Characterization of α-actinin as a member of the spectrin superfamily of proteins in "Neurospora crassa"

Thesis

Reference

Characterization of α-actinin as a member of the spectrin superfamily of proteins in "Neurospora crassa"

COTADO-SAMPAYO, Marta

Abstract

Nous avons étudié le rôle de l' α-actinine dans le développement de "Neurospora crassa". A l'aide des outils bioinformatiques, nous avons trouvé un gêne codant pour l'α-actinine puis nous avons caractérisé ce dernier biochimiquement. Nous avons déterminé sa localisation "in situ" et "in vivo" pendant la germination et pendant la croissance hyphale. Bien que le rôle exact de l'α-actinine n'ait pas pu être totalement élucidé, l'étude phénotypique du transformant ("knock-out" partiel de l'α-actinine) nous amène à penser que cette protéine participe avec l'actine à la coordination de la germination, la formation des septa et l'établissement des ramifications hyphales lors de la croissance du champignon. En outre, nous avons pu déterminer que l'α-actinine est la seule protéine appartenant à la superfamille des spectrines chez les champignons filamenteux, les levures et les Oomycètes et représente un membre

"primitif" de cette superfamille.

COTADO-SAMPAYO, Marta. Characterization of α-actinin as a member of the spectrin superfamily of proteins in "Neurospora crassa". Thèse de doctorat : Univ. Genève, 2008, no. Sc. 3976

URN : urn:nbn:ch:unige-18247

DOI : 10.13097/archive-ouverte/unige:1824

Available at:

http://archive-ouverte.unige.ch/unige:1824

Disclaimer: layout of this document may differ from the published version.

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UNIVERSITÉ DE GENÈVE FACULTÉ DES SCIENCES Département de botanique et biologie végétale Professeur Reto J. Strasser Laboratoire de bioénergétique et microbiologie Dr François Barja

Characterization of α-actinin as a member of the spectrin superfamily of proteins in Neurospora crassa

THÈSE

présentée à la Faculté des sciences de l’Université de Genève pour obtenir le grade de Docteur ès sciences, mention biologie

par

Marta COTADO-SAMPAYO de

Ourense (Espagne)

Thèse n° 3976

Genève

Repromail, Université de Genève 2008

Les minutes passées à réfléchir au contenu d’une erreur marquent plus profondément les esprits que les heures passées à ingurgiter des théorèmes exacts. Car ces minutes sont accompagnées d’une émotion, d’une révolte intérieure……..

Albert Jacquard « L’Équation du nénuphar »

REMERCIEMENTS

Je désire d’abord remercier les membres du Jury: le Dr. Roland Beffa, le Professeur William Broughton, le Professeur Reto Strasser et le Dr. Francisco Barja qui ont accepté de lire et d’évaluer ce travail. Je tiens à exprimer ma gratitude au Dr. R. Beffa qui a soigneusement veillé à la bonne organisation et rédaction de ce manuscrit et au Professeur R. Strasser pour son soutien.

L’aboutissement de cette thèse a été rendu possible par le soutien porté par mon superviseur de thèse, Francisco Barja. Arrivée à Genève comme étudiante Erasmus, un peu désorientée et avec un projet incertain, Francisco m’a ouvert les portes de son laboratoire et m’a donné sa confiance pour entreprendre une thèse. J’apprécie que malgré toutes les difficultés rencontrées, il m’ait toujours soutenue dans tous mes efforts et ma motivation pour mener à bien à ce travail. Ce projet de recherche qui se termine maintenant est le résultat de tout le

«groupe de Francisco». Grâce à la bonne entente de l’équipe propice à un travail fructueux, il est encore possible de nos jours de travailler dans une ambiance de convivialité et de confiance.

C’est donc avec une particulière gratitude que je remercie Malou Chappuis et Ariane Fehr pour leur aide efficace. Merci à Cristina Andrés pour tous les bons moments partagés et pour son si précieux soutien dans les moments difficiles.

Le mérite de cette thèse est à partager avec le Dr. Ruben Ortega qui m’a aidée par ses conseils judicieux et ses critiques constructives tant dans la réalisation pratique que dans la discussion.

Et encore plus important il ne m’a pas laissé « achicopalarme ».

Je remercie aussi vivement le Dr. Mukti Ojha pour son aide savante et amicale, et qui m’a sans cesse encouragée en me guidant et me donnant des conseils avisés.

Un grand merci aussi aux membres du groupe de Bioimagerie, le Dr. Christophe Bauer et Jérôme Bosset pour sa disponibilité. À Mike Parkan pour m’avoir auxilié avec beaucoup de patience avec mes « lacunes » dans le domaine d’informatique et bien sur pour sa bonne humeur.

Je tiens aussi à remercier tous mes collègues du laboratoire de Bioénergétique (Abdallah Oukarroum, Georgina Ceppi, Madeleine Fontana, Marie-France Blanc, Gert Schansker et Dina Hanggraini) ainsi que ceux du département de Biologie végétale, spécialement Christophe Dunand et Sonia Guimil, pour tout le soutien qu’ils m’ont apporté.

Enfin, ma reconnaissance va aussi aux nombreux membres du laboratoire, aussi bien anciens qu’actuels, qui ont également participé plus au moins directement à ce travail. Ainsi, je remercie Arlette Cattanéo, Pilar Okenve, Edurne Martinez, Idoia Alonso, Javier Remiro, Greta Rubio, Enrique Raposo, Loreto Naya, Lucia Soliño, Marta Alonso, Sibylle Baruchel (Schindhelm), Aurélia Weber, Catherine Wilson, Marco Dias, Géraldine Martinelli et Fabrizio Molino pour leur aide scientifique mais aussi pour tous les bons moments passés ensembles.

Bien sûr une pensée pour mes amis, spécialement Maria del Mar, Ana “pajaros”, Txema, Cristina “bailarina”, Urko, Ana “Barja”, Gorge Faustino et Sébastien qui ont suivi de très près l’aventure de ma thèse.

Un grand merci à Manu pour tout l’amour qu’il m’a donné malgré les heures que mon travail lui a volé.

Enfin, merci à ma famille pour leur soutien et leurs encouragements.

Con gran alegría y satisfacción, dedico este trabajo a mis padres.

CONTENTS

Résumé 3

Summary 5

List of original publications 6

Abbreviations 7

1. GENERAL INTRODUCTION 9

1.1. CYTOSKELETON 9

1.2. ACTIN CYTOSKELETON 11

1.2.1. Actin 11

1.2.2. Actin binding proteins 13 1.2.3. Spectrin superfamily 13

1.2.4. α-Actinin 17

1.2.4.1. Functions 1.2.4.2. Isoforms

1.2.4.3. “Atypical” α-actinins

1.3. WHY STUDY FUNGI AS A MODEL OF TIP GROWTH? 20 1.3.1. Filamentous fungi 21 1.3.2. Cytoskeleton in fungi 22

1.3.3. Actin cytoskeleton in fungi 23 1.3.3.1. Actin

1.3.3.2. Actin binding proteins

2. BACKGROUND AND AIMS OF THE STUDY 25 3. MATERIALS AND METHODS 27

4. RESULTS 37

4.1. Identity of anti-αβ-spectrin immunoreacting peptides 37 in fungi and Oomycetes (Publications I and II)

4.2. α-Actinin orthologs in fungi (Publication III) 51 4.3. Characterization of α-actinin from Neurospora crassa 53 (Publication IV)

5. DISCUSSION, CONCLUSIONS AND PERSPECTIVES 67

6. REFERENCES 75

ANNEXE 95

Résumé

Caractérisation de l’α-actinine, une protéine membre de la superfamille des spectrines chez Neurospora crassa

La superfamille des spectrines est composée par des protéines qui lient l’actine. Elles participent à l’organisation du cytosquelette et interagissent avec d’autres protéines ou structures comme la membrane plasmique. Les protéines de ce groupe possèdent trois domaines: N-terminal « CH- domain » (Calponin Homology domain), C-terminal « EF-hand motifs » et un domaine central composé d’un nombre variable de « spectrin repeats ». Avec ces caractéristiques, la spectrine, l’α- actinine et la dystrophine/utrophine ont toujours été considérées comme des membres de la superfamille des spectrines. Ces protéines ont été décrites dans la plupart des cellules eucaryotes.

Néanmoins, chez les plantes et les champignons, la présence de spectrine a été démontrée sur la base des résultats obtenus par des techniques immunologiques, souvent en utilisant des anticorps commerciaux polyclonaux. Ainsi, la présence des protéines «spectrin-like » chez les champignons filamenteux, les levures et les Oomycetes est mise en doute car ces protéines ont seulement été identifiées à l’aide d’anticorps dont la spécificité n’était pas suffisamment établie. De plus, chez ces organismes, après l’analyse des génomes complètement séquencés, aucun gène codant pour la spectrine n’a été trouvé. Par analyse de spectrométrie de masse, nous avons identifié la protéine

« spectrin-like » comme étant le facteur d’élongation 2 (EF 2) chez Neurospora crassa. Par ailleurs, nous avons aussi identifié une protéine correspondant à la protéine de choc thermique (Hsp70) chez l’oomycete Phytophthora infestans. La caractérisation du peptide reconnu chez N. crassa par l’anticorps anti-αβ-spectrine ainsi que la réactivité croisée de cet anticorps ont été amplement traités dans la partie 4.1 de cette thèse, ainsi que dans les publications I et II.

De plus, en utilisant les outils de bioinformatique (BLAST) nous avons trouvé dans la base de données génomique de N. crassa (http://www.broad.mit.edu/annotation/fgi/) un gène (ncu06429.4) qui code pour une protéine similaire à l’α-actinine. Nous avons utilisé la séquence de cette protéine pour chercher leurs orthologues chez les champignons. L’α-actinine semble être le seul membre de la superfamille des spectrines. Néanmoins, cette protéine n’est pas présente dans tous les champignons ; chez certaines levures du groupe des Saccharomycotina, il n’y a pas d’évidence prouvant l’existence d’α-actinine. Dans ces champignons cette protéine semblerait avoir été perdue lors de l’évolution du fait que d’autres protéines liant l’actine pourraient complémenter sa fonction. Ce sujet a été traité en détail dans la partie 4.2 de cette thèse et dans la publication III.

Suite à notre étude bioinformatique nous avons pu identifier un gène codant pour l’α-actinine, membre le plus « primitif » de la superfamille des spectrines. Cette protéine a été identifiée, localisée in situ et in vivo et caractérisée biochimiquement chez N. crassa.

Bien que le rôle exact de l’α-actinine chez N. crassa n’aie pas pu être totalement élucidé, l’étude phénotypique du transformant (« knock-out » partiel de l’α-actinine) nous amène à penser que l’α- actinine participe avec l’actine à la coordination d’activités telles que la germination, la formation des septa et l’établissement des ramifications hyphales lors de la croissance. En outre, à la différence d’autres organismes, chez N. crassa l’α-actinine est une protéine essentielle, car son absence (« knock-out » complet de l’α-actinine) est létale pour le champignon.

4

Summary

Cell shape, cell division, polarization and tip growth are processes that have been widely studied using fungi as models. The organization of the cytoplasm and the coordination of cell activity during the life cycle of fungi are poorly understood but are believed to depend on a cytoskeletal system. The cytoskeleton in fungi is composed principally of microfilaments and microtubules. The dynamics and function of these structures are regulated by associated proteins. These proteins were first described using immunological techniques but their existence can now be verified by the availability of a number of complete fungal and plant genome sequences. The existence of an actin-binding “spectrin- like” protein in plants, fungi, Oomycetes and lower eukaryotic organisms has been reported. However, these reports were solely based on immunological studies and evidence for a gene coding for spectrin was not presented. In fungi and Oomycetes another member of the spectrin superfamily, α-actinin, was first proposed by us to be the immunoreactive peptide and we gave this the name “spectrin- related” protein. However, further studies demonstrated that α-actinin is not related to the protein that is recognized by anti-spectrin antibodies in fungi and Oomycetes. Instead, this immunoreactive peptide turned out to be a cross-reacting protein not related to the spectrin superfamily.

Assessing the characteristics of α-actinin in fungi may provide further insights into the biology of fungi and also help to establish new links between developmental complexity and genome evolution.

In the work described here we studied the features of the α-actinin orthologous group in fungi. Using Neurospora as a model for filamentous fungi we studied the ability of this protein to bind actin and calcium. Our results on the localization of α-actinin and the phenotype of the α-actinin knock-out strain suggest that α-actinin is essential for conidial germination and septum formation during hyphal growth. The fact that α-actinin is also localized along the peripheral region suggests that this protein may have additional functions.

List of original publications

The present thesis is based on the following original articles (see annexe), which are referred to in the text by their Roman numerals:

I. Cotado-Sampayo M., Ojha M., Ortega Perez R., Chappuis M-L., Barja F., 2006. Proteolytic cleavage of a spectrin-related protein by calcium-dependent protease in Neurospora crassa. Curr.

Microbiol. 53: 311-316.

II. Cotado-Sampayo M., Okenve Ramos P., Ortega Perez R., Ojha M., Barja F., 2008. Specificity of commercial anti-spectrin antibody in the study of fungi and oomycetes: cross-reaction with proteins other than spectrin. Fungal Genet. Biol. 45 (6): 1008-1015.

III. Cotado-Sampayo M., 2008. Features of α-actinin in fungi and Oomycetes. Under revision.

IV. Cotado-Sampayo M., Ortega Perez R., Seum C., Ojha M., Barja F., 2008. Characterization of Neurospora crassa α-actinin. Under revision.

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Abbreviations

aa Amino acid

ABD Actin-binding domain

ABP Actin-binding protein

Aip1 Actin-interacting protein 1

Amp Ampicillin

ARP Actin related protein

ATP Adenosine-5'-triphosphate

BLAST Basic local alignment search tool BCIP Bromo-chloro-indolyl phosphate

bp Base pairs

BSA Bovine serum albumin

°C Degree Celsius

CBD Calcium-binding domain

cDNA Complementary DNA

CDP Calcium-dependent protease

GCY Glucose Casein hydrolase Yeast extract

GFP Green fluorescent protein

GST Glutathione S-transferase

CH Calponin homology

CIP Calf intestinal phosphatase

cm Centimeter

2D Two dimensional

Da Dalton

DAB 3,3'-Diaminobenzidine tetrahydrochloride

dH₂O Distilled water

dist. Distilled

DMSO Dimethylsulfoxide

DNA Deoxyribonucleic acid

DNAse Deoxyribonuclease

dNTP Deoxyribonucleotide triphosphate

DTT Dithiothreitol

EDTA Ethylenediaminetetraacetic acid

EF Elongation factor

e.g. “exempli gratia” (for example)

EGTA Ethyleneglycol-bis(2-aminoethylether)-N,N'-tetraacetic acid

EST Expressed sequence tag

et al. “et alii” (and others)

FGSC Fungal genetics stock center

Fig. Figure

FITC Fluorescein-isothiocyanate

g Gram

GFP Green fluorescence protein

GST Glutathione S-transferase

GTP Guanosin-5'-triphosphate

h Hour

HEPES 4-(2-Hydroxyethyl)-1-piperazineethanesulfonic acid

His Histidine

HMW High molecular weight

Hsp Heat shock protein

i.e. “id est” (that is)

IPTG Isopropyl-β-D-thiogalactopyranoside

k Kilo

kDa Kilodalton

KIF Kinesin family protein

KO Knock-out

l Liter

LB Luria Bertani

LMW Low molecular weight

M Mol/l

MALDI Matrix-assisted laser desorption/ionization

min Minute

μF Microfaraday

µm Micrometer

MIPS Munich center for protein sequences MOPS Morpholinopropanesulfonic acid

Mr Relative molecular mass

MS Mass spectrometry

MYA Million years ago

nm Nanometer

NMDA N-methyl-D-aspartate

OD Optical density

PAGE Polyacrylamide gel electrophoresis

PBS Phosphate buffer saline

PCR Polymerase chain reaction

PDA Potato Dextrose Agar

pH Potential of hydrogen

pI Isoelectric point

PMSF Phenylmethylsulfonyl fluoride

pNA para-Nitroaniline

PVDF Polyvinylidene difluoride

PVP Polyvinyl polypyrrolidone

RNA Ribonucleic acid

RNAse Ribonuclease

rpm Revolutions per minute

SDS Sodium dodecyl sulfate

SEM Simple and efficient method for transformation

sec Second

SR Spectrin repeat

siRNA Small interfering RNA

TBS Tris buffer saline

TCA Trichloroacetic acid

TOF Time-of-Flight

Tris Tris-hydroxymethyl-aminomethane

TRITC Tetramethylrhodamine isothiocyanate Triton X-100 tert-Octylphenoxypolyethoxyethanol Tween 20 Polyoxyethylene(20) sorbitan monolaureate

U Units

V Volt

v/v Volume per volume

w/v Weight per volume

WT Wild-type

YpSs Yeast protein Soluble starch

8

1. GENERAL INTRODUCTION

1.1. CYTOSKELETON

The cytoskeleton is fundamental to the intracellular organization, plays an important role in cell division and also in the communication of the cell with its environment. A large number of studies have been performed in recent years in order to identify proteins involved in the cytoskeleton and to understand its role in the physiology of eukaryotic cells. Processes such as the establishment of cellular shape, cell locomotion, endo- and exocytosis, signaling, intracellular transport and cell division depend on this complex network of protein filaments that extends throughout the cytoplasm (Schliwa, 1986; Lloyd, 1991; Qualmann and Kessels, 2002; Smythe and Ayscough, 2006; Lanzetti, 2007). A substantial amount of molecular, biochemical and physiological information has been obtained on the cellular organization of the cytoskeleton. Knowledge of cytoskeletal and associated proteins is also important to gain a better understanding of numerous human diseases that depend on cytoskeleton dysfunction. Abnormal phosphorylation of a microtubule associated protein, Tau, is associated with Alzheimer neurofibrillary pathology (Goedert et al., 1996; Strong et al., 2006; von Bernhardi, 2007) and point mutations in cardiac actin have been detected in familial hypertrophic cardiomyopathy (Mogensen et al., 1999; Vang et al., 2005; Monserrat et al., 2007).

The cytoskeleton is composed of three major types of protein filaments: actin filaments (microfilaments), microtubules and intermediate filaments (Figure 1). Each type of filament is formed as a chain of protein monomers and can be built into a variety of structures depending on its associated proteins.

Intermediate filaments are polymers of elongated fibrous protein monomers, such as vimentin, keratin and lamin, which belong to a family of structurally and genetically related proteins.

Intermediate filaments are found in metazoans. Different families of intermediate filaments are expressed in different cell types. One essential role of these filaments is to distribute tensile forces across cells in a tissue.

A B

Figure 1. Cytoskeleton in eukaryotic cell. A. Organization of actin, microtubules, and intermediate filaments within a cell. B. Confocal image of an endothelial cell where actin filaments are shown in red, microtubules in green, and the nuclei in blue (web images in the public domain).

Microtubules are hollow tubes formed by the assembly of heterodimers of α- and β-tubulin. The heterodimers are arranged in longitudinal rows called protofilaments. Thirteen protofilaments are assembled in a parallel fashion around a hollow core that is approximately 25 nm in diameter. The microtubules are polar structures with two distinct ends: a fast-growing plus end and a slow-growing minus end. This polarity is an important consideration in determining the direction of movement along microtubules. The “minus” end in the cell starts in the MTOC (microtubule organizing center). Oakley and Oakley (1989) identified a third type of tubulin, named γ-tubulin. Microtubules participate in chromosome segregation during cell division, transport of vesicles and organelles, and cilia and flagella movements.

Microfilaments are 4 to 7 nm wide filaments formed by actin monomers (G-actin), a globular protein of 43 kDa. Microfilaments (F-actin) are the principal components of the actin cytoskeleton. The actin cytoskeleton shows an organizational flexibility that allows cells to assume many shapes. In motile cells like fibroblasts actin participates in locomotion. In mammals and many other organisms, actin is required for muscle contraction, cytokinesis, cell-substrate interactions, endocytosis and secretion.

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1.2. ACTIN CYTOSKELETON 1.2.1. Actin

Microfilaments are polymers of actin monomers that, together with a large number of actin-binding proteins, form the actin cytoskeleton (dos Remedios et al., 2003; Dominguez, 2004).

Actin monomer structure

Monomeric G-actin has dimensions of ~67 × 40 × 37 Å and a molecular mass of about 43 kDa. About 40% of the structure consists of α-helices (Otterbein et al., 2001; dos Remedios et al., 2003). The actin monomer contains four subdomains (Figure 2), a central cleft contains a high-affinity binding site for a nucleotide (ATP or ADP) and a cation (usually Ca²⁺ or Mg²⁺). Many of the known actin-binding proteins bind to the same loci, in the hydrophobic cleft between subdomains 1 and 3, and therefore can be expected to compete for this binding site (Dominguez, 2004). Subdomain 2 contains a DNAse-I- binding loop that participates in the intra-strand interactionsbetween F-actin subunits. The function of the DNAase I loop is unknown. However, DNAse I is a valuable tool to measure the G-actin content of actin solutions. When DNAase I is bound by actin it no longer has the capacity to cleave DNA (Lazarides and Lindberg, 1974; Hitchcock, 1980) and it can therefore be used to titrate actin.

Actin isoforms

The actins are classified into three groups according to their isoelectric point: α-, β- and γ-isoforms. In mammals, there are at least six different actin isoforms, each encoded by a separate gene, and these differ by <10% of the amino acid sequence (Vandekerckhove and Weber, 1978). In plants the number of genes is higher; e.g. Petunia contains >l00 actin sequences in its genome (Baird and Meagher, 1987) and other plant species, such as soybean, tobacco, potato, rice, and lodgepole pine, also appear to have dozens of actin genes (Meagher, 1991; Thangavelu et al., 1993; Meagher and Williamson, 1994).

The presence of an ancestral actin protein in bacteria was first suggested by Bork et al. (1992). This protein, called MreB, was proposed to be the prokaryotic origin of the actin cytoskeleton (van den Ent et al., 2001). Although the amino acid sequence homology of the MreB to eukaryotic actin is limited to 15%, their overall size and shape are markedly similar (van den Ent et al., 2001). Furthermore, MreB has similar actin-like cytoskeletal roles (Jones et al., 2001; Carballido-Lopez, 2006; Pradel et al., 2006; Graumann, 2007; Vats and Rothfield, 2007).

Figure 2. Ribbon representation of the structure of uncomplexed actin in the ADP state (Otterbein et al., 2001).

Actin polymerization

Actin polymers assemble spontaneously through non-covalent interactions between monomeric subunits and thus form highly dynamic structures with turnover at both ends. Actin monomers polymerize under physiological conditions, i.e. high ionic strength (KCl concentrations >50 mM), neutral or slightly acidic pH, high Mg²⁺ levels and elevated temperature (Asakura et al., 1960; Grazi and Trombetta, 1985). The assembly of actin monomers (G-actin) into filaments (F-actin) occurs in three steps: (1) a slow initial association into a dimer, (2) the formation of a more stable trimer that represents the nucleus of polymerization, and (3) the elongation phase. Each actin monomer can bind ATP. When an actin monomer is incorporated into the polymer the ATP is hydrolyzed to ADP (Carlier, 1990, 1992; Carlier and Pantaloni, 1997; Romero et al., 2004; Zheng et al., 2007). The elongation rate is directly proportional to the concentration of free actin and only ATP-actin monomers are likely to participate in polymerization (Pollard, 1986). At the steady-state concentration, the rate of actin assembly is the same as the rate of actin disassembly and the actin filaments thus have a constant length. This phenomenon is known as treadmilling and it acts like a motor for cell motility and pathogen locomotion (Disanza et al., 2005).

Several actin-binding proteins exist in vivo and these regulate different aspects of actin dynamics and will be discussed in the next section.

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1.2.2. Actin-binding proteins

Monomer availability for polymerization is regulated by actin monomer binding and actin filament capping proteins (Weber et al., 1999; Pollard and Borisy, 2003). Capping proteins are F-actin-binding proteins that interfere with the growth of an actin filament by blocking one of its ends. The activities of these proteins are often regulated by signaling molecules and ions such as Ca²⁺ (Lehrer, 2002;

Oertner and Matus, 2005; Lange and Gartzke, 2006).

Actin-binding proteins can be grouped into three groups according to their function: (1) those that participate in the formation of filaments from G-actin and the subsequent stability of these filaments, (2) motor proteins that use F-actin for traction and (3) those that connect actin filaments to the cell membrane or cross-link actin filaments to form different structures such as bundles, branching filaments and three-dimensional networks. Examples of the latter group are all the members of the spectrin superfamily.

1.2.3. Spectrin superfamily

The spectrin superfamily is composed of spectrin, α-actinin, dystrophin and utrophin. These proteins are involved in the organization of the actin cytoskeleton. Proteins belonging to the spectrin superfamily have three characteristic domains. They contain an N-terminal actin-binding domain (ABD) and a C-terminal calcium-binding domain (CBD) linked by a rod domain. In spectrin, where the basic unit is a heterodimer of α- and β-spectrin, the actin- and calcium-binding domains are both in the N-terminal region of the dimer (Figure 3).

The actin-binding domain contains two calponin homology domains (CH1 and CH2 domains), both with a low sequence similarity and functional diversity despite the predicted similarity in secondary structure. Actin-binding studies on the isolated binding domain of α-actinin have shown that the two domains have different roles in actin-binding (Way et al., 1992; McGough et al., 1994; Lorenzi and Gimona, 2008). The CH1 domain by itself has a reduced affinity for F-actin. The CH2 does not have any intrinsic actin-binding activity but contributes substantially to the interaction of the complete actin-binding domain, perhaps by acting as a locator of low affinity docking sites on the actin filament (Djinovic-Carugo et al., 1997; Bañuelos et al., 1998; Gimona et al., 2002). The fact that the single calponin domain lacks actin-bindingactivity suggests the possibility that CH domainshave additional functions.

The calcium-binding domain is composed of EF-hand motifs located at the C-terminus (except for spectrin, where is absent on the β-subunit). The EF-hand motif found in the spectrin superfamily

EF-hand CH-domain

Septrin repeat

Plekstrin Homology

ZZ domain Src3 Homology WW domain

Cys-rich domain

β-spectrin (220 kDa) α-spectrin (240 kDa)

Dystrophin (420 kDa)

EF CH

CH EF

Utrophin (395 kDa)

shares structural homology with calmodulin. For this reason, the calcium-binding domain has also been named the calmodulin-like domain.

EF-hand regions are usually paired helix-loop-helix structures involved in the coordination of up to two divalent cations, usually calcium but occasionally magnesium (Tufty and Kretsinger, 1975). The binding of calcium to the EF-hands induces a conformational change that is implicated in the regulation of the actin- binding activity of the protein (Lundberg et al., 1992; Trave et al., 1995).

However, divergent evolution has led in some members of the spectrin superfamily to a set of EF- hands that no longer chelate calcium (Nakayama and Kretsinger, 1994).

Figure 3. Schematic representation of the different members of the spectrin superfamily (adapted from Broderick and Winder, 2002).

The rod domain is composed of several repeats called spectrin repeats because they were initially described in spectrin (Speicher and Marchesi, 1984). The folding of the spectrin repeats consists of three α-helices in a coiled-coil assembly, where the three helices wrap around each other (Pascual et al., 1996; Djinovic-Carugo et al., 1999, 2002; Broderick and Winder, 2005). The number of spectrin repeats varies within the spectrin superfamily and ranges from 2–4 in α-actinin to 17–24 repeats in spectrin, utrophin and dystrophin (Broderick and Winder, 2005). The repeat length has also been

14

found to differ between α-actinin, spectrin, and dystrophin, with 114–125, 106, and 109 residues, respectively. This is mainly due to differences in the inter-helical loop length (Parry et al., 1992).

Spectrin superfamily evolution

Since gene codings for any member of the spectrin superfamily of proteins have not been found in the bacterial and plant genomes available to date, it has been suggested that this superfamily appeared in a primitive unicellular organism belonging to the animal kingdom (Virel and Backman, 2004). The amino acid sequences and the structure analysis of spectrins, α-actinin and utrophin/dystrophin proteins suggest that all three protein families arose from a single common ancestral protein that was α-actinin-like (Byers et al., 1989 and 1992; Dubreuil, 1991).

Phylogenetic analysis indicated that the α-actinin-1 from the protozoan Entamoeba histolytica is the earliest diverging α-actinin, followed by the α-actinin of Encephalitozoon cunculi (Virel and Backman, 2004). These two proteins have the shortest rod domain, with only one spectrin repeat. The presence of five repeats in the rod domain of Trichomonas vaginalis α-actinin has been suggested (Addis et al., 1998; Bricheux et al., 1998), but only the first of these repeats shows some similarity with those found in other α-actinins (Bricheux et al., 1998). An intragenic duplication gave rise to two spectrin repeats (SR1 and SR4). This group of α-actinins is present in fungi and E. histolytica (α- actinin-2). A subsequent, second, intragenic duplication added two more repeats (SR2 and SR3) (Virel and Backman, 2004, 2007).

The most likely scenario for the evolution of the spectrin superfamily of proteins is that it began with the introduction of seven repeats between repeat 2 and 3 of α-actinin, producing an elongated “α- actinin”. At this point the dystrophin/utrophin lineage presumably diverged from the αβ-spectrin lineage (Figure 4). The next step consisted of a duplication of the seven-repeat block in the elongated

“α-actinin” and the insertion of a repeat between the two seven-repeat blocks. The protein was split into two parts at this inserted repeat (Figure 4A). Modern α- and β-spectrin evolved out of these two different fragments. One α-helix (a0) of the cleaved repeat became the N-terminal end of what is now known as α-spectrin and the two last α-helices (b17) became the end of what is now known as β- spectrin (Figure 4A). A triple helix bundle, which is the characteristic structure of an entire repeat, is completed when b17 binds to a0 to form the spectrin heterotetramer (Figure 4B) (Pascual et al., 1997;

Thomas et al., 1997; Viel, 1999).

The evolution of the spectrin superfamily can be divided into two phases. A first active phase is characterized by intragenic duplication and concerted evolution. In this phase gene duplication produced the α-actinin, dystrophin/utrophin and spectrin lineages. Just before arthropod/vertebrate

divergence the evolution of the spectrin superfamily entered a new, more stable phase (~500 MYA) (Thomas et al., 1997).

A

B

Figure 4. A. Model for the evolution of the spectrin superfamily of proteins (adapted from Pascual et al., 1997;

Thomas et al., 1997; Virel and Backman, 2004). B. Spectrin heterotetramer (the functional unit of spectrin). All the SR dark and light blue are similar.

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The three lineages of the spectrin superfamily of proteins are present in all metazoans. In sequenced bacterial and plant genomes, there is no evidence of genes coding for any member of the spectrin superfamily. In protozoans, fungi and Oomycetes the superfamily is represented by the α-actinin protein (Virel and Backman, 2004, 2007; Publications I and II). However, in plants, fungi and Oomycetes the presence of 240–220 kDa spectrin-like proteins has been reported based on studies using anti-spectrin antibodies (Michaud et al., 1991; Reuzeau et al., 1997; Kaminskyj and Heath, 1995; Holzinger et al., 1999; Degousée et al., 2000; Heath et al., 2003; Slaninová et al., 2003). This discrepancy between the bioinformatics data and the immunological studies will be discussed in more detail in Publications I and II.

The following section is focused on the basic principles of α-actinin structure and function. More information about this protein is given in Publications III and IV, which represent the main part of the thesis.

1.2.4. α-Actinin

α-Actinin is the smallest member of the spectrin superfamily. The functional unit is a homodimer (Blanchard et al., 1989; Viel, 1999; Ylänne et al., 2001). There is antiparallel binding of two α-actinin monomers, with the actin-binding domain (ABD) of one monomer facing the calcium-binding domains (CBD) of the other (Figure 3). This organization gives α-actinin the ability to cross-link actin filaments in a calcium-dependent manner (Tang et al., 2001). Another group of regulators, phosphoinositides, are also able to regulate the interaction of α-actinin with actin filaments but these can also regulate the association/dissociation of α-actinin with integrins, another class of α-actinin- binding proteins (Fukami et al., 1992; 1996; Fraley et al., 2005). α-Actinin can also be phosphorylated by kinases such as focal adhesion kinases. Focal adhesion plaques are an elaborate network of interconnecting proteins linking actin stress fibers to the extracellular matrix. The phosphorylation of α-actinin reduces its affinity for actin and prevents its localization to focal adhesion plaques (Izaguirre et al., 2001; von Wichert et al., 2003). Moreover, an increase in the level of α-actinin phosphorylation on tyrosine 12 of the ABD weakens the linkages formed between integrins and the cytoskeleton and alters the focal adhesion dynamics (Rajfur et al., 2002; von Wichert et al., 2003). These observations support the possibility that the phosphorylation of α-actinin may serve to modulate the coupling/uncoupling of integrins to the cytoskeleton (Zhang and Gunst, 2006).

1.2.4.1. Functions

α-Actinin has many biological functions. In striated muscle it is the major thin filament cross-linking protein in the muscle Z-disc (Suzuki et al., 1976; Fay et al., 1983; Luther, 2000), connecting actin filaments of adjacent sarcomeres. In non-muscle cells, α-actinin is a major component of stress fibers, a contractile structure analogous to the more organized units found in striated muscle cells (Otey and Carpen, 2004).

α-Actinin is also found in adhesion sites close to the plasma membrane, where it cross-links cortical actin to different adhesion and trans-membrane proteins, such as catenin (Knudsen et al., 1995) and integrins (Otey and Carpen, 2004), and serves as a linker between trans-membrane receptors and the cytoskeleton. In synapsis α-actinin-2 may play a role in the localization of the neurotransmitter receptor NMDA and its modulation by Ca²⁺ (Wyszynski et al., 1997; Rycroft and Gibb, 2004; Franzot et al., 2005).

It has been proposed that α-actinin participates in cytokinesis in fission yeast (Wu et al., 2001).

In addition, a number of important human diseases are caused by α-actinin dysfunction. Mutations in α-actinin-4 cause a form of familial focal segmental glomerulosclerosis (FSGS) (Kaplan et al., 2000), which is a common nonspecific renal lesion characterized by regions of sclerosis in some renal glomeruli and often results in loss of kidney function. Different studies have found a relationship between α-actinin dysfunction and different types of cancer, such as colorectal cancer (Honda et al., 2005; Craig et al., 2007), lung cancer (Honda et al., 2004), breast cancer (Guvakova et al., 2002) and neuroblastoma (Nikolopoulos et al., 2000).

1.2.4.2. Isoforms

Four isoforms of α-actinin exist in almost all vertebrate organisms: α-actinins-1, -2, -3 and -4 (MacArthur and North, 2004) classified on the basis of their Ca²⁺ affinity. α-Actinins-1 and -4 are generally located in non-muscle cells and they have conserved EF-hand motifs that bind Ca²⁺. In contrast, the muscle-isoforms α-actinins-2 and -3 have lost their Ca²⁺-bindingability. Furthermore, alternative splicing generates additional isoforms in some vertebrates (Parr et al., 1992;

Kremerskothen et al., 2002; Honda et al., 2004). Only one gene coding for α-actinin has been found in the invertebrate organisms studied to date. Therefore, the three isoforms described for Drosophyla are probably due to the occurrence of alternative splicing (Roulier et al., 1992).

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1.2.4.3. “Atypical” α-actinins

The rod domain of “atypical” α-actinins reported for some protozoa (Trichomonas vaginales and Entamoeba histolytica), fungi and Oomycetes (Addis et al., 1998; Bricheux et al., 1998; Wu et al., 2001; Virel and Backman, 2004, 2006, 2007; Virel et al., 2007; Publication I) is composed of one or two spectrin repeats instead of the four observed in classical α-actinins (Virel and Backman, 2007) (Figure 5). In these organisms there is generally only one α-actinin gene; the presence of different isoforms due to alternative splicing has not been reported. However, in the protozoa Entamoeba histolytica two proteins coded by two different genes have been characterized (Virel and Backman, 2006, Virel et al., 2007). “Atypical” α-actinin seems to represent the only member of the spectrin superfamily in protozoa, fungi and Oomycetes (Virel and Backman, 2004; Publication I). It has been proposed that this is the ancestor from which dystrophin, utrophin and spectrin evolved in two phases (see section 1.2.3).

Figure 5. Diagrammatic representation of the “atypical” α-actinins. (CH: Calponin homology; SR: spectrin repeat; CC: coiled coil region; CBD: Calcium-binding domain). The CC region from Phytophthora infestans is shorter than that in T. vaginalis, corresponding to the accommodation of three and four putative SR, respectively.

The controversial reports of spectrin-like proteins in fungi and the relevance of the atypical fungal α- actinins for the understanding of the evolutionary history of this superfamily make these organisms an excellent model to obtain further insights into the evolution of this superfamily. The study of the function of α-actinin in tip growth could reveal new roles for this actin-binding protein.

1.3. WHY STUDY FUNGI AS A MODEL OF TIP GROWTH?

Fungi represent the second largest group of organisms after insects, with about 1.5 million species – most of which are filamentous fungi (Hawksworth, 2001). Fungi play an important role in the decay of organic material and nutrient recycling.

The relationship between plants and fungi is sometimes positive, leading to the formation of symbiotic structures inside plant roots. The mycorhizae form a group of organisms that are capable of such a positive interaction. In other cases, fungi can be pathogenic agents and cause significant damage to agricultural crops. Plant diseases caused by fungi include rusts, smuts, and leaf, root, and stem rots.

Fungi are also agents of animal diseases. From evolution point of view, fungi are more chemically and genetically similar to animals than other organisms, making fungal diseases difficult to treat.

Fungi are also important in the pharmaceutical and food industries. They are used to produce enzymes and secondary metabolites such as antibiotics and also participate in beer, champagne, cheese and bread production (Hesseltine, 1965; Hersbach et al., 1984; Mapari et al., 2005; Schuller and Casal, 2005; Menacho-Márquez and Murguía, 2007; Wang and Lin, 2007).

The experimental tractability of fungi makes these organisms among the most important models in fundamental research. Important knowledge in biochemistry, genetics and molecular biology has been acquired from studies on fungi. Fungal cellular physiology and genetics share key components with animal cells, including multicellularity, cytoskeletal structures, development and differentiation, sexual reproduction, cell cycle, intercellular signaling, circadian rhythms, DNA methylation, regulation of gene expression through modifications of the chromatin structure, and programmed cell death (Colot and Rossignol, 1999; Borkovich et al., 2004; Dunlap and Loros, 2004; Galagan et al., 2003; Galagan and Selker, 2004). The shared origins of the genes responsible for these fundamental biological functions in humans and fungi continue to make the study of the fungal genes of vital interest to human biology.

Hyphae of filamentous fungi belong to the most polarized cellular structures found in nature. The study of fungal tip growth has been widely used to increase our understanding of the physiology of other highly polarized cells, such as root hairs, pollen tubes and neurons. Common aspects of, as well as differences in, cytoskeleton organization and function between these types of cells have provided important insights into the relationship between the cytoskeleton and cell growth.

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1.3.1. Filamentous fungi

In filamentous fungi polar extension is needed for vegetative growth and the development of complex tissues. For several reasons, tip-growing cells represent an ideal system to study cell expansion. Fungi concentrate their growth machinery at one cell surface site and only at this site does robust growth occur. Localized growth implies that all material required for growth has to be present at, or delivered to, the surface area where cell expansion takes place. Furthermore, the growth rate of the expanding cell surface area in a tip growing cell is much higher than the growth rate of a cell that distributes the growth machinery more or less evenly over its surface. This allows changes in growth rate and direction to be observed more easily.

When the spores of filamentous fungi germinate, nuclear division is accompanied by a series of ordered morphological events, including the switch from isometric to polar growth. As growth continues, the hyphae become compartmentalized with the addition of more septa, and lateral branches emerge from basal compartments.

Most of the studied filamentous fungi are of industrial interest, e.g. Aspergillus nidulans and Ashbya gossypii (Steiner et al., 1995; Harris, 1997; Wendland et al., 1999; Momany and Taylor, 2000;

Kaminskyj, 2001; Goldman and Kafer, 2004; Guest et al., 2004; Oakley, 2004; Gattiker et al., 2007).

Other examples are plant pathogens such as Uromyces appendiculatus, Magnaporthe grisea, Botrytis cinerea and Ustilago maydis (Barja et al., 1998; Dijksterhuis, 2003; Hamer and Talbot, 1998; Banuett and Herskowitz, 2002; Silva et al., 2006; Ebbole, 2007; Klosterman et al., 2007). Examples of non- pathogenic fungi include the ascomycete N. crassa (Borkovich et al., 2004; Galagan and Selker, 2004;

Dunlap, 2006; Dunlap et al., 2007) and the chytridiomycete Allomyces arbuscula (Ojha, 1996; Ojha and Barja, 2003).

The ascomycete N. crassa is the main experimental model used in this work, but other members of this “phylum” have also been studied (M. grisea and B. cinerea). The complete sequence of these three fungal genomes has been annotated by the Broad Institute (http://www.broad.mit.edu/annotation/fgi/).

N. crassa is an ascomycete that grows on semisolid media by forming colonies that spread. This fungus was first described as an infectious agent in French bakeries and today is used in diverse research programs.

The ascomycete Magnaporthe grisea is a plant pathogenic fungus that is responsible for an important disease in rice, rice blast, but can also infect other agriculturally important cereals. This fungus has been used as a model to understand plant-pathogen interactions.

Botrytis cinerea is another plant pathogen that affects many plant species, although its most notable hosts are wine grapes. The fungus is usually referred to by the name of its asexual (anamorph) form

because the sexual phase is rarely observed. The sexual form (teleomorph) is called Botryotinia cinerea.

In the work described here we also studied the chytridiomycete Allomyces arbuscula, which is a water fungus that is distributed throughout the world and is particularly abundant in warm climates. The filamentous cells of Allomyces divide in a characteristic dichotomous pattern. The life cycle can alternate between two stages, gametophytic and sporophytic, reflecting the capacity for both sexual and asexual reproduction. The reproductive structures are located at the end of the hyphae and are separated by complete septa.

Phytophthora infestans, an oomycete in the kingdom of Stramenopila, has a filamentous phenotype but is no longer considered as a fungus. In contrast to the fungi, Oomycetes are more closely related to plants than to animals. Whereas fungal cell walls are made primarily of chitin, the cell walls of Oomycetes are built mainly of cellulose. Ploidy levels and biochemical pathways are different between these two kingdoms. P. infestans is an important potato pathogen that causes late blight and has considerable economic impact.

The mode of growth of filamentous fungi is supported by the extreme polarization of the cytoskeleton and endo-membrane network, which allows the long-range transport of vesicles containing precursors required for apical extension to the tip region (Bourett and Howard, 1991; Roberson and Vargas, 1994).

1.3.2. Cytoskeleton in fungi

The principal components of the fungal cytoskeleton are microtubules and microfilaments. The presence of intermediate filaments is more controversial and has only been reported in a few studies (May and Hyams, 1998; McConnel and Yaffe, 1993; Geitmann and Emons, 2000). These cytoskeletal elements are required for a variety of cellular processes including polarized growth, organelle movement and positioning, secretion, endocytosis, cell division and chromosome segregation. As in other eukaryotic cells, fungal microtubules are known to function in mitosis and chromosome segregation (Morris and Enos, 1992; Thaler and Haimo, 1996; Jung et al., 1998). The importance of microtubules for the establishment of polar growth and tip growth varies depending on the fungal species (Oakley and Morris, 1980; Caesar-Ton That et al., 1988; Barja et al., 1993; Sawin and Nurse, 1998; Heath et al., 2000; Horio, 2007). Microtubules cooperate with microfilaments to control cell shape, cell division and intracellular transport (Momany and Hamer., 1997; Schott et al., 2002;

Bretscher, 2005).

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1.3.3. Actin cytoskeleton in fungi 1.3.3.1. Actin

Fungi contain either one or only a few actin genes (Tarkka et al., 2000; Helgason et al., 2003). In N.

crassa the presence of three isoforms, α- β- and γ-actins, has been described by Barja et al. (1991).

Nevertheless, as in other filamentous fungi and yeast there is only one bona fide actin gene, indicating that the three isoforms may be products of post-transcriptional modifications. However, actin-related proteins such as ARP 1, ARP 3 and RO 4 have recently been identified in Neurospora. Some of these proteins share the same molecular weight and sequence similarity as actin (Robb et al., 1995; Tinsley et al., 1998; Lee et al., 2001). Therefore, the reported three isoforms found in Neurospora could be the result of a cross-reaction of anti-actin antibodies with these actin-related proteins.

F-Actin appears in fungi in two principal forms: patches and cables (Heath et al., 2000; Walker and Garrill, 2006). Patches are usually localized in growing regions of yeast or filamentous fungi. Cables are described in yeast in a polarized manner from the bud along the mother cell. The organization observed in yeast is less evident in filamentous fungi such as N. crassa or Aspergillus (Xiang and Plamann, 2003), where the actin cytoskeleton appears principally as patches associated with the hyphal tip and cell cortex (Barja et al., 1991, 1993; Heath et al., 2000). The absence of actin bundles in some filamentous fungi has been explained in terms of the difficulty of preserving these structures during the preparation of samples used for immunofluorescence microscopy (Heath, 1987; Harold and Harold, 1992; Kaminskyj and Heath, 1994). However, in Aspergillus short actin cables were described in the tip region of the hyphae (Pearson et al., 2004; Virag and Harris, 2006). In the basidiomycete Ustilago maydis and the Oomycetes both structures, cables and patches, form the actin cytoskeleton (Bachewich and Heath, 1998; Banuett and Herskowitz, 2002; Walker et al., 2006).

Actin is also localized in the cytoplasm and forms a diffuse network of microfilaments in the septa of filamentous fungi (Harris et al., 1994; Capelli et al., 1997; Rasmussen and Glass, 2005, 2007) and in the contractile actin rings of yeast (Pruyne and Bretscher, 2000; Schott et al., 2002; Kamasaki et al., 2007).

The actin-binding proteins (ABP) form part of the actin cytoskeleton. These regulate its dynamics and mediate the interaction with other proteins. The actin cytoskeleton also contains actin related proteins (ARPs).

1.3.3.2. Actin-binding proteins

ABPs have been described principally in yeast. There are, e.g., twinfilin and cofilin/ADF, which sever actin filaments (Moon et al., 1993; Okada et al., 2006; Moseley et al., 2006); profilin and formin, which regulate the assembly of G-actin into filaments (Haarer et al., 1990; Sagot et al., 2002;

Evangelista et al., 2003; Kovar et al., 2005; Takaine and Mabuchi, 2007). In addition, there are capping proteins, which regulate microfilament length (Amatruda and Cooper, 1992; Sizonenko et al., 1996; Nakano and Mabuchi, 2006), fimbrin and α-actinin, which promote the bundling of microfilaments (Adams et al., 1989; Wu et al., 2001; Goodman et al., 2003), and myosin, which is involved in organelle movement (Watts et al., 1987; Matsui, 2003). Some ABPs were first described in Saccharomyces as Aip1 (Amberg et al., 1995) and were shown to collaborate with cofilin, capping the barbed ends of filaments severed by cofilin (Okada et al., 2006; Okreglak and Dubrin, 2007).

As far as filamentous fungi are concerned, there are some reports on proteins involved in the assembly and stability of actin filaments such as formin. A. nidulans and N. crassa encode a single formin (Xiang and Plamann, 2003). The A. nidulans formin, SepA, localizes to both septation sites and hyphal tips, suggesting that filamentous fungi use site-specific regulatory mechanisms to control formin- mediated actin polymerization (Harris et al., 1997; Sharpless and Harris, 2002). Genes encoding for myosin proteins have been found in the genomes of N. crassa and A. nidulans (Xiang and Plamann, 2003; Steinberg, 2007) and has been characterized in these two organisms (van Tuinen et al., 1986;

McGoldrick et al., 1995; Osherov et al., 1998; Yamashita et al., 2000; Takeshita et al., 2002) and in other filamentous fungi (Woo et al., 2003; Schuchardt et al., 2005; Weber et al., 2006). In N. crassa, an actin-binding protein of 47 kDa with the same intracellular distribution as actin has been reported (Capelli et al., 1997). This protein, which is named p47 and identified as EF1α (elongation factor 1α) (Taillefert, 1988), also binds to calmodulin (CaM). The actin-p47-CaM complex may participate in the relationship between the actin cytoskeleton and protein synthesis machinery.

So far, only a relatively small number of ABPs has been identified in filamentous fungi. However, the completion of their genome sequence will allow the identification of a large number of the ABPs in these organisms. For example, most of the Saccharomyces genes coding for the actin cytoskeleton, including the actin-binding proteins, have orthologs in N. crassa (Borkovich et al., 2004).

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2. BACKGROUND AND AIMS OF THE STUDY

The cytoskeleton in fungi is not yet well understood. The functions of actin and tubulin have been established to some extent by observing the effects of anti-cytoskeleton drugs on cell phenotype or on the in situ localization of these proteins (Caesar-Ton That et al., 1988; Hoang-Van et al., 1989; Barja et al., 1993; Riquelme et al., 1998; Torralba et al., 1998; Czymmek et al., 2005). Tools, such as constructs of green fluorescence protein (GFP)-fusion proteins, improvements in mutant constructs and bioinformatics have revealed more details on the composition and dynamics of the cytoskeleton network. Nowadays, we try to understand the complexity of this structure through the interaction that occurs between cytoskeletal proteins and other cellular structures, a process that is mediated by a plethora of associated proteins.

We were interested in a group of actin-binding proteins, the spectrin superfamily, that participate in the organization of the actin cytoskeleton and its connection with the plasma membrane in metazoans.

In the case of neurons, several studies report the essential role of spectrin in tip growth (Morris, 2001;

Spira et al., 2003). Initial studies on “spectrin-like” proteins in other highly polarized cells like root hairs, pollen tubes and oomycetes and fungal hyphae assign “spectrin-like” roles in the process of apical growth (Michaud et al., 1991; Kaminskyj and Heath, 1995; Bisikirska and Sikorski, 1997;

Reuzeau et al., 1997; De Ruijter et al., 1998, 2000; Holzinger et al., 1999; Degousée et al., 2000;

Braun, 2001; Heath et al., 2003; Slaninová et al., 2003; Toquin et al., 2006). In this thesis I provide strong evidence for the absence of spectrin genes in any of the completed plant and fungal sequenced genomes. Furthermore, the antibody used to report the presence of spectrin in these organisms cross- reacts with proteins other than spectrin. α-Actinin is the only member of the spectrin superfamily in fungi and this represents an early step in the present model for the evolution of this superfamily.

The goal of this thesis was to characterize the α-actinin protein in Neurospora crassa and to establish the relationship between α-actinin and actin. This would allow a better understanding of the function of the actin cytoskeleton in processes such as spore germination or growth of the mycelium tip. In addition, elucidation of the structure and function of α-actinin in N. crassa will allow a comparison with other better characterized metazoan models and lead to the possible discovery of new functions for this protein in fungi. The first target of the study was to determine the veracity of the presence of a

“spectrin-like” protein in fungi and Oomycetes and to identify the peptide detected by the commercial anti-spectrin antibody (commonly used so far in the scientific comunity) in these organisms.

For this purpose we planned to:

• Look for the gene coding for a putative spectrin in the completed fungal genome databases.

• Perform mass spectrometric analysis on the peptide reacting with the anti-αβ-spectrin antibody in N. crassa and P. infestans.

• Use bioinformatics tools to study the features of the α-actinin protein in fungi and Oomycetes.

• Construct a GST Neurospora α-actinin fusion protein for biochemical characterization.

• Produce polyclonal anti-α-actinin using the GST-fusion protein as an immunogen for further biochemical characterization of the fungal α-actinin.

• Localize the protein in situ by immunodetection.

• Construct a GFP-α-actinin fusion protein for localization of the protein in vivo.

• Study of α-actinin knock-out Neurospora strain to provide an insight into protein function.

In this thesis I provide strong evidence for the absence of spectrin genes in any of the completed plant and fungal sequenced genomes. Furthermore, the antibody used to report the presence of spectrin in these organisms cross-reacts with proteins other than spectrin. α-Actinin is the only member of the spectrin superfamily in fungi and this represents an early step in the present model for the evolution of this superfamily. Studies on this fungal protein may help to provide more details of the evolutionary history of the spectrin superfamily as well as to give a greater insight into the function of the actin cytoskeleton in fungi.

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3. MATERIALS AND METHODS Materials

Unless otherwise indicated, chemicals were obtained from Sigma-Aldrich/Fluka, Merck, Bio-Rad, Amersham, and Roche Diagnostics (Mannheim). Restriction enzymes and buffers as well as other DNA-modifying enzymes were purchased from New England Biolabs (Frankfurt am Main), Promega and Roche. For the PCR-reactions the Primus 25 advanced PCRSystem (PeqLab) was used.

Antibodies were obtained from Sigma-Aldrich. Oligonucleotides were provided by Microsynth (http://www.microsynth.ch/).

Fungal strains and cell culture Neurospora crassa

Wild type N. crassa (FGSC 262, strain St. Lawrence STA 4) was obtained from the Fungal Genetics Stock Center, School of Biological Sciences, Kansas City, MO. In order to produce large quantities of macroconidia, the fungus was first grown on solid “Davis and De Serres” medium (Davis and De Serres, 1970), for 3 days at 33°C in the dark and then at 25°C in artificial light for 4 days. Conidia were harvested and inoculated at an inoculum density of 5x10⁶ conidia/ml of Vogel’s minimal medium (Vogel, 1956) enriched with 1.5% or 2% sucrose. The culture was incubated at 30°C for 0, 6, 12, 18 hours on a rotary shaker at 150 rpm.

“Davis and De Serres” medium with slight modification (1 liter)

5 g Na and K Tartrate, 3 g NaNO3, 0.5 g MgSO4.7H2O, 0.1 g CaCl2.2H2O, 3 g KH2PO4, 0.1 g NaCl, 10 g Sucrose, 10 ml Glycerol, 0.1 ml Oligoelements*¹, 0.1 ml FeCl₃ (sol. 0.5%), 0.05 ml Biotine*².

The pH value was adjusted to 5.6 with 1N KOH. Medium was dispensed in 150 ml Erlenmeyer (20 ml/Erlemeyer) and solidified with 2% agar.

Vogel’s miminal medium (1 liter)

2 g NH₄NO₃, 2.5 g Citrate de Na.2H₂O, 5 g KH₂PO₄, 0.2 g MgSO₄.7H₂O, 0.1 g CaCl₂.2H₂O, 0.1 ml Oligoelements*¹, 50 μl Biotin*² .

*¹Solution of oligoelements (100 ml)

5.0 g Citric Acid, 5.0 g ZnSO₄, 1,0 g Fe(NH₄) ₂ (SO₄) ₂.6 H₂O, 5 mg H₃BO₃, 0.25 g CuSO₄. H₂O, 0.05 g MnSO₄.2H₂O, 0,05 g Na₂MoO₄.H₂O.

*²Solution of Biotin

Biotin 1 mg, in 10 ml 80% Ethanol (stock 4°C, during 6 months)

Magnaporthe grisea

M. grisea (wild type P1.2 strain) was kindly donated by Dr. M-H. Lebrun (Unité Mixte de Recherche, Centre National de la Recherche Scientifique/ BayerCropScience, Lyon). M. grisea was grown on solid rice medium with small pieces of filter for a few days until the filters were colonized and white mycelium appeared.

For storage the colonized filters in solid rice medium containing M. grisea were put into a sterile container and placed for at least 3 days in an oven at 37°C until they were dried, then stored at –20°C.

Mycelia for protein extraction were obtained from liquid culture in TNK medium containing 0.2%

Yeast extract and 1% Glucose (Ou, 1985). The cultures were allowed to grow for 48 hours in the dark in a shaker (150 rpm). The average weight of mycelium collected per liter of liquid medium was 2.5 g.

Solid Rice Medium (200 ml)

4 g Rice powder, 0.4 g Yeast extract, 4 g Agar.

TNK Medium (1 liter)

2.0 g NaNO3, 2.0 g KH2PO4, 0.5 g MgSO4.7H2O, 0.1 g CaCl2.2H2O, 4 mg FeSO4.7H2O, 1 ml Oligoelements*.

*Solution of Oligoelements (100 ml)

0.79 g ZnSO₄.7H₂O, 60 mg CuSO₄.5H₂O, 10 mg H₃BO₃, 20 mg MnSO₄.2H₂O, 14 mg NaMoO4.2H2O.

Botrytis cinerea

B. cinerea (BO47) was kindly donated by Dr. R. Beffa (BayerCropScience, Lyon). Mycelia were grown in solid potato dextrose agar medium (Difco) for one week on the bench with day periods of light. Spores were then harvested by flooding the culture with water and separated from the mycelial fragments by filtration. Spores obtained from a Petri dish (9 cm diameter) culture were used to inoculate 200 ml of liquid potato dextrose broth medium (24 g/l, Difco). Mycelia were harvested after 24 hours of growth with agitation (150 rpm) at 25°C.

Allomyces arbuscula

A. arbuscula strain Burma LD was grown on filter pads placed on solid YpSs medium (Emerson, 1941). Zoospores were induced by shifting the filters in sterile Petri-plates containing 30-40 ml sterile

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distilled water (water active liberation of zoospores) according to a procedure described by Ojha and Turian (1981). Zoospores were inoculated in CGY medium (Turian, 1963) and grown for periods of 6, 12 and 18 hours with forced aeration.

Solid YpSs (1 liter) (Emerson, 1941)

7.5 g Soluble starch, 2 g Yeast extract, 0.5 g K₂HPO₄, 0.25 g MgSO₄.7H₂O, 15 g Agar.

GCY Medium (1 liter)

1.0 g K₂HPO₄ *, 0.2 g MgSO₄.7H₂O, 0.1 g NaCl, 0.1 g CaCl₂.2H₂O, 0.02 g FeCl₃.6H₂O, 3.0 g Casein Hydrolysate, 5.0 g Sucrose, 0.1 g Yeast extract.

The pH was adjusted to 6.8 with 125 μl/l of 12 N HCl. *K₂HPO₄ was dissolved in water, sterilised separately and added to medium before use.

Phytophthora infestans

P. infestans (PT78) isolate was kindly donated by Dr. R. Beffa (BayerCropScience, Lyon). Mycelium was first grown on pea-agar (125 g of pea cooked and grinded in 1 liter of H2O, 2% agar). Sporangial inoculum was prepared from a 8 to 12 days culture. The sporangia were detached from the mycelia by flooding the culture with water and separated from the mycelial fragments by filtration. Sporangia were inoculated in V8 liquid medium (50 ml tomato juice/liter of distilled water, the pH was adjusted to 5 if necessary) at a final concentration of 10⁵ sporangia/ml. Mycelia were harvested after 72 h of growth in the dark at 20°C without agitation.

Protein extraction

To optimise the conditions for spectrin extraction, we tested different experimental conditions and protocols for cell disruption including:

a) several detergents: Triton-X-100, Triton-X-114, Empigen 1%, Octyl β-D glucopyranoside,

b) Sodium Dodecyl Sulfate (SDS) extraction. Dry fungal powder was resuspended in buffer containing 4% SDS, 5% 2-β−mercaptoethanol, 5% sucrose, 10 mg of insoluble polyvinil polypirrolidone (PVP) and boiled for 3 min before centrifugation (13000 x g;

20 min),

c) Trichloroacetic Acid (TCA)-Acetone extraction (Granier, 1988). The dry powder was resuspended in buffer containing 10% TCA, 0.07% 2-β−mercaptoethanol in cold acetone and kept at -18°C for 1h. After 15 min centrifugation at 20000 x g the supernatant was removed and the pellet was rinsed for 1 h at -18°C with cold acetone

containing 0.07% 2-β−mercaptoethanol. The rinsing solution was removed with caution and the pellet vacuum-dried for 1 h. The pellet was resuspended in extraction buffer [5 mM Hepes-KOH pH 7.5; 2 mM EGTA, 2 mM dithiothereitol (DTT)],

d) different concentrations of NaCl (0 to 100 mM) in 20 mM Tris-HCl pH 7.5, e) cells fixed in 3% paraformaldehyde before extraction,

f) cell wall digestion with Lysing enzyme (Sigma 5 mg/ml) for 20 min before extraction, g) buffer with Vanadate, a phosphatase inhibitor, in order to reduce the susceptibility of

spectrins to cleavage by μ-calpain (Nedrelow et al., 2003),

h) protein extraction from the cell wall-less mutant Neurospora crassa (FGSC 1118 fz;sg;os-1).

The standard procedure adopted to study the evolution of spectrin in growing mycelia was the following: cells from different stages of development were harvested by filtration through Millipore filters (pore size 0.5-1 μM; Millipore Corporation, Bedford, MA /USA), washed twice with distilled water, frozen in liquid nitrogen and ground in a mortar kept at a low temperature using liquid nitrogen.

The frozen powder was suspended in cytoskeleton stabilizing buffer (2-4 ml per mg powder) of Abe and Davies (1995) with slight modifications, containing 5 mM Hepes-KOH, pH 7.5, 250 mM sucrose, 15 mM Mg(OAc)₂, 2 mM EGTA, 2 mM dithiothreitol (DTT), 25 mM K₂O₅S₂, 10% glycerol and 0.5%

polyvinylpyrrolidone and the following protease inhibitors: 1 mM benzamidine, 2 mM phenylmethylsulfonyl fluoride (PMSF), 2 μg/ml each of leupeptin, chymostatin, trasylol; or Complete EDTA-free Protease Inhibitor Cocktail Tablets (Roche Applied Science). The homogenates were incubated at 4°C with gentle agitation for 20 min and then centrifuged at 6000xg for 15 min at 4°C.

The supernatant was recovered for in vitro proteolysis, analysis by SDS-PAGE and immunoblotting.

The protein concentration in supernatants was measured according to Bradford (1976) with Bovine Serum Albumine (BSA) as a standard.

One-dimensional gel electrophoresis

The proteins in the crude extract were separated on SDS-PAGE and native gels, according to Chrambach and Rodbard (1971) and Laemmli (1970), respectively. The gels were stained with Coomassie brilliant blue R-250. Samples of 50 μg or 10 μg of protein were loaded on each lane of a 15x10 cm gel or mini-gels respectively. Various molecular weight markers were utilized (for SDS- PAGE gels: Precision Plus Protein Standards from BioRad, LMW-SDS Marker Kit from Amersham Biosciences and Prestained Protein Marker IV from PeqLab. For non-denaturing gels: HMW Native Marker Kit from Amersham Biosciences).

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