC-8066; Santa Cruz Biotechnology, Santa Cruz, CA, USA) in 1% NHS and 0.4% Triton X-100in KPBS at 4°C overnight. Sections were washed (2% NHS, prepared in 0.02 M KPBS) andincubated in a secondary antibody solution containing biotinylated horse anti-goat IgG(1:300; BA-9500; Vector laboratories), 1% NHS, and 0.4% Triton X-100 in KPBS for 2 h atRT. After washing (0.02 M KPBS) they were further incubated in 1% avidin-biotin (PK-4000; Vector Laboratories) in KPBS for 1 h at RT. Then were washed (0.02 M KPBS) andrecycled back to the secondary antibody solution for 45 min, and then to the avidin-biotinsolution for 30 min. The secondary antibody was visualized by incubating in a solutioncontaining 0.05% DAB (Pierce Chemical) and 0.04% H2O2 in KPBS for 2 min at RT.

Analysis of neurogenesisTo estimate total number of DCX-positive cells, the optical factionator method was usedwith the aid of the stereoinvestigator software as described above for hilar cells. In rats,analysis was performed in the whole dentate gyrus. A section sampling fraction (ssf) of1:15 was used and cells were counted with an x-y step of 100 μm by 100 μm and acounting frame of 22 μm by 22 μm.

In mice, in addition to the entire dentate gyrus, analysis was performed in the threeseparate areas; hilus, subgranular zone and granule cell layer. Sampling was performed asdescribed above for hilar cells. An x-y step of 40 μm by 40 μm and a counting frame of 19μm by 19 μm were used. The total number of DCX- positive cells was estimated usingsame formula as described for hilar cells.

4.9.4 IgG immunohistochemistry and analysis of blood-brain barrier damage (I, III)

In order to investigate post-SE disruption of blood-brain barrier (BBB) sections wereimmmuno-stained for IgG leakage into the brain. First, sections were washed andendogenous peroxidase was removed as described for DCX staining. In rats, sectionswere subsequently incubated in a rabbit-derived biotinylated anti-rat IgG antibody (1:200;#BA- 4000, Vector laboratories) and in mice in horse-derived anti-mouse IgG antibody(1:300, BA2000, Vector Laboratories) prepared in a solution containing 4% NHS and 0.4%Triton-X100 at 4°C overnight. Subsequently, sections were washed (0.02 M KPBS) andincubated in a solution containing avidin-biotin-complex prepared according to

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instructions provided by the manufacturer, for 1 h. The peroxidase activity was visualizedas described for DCX staining but incubated for 1 min.

Quantification of blood-brain barrier damageThe magnitude of BBB damage was quantified by measuring the intensity of IgGimmunoreactivity in stained hippocampal sections by using the NIH ImageJ® software.Briefly, digital photomicrographs of the hippocampus were captured using Leica DMRDmicroscope at 5x. The ROI was outlined, the image was converted to gray scale, and thegray scale value corresponding to relative optical density (ROD) (immunoreactivity) of theIgG staining was measured. In rats, the background value was measured from theundamaged region of the corpus callosum and for mice, the thalamus. In rats, the degreeof IgG leakage was calculated as a ratio of the mean hippocampal grey scale value to thatof the corpus callosum. In mice, IgG reactivity was calculated as the difference betweenmeasure ROD in the ROI and background staining in the thalamus.

4.9.5 Thrombosis immunohistochemistry and assessment of thrombocyte (I)

To study the formation of thrombus post-SE, free floating sections were immunostainedfor thrombus. The protocol was essentially same as for DCX staining except the primaryantibody was a rabbit-derived antibody raised against rat thrombocyte (1:1000, #CLA41440, Cedarlane laboratories, Hornby, Ontario, Canada) and the secondary antibodywas biotinylated goat-derived antibody raised against rabbit-IgG (1:200; #BA-100, Vectorlaboratories). Antigen was visualized using Vector VIP substrate kit (#SK-4600, Vectorlaboratories) according to instructions provided by the manufacturer.

Assessment of thrombocyte formationThe total number of thrombi was analyzed from consecutive sections from the septalhippocampus (1-in-5, 150 �m apart from each other), beginning at the level where thesuprapyramidal and infrapyramidal blades of the granule cell layer form a continuousband of cells [-3.3mm posterior from bregma, Paxinos and Watson (1998)]. Using theStereoInvestigator software, a grid of dimension 400 μm (x) by 400 μm (y) was laid on thesection and a counting frame of 20 μm by 20 μm was used to count the thrombi. The totalnumber of thrombi was estimated by multiplying the total number of counted thrombifrom the 3 sections by the area sampling fraction (asf) [Ntot = �Q · 1/asf] (where �Q is thetotal number of counted thrombi).

4.9.6 CD68 immunohistochemistry and assessment of macrophage response (III)

To evaluate the effect of an injury-induced macrophage response, sections were stained forCD68, which is a lysosomal marker in phagocytotic macrophages. The protocol was

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essentially the same as for DCX staining except the primary antibody was a rat-derivedanti-mouse CD68 antibody (1:500, MCA 1957, Serotec) and the secondary antibody arabbit-derived biotinylated anti-rat IgG antibody (1:300, BA4001, Vector Laboratories).

Assessment of macrophage responseSection sampling for analysis was performed as previously described for mice hilar cellsanalysis. To study the progression of CD68 reactivity a time course analysis wasperformed at 4 d and 30 d post-SE. Analysis was first performed in the entirehippocampus, and then, separately in the CA1 and CA3 subfields of the hippocampusproper and the hilus of the dentate gyrus. Quantification of immunoreactivity was doneby using NIH ImageJ® software as described for IgG (see above). In addition, CD68immunoreactivity in extra-hippocampal regions (amygdala and hypothalamus) wasanalyzed using a semiquantitative scoring scheme. Scoring was done as follows: 0 = noreactivity, 1 = very weak reactivity, 2 = weak reactivity, 3 = moderate reactivity, and 4 =strong reactivity.

4.9.7 CD3 immunohistochemistry and assessment of T cell response (III)

To further assess the inflammatory response post-SE, sections were stained for CD3�, amarker for T-lymphocytes. A similar staining protocol as described for DCX staining wasemployed, except that the primary antibody was a hamster-derived anti-mouse CD3�antibody (1:1000, #553058, BD Pharmingen) and the secondary antibody was a goat-derived anti-hamster IgG (1:300, BA9100; Vector Laboratories).

Assessment of T cell responseThe total number of CD3-positive cells was estimated in the ipsilateral CA1 and CA3subfields of the hippocampus, and the hilus of the dentate gyrus, with the aid of theStereoInvestigator� software. The contralateral hippocampus was not included in theanalysis due to the relatively sparse number of T-lymphocytes observed. A similarsampling scheme as described earlier for the estimation of the total number of hilar cells inmice was employed. The number of immunopositive T-cells in each region of interest wascounted. The number of cells per mm3 (T) was estimated using the formula: T = �Q /Ai · 12· 0.025, where �Q is the total number of cells (Q) counted in all sections, Ai is the sum ofareas of ROIs (mm2), 12 is the sampling fraction, and 0.025 is the section thickness (mm).

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4.9.8 Blood vessel immunohistochemistry (I, II, III)

4.9.8.1 Rat endothelial cell antigen-1 (RECA-1) and endothelial barrier antigen (EBA)immunohistochemistry (I)To study the vascular morphometry and spatiotemporal evolution blood vessels post-SEin rats, sections were immune-stained with either mouse derived anti-rat EBA (SMI71, 1:10000; Sternberger Monoclonals Inc; Baltimore, MD) or mouse derived anti-RECA-1 (1:5000;#MCA970R, Serotec, Oslo, Norway) as primary antibodies. A biotinylated horse-derivedanti-mouse IgG (1:200; #BA-2000, Vector Laboratories, Burlingame, CA, U.S.A.) was usedas the secondary antibody. The protocol was basically the same as described for DCXstaining.

4.9.8.2 Lycopersicon esculentum (tomato) lectin staining (II, III)Biotinylated Lycopersicon esculentum (tomato) lectin (1:1000; B-1175; Vector Laboratories,Burlingame, CA, USA) was used to visualize blood vessels in mice brain sections. Theprotocol was essentially the same as described for IgG staining.

4.9.8.3 Assessment of total blood vessel length (I, II, III)The total blood vessel length was estimated using unbiased stereology with the aid ofstereoinvestigator® software. The virtual sphere method (Mouton et al., 2002) was used. Athree-dimensional sampling hemisphere (“space ball”) of known radius and height wasplaced in a box of known dimensions and focused through the section thickness. Forevery x-y step, the number of vessels intersecting the sphere was counted with the aim ofgenerating counts of about 300 intersections per animal.

In rats, the total blood vessel length was estimated unilaterally (left hippocampus)in the CA1 and CA3 subfields of the hippocampus and the molecular layer of the dentategyrus. Every third section from a 1-in-5 series was used (ssf = 1/15). A samplinghemisphere ("space ball") with a radius of 20 μm and height of 10 μm was placed within asampling box of known dimensions (x = 69.5 μm, y = 69.5 μm and z = 10 μm). The x-ysteps giving the distance between sampling areas was 400 μm (x-axis) by 400 μm (y-axis).In order to calculate the total length of blood vessels, the following equation was used: Ltot

= �Q · 2 · 1/ssf · 1/asf · 1/tsf · [v/a], where ssf is 1/15, asf is 0.03, tsf is 1, and v/a 19.2 μm[defined as the ratio of the volume of the counting frame (v) (sampling box) to the surfacearea of the hemisphere probe].

In mice, the total blood vessel length was estimated bilaterally using a similarsampling scheme as described for hilar cell count in mice. A hemisphere probe of diameter20 μm and height 10 μm was placed within a counting frame 40 μm wide (x-axis), 40 μm

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high (y-axis), and 10 μm deep (z-axis). An x-y step of 300 μm (x-axis) by 300 μm (y-axis)was used. The total blood vessel length was estimated using the same formula asdescribed above for rats.

4.9.8.4 Assessment of Blood vessel diameter (I)Blood vessel diameter was measured in sections stained for RECA-1. Every third sectionfrom a 1-in-5 series was sampled (ssf = 1/15). The hippocampal fissure was analyzedseparately. The analysis was performed using the StereoInvestigator® software. A grid of380 μm (x) by 380 μm (y) (hippocampus) and 150 μm (x) by 150 μm (y) (for hippocampalfissure) was superimposed on the section. A counting frame (40 μm x 40 μm) was used inthe analysis. The diameter of all vessels within the counting frame was measured usingthe “quick line” measurement tool.

4.9.9 BrDU-RECA-1 double staining and analysis of endothelial cell proliferation (I)Double staining of BrdU-positive cells and blood vessels with RECA-1 permitted the studyof newly formed blood vessels post-SE. First, free-floating sections were treated with 2 MHCl for 30 min at 37°C to denature DNA (enabling BrdU to be labeled), and after thatwashed with 0.1 M boric acid (pH 8.5) to remove residual acid. The subsequent part of theprotocol is essentially the same as that used for DCX staining except that two primaryantibodies were used, mouse-derived anti-RECA-1 (1:5000) and rat-derived anti-BrdU(1:10 000; #MCA2060, Serotec, Oslo, Norway). The secondary antibodies were applied intwo phases: the first was rabbit-derived biotinylated anti-rat IgG (1:200; #BA-4000, VectorLaboratories). Antigen was visualized using glucose-oxidase-DAB solution, containing 1%nickel ammonium sulphate, 1 mg/ml DAB (Pierce), 20% �-D(+)-glucose, 0.4% ammoniumchloride, and 1 μl/ml glucose oxidase (#G-6891, Sigma). After extensive washing steps, inthe second phase, goat-derived biotinylated anti-mouse IgG (1:200; #BA-2000, Vectorlaboratories) was applied and antigen was visualized using DAB.

Analysis of proliferating endothelial cells (I).The total number of proliferating endothelial cells was determined using the opticalfractionator method (West MJ et al., 1991). Every third section from a 1-in-5 series wassampled (ssf = 1/15). Proliferating endothelial cells were defined by the following criteria:BrdU positive (BrdU+) cell nucleus with flattened or slightly cupped appearance foundwithin RECA-1 positive stained vessels (RECA-1+ / BrdU+). A virtual counting grid 213.9μm (x-axis) by 213.9 μm (y-axis) was laid on the sections. Then, a three-dimensionalcounting frame of 18.3 μm (x-axis) by 18.3 μm (y-axis) with height (z-axis) of 10 μm wasfocused through a known depth of the section. Every cell that came into focus wascounted. Total numbers of cells were estimated using the same formula as described forhilar cells.

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4.10 STATISTICAL ANALYSIS (I, II, III)

Data were analyzed using SPSS for Windows (version 17.0). Data from spike countingwere analyzed using Repeated Measure Anova (RMA) with the Greenhouse-Geissercorrection factor. Differences between groups following semiquantitative scoring wereanalyzed using the Chi-Square test. Differences between groups in other measurementswere analyzed using the Kruskall-Wallis test. If differences were found, post hoc analysiswas performed using the Mann-Whitney U test. Correlations were analyzed using theSpearman rank correlation test. Data are presented as mean ± standard error of the mean(SEM). A P-value less than 0.05 was considered significant.

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5. Results

5.1 STATUS EPILEPTICUS AND SEIZURE PHENOTYPE IN uPA-/- ANDuPAR -/- MICE (II, III)

In order to establish whether uPA and uPAR influence the severity of an epileptogenicbrain insult such as status epilepticus (SE) and the subsequent epilepsy outcome, theseverity of SE and the development of epilepsy was assessed in uPA and uPAR deficientmice after intrahipocampal (i.h.) kainate-induced SE.

5.1.2 Severity of status epilepticus (SE) (II, III)The severity of SE was investigated in mice lacking uPA (uPA-/-) or uPAR (uPAR-/-) andwild type (Wt) controls. Preliminary analysis revealed that SE resolved 48 hr after kainateinjection; hence, the total number of spikes generated in EEG during the first 48 hr wasused to assess the severity of SE.

The total number of spikes was the same in uPA-/- mice and their Wt littermates (16553 ± 1876 vs. 19 173 ± 3474) (p>0.05) (II, Fig. 7). Similarly, the evolution of SE and the totalnumber of spikes did not differ between uPAR-/- mice and Wt mice (40 626 ± 14 035 vs. 58894 ± 11 426) (p>0.05) (III, Fig. 1).

5.1.2 Seizure phenotype characteristics (II, III)uPA-/- and Wt mice showed no difference in latency to first spontaneous seizure (p>0.05),seizure frequency (uPA-/-, 2.6 ± 0.49 sz/ day; Wt, 2.2 ± 0.4 sz/ day) (p>0.05) or seizureduration (uPA-/-, 44 ± 1 s vs. Wt, 37 ± 5 s) (p>0.05) (II, Fig. 7).

uPAR-/- and Wt mice showed no difference in latency to first spontaneous seizure(p>0.05), and seizure frequency (uPAR-/-, 1.1 ± 0.3 sz/ day; Wt, 1.5 ± 0.2 sz/ day) (p>0.05)(III, Fig. 1). However, in contrast to uPA-/- mice, the seizure duration was longer in uPAR-/- mice when compared to Wt mice (50.3 ± 1.8 s vs. 43.0 ± 2.6 s) (p<0.05) (III, Fig. 1).Furthermore, increased seizure frequency was associated with the severity of SE (r=0.507,p<0.05). However, this association was not seen in specific genotypes (uPAR-/- or Wtmice).

5.1.3 Severity of behavioral seizures (III)Behavioral analysis of seizures was performed only in uPAR-/- mice due to technicalissues related to our analogue video recording system. The analysis revealed that 74% of

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all seizures in uPAR-/- mice were scored at 3-5 (secondary generalized seizure) comparedto only 63% in Wt mice (p<0.01) (III, Fig. 1).

5.2 NEURODEGENERATION IN THE HIPPOCAMPUS (I, III)

5.2.1 Degeneration of principal cellsIn the pilocarpine induced-SE rat model, the severity of neurodegeneration was assessedat 2 d, 4 d, and 14 d post-SE. At all time points, severe neurodegeneration was observedbilaterally in pilocarpine injected animals with SE in the CA1 (p<0.05) and CA3 (p<0.05)subfields of the hippocampus and hilus (p<0.05) of the dentate gyrus when compared tosaline injected controls (I, table 1).

In order to establish whether uPA-uPAR systems play a role in neurodegenerationduring epileptogenesis, the severity of neurodegeneration was assessed using uPA-/- anduPAR-/- mice. The results of neurodegeneration and GCL dispersion in uPA-/- mice arealready published (for details see Lahtinen, 2010). Analysis was performed in uPAR-/- andWt mice at 4 d and 30 d following kainate-induced SE. Both genotypes experienced moresevere neurodegeneration in the ipsilateral more than contralateral hippocampus (III, Fig.3). At 4 d, neurodegeneration was already more severe contralaterally in uPAR-/-compared to Wt in the CA3 subfield (p<0.05) (III, Fig. 3). At 30 d neurodegenerationprogressed ipsilaterally in uPAR-/- and Wt mice in the CA1 subfield (p<0.05, compared to4 d). Also, progression of neurodegeneration was observed in both genotypes in thecontralateral CA1 subfield (p<0.05, compared to 4 d) but with a more robust effect inuPAR-/- mice than Wt mice (p<0.05) (III, Fig. 3).

5.2.2 Degeneration of hilar neuronsThe severity of degeneration of hilar neurons following pilocarpine-induced SE in rats wasassessed semiquantitatively at 2 d, 4 d, and 14 d after SE. The results demonstrated a >50%cell loss at all time points when compared to saline injected animals (I, table 1).

To investigate whether uPAR is involved in the degeneration of hilar neurons, thetotal number of hilar cells was counted in uPAR deficient mice following kainate-inducedSE. The results showed that at 4 d after i.h. kainate injection uPAR-/- and Wt miceexperienced 63% and 90% ipsilateral hilar cell loss respectively when compared to salineinjected controls (p<0.01). The magnitude of hilar cell loss in kainate-injected animals wassimilar in both genotypes (p>0.05). Contralaterally, kainate-injected uPAR-/- and Wt miceshowed a 39% and 43% cell loss respectively, compared to saline injected controls (p<0.01).The magnitude of contralateral hilar cell loss in kainate-injected animals was similar inboth genotypes (p>0.05). At 30 d, kainate injected uPAR-/- and Wt mice revealed 89% and

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91% ipsilateral hilar cell loss respectively, compared to saline injected controls (p<0.001).Contralaterally, uPAR-/- had 33% cell loss (p<0.05) while Wt mice showed no differencewhen compared to saline controls (III, Fig. 3).

5.2.3 Degeneration and dispersion of granule cellsThe effect on granule cell degeneration and dispersion of pilocarpine-induced SE, in rats,was assessed semiquantitatively at 2 d, 4 d and 14 d post-SE. Less than 10% damage wasnoticed at all time points. This was not statistically significant when compared to controls(p>0.05) (I, table 1). However, no dispersion of granule cell layer was observed at all timepoints analyzed after SE.

The role of uPAR in granule cell degeneration and dispersion was investigated inuPAR-/- and Wt mice by measuring the volume of the granule cell layer at 30 d after i.h.kainate-induced SE. The volume of the ipsilateral granule cell layer in uPAR-/- and Wtmice was increased by 2-fold in both genotypes when compared to saline injected controls(p<0.01). Contralateral granule cell layer volume did not vary between kainate and salineinjected uPAR-/- and Wt mice (p>0.05). Granule cell layer volume did not vary betweenkainate injected uPAR-/- and Wt mice ipsilaterally or contralaterally (p>0.05) (III, Fig. 4).About 5-10% damage in contralateral granule cells was noted in uPAR-/- (n=2) and Wt(n=1) mice.

5.3 NEUROGENESIS IN THE DENTATE GYRUS (I, III)

The magnitude of neurogenesis after the induction of SE was assessed in the dentate gyrusby counting the number of DCX-positive cells. In pilocarpine-induced SE rats, themagnitude of neurogenesis was assessed at 2 d, 4 d and 14 d Post-SE. No difference wasseen in the total number of DCX-positive cells at 2 d and 4 d when compared to salineinjected controls. However, at these time points, DCX-positive cells in SE animalsappeared scattered along the subgranular zone with their dendrites seen within thevicinity of soma as opposed to controls where there was a linear arrangement of cellsalong the subgranular zone with dendrites extending into the inner molecular layer. At 14d the number of DCX-positive cells was increased by 156% in SE animals compared tocontrols (p<0.01) (I, Fig. 11).

The effect of uPA or uPAR deficiency on post-SE neurogenesis was analyzed inuPA-/- and uPAR-/- mice at 20 and 30 d after i.h. kainate injection respectively. The resultsfrom the uPA-/- mice are already published (for details see Lahtinen, 2010). Kainate-injected uPAR-/- and Wt mice experienced a 142% (p<0.01) and 234% (p<0.01) increase inDCX-positive cell number respectively, compared to saline treated controls. The total

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number of DCX-positive cells in the dentate gyrus did not vary between kainate injecteduPAR-/- and Wt mice (p>0.05) nor did it vary in any of the sub-regions analyzed (hilus,subgranular zone and granule cell layer). Many of the DCX-positive cells in uPAR-/- micewere seen scattered and disorientated in the inner molecular layer of the dentate gyrus (III,Fig. 5).

5.4 MOSSY FIBER SPROUTING IN THE DENTATE GYRUS (I, III)

The volume of mossy fiber in the dentate gyrus of rats after pilocarpine-induced statusepilepticus was assessed at 30 d and 60 d post-SE. There was already a significant increasein mossy fiber sprouting at 30 d when compared with controls, with a median score of 3(p<0.05). At 60 d, the density of mossy fiber sprouting had increased even more whencompared with controls, with a median score of 5 (p<0.01).

The effect of uPAR deficiency on mossy fiber sprouting in the dentate gyrus wasassessed in uPAR-/- and Wt mice at 30 d after i.h. kainate injection. Analysis revealed a 7-fold increase in mossy fiber volume in kainate injected uPAR-/- and Wt mice whencompared to saline injected controls. However, the volume of mossy fiber was notdifferent between kainate-injected uPAR-/- and Wt mice (0.014 ± 0.004 mm3 vs. 0.014 ±0.009 mm3, p>0.05) (III, Fig. 6).

5.5 BLOOD-BRAIN BARRIER IMPAIRMENT IN THE HIPPOCAMPUS (I,III)

Blood-brain barrier (BBB) disruption in the hippocampus following pilocarpine-inducedSE in rats was assessed at 2 d, 14 d and 60 d after post-SE by measuring the degree of IgGleakage into the brain. At 2 d, there was a 3.9-fold increase in IgG immunoreactivity in thehippocampus when compared to controls (p<0.001). By 14 d post-SE, it dropped by 22%from its 2 d values but was still about 3-fold above control values (p<0.01). IgGimmunoreactivity at 60 d was the same as at 14 d and 2.9-fold above control levels (p<0.01)(I, Fig. 8).

To determine whether uPAR is involved in post-SE BBB impairment, the degree ofIgG leakage into the hippocampus was assessed in uPAR-/- and Wt mice at 4 d after i.h.kainate injection. Kainate-injected uPAR-/- and Wt mice experienced a significant increasein IgG immunoreactivity ipsilaterally and contralaterally when compared to saline treatedcontrols. However, no genotype difference was observed in the magnitude of IgGimmunoreactivity ipsilaterally or contralaterally (p>0.05) (III, Fig. 7). It was observed thatthe degree of contralateral IgG immunoreactivity was associated with the severity of

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hippocampal damage (r=0.0675, p<0.05). This association was lacking in the ipsilateralkainate-injected hippocampus.

5.6 INFLAMMATORY RESPONSE IN THE HIPPOCAMPUS (III)

In order to determine whether uPAR is involved in post-SE inflammatory response, themagnitude of macrophage and T-cell response was assessed in uPAR-/- and Wt mice at 4 dand 30 d post-SE. Saline injected animals demonstrated very mild microglia response andno T-cell response at 4 d and 30 d after injection.

Macrophage immune responseAt 4 d post-SE, ipsilateral macrophage response in kainate-injected uPAR-/- was mildcompared to in Wt (p<0.05). The difference was prominent in the CA1 and CA3 subfieldsof the hippocampus (p<0.05) but not in the hilus of the dentate gyrus (p>0.05).Contralateral macrophage response was similar in both genotypes. At 30 d, the magnitudeof ipsilateral macrophage response was similar in uPAR-/- and Wt mice. However,contralaterally, macrophage response was more prominent in uPAR-/- mice compared toWt mice (p<0.05). This was seen particularly in CA1 and hilus (p<0.05) but not in CA3(p>0.05) (III, Fig. 8, 9).

T-cell immune responseT-cell response at 4 d was mild in uPAR-/- mice compared to Wt mice, especially in theCA3 subfield (p<0.05). At 30 d, the magnitude of T-cell response in uPAR-/- and Wt miceincreased exponentially compared to at 4 d. However, uPAR-/- and Wt mice showed nodifference in T-cell response at this time point (III, Fig. 10).

5.7 HIPPOCAMPAL VASCULAR MORPHOMETRY (I, II, III)

5.7.1 Endothelial cell proliferation, blood vessel length and diameter change (I, II, III)In order to establish the occurrence of angiogenesis during epileptogenesis and whetherthe blood vessels are affected by post-SE events, the number of proliferating endothelialcells, blood vessel length and diameter were assessed in pilocarpine-induced SE rats.

Endothelial cell proliferation and total blood vessel length in the hippocampus wasassessed at 2 d, 4 d and 14 d post-SE. Endothelial cell proliferation was 3-fold at 2 d(p<0.01), 10-fold at 4 d (p<0.001) and 3-fold at 14 d (p<0.05) when compared to controls.Subfield analysis in the CA1, CA3 and molecular layer of the dentate gyrus revealed asimilar pattern, being elevated at 2 d, peaking at 4 d and returning to control levels by 14 d(I, Fig. 6).

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The total blood vessel length reduced to 83% of control values at 2 d post-SE(p<0.05). It returned to control values at 4 d post-SE. However, by 14 d post-SE, it was 6%above control levels (p>0.05). At 2 d, the most significant drop in blood vessel length wasobserved in the CA1 subfield (88% of control values) (p<0.05). At 4 d, vessel length in allsubfields was similar to controls. At 14 d post-SE, the most pronounced increase in vessellength was in the CA3 subfield (128% above control levels) (p<0.05) (I, Fig. 5).

The blood vessel diameter increased by 125% at 2 d and 111% at 14 d whencompared to controls. Increased vessel diameter was much more pronounced in thehippocampal fissure where it was 125% at 2 d and 122% at 4 d when compared to controls(I, Fig. 9).

5.7.2 Association of angiogenesis and hippocampal CBV and CBF (I)To establish whether there are any functional changes associated with the observed bloodvessel length changes, the CBV and CBF were measured using MRI at 2 d and 14 dfollowing pilocarpine-induction in rats. The results revealed a 178% increase in CBV from2 d to 14 d post-SE (p<0.05) in the same animal and a 124% increase when compared tocontrols (p=0.88). This was associated with the increased vessel length observed from 2 dto 14 d post-SE. No changes were observed in CBF from 2 d to 14 d post-SE in the sameanimal (p=0.075), or when compared to controls (p=0.055) (I, Fig. 10).

5.7.3 Association of angiogenesis with neurodegeneration, neurogenesis and mossyfiber sprouting (I)In order to establish whether there is any interaction between the observed vessel lengthchanges and neuronal plastic changes during epileptogenesis, we assessed the correlationbetween blood vessel length changes and neurodegeneration, neurogenesis, and mossyfiber sprouting. No correlation was found between vessel length and neurodegenerationin all hippocampal subfields analyzed (CA1 and CA3) at 2 d, 4 d and 14 d post-SE (p>0.05).However, at 14 d, a strong association was seen between endothelial cell proliferation andneuronal damage score (r=0.833, p<0.05) (I, Fig. 6). No correlation was seen between vessellength or proliferating endothelial cells and neurogenesis at 2 d, 4 d and 14 d post-SE(p>0.05). Neither was there any correlation between vessel length and mossy fibersprouting at 14 d and 60 d post-SE (p>0.05).

5.7.4 Effect of uPA or uPAR deficiency on vascular morphometry (II, III)In an effort to establish whether uPA and uPAR play a role in vascular remodeling in thehippocampus, during epileptogenesis, the blood vessel length was assessed in uPA oruPAR deficient mice after i.h. kainate-induced SE.

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In uPA-/- mice, the blood vessel length was assessed at 20 d post-SE. Saline andkainate-injected uPA-/- and Wt mice showed no difference in total blood vessel length,ipsilaterally and contralaterally. Ipsilateral and contralateral blood vessel length wassimilar in kainate-injected uPA-/- and Wt mice (II, Fig. 6).

On the other hand, in uPAR-/- mice the blood vessel length was assessed at 30 dafter i.h. kainate-induced SE. Saline-injected uPAR-/- and Wt mice showed no difference intotal blood vessel length bilaterally. Total blood vessel length in kainate-injected uPAR-/-and Wt mice was similar to saline-injected controls bilaterally. Kainate-injected uPAR-/-and Wt mice showed similar ipsilateral blood vessel length. However, contralaterally, theblood vessel length was shorter in uPAR-/- compared to Wt mice (p<0.05) (III, Fig. 11).Septo-temporal profiling of vessel length changes in the hippocampus indicated asignificant increase in blood vessel length at 2 mm caudal to injection site in kainate-injected Wt mice when compared to saline-injected controls. However, similar profiling inkainate-injected uPAR-/- mice revealed no change when compared to their saline-injectedcontrols (III, Fig. 11).

Figure 3. Epileptogenic network alterations in uPA and uPAR deficient mice afterintrahippocampal kainate-induced status epilepticus (SE). (A) In uPA deficient mice theepilepsy phenotype did not vary with wild type mice. However, they demonstrated severeneurodegeneration and impaired neurogenesis. (B) In uPAR deficient mice, the epilepsyphenotype was severe compared to Wt mice. This associated with severe neurodegeneration,impaired angiogenesis, impaired neuronal migration and delayed acute macrophage and T cellresponse. Adapted from Crittenden et al., 2007

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Table 1. Summary of the epilepsy phenotype and epileptogenic network alterations in thehippocampus of uPA and uPAR deficient mice following intrahippocampal injection of kainicacid.

uPA uPAR

uPA-/- Wt uPAR-/- Wt

SE (total spikes/ 48h) 16553 ± 1876 19173 ± 3474 40626 ± 14035 58894 ± 11426

Epilepsy phenotypesz latency (hr)sz frequency/ daysz duration (sec)total number of sz (30 d)median sz score

52.2 ± 20.4 49.6 ± 11.4 88.0 ± 17.9 56.6 ± 11.72.06 ± 0.49 2.18 ± 0.44 1.1 ± 0.3 1.5 ± 0.244.0 ± 1.0 37.0 ± 5.0 50.3 ± 1.8 * 43.0 ± 2.661 ± 15 65 ± 13 33.0 ± 7.5 43.8 ± 6.3NA NA 5 * 3

NeurodegenerationContra (% damage)

acutechronic

Hilar cell numberipsicontra

GCL volume (mm3)ipsicontra

25% (6 d) 33% (6 d) 33% * (4 d) 17% (4 d)67% * (20 d) 33% (20 d) 50% * (30 d) 17% (30 d)

1308 ± 273 1071 ± 131 758 ± 232 680 ± 2124555 ± 533 * 7515 ± 556 5033 ± 654 7068 ± 576

1.11 ± 0.11 1.27 ± 0.06 0.97 ± 0.09 1.06 ± 0.090.43 ± 0.03 0.37 ± 0.02 0.53 ± 0.02 0.53 ± 0.02

Neurogenesis (DCX)(uPA, 20 d; uPAR, 30 d)

totalhilusSGZGCL

6221 ± 878 * 10020 ± 459 16820 ± 2706 22920 ± 2893556 ± 109 975 ± 175 417 ± 202 208 ± 865224 ± 775 * 8117 ± 488 13830 ± 2118 20030 ± 2483442 ± 94 929 ± 225 2566 ± 583 2673 ± 476

BBB impairment (4 d)ipsicontra

NA NA 23.29 ± 2.28 24.03 ± 46.1NA NA 7.19 ± 10.21 23.05 ± 5.17

Blood vessel length (m)(uPA, 20 d; uPAR, 30 d)

ipsicontra

2.39 ± 0.18 2.48 ± 0.09 1.96 ± 0.10 2.19 ± 0.062.53 ± 0.13 2.63 ± 0.09 1.89 ± 0.12 * 2.31 ± 0.09

Mossy fiber (mm3) (30 d) NA NA 0.014 ± 0.004 0.014 ± 0.009

InflammationMacrophage- 4 d

ipsicontra

Macrophage - 30 dipsicontra

T-cell (cells/mm3)4 d30 d

NA NA

25.59 ± 2.31 * 30.97 ± 0.92NA NA 15.93 ± 3.66 19.33 ± 1.94NA NA

19.12 ± 1.19 18.93 ± 0.64NA NA 10.49 ± 1.35 * 5.98 ± 0.96NA NANA NA 54 ± 14 * 134 ± 29NA NA 185 ± 15 154 ± 15

Hippocampal damage is indicated as percentage of total cell loss. Hilar cell number, GCLvolume, neurogenesis and blood vessel length was assessed in uPA-/- mice at 20 d post-SEand in uPAR-/- at 30 d post-SE. BBB impairment and macrophage response are measured inarbitrary units. Abbreviations: BBB, blood-brain barrier; contra, contralateral; DCX,doublecortine; GCL, granule cell layer; hr, hour; ipsi, ipsilateral; m, meter; sec, second; SGZ,subgranular zone; sz; seizure; NA, not analyzed. *p<0.05, as compared to Wt mice. Data arepresented as mean ± SEM

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6. Discussion

The aim of this thesis was first to investigate the interaction between the cellularreorganization processes known to occur during epileptogenesis, particularly exploringthe interaction between angiogenesis and neuronal plastic events, since common trophicstimuli are involved in vascular network and nerve wiring (Carmeliet and Tessier-Lavigne, 2005). The second part of the thesis focused on studying the mechanisms thatunderlie the cellular reorganization processes occurring during epileptogenesis. The uPA-uPAR system was chosen as a possible mechanism, since its expression is increased in thehippocampus during epileptogenesis (Lahtinen et al., 2006, 2009; Iyer et al., 2010).Furthermore, uPA and uPAR is implicated in several physiological and pathologicalprocesses involving tissue remodeling (Sumi et al., 1992; Meiri et al., 1994; Irigoyen et al.,1999; Wu YP et al., 2000). This suggests that uPA-uPAR may be a regulatory notch inepileptogenic network reorganization.

The results presented in this thesis demonstrate the following: first, angiogenesisrestores hippocampal blood vessel length during epileptogenesis and correlates withcerebral blood volume increase in hippocampus. Second, angiogenesis in thehippocampus lacks association with regions of severe neurodegeneration as well as withneurogenesis or axonal sprouting during epileptogenesis. Third, the epilepsy phenotype issevere in uPAR deficiency but not uPA deficiency. Fourth, uPA or uPAR deficiency resultsin severe neurodegeneration and impaired neurogenesis. Fifth, the process of angiogenesisis impaired in uPAR deficiency, whereas it is not affected in uPA deficiency. Sixth, uPARdeficiency results in a delayed acute inflammatory response which subsequently becomeschronic. Lastly, uPAR deficiency does not affect post-SE BBB impairment or axonalsprouting.

6.1 METHODOLOGICAL CONSIDERATIONS

Two SE animal models of human temporal lobe epilepsy (TLE) were used in the presentstudy, the pilocarpine-induced SE rat model and the intrahippocampal kainate-induced SEmouse model. The reason why the pilocarpine-induced SE rat model was chosen is, firstly,this model replicates some of the behavioral, electrographic and neuropathologicalfeatures of human TLE (Turski et al., 1984). Secondly, the model produces very severeneurodegeneration, robust neurogenesis and mossy fiber sprouting, which suits the

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purpose of investigating the interaction between these processes and angiogenesis(Cavalheiro et al., 2006).

The intrahippocampal kainate-induced SE mouse model shares someelectrographic, histopathologic and synaptic reorganization similarities with human TLE(Bouilleret et al., 1999, Riban et al., 2002). This model was used because the C57BL/6 mousebackground is resistant to i.p. kainate or pilocarpine induced cell death (Schauwecker,2002). To minimize variability in kainate response, appropriate control animals werechosen. For uPA deficient mice, Wt littermates were used as controls. For uPAR deficientmice, Wt mice of the C57Bl/6 genetic background produced from the same colony as theknock-out were used as controls. Littermates were not used for uPAR-/- mice experimentsdue to difficulties encountered during heterozygote breeding.

6.2 EPILEPSY PHENOTYPE IN uPA AND uPAR DEFICIENT MICE

There is increased expression of uPA and its receptor uPAR in humans and animal modelsof temporal lobe epilepsy (Lahtinen et al., 2006, 2009; Iyer et al., 2010; Liu et al., 2010).However, knowledge on the functional role of uPA and uPAR during epileptogenesis andthe subsequent epilepsy outcome remains poor. In order to address this question SE wasinduced by intrahippocampal injection of kainate into mice lacking either uPA (uPA-/-) oruPAR (uPAR-/-) and the development of epileptogenesis was followed by continuous(24/7) video-EEG monitoring.

Our results indicate that in uPA-/- mice the severity of the initial insult (i.e. SE) didnot differ from Wt mice. Analysis of the epilepsy phenotype showed no differencebetween uPA-/- and Wt mice. Similarly, the severity of SE did not differ between uPAR-/-mice and their Wt controls. uPAR-/- and Wt mice showed no difference in latency to firstspontaneous seizure and seizure frequency. However, unlike uPA-/- mice, the seizureduration was 16% longer in uPAR-/- mice when compared to Wt mice. Furthermore,uPAR-/- mice demonstrated severe behavioral seizures, typically secondary generalized asopposed to partial seizures in Wt mice. This indicates that the epilepsy phenotype is moresevere in uPAR-/- than in uPA-/- mice. However, it was interesting to note that the averageseizure frequency in uPA-/- and their Wt control mice was two times higher than that inuPAR-/- and their Wt control mice, even though their (uPA-/- mice) total number of spikeswas only about half of that in uPAR-/- mice (see table 2). Whether this is related to theexperimental setting, since experiments were performed at different times, or geneticbackground, remains to be investigated.

Previous studies have reported that uPAR-/- mice have a lowered threshold forpentyleneterazol (PTZ)-induced seizures, showed epileptiform spiking in EEG, and about

49

5.9% of them express spontaneous seizures during handling (Powell et al., 2003). Howeverthe behavioral manifestations as reported by Powel and colleagues (2003), are differentfrom ours. They associated the seizure susceptibility and spontaneous seizures with areduced number of parvalbumin cells in the fronto-parietal cortex. Our preliminary datadid not reveal any major difference in the number of GAD67 or parvalbumin positive cellsbetween uPAR-/- and Wt mice. The differences between the laboratories can relate todifferences in the genetic background of uPAR-/- mice.

Taken together, our data suggest that lack of uPAR but not uPA can worsen theepilepsy phenotype of acquired epilepsy. To understand the mechanisms associated withthe epilepsy phenotype in uPA-/- and uPAR-/- we next investigated the epileptogeniccircuitry reorganization at the site of primary focus, that is, the hippocampus.

6.3 NEURODEGENERATION IS AFFECTED BY uPA AND uPAR

Neuronal loss in the hippocampus and hilus of the dentate gyrus are amongst the mostprominent histopathological findings in the resected hippocampus of patients withtemporal lobe epilepsy (TLE) (Honavar and Meldrum, 1997). In the current study, we seekto elucidate the evolution of neurodegeneration in the hippocampus and the dentategyrus, and whether the uPA-uPAR system is the underlying mechanism in this process.

Pilocarpine-induced SE resulted in severe cell loss in all hippocampal subfields andin the hilus of the dentate gyrus as early as 2 days post-SE. However, very little damagewas seen in the granule cell layer. These findings are in line with previous studies in thepilocarpine-induced SE model (Turski et al. 1983, de Lanerolle et al., 1989; Obenaus et al.,1993; Houser and Esclapez, 1996). Analysis of neurodegeneration in uPA deficient micerevealed that genotype had no effect on acute pyramidal cell loss in the hippocampus.These data are in line with observations by Tsirka et al. (1997) who found no difference inhippocampal neurodegeneration between uPA and Wt mice at 5 days after i.h. KA.Thus, uPA does not have a significant effect on acute post-injury neurodegeneration.However at chronic time points, neurodegeneration had progressed significantly in uPA-/.On the other hand, acute pyramidal cell loss was severe in uPAR-/- and progressedsignificantly by 30 days post-SE.

The robust delayed cell death in uPA-/- mice suggests that uPA deficiency has anunfavorable effect on recovery by compromising the survival of remaining neurons. Thisis supported by studies demonstrating poor recovery of uPA-/- mice after TBI (Morales etal., 2006) or sciatic nerve crush (Siconolfi and Seeds, 2001). Also, previous in vivo and invitro studies in glioma models have suggested that blocking uPAR either by mRNAinterference or genetic deletion favors the expression of pro-apoptotic molecules like BAD,

50

Bax and Bal over anti-apoptotic molecules, resulting in increased cell death (Besch et al.,2007, Malla et al., 2012). Thus the increased cell death in uPA-/- and uPAR-/- mice couldpartly be explained by the fact that uPA or uPAR deficiency reduces interaction of theuPA-uPAR complex with various transmembrane receptors that initiate intracellularsignaling cascades for cell survival, proliferation, and growth (Blasi and Carmeliet, 2002;Alfano et al., 2005; Hildenbrand et al., 2008). Another likely explanation for the facilitatedneurodegeneration in uPAR-/- mice is an augmented activation of pro-apoptoticmechanisms during SE, as discussed below, compared to Wt mice.

Taken together, our data demonstrate that only uPAR deficiency has an effect onacute neurodegeneration whereas uPA or uPAR deficiency enhances delayed post-injuryneurodegeneration. This suggests that uPA and uPAR are involved in post-SEneurodegeneration.

6.4 uPA AND uPAR ARE INVOLVED IN NEUROGENESIS

Status epilepticus induces remarkable neurogenesis in the dentate gyrus (Parent et al.,1998; Scott et al., 1998; Kralic et al., 2005). Similarly, a significant increase in DCX positivecells was noted at 14 d following pilocarpine-induced SE. We further investigated whetherthe uPA-uPAR system is implicated in post-SE neurogenesis in the intrahippocampalkainate-induced SE model. In this model, the injected hippocampus contained only veryfew DCX positive cells, suggesting that either kainic acid itself or seizure-inducedexitotoxicity had killed the neuronal precursor cells. Therefore, analysis was limited to thecontralateral hippocampus. Considering the role of uPA and uPAR in cell proliferationand migration, especially as depicted with cancer cells (Sumi Y. et al., 1992; Meiri N. et al.,1994; Irigoyen et al., 1999; Wu YP et al., 2000), and in particular the role of uPAR-mediatedsignaling in the activation of mitogenic activity of the hepatocyte growth factor (Moriyamaet al., 1999; Powell et al., 2001, Levitt, 2005), we expected to see a change in the magnitudeof post-SE neurogenesis and migration of newly-born cells in the dentate gyrus in uPA-/-or uPAR-/- mice.

The number of DCX positive cells in uPA-/- mice was unchanged when comparedto saline treated controls, whereas, in Wt mice, it increased by 18% when compared tocontrols. In addition, subfield analysis showed less DCX-positive cells in the subgranularzone and granule cell layer in uPA-/- compared to Wt mice. This indicates compromisedgeneration and migration of DCX-positive cells within the dentate granule cell layer ofuPA-/- mice. On the other hand, uPAR-/- and Wt mice showed a 1.4 and 2.3-fold increasein DCX-positive cells respectively with no apparent genotype differences. However, manyof the DCX positive cells in uPAR were scattered and disorientated within the granule cell

51

layer and inner molecular layer, indicating the loss of directional or cued proteolyticmigration.

Previous studies have demonstrated that animals with severe hippocampal injuryhave a more dramatic decrease in neurogenesis than animals with mild injury(Hattiangady et al., 2004; Heinrich et al., 2006). However, this reason alone cannot justifythe mild neurogenesis in uPA-/- mice, since uPAR-/- also experienced severeneurodegeneration but did not experience a poor magnitude of neurogenesis.

Mossy fiber sprouting, one of the most studied types of axonal plasticity inexperimental and human epilepsy (Sutula and Dudek, 2007), was noted in the dentategyrus in rats following pilocarpine-induced SE. A significant amount of mossy fibersprouting was observed at 30 d and 60 d post-SE. We next assessed whether uPARdeficiency affects axonal spouting. As expected based on the meagerness of newly-borncells ipsilaterally in the septal hippocampus, there was no remarkable mossy fibersprouting ipsilaterally in uPAR-/- mice. Contralaterally, we found substantial sproutingalong the entire septotemporal axis without any difference between the genotypes. Thisindicates that uPAR has no effect on axonal plasticity.

Taken together these data suggest that while uPA may modulate the course ofproliferation and maturation of neuronal precursors during neurogenesis, uPAR mayinfluence connectivity of the dentate gyrus by affecting the migration of newly-born cells.

6.5 ANGIOGENESIS IS AFFECTED BY uPAR BUT NOT BY uPA

There are several reports demonstrating increased vascular density in the resectedhippocampus of patients with temporal lobe epilepsy and in animal models (Sokrab et al.,1989; Seiffert et al., 2004). In this study, we assessed the spatiotemporal evolution ofangiogenesis in the hippocampus after status epilepticus and studied whether the uPA-uPAR system is involved in post-SE vascular remodeling.

At 2 d after pilocarpine-induced SE, the total blood vessel length as revealed byRECA-1 staining reduced to 80% of control values but progressively recovered to controlvalues by 4 d and was even 6% above control level by 14 d. Similarly, endothelial cellproliferation increased from 3-fold at 2 d to 10-fold at 4 d and reduced to 3-fold by 14 d.Interestingly, endothelial cell proliferation correlated positively with blood vessel length at4 d, suggesting that endothelial cells do actually contribute to vessel build-up. A similarpattern of vessel length changes and endothelial cell proliferation was observed in themolecular layer of the dentate gyrus and the CA1 and CA3 subfield of the hippocampus. Itwas also revealed that at 2 d and 14 d post-SE, the blood vessel diameter increased in thehippocampus and especially in the hippocampal fissure. This was probably as a result of

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clot formation since a number of clots were found at 2 d post-SE, and these clots werepresent up until 14 d post-SE, though in lower numbers.

Rigau et al. (2007) reported a 200% increase in hippocampal vessel density atchronic phase in Li-pilocarpine treated rats. Major discrepancies between this study andours are that blood vessels were detected using either endogenous peroxidase or vonWillebrand factor. Also, they did not use unbiased stereological analysis to quantify bloodvessel length. Similar to our findings is the study by Hellsten et al. (2005) which showed a16% increase in hippocampal vessel length after 10 daily electroshocks when assessed at21 d using unbiased stereology in RECA-1 stained sections.

In order to demonstrate whether the uPA-uPAR system is involved in post-SEvascular remodeling, we analyzed the blood vessel length in uPA and uPAR deficientmice after SE. Studies in the cancer field show that uPA-uPAR plays a role in endothelialcell proliferation, adhesion, and migration during cancer angiogenesis (Deng et al., 1996,Fibbi et al., 1998). Hence one will expect that post-SE vascular remodeling will be affectedin uPA or uPAR deficiency.

Stereological measurements of total blood vessel length revealed no differencebetween uPA-/- and Wt mice at 20 d post-SE, ipsilaterally and contralaterally. Also, thetotal blood vessel length in kainate-injected uPA and Wt mice did not return to controllevels or beyond, which would have suggested ongoing angiogenesis. Whether or not thisis due to a short follow-up time warrants further studies. The finding that uPA deficiencydid not affect vessel length after kainate injection is in agreement with observations byLund et al. (2006) showing that wound healing in uPA-/- mice is moderately delayed butnot completely blocked.

On the other hand, there were no major changes in the total blood vessel lengthipsilaterally or contralaterally in uPAR-/- and Wt mice. However, when the vessel lengthwas estimated at different levels of the septotemporal axis of the hippocampus, Wt micewith SE showed an increased vessel length over 2 mm caudal to the injection site. This wasnot seen in uPAR-/- mice with SE, suggesting compromised post-SE vascular remodeling.

Taken together, these results demonstrate that uPAR seems to be mandatory forangiogenesis since deficiency of uPA did not affect angiogenesis, which is in line withobservations in cancer studies (Deng et al., 1996, Fibbi et al., 1998, Bu et al., 2004, Lakka etal., 2005).

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6.6 ASSOCIATION BETWEEN ANGIOGENESIS AND NEURONAL PLASTICEVENTS

Very few studies have demonstrated that vascular damage and consequent angiogenesisinduced by epileptogenic brain insults is prominent in the region of most severeneurodegeneration, that is, in areas where most of the oxygen and glucose is needed forthe repair process. Along the same line of thought, no studies have demonstrated anassociation between angiogenesis and neurogenesis or axonal sprouting to support theidea of common trophic stimuli at neurovascular niches during epileptogeneis (Carmelietand Tessier-Lavigne, 2005). In an attempt to resolve these questions, we investigated theassociation between blood vessel length changes and neurodegeneration, neurogenesisand mossy fiber sprouting after pilocarpine-induced status epilepticus.

Correlation analysis revealed no clear association between blood vessel lengthchanges and neuronal damage score. In addition, contrary to our hypothesis thatangiogenesis would be most robust in those regions with severe neurodegeneration, thenumber of proliferating endothelial cells did not correlate with the injury score in any ofthe hippocampal subfields at early post-SE. These observations suggest that the milieu atthe site of neurodegeneration, including inflammatory response and cytokine release aswell as activation of extracellular proteinases, does not stimulate endothelial cellproliferation but does not suppress it either in the model used (see Marchi et al., 2007;Pitkänen and Lukasiuk, 2009).

In the present study, we expected the creation of an association betweenangiogenesis and the degree of neurogenesis and axonal sprouting to support the idea thatcommon trophic stimuli are involved in nerve and blood vessel wiring (Carmeliet andTessier-Lavigne, 2005). Endothelial cell proliferation was observed as early as 2 d post-SE,peaking at 4 d and dropping down to control levels by 14 d. On the other hand, like inprevious studies, a significant increase in DCX-positive cells was not observed earlier than14 d (Hattiangady et al., 2004; Rao et al., 2008). This suggests that the time course ofendothelial cell proliferation after injury is much faster than neurogenesis, supporting thefinding that no association was observed between neurogenesis and angiogenesis duringearly post-SE. However, this does not rule out the possibility that mechanisms drivingthese processes could be overlapping. Such overlaps have been documented in works byPalmer et al. (2000) where they showed neural progenitor cells in close contact withendothelial cells in the infragranular layer of the dentate gyrus. In addition, studies havedemonstrated close association between newly-born immature neurons and remodelingvasculature in the subventricular zone (SVZ), recruited by trophic action of stromal-derived factor (SDF-1) and angiopoietin 1 (Ang1) upregulated by endothelial cells (Ohab et

54

al., 2006). Similar cross-talk between these two processes has been demonstrated inhumans (Chairetti et al., 2008).

Previous studies have suggested that endothelial cell could secrete moleculesfacilitating axonal growth (Carmeliet and Tessier-Lavigne, 2005; Ozdinler et al., 2006;Chiaretti et al., 2008; Zacchigna et al., 2008). For example, Honma and colleagues (2002)were able to show using artemin (ARTM) deficient mice that ARTM, a vascular-derivedneurotrophic factor, promotes migration and axonal projections of sympatheticneuroblasts. After SE, a condition known to trigger a remarkable growth of granule cellaxons or mossy fibers, we did not, however, find any association between the number ofproliferating endothelial cells or the vessel length and density of mossy fiber sprouting inthe dentate gyrus. Also, the time courses of endothelial cell proliferation and mossy fibersprouting were different. Sprouting continues for several weeks to months. These datasuggest that, after SE, the mechanisms maintaining angiogenesis and axonal sproutingmay be separate. Whether there is overlap in the molecular mechanisms that trigger bothphenomena at early post-SE phase remains to be explored.

6.7 INFLAMMATION IS MODULATED IN uPAR DEFICIENT MICE

A growing amount of evidence suggests that inflammation plays a role in epileptogenesisand epilepsy (Ravizza et al., 2008; Maroso et al., 2010; Vezzani et al., 2009). Microglialactivation and leukocyte infiltration in sclerotic tissue of patients with mesial temporallobe epilepsy (TLE) and animal models of TLE contributes to the development of epilepsy(Fabene et al., 2008). uPAR is expressed by activated microglia/macrophages after variousbrain injuries, including SE, but its function in these cells is unknown (Beschorner et al.,2000). In addition, previous studies have shown that uPAR is expressed in stimulated Tcells, and it is involved in their migration, differentiation, and phagocytosis via itsinteractions with integrins on the cell surface (Gyetko et al., 1994; Simon et al., 2000; Gyetkoet al., 2001; Smith and Marshall, 2010).

Here, we show using a CD68 antibody, which stains both the phagocytoticperipheral macrophages and resident microglia, that at 4 d post-SE, the acute hippocampalmacrophage response in uPAR-/- mice was milder than that in Wt mice. Interestingly, theCD68 response in uPAR-/- mice was also milder in the extrahippocampal areas like theamygdaloid complex and the hypothalamus. Even though the overall macrophageresponse became alleviated over the 30-d follow-up, dropping to 20-30% of that at 4 dpost-SE, the density of immunopositive CD68 cells in uPAR-/- mice remained 2-foldhigher in many hippocampal subfields, including CA1 as compared to that in Wt mice.

55

We show that, at 4 d post-SE, there was an invasion of T cells into thehippocampus, which is in line with previous studies using the intrahippocampal kainatemodel (Chen et al., 2004). However, the acute T-cell counts in the CA3 subfield of thehippocampus in kainate-injected uPAR-/- mice were only about 40% of that in Wt mice.This is in line with previous studies in experimental lung infection, demonstrating morethan 50% alleviation of acute T cell response in uPAR-/- mice (Gyetko et al., 2001). As weshow here the difference in T cell number was not related to the difference in blood-brain-barrier damage as IgG immunoreactivity at 4 d post-SE was comparable between thegenotypes. A more likely explanation for the difference is the compromised migratorycapacity of T cells from peripheral blood to brain parenchyma in uPAR-/- mice.Interestingly, contrary to the CD68 cells which decreased during the follow-up, T cellcounts in uPAR-/- mice showed a 10-fold increase over time, and reached the same level oreven beyond that in Wt mice during the 30-d follow-up.

These data demonstrate that uPAR is involved in post-SE inflammatory response,by affecting the recruitment of inflammatory cells. Furthermore, this data shows uPAR isnot involved in post-SE blood-brain barrier impairment.

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7. Conclusions

The aim of this thesis was to study the interaction between the cellular reorganizationprocess known to occur during epileptogenesis, their functional implications and themolecular mechanism that underlie these processes. Special focus was placed on theinteractions between angiogenesis and neuronal plastic event. The uPA-uPAR system wasinvestigated as a possible candidate mechanism in epileptogenic network alterations. Themain findings of this thesis can be summarized as follows:

1. uPAR but not uPA is implicated in the severity of the epilepsy phenotype followinga brain injury such as SE

2. Epileptogenic network alterations following SE, such as neuronal cell loss andneurogenesis, are regulated by uPA and uPAR

3. Vascular network changes post-SE correlates with functional changes in cerebralblood volume in the hippocampus but lacks association with neuronal plasticityduring epileptogenesis.

4. uPAR plays a stronger role than uPA in the progressive vascular buildup in thehippocampus during epileptogenesis.

5. uPAR is implicated in the severity of the inflammatory response duringepileptogenesis

In summary, this work provides novel information on the interaction of epileptogenicnetwork alterations. Furthermore, it extends our knowledge on the mechanisms involvedin these network alterations that are a potential target for therapeutic interventions.

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Appendix

ORIGINAL PUBLICATIONS (I – III)

I

Vascular changes in epilepsy: Functional consequences and association with network

plasticity in pilocarpine-induced experimental epilepsy.

Ndode-Ekane XE, Hayward N, Grohn O, Pitkanen A

Neuroscience. 166:312-332, 2010

Reprinted with kind permission from Elservier

II

Urokinase-type plasminogen activator regulates neurodegeneration and neurogenesis

but not vascular changes in the mouse hippocampus after status epilepticus.

Lahtinen L, Ndode-Ekane XE, Barinka F, Akamine Y, Esmaeili MH, Rantala J, Pitkanen A

Neurobiol. Dis. 37:692-703, 2010

Reprinted with kind permission from Elservier

III

Urokinase-type plasminogen activator receptor modulates epileptogenesis

in mouse model of temporal lobe epilepsy

Ndode-Ekane XE and Pitkänen A

Submitted

Publications of the University of Eastern Finland

Dissertations in Health Sciences

isbn 978-952-61-0990-9

Publications of the University of Eastern FinlandDissertations in Health Sciences

dissertatio

ns | 14

4 | Xav

ier Ek

olle N

do

de-E

ka

ne | D

evelopm

ent of Epilep

togenic Netw

ork Alteration

s in Rodent M

odels of Statu

s...

Xavier Ekolle Ndode-EkaneDevelopment of Epileptogenic

Network Alterations in Rodent Models of Status Epilepticus:

Role of the urokinase-type plasminogen

activating system


Development of Epileptogenic Network Alterations in Rodent Models of Status Epilepticus:Role of the urokinase-type plasminogen activating

system

Temporal lobe epilepsy is the

most common form of acquired

epilepsy. This thesis explored the

interaction between the cellular

reorganization processes occurring

during epileptogenesis, their

functional implications, and the role

of the urokinase-type plasminogen

activator (uPA) and its receptor

(uPAR) in epileptogenic network

alterations. It shows that components

of epileptogenic network alterations

may have different temporal

profiles, and the Plau and Plaur

genes encoding for uPA and uPAR

respectively are modifier genes for

acquired epileptogenesis.

dissertations xavier ekolle ndode-ekane development of...

Documents