STRUCTURAL AND FUNCTIONAL CHARACTERIZATION OF THE FOCAL ADHESION PROTEIN FAP52

MARKONIKKI

Faculty of Medicine,Department of Pathology,

University of Oulu;Department of Pathology,

University of Helsinki

OULU 2004

MARKO NIKKI

STRUCTURAL AND FUNCTIONAL CHARACTERIZATION OF THE FOCAL ADHESION PROTEIN FAP52

Academic Dissertation to be presented with the assent ofthe Faculty of Medicine, University of Oulu, for publicdiscussion in the Auditorium 101 A of the Faculty ofMedicine (Aapistie 5 A), on December 11th, 2004,at 12 noon.

OULUN YLIOPISTO, OULU 2004

Copyright © 2004University of Oulu, 2004

Supervised byProfessor Veli-Pekka Lehto

Reviewed byDocent Pekka LappalainenDocent Jari Ylänne

ISBN 951-42-7454-7 (nid.)ISBN 951-42-7455-5 (PDF) http://herkules.oulu.fi/isbn9514274555/

ISSN 0355-3221 http://herkules.oulu.fi/issn03553221/

OULU UNIVERSITY PRESSOULU 2004

Nikki, Marko, Structural and functional characterization of the focal adhesionprotein FAP52 Faculty of Medicine, Department of Pathology, University of Oulu, P.O.Box 5000, FIN-90014University of Oulu, Finland, Department of Pathology, University of Helsinki, P.O.Box 21, FIN-00014 University of Helsinki, Finland 2004Oulu, Finland

AbstractFAP52 (focal adhesion protein, 52 kDa) is a focal adhesion-associated protein composed of a highlyα-helical NH2-terminus containing a poorly characterized FCH (Fes/CIP4 homology) domain,unstructured linker region and the COOH-terminal SH3 domain. FAP52 is also known as PACSIN 2or syndapin II. Together with other PACSINs and syndapins FAP52 shares a common domainarchitecture.

The aim of this study was to characterize FAP52 in structural and functional terms. The functionwas pursued by identifying binding partners for FAP52, and by overexpressing the recombinantFAP52 in cultured cells. For the structural studies, various physico-chemical methods, such aschemical cross-linking, gel filtration chromatography, circular dichroism and X-ray crystallographywere applied. In addition, the histological distribution of FAP52 in chicken tissues was explored.

FAP52 binds filamin, a protein that regulates the dynamics of the cytoskeleton by crosslinkingactin filaments. The binding site in FAP52 was mapped to the NH2-terminal 184 amino acids, ofwhich the residues 146–184 form the core of the binding. In filamin, the binding site resides in therepeats 15–16 in the rod-like molecule encompassing 24 such repetitive domains. Overexpression ofFAP52 or its filamin-binding domain in chicken embryo heart fibroblasts induced the formation offilopodial extensions on the cell surface and reduced the number of focal adhesions, suggesting a rolein the organization of the cellular cytoskeleton and in cell adhesion machinery.

Experiments utilizing surface plasmon resonance analysis, size exclusion chromatography andchemical cross-linking showed that FAP52 self-associates in vitro and in vivo. The region responsiblefor the self-association was mapped to the amino acids 146–280, which is predicted to fold into acoiled-coil arrangement.

FAP52 was crystallized by using the hanging-drop vapor-diffusion method and ammoniumsulfate grid screen. Native dataset was collected from two crystals, which diffracted to 2.8 Å and 2.1Å resolution. For one form of crystals, phasing was performed using the native dataset and thedatasets from two xenon-derivatized crystals. X-ray crystallography studies revealed a dimer inasymmetric unit.

Histological and in vitro studies showed that, in liver, FAP52 is preferentially expressed in bilecanaliculi. In other tissues, FAP52 showed a specific staining pattern in gut, kidney, brain andgizzard.

Together, these data show that FAP52 self-associates in vivo and, probably via its interaction withits binding partner filamin, participates in the organization of the cytoskeletal architecture, especiallyof the cell surface protrusions, such as filopodia and microvilli of bile canaliculi.

Keywords: bile canaliculi, cytoskeleton, dimerization, filamin, focal adhesions

To Marja, Topias and Eelis

Acknowledgements

My thesis was carried out at the Department of Pathology, Faculty of Medicine, Universityof Oulu, during the years 1997–2004. It is my pleasure to thank the following persons whohave supported me in my work:

My great supervisor, professor Veli-Pekka Lehto, M.D., Ph.D., for guidance of my work.Vepeli has some features that make him an excellent scientist and an appreciated group lea-der. His vast knowledge of the literature and unbelievable ingenuity provide an excellentfacilities for his group to make high-quality science on “hot” research areas. His positiveand optimistic attitude of life, and always patient and emphatic stance towards students hasresulted in a joyful research atmosphere.

Professor Timo Paavonen, M.D., Ph.D., for his mental contribution to my work and forhis invaluable advice on the way to the dissertation.

Professor Frej Stenbäck, M.D., Ph.D., for both mental and financial contribution to mywork, for practical guidance in various matters, and for providing me with good researchfacilities.

Jari Ylänne, Ph.D., and Pekka Lappalainen, Ph.D., for their careful review of themanuscript for my thesis, and for their critical and professional comments.

Docents Helena Autio-Harmainen, M.D., Ph.D., Tuomo Karttunen, M.D., Ph.D., Mar-kus Mäkinen, M.D., Ph.D., Paavo Pääkkö, M.D., Ph.D., and Ylermi Soini, M.D., Ph.D., fortheir valuable support to my medical studies and research. In addition, Helena is, as anexpert pathologist, an important co-author in one of my works.

My colleagues Jari Meriläinen, Ph.D., Olli Lohi, M.D., Ph.D., Anssi Poussu, M.D.,Ph.D., Karoliina Tuppurainen, M.Sc., and Sari Vaittinen, M.Sc., who gave valuable helpand advice concerning the practical problems in the beginning of my research, and, who,by their own examples, have encouraged me to work towards the thesis. By cloning andpreliminary characterization of FAP52, Jari gave me fruitful facilities to start my work.

Kristina Djinovic Carugo, Ph.D., Kalervo Hiltunen, M.D., Ph.D., Tuomo Glumoff,Ph.D., Mikko Järvinen, Ph.D., Sinikka Eskelinen, M.D., Ph.D., Riitta Palovuori, M.Sc.,Raija Sormunen, Ph.D., Anthony Heape, Ph.D., Petri Kursula, Ph.D., Vimal Parkash,M.Sc., Annikki Huhtela, Marja-Liisa Martti, Anna-Liisa Oikarainen, Tarja Piispanen,Marja Tolppanen, Taina Uusitalo, Mirja Vahera, Riitta Vuento, and Marjaana Vuoristo, forinvaluable aid by their special skills, and for creating pleasant working conditions.

Hannu Wäänänen and Tapio Leinonen, technical specialists of our department, for assis-tance in various matters.

Hilkka Penttinen, Kati Hietala, Arja Jumpponen and Marja Korkatti, for helping inmany practical matters.

My parents Hilkka and Unto, for their encouraging attitude towards my studies and mylife in general. My father has, by his example as an entrepreneur, taught me to work hardto reach the targets of my life. My whole family for a happy and colorful childhood in Toho-lampi. My parents-in-law, Anneli and Pertti, for their attitude which is always so warm andfriendly, and especially for the great encouragement they have given me in my efforttowards the thesis. I have had numerous fruitful scientific discussions with Pertti, who ishimself a Doctor of Philosophy.

Marja, my dear wife, for her love and faithful encouragement during these years. Herlovely smile and lively eyes have welcomed me home after numerous lonely, dark eveningswhich I have spent with my work or hobbies. Topias and Eelis, our dear sons, for numerousdelightful moments with their play and everyday life.

The following foundations for the financial contribution of my studied during the years:the Alfred Kordelin, Emil Aaltonen, Farmos, Ida Montin, and Oskar Öflund foundations,Cancer Organisations, Finnish Medical foundation, Finnish Cultural foundation, Instru-mentarium Scientific foundation, Oulu University supporting foundation, Pohjois-SuomenSyöpäyhdistys r.y.

Oulu, September 2004 Marko Nikki

Abbreviations

aa amino acid(s)ABD actin-binding domainABP-280 actin-binding protein, 280 kDaBSA bovine serum albuminCEHF chicken embryo heart fibroblastECM extracellular matrixECL enhanced chemiluminescenceEGF epidermal growth factorEMBL European Molecular Biology LaboratoryFA(s) focal adhesion(s)F-actin fibrous actinFAP52 focal adhesion protein, 52 kDaFCH Fes-CIP4 homologyFPLC fast protein liquid chromatographyFPS FAP52-PACSIN-syndapinG-actin globular actinGP-Ibβ glycoprotein- IbβGST glutathione S-transferaseHA hemagglutininHIP1R huntingtin-interacting protein 1-related proteinHRP horseradish peroxidaseIPTG isopropyl-β-D-thiogalactopyranosideLPA lysophosphatidic acidMAPK mitogen-activated protein kinaseMAYP macrophage-associated tyrosine-phosphorylated proteinmSos mammalian son-of-sevenlessNCBI National Center of Biotechnology InformationN-WASP neural Wiskott Aldrich syndrome proteinPBS phosphate-buffered salinePCH pombe Cdc15 homologyPCR polymerase chain reaction

PEST proline, glutamine/aspartic acid, serine, threoninePIP2 phosphatidylinositol-4,5-bisphosphateSAPKs stress-activated protein kinasesSDS-PAGE SDS-polyacryl amide gel electrophoresisSH2 Src-homology 2SH3 Src-homology 3SPR surface plasmon resonanceVASP vasodilator-stimulated phosphoprotein

List of tables

Table 1 The currently identified FPS proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23Table 2 Currently characterized binding partners of FPS proteins. . . . . . . . . . . . . 25Table 3 Consensus sequences flanking the NPF motifs. . . . . . . . . . . . . . . . . . . . . . 27Table 4 List of known focal adhesion proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . 36

List of figures

Figure 1. Domain structure of FPS family proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24Figure 2. The phylogenetic tree of the FPS proteins of man, mouse, rat and chicken. . . 26Figure 3. Presence of the alternative splicing sites in the PACSIN 2 isoforms in

humans, mouse, rat and chicken. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27Figure 4. Suggestions for the regulation of the function of filamin. . . . . . . . . . . . . . . . . 43Figure 5. FPS proteins and endocytosis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 46Figure 6. Actin microfilament architecture in pericanalicular web. . . . . . . . . . . . . . . . . 68

List of original articles

The present thesis is based on the following separate articles, which are referred to in thetext by their Roman numerals:

I Nikki M, Meriläinen J & Lehto V-P (2002) FAP52 regulates actin organization viabinding to filamin. J Biol Chem 277: 11432–11440.

II Nikki M, Meriläinen J & Lehto V-P (2002) Focal adhesion protein FAP52 self-asso-ciates through a sequence conserved among the members of the PCH family pro-teins. Biochemistry 41: 6320–6329.

III Törö I, Nikki M, Glumoff T, Lehto V-P & Djinovic K (2004) Crystallization andphasing of Focal Adhesion Protein 52 from Gallus gallus. Acta Crystallogr D BiolCrystallogr 60: 539–541.

IV Nikki M, Meriläinen J, Autio-Harmainen H, Virtanen I, & LehtoV-P. PreferentialExpression of the Focal Adhesion Protein FAP52 in Bile Canaliculi. Manuscript.

Contents

AbstractAcknowledgementsAbbreviationsList of tablesList of figuresList of original articles1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 212 Review of the literature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22

2.1 The FAP52/PACSIN/syndapin family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 222.1.1 The PCH family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 232.1.2 Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24

2.1.2.1 Domains and the sequence analysis of FPS family proteins . . . 242.1.2.2 Sequence comparison . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25

2.1.3 Tissue expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 282.1.4 Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28

2.1.4.1 FPS family proteins in neurons . . . . . . . . . . . . . . . . . . . . . . . . . 282.1.4.2 Function in endocytosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 292.1.4.3 Actin reorganization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30

2.2 Actin microfilaments and focal adhesions . . . . . . . . . . . . . . . . . . . . . . . . . . . . 312.2.1 Features of actin microfilaments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 312.2.2 Proteins regulating actin polymerization . . . . . . . . . . . . . . . . . . . . . . . . . 322.2.3 Focal adhesions – structure and dynamics . . . . . . . . . . . . . . . . . . . . . . . 34

2.2.3.1 Structure of focal adhesions . . . . . . . . . . . . . . . . . . . . . . . . . . . . 342.2.3.2 Dynamics of focal adhesions . . . . . . . . . . . . . . . . . . . . . . . . . . . 38

2.3 Filamin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 392.3.1 Splice variants, domain structure and tissue expression . . . . . . . . . . . . . 392.3.2 Functions of filamin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 40

2.3.2.1 Actin cross-linking and organization . . . . . . . . . . . . . . . . . . . . . 402.3.2.2 Filamin as an anchoring protein for transmembrane receptors . 402.3.2.3 Filamin as a scaffolding protein for signaling molecules . . . . . 412.3.2.4 Regulational aspects of filamin function . . . . . . . . . . . . . . . . . . 42

2.4 Endocytosis and the cytoskeleton . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 432.4.1 Actin and endocytosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 442.4.2 Proteins acting at the interface between endocytosis and the actin

cytoskeleton . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 443 The aims of the present study . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 484 Materials and methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49

4.1 General procedures and computer programs (I–IV) . . . . . . . . . . . . . . . . . . . . . 494.2 Materials (I–IV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 494.3 Antibodies (I, II, IV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 504.4 cDNA constructs (I–III) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 504.5 Expression and purification of the GST fusion proteins (I–III) . . . . . . . . . . . . 514.6 Cell cultures, transfections and immunofluorescence

microscopy (I, II, IV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 514.7 Immunoelectron microscopy (I, IV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 524.8 Cell lysates and immunoprecipitation (I, IV) . . . . . . . . . . . . . . . . . . . . . . . . . . 524.9 Affinity chromatography and pull-down experiments (I) . . . . . . . . . . . . . . . . 524.10 Immunoblotting and blot overlay assay (I, II, IV) . . . . . . . . . . . . . . . . . . . . . . 534.11 Electrospray ionization mass spectrometry (I) . . . . . . . . . . . . . . . . . . . . . . . . . 534.12 Surface plasmon resonance analysis (I, II) . . . . . . . . . . . . . . . . . . . . . . . . . . . . 534.13 Chemical cross-linking (II) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 544.14 Gel filtration chromatography (I, II) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 544.15 Far UV circular dichroism (II) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 544.16 Crystallization of FAP52 (III) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55

5 Results . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 565.1 Interaction of FAP52 with filamin (I) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 56

5.1.1 Association between FAP52 and filamin in cells in vivo . . . . . . . . . . . . 565.1.2 Direct binding of FAP52 to filamin . . . . . . . . . . . . . . . . . . . . . . . . . . . . 575.1.3 Filamin-binding site in FAP52 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 575.1.4 FAP52-binding site in filamin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 585.1.5 Colocalization of FAP52 with filamin in cultured cells . . . . . . . . . . . . . 58

5.2 Overexpression of FAP52 in cultured cells (I, II) . . . . . . . . . . . . . . . . . . . . . . 595.3 Self-association of FAP52 (II) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60

5.3.1 Computational analysis and circular dichroism . . . . . . . . . . . . . . . . . . . 605.3.2 Self-association in vitro and in vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . 605.3.3 Kinetics of self-association . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 615.3.4 Mapping of the self-association site . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61

5.4 Crystallization and phasing of FAP52 (III) . . . . . . . . . . . . . . . . . . . . . . . . . . . 615.5 FAP52 in bile canaliculi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62

6 Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 636.1 FAP52 as a multidomain protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 636.2 FAP52 self-associates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 646.3 FAP52 as a filamin-binding protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 656.4 FAP52 as a regulator of the cytoskeleton . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 666.5 FAP52 as a component of the pericanalicular web of the bile canaliculi . . . . 67

7 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69References

1 Introduction

Details of the regulation of many central cellular processes, such as cell division, migra-tion and differentiation, are still only partially known. In order to clarify the molecularmechanisms of the processes, various schematic models are built with known proteinsand arrows marking the interactions between them. However, the models still have gaps:proteins are missing and many interactions and functions unrevealed. The aim of basicresearch is to fill those gaps by completing the catalog of all the proteins and theirdomains, and by identifying their interactions and functions. Only detailed knowledge ofthe mechanisms of different cellular events enables us to understand the workings of thehuman body and, in the future, to develop therapeutic strategies and prophylactic mea-sures against diseases.

Signaling proteins are typically constructed of distinct modules, domains, which occurin different combinations in different proteins. A domain can either have enzymatic activ-ity, or it may be nonenzymatic and mediate interactions with specific sequences in spe-cific proteins or other macromolecules. Currently, roughly 150 signaling domains havebeen identified (domain search by SMART; EMBL computational service at the networkserver http://smart.embl-heidelberg.de). A domain may be highly conserved among vari-ous proteins and between different species, such as SH3, which reveals a high degree ofsequence similarity throughout the domain, or it may contain only a few conserved coreresidues, such as the WW domain, a 33-amino acid (aa)-long domain, which only has twoconserved tryptophans. Examples of typical domains in signaling proteins are SH2- andSH3-domains. The SH2 domain binds to phosphotyrosine residues, and SH3 to specificproline-rich sequences (Cohen et al. 1995).

Focal adhesions are specialized structural entities that mediate adhesion of a cell to itssubstratum. Molecularly, they are composed of a large number of both structural and sig-naling proteins, which are linked together and coordinate their functions via specificdomain-domain interactions.

This study focuses on the structure and function of the focal adhesion protein FAP52.FAP52 is an SH3 domain-containing, serine-phosphorylated protein, which in culturedfibroblasts localizes at focal adhesions (Meriläinen et al. 1997). It is expressed in mosttissues.

2 Review of the literature

2.1 The FAP52/PACSIN/syndapin family

FAP52 (Meriläinen et al. 1997), PACSINs (Plomann et al. 1998) and syndapins (Qual-mann et al. 1999) are closely related proteins that share a common domain architectureand are involved in actin organization and vesicular trafficking (Meriläinen et al. 1997,Ritter et al. 1999, Qualmann & Kelly 2000). FAP52 has been characterized from chicken(Gallus gallus). Its orthologs in other species have been named PACSIN 2 or syndapin II.In this thesis, the FAP52/PACSIN/syndapin proteins except for chicken FAP52 are calledPACSINs. Three PACSIN genes have been identified. They encode the proteins calledPACSIN 1, PACSIN 2 and PACSIN 3 (Ritter et al. 1999, Modregger et al. 2000). Basedon the genebank search (program BLAST, NCBI network server at http://www.ncbi.nlm.nih.gov/BLAST/), all three PACSINs are found in all vertebrates, includ-ing mammals, birds, frogs and fishes. On the contrary, only PACSIN 2 is found in wormsand insects. On the basis of sequence similarity, these related proteins form a novel fam-ily of proteins called the FAP52/PACSIN/syndapin (FPS) family. The known FPS pro-teins are listed in Table 1. FPS family is part of a larger protein family known as PCH, thename derived from their homology to the cdc15-protein of Schizosaccharomyces pombe(PCH= pombe cdc15 homology, Lippincott & Li 2000, see below).

23

Table 1. The currently identified FPS proteins.

2.1.1 The PCH family

The PCH family is an emerging group of proteins, which characteristically have a typicaldomain architecture, an NH2-terminal coiled-coil and COOH-terminal SH3 domain, andwhich participate in actin-based cellular events, cytokinesis and vesicular traffic. How-ever, the members show rather low sequence similarity (BLAST search). The currentmembers of the PCH superfamily are Cdc15 (Fankhauser et al. 1995) and Imp2 (Demeter& Sazer 1998) of Schitzosaccharomyces pombe, Hof1/Cyk2 (Kamei et al. 1998,Lippincott & Li 1998) of Saccharomyces cerevisiae, FPS proteins (Meriläinen et al.1997, Plomann et al. 1998, Qualmann et al. 1999), and the mammalian proteins PSTPIP1, a proline-serine-threonine phosphatase-interacting protein (Spencer et al. 1997),CD2BPI, a CD2 cytoplasmic tail-binding protein I (Li et al. 1998), and CIP4, aCdc42-interacting protein 4 (Aspenström 1997). Mouse protein MAYP, the macrophageactin-associated tyrosine-phosphorylated protein (Yeung et al. 1998) and its humanortholog PSTPIP 2 (Wu et al. 1998) are similar to PSTPIP 1 except for the lack of theSH3 domain.

Source Protein Reference (if available) Accession nrHomo sapiens PACSIN 1

PACSIN 2PACSIN 3

Sumoy et al. 2001Ritter et al. 1999Sumoy et al. 2001

AF242529NM_007229AF242530

Mus musculus PACSIN 1PACSIN 2PACSIN 3

Plomann et al. 1998Ritter et al. 1999Modregger et al. 2000

BC014698NM_011862AF242531

Rattus norvegicus Syndapin ISyndapin IIPACSIN 3

Qualmann et al. 1999Qualmann & Kelly 2000

NM_017294NM_130740

Gallus gallus PACSIN 1FAP52PACSIN 3

Meriläinen et al. 1997 GGDP52MR

Xenopus laevis PACSIN 1PACSIN 2PACSIN 3

Cousin et al. 2000 NP_035992

Tetraodon nigroviridis PACSIN 1PACSIN 2PACSIN 3

Drosophila PACSIN 2C. elegans PACSIN 2Anopheles gambiae PACSIN 2

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2.1.2 Structure

2.1.2.1 Domains and the sequence analysis of FPS family proteins

The FPS proteins share a common order of domains as schematically presented in Fig. 1.In the NH2-terminus, there is an 89-aa long, highly α-helical Fes-CIP4 homology (FCH)domain (Lippincott & Li 2000), followed by α-helical coiled-coil domain, linker region,and the COOH-terminal SH3 domain. Additionally, FPS proteins contain consensussequences for phosphorylation by serine/threonine kinases, PKA, PKC and casein kinase(Plomann et al. 1998).

Fig. 1. Domain structure of FPS family proteins. The aa scaling is according to chicken FAP52.The structure is rather uniform, except for the linker region, which displays notable heteroge-neity. In PACSIN 3, instead of two NPF motifs (asparagine-proline-phenylalanine sequences),it contains a proline-rich region. On the other hand, mouse PACSIN 2 and rat PACSIN 2 iso-forms IIaa and IIba contain an additional 41-aa insert, which includes the third NPF motif.

The FCH domain contains a 20 aa-long RAEYL motif, named according to the con-served amino acid residues in its sequence (Plomann et al. 1998). The FCH domain istypically found in the NH2-termini of proto-oncogene Fes/FES-related tyrosine kinases,RhoGAP-type GTPase activating proteins and in PCH family members, ranging fromviruses and yeast to protozoans and mammals (domain search with the program SMART).The function of the FCH domain is largely unknown. Some recent results suggest, how-ever, that it mediates interaction with microtubules (Tian et al. 2000, Fujita et al. 2002).

Most FCH domain-containing proteins also have a coiled-coil domain in theirNH2-terminal halves (analysis with the SMART program). The coiled-coil domain of FPSproteins has a propensity to mediate self-association, judged by computational analysesbased on PairCoil- and Coils-programs (network servers at http://nightingale.lcs.mit.edu/cgi-bin/score and http://www.ch.embnet.org/software/COILS_form.html). The NH2-ter-

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minal region containing the FCH and coiled-coil domains is also called Cdc15-NTdomain, where Cdc15 refers to S. pombe protein Cdc15 and NT to its NH2-terminus(Ritter et al. 1999).

The linker region, which separates the NH2-terminal FCH and coiled-coil domainsfrom the COOH-terminal SH3 domain, is approximately 110 aa-long. It does not displayany regular secondary structure or any specific function (Meriläinen et al. 1997). It does,however, contain a PEST sequence, a proline, glutamic/aspartic acid, serine and threo-nine-rich region, which is considered a signal for rapid degradation of proteins (Rogers etal. 1986, Rechsteiner & Rogers 1996). It also contains 2 or 3 asparagine-proline-phenyla-lanine (NPF) motifs (see Chapter 2.1.2.2). The NPF motif is a binding site for Eps15homology domains (Paoluzi et al. 1998). Currently, no interactions mediated by NPFmotifs of any FPS proteins have been described.

The SH3 domain is a widely occurring domain in a large variety of different proteins.Via binding to proline-rich target sequences, it participates in a wide variety of cellularevents by mediating specific interactions between the components of signaling pathwaysand protein assemblies. One of its functions is to direct the localization of molecules totheir specific compartments in the cell (for reviews, see Yu et al. 1994, Cohen et al. 1995,Mayer 2001). The SH3 domain of PACSIN isoforms has been found to bind to the follow-ing proteins (for references, see table II): dynamin I, synaptojanin, and synapsin I, anactin polymerization nucleating factor N-WASP, a cell surface metalloproteases MDC15,ADAM12 and ADAM13, the mammalian Son-of-sevenless (mSos), an apoptotic media-tor CD95L and huntingtin. The known interactions are listed in Table 2.

Table 2. Currently characterized binding partners of FPS proteins.

2.1.2.2 Sequence comparison

Members of the FPS family are closely related proteins a with a high aminoacid-sequence similarity. The identities among the mammalian and chicken orthologs are81–98% for PACSIN 1, 85–98% for PACSIN 2, and 65–97% for PACSIN 3 (sequencecomparison with BLAST). The relationship between FPS proteins is presented by a phy-logenetic tree in Fig. 2.

Binding partner ReferenceDynamin I Qualmann et al. 1999, Modregger et al. 2000synaptojanin Qualmann et al. 1999, Modregger et al. 2000Synapsin I Qualmann et al. 1999, Modregger et al. 2000N-WASP Qualmann et al. 1999, Modregger et al. 2000MDC15 Howard et al. 1999ADAM13 Cousin et al. 2000mSos Wasiak et al. 2001CD95L Ghadimi et al. 2002huntingtin Modregger et al. 2002ADAM12 Mori et al. 2003

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Fig. 2. The phylogenetic tree of the FPS proteins of man, mouse, rat and chicken.

The NH2-terminus (aa 1–340 in FAP52) and the SH3 domain (aa 390–448) are highlyconserved regions in the members of the FPS family, typically having an identity of morethan 90% among these proteins. One outstanding feature is the variation/alternative splic-ing in the region of four acidic residues EDDE/EDEE in the “linker” region (aa 302–305in chicken FAP52). In addition to FAP52, PACSIN 2 isoforms IIaa and IIab in the rathave these four residues intact, whereas the isoforms IIba and IIbb, rat PACSIN 1 and allmouse PACSINs lack the first two residues (Qualmann & Kelly 2000; sequence analysiswith the BLAST program). Alternative splicing of PACSIN 2 orthologs in humans,mouse, rat and chicken is schematically presented in Fig. 3.

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Fig. 3. Presence of the alternative splicing sites in the PACSIN 2 isoforms in humans, mouse,rat and chicken. There are two alternative splicing sites in the polypeptides, the two acidic re-sidues ED, and the 41-aa insert in the linker region. Two different splice variants have beenfound in humans, and four in rat. In mouse and chicken, only one form has been described.

The highest variation in the sequence of the FPS proteins is seen within the linkerregion. The two or three NPF motifs are fully conserved among the family, with theexception of PACSIN 3. Instead of NPF motifs, the linker region of Pacsin 3 contains twoproline-rich sequences PxxP (P, proline; x, whatever aa), which are putative binding sitesfor SH3 domains. A distinguishing feature of the linker region of PACSIN 2 and syndap-ins IIaa and IIba is an approximately 40 aa-long insert (see Fig. 3), which is not present inthe other members. The insert also contains an additional NPF motif (Qualmann & Kelly2000, analysis with BLAST). Interestingly, the NPF motifs throughout the FPS family areclosely associated with acidic aspartic and glutamic acid residues. The consensussequences for the three NPF regions are ZZZxxNPFxxxZ, NPFZZZ and NPFZZZZZ (Z,either aspartic or glutamic acid; x, whatever residue), as schematically presented in Table3. Two of them contain the sequence NPFxD/E; NPFxD has been reported to serve as anendocytotic signal (Tan et al. 1996).

Table 3. Consensus sequences flanking the NPF motifs.

Protein I (insert) II IIIFAP52 deesnNPFsstd* NPFdedPACSIN 1 ddesgNPFggne NPFeddPACSIN 2 NPFededd ddesnNPFsstd NPFdedSdp I ddesgNPFggne NPFeddSdp IIaa/ba NPFededd ddesnNPFsstd NPFdedSdp IIab/bb ddesnNPFsstd NPFdedConsensus NPFZZZZZ ZZZxxNPFxxxZ NPFZZZ* n, asparagine; p, proline; f, phenylalanine; e, glutamic acid; d, aspartic acid; Z, either e or d; x, whatever aa

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2.1.3 Tissue expression

PACSIN 1 is expressed in the brain, as studied by Northern and Western blotting of vari-ous tissues, and by immunoelectron and immunofluorescence microscopy of mouse braintissue and cultured rat forebrain neurons, respectively (Plomann et al. 1998, Qualmann etal. 1999). PACSIN 1 has also been found expressed in dorsal root ganglia neurons (Plo-mann et al. 1998). Human, but not mouse PACSIN 1 is also seen expressed in the heart,pancreas and liver (Sumoy et al. 2001). FAP52/PACSIN 2 is ubiquitously expressed inchicken, rat and mouse tissues (Meriläinen et al. 1997, Ritter et al. 1999, Qualmann &Kelly 2000). In rat, the PACSIN 2 splice variants IIaa and IIba have been found specifi-cally in the heart when studied by Western blotting of a variety of rat tissues (Qualmann& Kelly 2000). PACSIN 3 has been detected in all examined tissues, but is epressed par-ticularly strongly in the lung and muscle (Modregger et al. 2000, Sumoy et al. 2001).

2.1.4 Function

PACSINs have been implicated in endocytosis and actin organization (Plomann et al.1998, Qualmann et al. 1999, Modregger et al. 2000, Qualmann & Kelly 2000). Overex-pression of PACSINs induces a rearrangement of cortical actin cytoskeleton (Qualmann& Kelly 2000), and, on the other hand, inhibits endocytosis (Simpson et al. 1999,Modregger et al. 2000, Qualmann & Kelly 2000). FAP52 is phosphorylated in serine resi-dues and associates with focal adhesions together with several other signaling proteins(Meriläinen et al. 1997).

2.1.4.1 FPS family proteins in neurons

PACSIN 1 is a highly brain-specific protein (Plomann et al. 1998, Qualmann et al. 1999).All PACSINs bind to the neuron-specific synaptic proteins dynamin I and synaptojanin(Qualmann et al. 1999, Modregger et al. 2000, Qualmann & Kelly 2000). In rat, PAC-SINs also bind to synapsin I, a protein implicated in the regulation of neurotransmitterrelease (Ferreira & Rapoport 2002). Expression of PACSIN 1 in the brain is developmen-tally regulated, so that the level of expression is increased from the 17th embryonal day toa fully differentiated, adult mouse. On the other hand, its expression is decreased withinentorhinal-cortex lesion, suggesting that it negatively regulates self-repair system andregeneration (Plomann et al. 1998). Association of PACSIN 1 with neural differentiationis implicated by the finding that it is phosphorylated by casein kinase II, an enzymeknown to play a role in neurogenesis (Diaz-Nido et al. 1994, Lim & Zaheer 1995). Histo-logically, PACSIN 1 is localized to the large neurons of the cortex and the brain stem, thepyramidal cells and astrocytes of the hippocampus. It is also seen in purkinje cells and inthe molecular, but not the granular, layer of the cerebellum (Plomann et al. 1998).

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A recent study has implicated PACSIN 1 in the pathogenesis of Huntington’s disease(Modregger et al. 2002) in that the SH3 domain of PACSIN 1 binds to mutated, but not tonon-mutated, huntingtin. Huntington’s disease is a neurodegenerative disease, in whichthe gene encoding huntingtin is mutated to encode an abnormally long polyglutamatestretch. This is accompanied by neuronal loss in the striatal cortex, and, as a clinical man-ifestation, by progressive motor, psychiatric and cognitive dysfunctions (Young 2003). Inthe brain tissue in Huntington’s disease, the distribution PACSIN 1 is altered so thatinstead of axons, it is seen in the perinuclear cytoplasm already at the early stages of thedisease (Modregger et al. 2002).

Also PACSINs 2 and 3 have recently been linked to neuronal differentiation via theirbinding to ADAM13. PACSIN 2 of Xenopus laevis (X-PACSIN 2) is suggested to serve asa downregulator of metalloprotease disintegrin ADAM13 (Cousin et al. 2000). ADAMs(also known as MDCs) are multifunctional transmembrane metalloglycoproteases, whichare involved in a variety of biological processes, such as fertilization, neurogenesis, myo-genesis, embryonic transforming growth factor-α release and the inflammatory response(Primakoff & Myles 2000). X-PACSIN 2 colocalizes with ADAM13 in migrating cepha-lic neural crest cells in the developing frog embryo, and, via its SH3 domain, binds to theproline-rich regions in the cytoplasmic domain of ADAM13 in vitro. In cultured XenopusXTC cells, X-PACSIN 2 colocalizes with ADAM13 in membrane ruffles and cytoplas-mic vesicles. Overexpressed X-PACSIN 2 also inhibits the developmental defects causedby ADAM13 overexpression, indicating that X-PACSIN downregulates the function ofADAM13 (Cousin et al. 2000).

2.1.4.2 Function in endocytosis

Endocytosis can be subdivided into five sequential steps: membrane invagination, coatedpit formation, coated pit sequestration, detachment of the newly formed vesicle, andmovement of the vesicle away from the plasma membrane. FPS proteins have beenclosely associated with the regulation of endocytosis via several interactions. They bind,via their SH3 domains, to dynamin I, synaptojanin and synapsin I, proteins that regulatevesicular traffic on plasma membrane (Modregger et al. 2000, Qualmann & Kelly 2000).Dynamin is a GTPase and plays an important role at the fission step of nascent clath-rin-coated vesicles from plasma membrane (Schmid 1997). Synaptojanin, on the otherhand, is a phosphatase and is present in endocytotic coated intermediates (McPherson etal. 1996) via interactions with various phosphoinositides (Guo et al. 1999). Synapsin I isa neuronal vesicle-associated phosphoprotein involved in exocytosis. It also regulatesarchitecture of the actin cytoskeleton in the presynaptic nerve terminal (Sudhof et al.1989, Greengard et al. 1993). Moreover, PACSINs bind to N-WASP and have been sug-gested to serve as a molecular link between the actin cytoskeleton and vesicle transport inpresynaptic nerve terminal (see chapter 2.4; Qualmann et al. 1999, Modregger et al.2000, Qualmann & Kelly 2000).

In cultured hippocampal neurons, PACSIN 1 has been shown to partially colocalize invesicular structures with dynamin I, but not with clathrin (Modregger et al. 2000). Similarresults were also obtained for PACSINs 2 and 3 in cultured NIH 3T3 fibroblasts and

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C2F3 myotubes, respectively (Modregger et al. 2000). PACSIN 2 shows vesicular-likecytoplasmic staining also in migrating neural crest cells in the developing frog embryo(Ritter et al. 1999). Colocalization with dynamin I has also been seen for rat PACSIN 1 inthe synaptic vesicles and growth cones of cultured neurons (Qualmann et al. 1999, Kes-sels & Qualmann 2002), and for rat PACSIN 2 in cultured rat PC12 cells (Qualmann &Kelly 2000). Thus, FPS proteins seem to interact with dynamin I in the dynamics of vesi-cle transport rather than to serve as structural components of clathrin-coated vesicles.

The function of FPS proteins in endocytosis has been studied in overexpression experi-ments in cultured cells. For instance, the SH3 domain of PACSIN 1 was transiently over-expressed in 3T3-L1 adipocytes, and endocytosis was monitored by detecting the inter-nalization of transferrin (Simpson et al. 1999). The SH3 domain inhibited endocytosis atthe fission step in a dose-dependent manner. In another assay, in which PACSIN isoformswere overexpressed in cultured cells, PACSINs were shown to block the endocytosis oftransferrin (Modregger et al. 2000). The inhibitory effect was abolished when the func-tion of the SH3 domain was abrogated by a specific point mutation. Several other SH3domains also block endocytosis in vitro. For instance, overexpression of the NH2-termi-nal SH3 domain of intersectin inhibits the intermediate phase of endocytosis, while that ofendophilin I and amphiphysin II blocks the fission step (Simpson et al. 1999). Recently, anovel regulatory aspect in endocytosis was discovered involving increased association ofPACSIN 1 with dynamin I upon phosphorylation of PACSIN 1 on serine by an inositolhexakisphosphate-regulated protein kinase (Hilton et al. 2001). The phosphorylatedserine resides outside the SH3 domain that mediates the binding to dynamin I. Inositolhexakisphosphate is a phosphoinositol, which has been suggested to be involved in a widevariety of cellular processes, such as regulation of mRNA transport from nucleus, DNArepair, and control of phosphatase activity in pancreatic β cells (Shears 2001).

In addition to the interactions between FPS proteins and the endocytic proteins dis-cussed above, the FPS protein-mediated inhibition of endocytosis may also be relayed viaRho proteins, multifunctional proteins that affect actin cytoskeleton, membrane traffick-ing, transcription, cell adhesion and cell cycle regulation (Hall 1998, Ridley 2001). Tothat effect, a recent study reveals direct SH3 domain-mediated binding of PACSINs tomSos, a guanine nucleotide exchange factor and activator of the Rho proteins Ras andRac (Wasiak et al. 2001). Activated Rac, on the other hand, inhibits clathrin-coated vesi-cle formation and transferrin receptor-mediated endocytosis (Lamaze et al. 1996).

A recent study describes an interaction of PACSIN 2 with CD95 ligand (CD95L), atransmembrane receptor, which mediates cell death. PACSIN 2 has been suggested to reg-ulate the expression of CD95L on plasma membrane by vesicle trafficking (Ghadimi etal. 2002).

2.1.4.3 Actin reorganization

FPS proteins have been suggested to regulate actin organization via two specific SH3domain-mediated interactions. They include, first, the actin polymerization proteinN-WASP, and, second, the ras- and rac-regulating protein Sos. FPS proteins bind to thepolyproline region of the COOH-terminal half of N-WASP (Qualmann et al. 1999,

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Modregger et al. 2000, Qualmann & Kelly 2000). N-WASP has an essential role in actinpolymerization, which is the basis of the formation of filopodia and microspikes (Miki etal. 1998). In cultured HeLa cells and NIH 3T3 fibroblasts, overexpression of rat PAC-SINs 1 and 2 induced a rearrangement of the cortical actin and formation of long filopo-dia. In an experiment in which the COOH-terminus of N-WASP, containing the VCAregion that is responsible for Arp2/3 binding, was overexpressed, these effects on actinorganization were suppressed (Qualmann & Kelly 2000). Thus, the actin regulation isthought to be mediated via the Arp2/3 complex. N-WASP acts downstream of Cdc42 andphosphatidylinositol-4,5-bisphosphate (PIP2) (reviewed by Tekenawa & Miki, 2001). Inresting cells, N-WASP is in autoinhibited state, where the VCA region is masked by anintramolecular interaction involving VCA and the Cdc42-binding domain, GBD/CRIB.Binding of Cdc42 brings about conformational changes in N-WASP, which unmasks theCOOH-terminus and allows its binding to the Arp2/3 complex (Kim et al. 2000).

The route by which FPS proteins induce Arp2/3 activation is unclear. Several poten-tial mechanisms have been suggested (Qualmann & Kelly 2000). They may work asupstream activators of the N-WASP activator Cdc42, behave like Cdc42 and cause theconformational change, recruit N-WASP to membrane, or directly activate the Arp2/3complex. For further discussion of Arp2/3, see chapter 2.2.2.

There is also accumulating evidence that FPS proteins could regulate the dynamics ofthe cytoskeleton via mitogen-activated protein kinases (MAPK) (Wasiak et al. 2001).PACSINs 1 and 2 bind to the proline-rich sequence of mSos, activator for Ras and Rac,which regulate MAPKs and actin dynamics (Hall 1998). Activation of Rac has multipleeffects, such as membrane ruffling and lamellipodia formation (Hall 1998, Ridley 2001).The interaction is mediated by the SH3 domain of PACSINs, and it is abolished by domi-nant negative proline-to-leucine mutation (P434L for PACSIN 1; P478L for PACSIN 2)in the SH3 domain (Wasiak et al. 2001).

A coupling of FPS proteins to actin dynamics is also suggested by the recent findingthat PACSIN 3 binds to ADAM12 (Mori et al. 2003), an enzyme involved in theectodomain shedding of the growth factor HB-EGF, and, thus, in the EGF receptor trans-activation. ADAM12 has also been implicated in actin reorganization and muscle devel-opment and regeneration (Gilpin et al. 1998, Bornemann et al. 2000, Kawaguchi et al.2003).

2.2 Actin microfilaments and focal adhesions

2.2.1 Features of actin microfilaments

Actin microfilaments are responsible for the maintenance and regulation of cell shape,but they also play an essential role in dynamic processes, such as cell division, spreadingand migration (Welch & Mullins 2002). Moreover, they participate in the subcellular pro-cesses, such as vesicular transport and cell organelle movement (Taunton 2001). Actin fil-aments display architectural polymorphism in cells. They can bind together parallelly to

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form tight bundles (stress fibers), and thus provide the cell the ability to resist mechani-cal stress, but they can also form a loosely oriented network, as is the case in lamellipo-dia (Stryer 2000).

Actin filaments consist of fibrous actin (F-actin), which is a polymer of globular actin(G-actin) monomer. A single actin filament has the appearance of a two-stranded helix.They are polarized and dynamic structures, consisting of a barbed end (fast-growing end,positive end) facing the plasma membrane, and pointed end (slowly growing end, nega-tive end) facing the cell center. Polymerization and depolymerization processes happensimultaneously in both ends. The barbed end favors polymerization, whereas depolymer-ization takes place in the pointed end. Actin is an ATPase, and in the cell the dynamics ofactin filaments are partly driven by ATP hydrolysis. The speed of polymerization/depoly-merization depends on the concentration of free G-actin molecules, on their bound ATPlevel, and on the presence of various regulatory proteins (see below). New actin subunitscan be rapidly added to or removed from membrane-facing ends of the filaments intightly regulated biological processes, such as cell migration and spreading. Rapid actinpolymerization or depolymerization can be a response to an extra- or intracellular signal(Welch & Mullins 2002).

2.2.2 Proteins regulating actin polymerization

Numerous actin-binding proteins possess a propensity to modulate the actin organizationin a cell. They cross-link actin filaments together, cap F-actin, sever the filaments to pro-duce new barbed ends, bind free G-actin or link other proteins to F-actin (Hubberstey &Mottillo 2002, Remedios et al. 2002). The best-characterized and most important actorsare discussed here.

Actin-binding proteins are a heterogenous group of proteins, which do not share anindividual actin-binding domain (ABD) with a common consensus sequence. However,certain subfamilies with a structural homology in ABDs have been identified. Forinstance, ABD of calponin, filamin, fimbrin and α-actinin consists of one to four calponinhomology domains. They display a similar 3-D fold with 7 to 8 α-helical stretches andrandomly coiled stretches between them, but have low sequence similarity (Winder2003). The ABD of gelsolin superfamily proteins, which include e.g. gelsolin, villin andseverin, on the other hand, is folded into several sandwiches consisting of numerousα-helices and β-sheets (Winder 2003).

Capping protein (Cap Z) is an important negative regulator of actin polymerization.Almost all barbed ends are capped with Cap Z. Dissociation of Cap Z is a prerequisite forpolymerization or depolymerization to take place. (Cooper & Pollard 1985, Casella et al.1987, Caldwell et al. 1989, Carlier & Pantaloni 1997). However, Cap Z is necessary forcell motility (Hug et al. 1995, David et al. 1998, Loisel et al. 1999). Binding of PIP2 toCap Z leads to its dissociation from the barbed ends (Heiss & Cooper 1991).

Profilin is a small protein that binds to G-actin (Carlsson et al. 1977), PIP2 (Lassing &Lindberg 1985) and poly-L-proline containing proteins (Tanaka & Shibata 1985, Lind-berg et al. 1988). Vasodilator-stimulated phosphoprotein (VASP; Reinhard et al. 1995),and possibly radixin (Funayama et al. 1991), zyxin (Sadler et al. 1992) and CAP-like pro-

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teins (Vojtek et al. 1991) are the poly-L-proline-containing ligands of profilin. Profilinassociates with microfilaments in the most dynamic areas of the cell, such as leadinglamellae and newly formed attachment sites (Buss et al. 1992). The mechanism by whichprofilin regulates the dynamics of actin microfilaments is complex. By sequestratingG-actin, profilin decreases the accessibility of the building blocks of F-actin, thus leadingto inhibition of actin polymerization and induction of depolymerization (Sohn & Gold-schmidt-Clermont 1994). On the other hand, profilin has also been suggested to catalyzeactin polymerization by delivering monomers to F-actin (Pring et al. 1992), and by con-verting ADP-actin to ATP-actin (Goldschmidt-Clermont et al. 1992). The profilin-G-actincomplex can interact directly with actin filaments, and in that way counteract polymeriza-tion (Goldschmidt-Clermont et al. 1992, Pantaloni & Carlier 1993, Theriot & Mitchison1993, Sohn & Goldschmidt-Clermont 1994). PIP2, upon binding to profilin, inhibits itsbinding to actin. On the other hand, binding of PIP2 to actin-bound profilin causes the dis-sociation of the complex (Katakami et al. 1992).

Gelsolins are a large family of actin-binding proteins (Janmey 1993) that have thecapacity to bind to barbed ends of actin, nucleate new filament assembly, and bind tosides of filaments and sever them into smaller fragments (Lind et al. 1987, Hartwig 1992,Lamb et al. 1993, Allen & Janmey 1994). The capping and severing activity of gelsolin isactivated by elevated concentration of Ca2+-ions or a decrease of pH (Lamb et al. 1993),and inhibited by binding of PIP2 (Janmey et al. 1987, Janmey & Stossel 1989). Severingproduces new barbed ends, which can, after dissociation of gelsolin, serve as new foci ofactin polymerization. In transformed fibroblasts, gelsolin has been found to be associatedwith podosomes (Wang et al. 1984), which are transient, highly dynamic types of focaladhesions along the edges of spreading cells (Petit & Thiery 2000). Due to the strongerbinding of gelsolin than profilin to actin, its capping activity surpasses that of profilin(Ampe & Vandekerckhove 1994, Rozycki et al. 1994). Gelsolin is necessary forRac-induced formation of filopodia (Azuma et al. 1998). Activation of Rac results in dis-sociation of gelsolin from F-actin (Arcaro 1998).

Adseverin/scinderin, fragmin, severin, villin and Cap G are close relatives of gelsolin.They show structural and functional similarity to gelsolin, with the exception that villinhas the additional ability to merge actin filaments to bundles and that Cap G does notsever F-actin (Cant et al. 1998, Friederich et al. 1999, Hubberstey & Mottillo 2002). Twoadditional structural and functional relatives of villin, advillin (Marks et al. 1998) andsupervillin (Wulfkuhle et al. 1999) have also been described. Tensin and its proteolyticfragment insertin bind barbed ends and decrease the polymerization rate of F-actin (Weigtet al. 1992, Lo et al. 1994a, Chuang et al. 1995). Tensin has also been shown to bind tothe sides of actin filaments (Lo et al. 1994b).

Arp2/3 is a seven-subunit complex, which, in response of extracellular stimulus nucle-ates actin polymerization. The Arp2/3 complex consists of two actin-related proteins,Arp2 and Arp3, and of five additional proteins, named ARPC1-5 (Welch & Mullins2002). Depending on the upstream signaling molecule, activation of the Arp2/3 complexcan lead to the formation of either lamellipodia/membrane ruffling, or filopodia/microspikes. The former is a result of mesh-like actin filament organization, the latter ofthe formation of straight actin bundles (Takenawa & Miki 2001). Activation of the Arp2/3complex by N-WASP directs actin polymerization to filopodia formation (Rohatgi et al.1999). The mechanism behind this is unclear. One putative regulator could be the

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actin-bundling protein filamin, which plays an important role in Cdc42-induced formationof filopodia (Ohta et al. 1999). Two mechanisms for Arp2/3 complex to activate actinpolymerization have been suggested. First, the dendritic nucleation model, in which theactivated Arp2/3 complex binds to the sides of existing filaments and nucleates theassembly of new filaments (Mullins et al. 1998, Blanchoin et al. 2000, Pollard et al. 2000,Millard et al. 2004). Second, the barbed end branching model, in which activated Arp2/3nucleates branched filaments at existing barbed end of filament (Pantaloni et al. 2000,Millard et al. 2004). Arp2/3-induced actin polymerization is also important in the intracel-lular motility of Listeria monocytogenes, which utilizes its surface protein ActA as anactivator of the Arp2/3 complex (Welch et al. 1998).

Several proteins are known to bind to sides or pointed ends of actin microfilamentsand in this way to regulate actin polymerization or depolymerization. Tropomyosin, forinstance, binds multiple subunits along the side of F-actin, and, when bound near thepointed end, prevents them from dissociation (Broschat 1990). Tropomodulin caps thepointed ends in striated muscle and participates in the assembly of sarcomers (Littlefield& Fowler 1998). Actin depolymerizating factor/cofilin-family members sever actin fila-ments into small pieces (Maciver et al. 1991, Du & Frieden 1998, Maciver et al. 1998,Blanchoin & Pollard 1999) and increase the loss of actin subunits from the pointed end(Carlier et al. 1997, Maciver et al. 1998).

Ezrin/radixin/moesin family members couple actin fibers to plasma membrane, proba-bly by binding to the sides of the filaments (Pestonjamasp et al. 1995). Purified radixinalso binds to the barbed ends in vitro (Tsukita et al. 1989). An integral membrane protein,ponticulin, also couples F-actin to plasma membrane by binding to the sides of microfila-ments (Hill et al. 1994). In addition, it has the ability to nucleate actin assembly and cre-ate new barbed and pointed ends (Chia et al. 1993).

Fimbrin binds to the sides of microfilaments, and it is the major cross-linker of the fil-aments in the core of microvilli (Bretscher & Weber 1980). It contains two adjacentactin-binding domains and, thus, couples individual filaments to tight bundles (Matsu-daira et al. 1983). In inner ear hair cell stereocilia it serves as an actin-bundling protein(Tilney et al. 1989).

2.2.3 Focal adhesions – structure and dynamics

2.2.3.1 Structure of focal adhesions

The major adhesion sites between the cell and the extracellular matrix (ECM), termedfocal adhesions (FAs), were initially observed by interference-reflection and electronmicroscopy as spots along the ventral plasma membrane of cultured fibroblasts (Aber-crombie et al. 1971, Abercrombie & Dunn 1975, Izzard & Lochner 1976, Izzard &Lochner 1980). F-actin is present at early stages of FA-formation in chicken embryofibroblasts (Depasquale & Izzard 1987), suggesting its important role in FA assembly. Infully developed FAs, the associated actin filaments merge to form tight parallel bundles.

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Their barbed ends face FAs. New actin subunits can be rapidly added to or removed frommembrane-facing ends of the filaments in tightly regulated biological processes, such ascell migration and spreading.

FAs couple the actin cytoskeleton of the cell to ECM, or, in cell cultures, to the growthsubstratum, and, thus, fix the cells to their environment. In addition to the role of FAs inconnecting cells to the proper position in relation to their neighborhood and mediatingmechanical force through plasma membrane, FAs serve as machinery for various extra-cellular stimuli to penetrate plasma membrane and to trigger and regulate various signal-ing cascades. Conversely, the cytoplasmic regulatory molecules also mediate the retro-grade flow of signaling by modulating the function of FAs and e.g. the tenacity of theadhesion. FAs mediate numerous signals, such as those regulating cell adhesion, migra-tion, proliferation and differentiation, apoptosis and gene expression. They play an essen-tial role in tissue formation during embryogenesis and in biological regeneration pro-cesses, such as wound healing (for reviews of FAs, see Jockusch et al. 1995, Burridge &Chrzanowska-Wodnicka 1996, Petit & Thiery 2000, Juliano 2002).

FAs consist of transmembrane receptors and cytoplasmic proteins. Integrins are themain group of the transmembrane receptors segregated in FAs. The intracellular proteinsinclude structural components, regulatory proteins, such as protein kinases and phos-phatases and their substrates, as well as adapter proteins. Clear and exact distinctionsbetween these groups cannot be made. The adapter protein paxillin, for instance, has alsostructural and regulatory features. Numerous FA components have been described, aslisted in Table 4. The most important and best-characterized proteins with direct interac-tion with actin microfilaments are discussed in the following. For a detailed scrutiny, seethe excellent reviews by Jockusch et al. (1995), Burridge and Chrzanowska-Wodnicka(1996), Petit and Thiery (2000), Zamir and Geiger (2001) and Juliano (2002).

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Table 4. List of known focal adhesion proteins.Transmembraneproteins

Regulatory proteins Structural proteins Adapter proteins or unclassified

Caveolin-1 (1)CD98/4F2HC (2)EGF-receptor (3)insulin receptor (4)integrins (18 α- and 8 β-sub-units) (5)IAP/CD47 (6)LAR-PTP (7)layilin (8)PDGF-receptor (9)polycystin-1 (10)proteoglycans (11)SHPS-1 (12)syndecan4/ amphiglycan (13)TM4SF proteins/ tet-raspanins (14)uPA-receptor (15)

Abl (16)AND-34 (17)ASAP1 (18)β3-endonexin (19)calnexin (20)calpain II (21)calreticulin (22)CASK (23)Csk (24)cytohesin-1 (25)Dbl (26)FAK (27)Fyn (28)gelsolin (29)Graf (30)ICAP-1 (31)ILK (32)PAK (33)PIX (34)PI3K (35)PKA (36)PKB/AKT (37)PKC (38)PKG (39)PKL (40)PLCγ1 (41)PP2A (42)profilin (43)PSGAP (44)PTEN (45)PTP-PEST (46)PTP1B (47)Pyk2/CAKβ/ Raftk/Cadtk (48)Rac (49)Rack1 (50)radixin (51)RhoA (52)Shp-2 (53)Sos (54)Src (55)TAP20 (56)Trio (57)

actin (58)α-actinin (59)filamin (60)(acto)parvin (61)ponsin (62)talin (63)tensin/insertin (64)tenuin (65)vinculin (66)vinexin (67)zyxin (68)

CAP (69)Cas (70)Cbl (71)CH-ILKBP (72)CIB (73)Crk (74)CRP-1/bombesin (75)C3G (76)DOCK180 (77)DRAL (78)EAST (79)FAP52 (80)fimbrin (81)Grb2 (82)Grb7 (83)Hic-5 (84)IRS-1 (85)LIP.1 (86)MARCKS (87)melusin (88)Nck-2 (89)nexilin (90)palladin (91)paxillin (92)pE6 protein (93)PINCH (94)PKL (95)PST-PIP (96)Shc (97)syndesmos (98)synectin (99)syntenin (100)VASP/ENA (101)vimentin (102)

1. Glenney Jr & Soppet 1992, 2. Teixeira et al. 1987, 3. Fox et al. 1980, 4. Raizada & Perdue 1967, 5. Hynes 2002, 6. Brown et al.1990, 7. Pulido et al. 1995, 8. Borowsky & Hynes 1998, 9. Heldin et al. 1981, 10. Harris et al. 1995, 11. Heinegard & Gardell1967, 12. Fujioka et al. 1996, 13. David et al. 1992, 14. Maecker et al. 1997, 15. Del Rosso et al. 1985, 16. Goff et al. 1980, 17.Cai et al. 1999, 18. Brown et al. 1998, 19. Shattil et al. 1995, 20. Wada et al. 1991, 21. Murachi et al. 1980, 22. Ostwald & Mac-Lennan 1974, 23. Hata et al. 1996, 24. Okada et al. 1991, 25. Kolanus et al. 1996, 26. Srivastava et al. 1986, 27. Schaller et al.1992, 28. Popescu et al. 1987, 29. Yin & Stossel 1979, 30. Hildebrand et al. 1996, 31. Chang et al. 1997, 32. Hannigan et al. 1996,33. Manser et al. 1995, 34. Bagrodia et al. 1998, 35. Whitman et al. 1987, 36. Miyamoto et al. 1968, 37. Coffer & Woodgett, 1991,38. Inoue et al. 1977, 39. Kuo & Greengard, 1970, 40. Bagrodia et al. 1999, 41. Ryu et al. 1986, 42. Jakes et al. 1986, 43. Carlssonet al. 1977, 44. Ren et al. 2001, 45. Li et al. 1997, 46. Yang et al. 1993, 47. Charbonneau et al. 1989, 48. Lev et al. 1995, 49. Pola-kis et al. 1989, 50. Mochly-Rosen et al. 1991, 51. Tsukita et al. 1989, 52, Ridley & Hall 1992, 53. Adachi et al. 1992, 54. Rogge etal. 1991, 55. Duesberg et al. 1976, 56. Tang et al. 1999, 57. Debant et al. 1996, 58. Ivanov & Asmolova, 1950, 59. Maruyama &Ebashi, 1965, 60. Wang et al. 1975, 61. Olski et al. 2001, 62. Mandai et al. 1999, 63. Collier & Wang 1982, 64. Davis et al. 1991,65. Tsukita et al. 1989, 66. Geiger et al. 1980, 67. Kioka et al. 1999, 68. Crawford & Beckerle 1991, 69. Fedor-Chaiken et al.1990, 70. Sakai et al. 1994, 71. Langdon et al. 1989, 72. Tu et al. 2001, 73. Naik et al. 1997, 74. Mayer et al. 1988, 75. Anastasi1971, 76. Tanaka et al. 1994, 77. Hasegawa et al. 1996, 78. Genini et al. 1997, 79. Lohi et al. 1998, 80. Merilainen et al. 1997, 81.Bretscher & Weber 1980, 82. Lowenstein et al. 1992, 83. Margolis et al. 1992, 84. Shibanuma et al. 1994, 85. Sun et al. 1991, 86.Serra-Pages et al. 1995, 87. Stumpo et al. 1989, 88. Brancaccio et al. 1999, 89. Tu et al. 1998, 90. Ohtsuka et al. 1998, 91. Parast& Otey 2000, 92. Turner et al. 1990, 93. Androphy et al. 1985, 94. Rearden 1994, 95. Turner et al. 1999, 96. Spencer et al. 1997,97. Pelicci et al. 1992, 98. Baciu et al. 2000, 99. Gao et al. 2000, 100. Grootjans et al. 1997, 101. Halbrugge & Walter 1989, 102.Franke et al. 1978.

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Alpha-actinin is an antiparallelly organized rod-like homodimer, which cross-linksactin filaments and serves as a spacer between actin filaments (Jockusch & Isenberg1981, Blanchard et al. 1989, Meyer & Aebi 1990). It mediates the binding of actin fila-ments to FAs by binding to β1, β2 and β3 integrins (Otey et al. 1990, Burridge & Chrza-nowska-Wodnicka 1996). It also binds to vinculin (Kroemker et al. 1994) and zyxin(Crawford et al. 1992), other important structural components of FAs. Overexpression ofα-actinin in cultured cells causes the formation of more stable cell attachment sites,whereas suppression results in more motile cells (Gluck et al. 1993, Gluck & Ben-Ze'ev1994). Its expression is quickly increased upon accelerated FA assembly (Gluck et al.1992).

Vinculin, an abundant structural protein at FAs, regulates, together with α-actinin, cellmotility and spreading. Fibroblasts overexpressing vinculin cannot move, whereas thecells with suppressed vinculin expression are hypermotile and invasive (RodriguezFernandez et al. 1992, Varnum-Finney & Reichardt 1994, Coll et al. 1995, Eimer et al.1993). Stimulation of cell proliferation induces vinculin synthesis (Ungar et al. 1986).Vinculin consists of a globular head and a rod-like tail (Milam 1985, Molony & Burridge1985), interspersed by a proline-rich hinge (Coutu & Craig 1988, Price et al. 1989). Thehead binds to talin and α-actinin, whereas the proline-rich region binds to VASP and vin-exin, and the rod domain to paxillin, fibrillar actin, lipid bilayer and PIP2 (reviewed byPetit & Thiery 2000). Binding of actin, α-actinin, talin and VASP to vinculin is preventedby intramolecular association between the head and tail of vinculin (Kroemker et al.1994, Johnson & Craig 1994, Johnson & Craig 1995, Huttelmeier et al. 1998). Binding ofPIP2 to vinculin leads to the dissociation of the self-association and unmasking of thebinding sites (Gilmore & Burridge 1996, Weekes et al. 1996).

Talin is an FA-specific homodimer, constructed of two antiparallelly organized270-kDa polypeptides. The dimer is a rod-shaped molecule with globular ends, which areformed of the NH2-termini of the subunits (Rees et al. 1990, Goldmann et al. 1994). TheNH2-terminus binds to the focal adhesion kinase and β integrins, and interacts with phos-pholipids and membranes, whereas the COOH-terminus binds actin, β integrin and vincu-lin (reviewed by Petit & Thiery 2000). In vitro studies reveal that talin can cap andcross-link actin filaments and nucleate the formation of new filaments (Muguruma et al.1990, Muguruma et al. 1992, Kaufmann et al. 1991, Goldmann et al. 1992), and possiblylink actin filaments to plasma membrane (Goldmann et al. 1992, Niggli et al. 1994).Reducing the level of active talin-1 in fibroblasts, HeLa cells or undifferentiated embry-onic stem cells by different techniques results in the inhibition of cell spreading andmigration as well as disassembly of FAs and stress fibers (Nuckolls et al. 1992, Albi-ges-Rizo et al. 1995, Bolton et al. 1997, Priddle et al. 1998). However, a similar effectwas not observed in differentiated stem cells (Priddle et al. 1998). During platelet activa-tion, talin undergoes an increase in its phosphorylation state (Bertagnolli et al. 1993) andredistribution to newly formed adhesion sites (Beckerle et al. 1989).

Tensin is a homodimeric actin filament-capping protein, which has three actin-bindingsites per subunit (Lo et al. 1994a). This gives the protein a propensity to cap or cross-linkactin microfilaments. By capping of barbed ends, tensin regulates actin polymerization(Lo et al. 1994a,b). In addition, tensin has a vinculin-binding domain (Lo et al. 1994b).This enables tensin to anchor the microfilaments to FAs and the SH2 domain (Davis et al.

38

1991), which gives regulatory features to tensin. Recently, participation of tensin in theregulation of cell migration has been described (Chen et al. 2002).

VASP is an F-actin binding protein, which has been shown to localize at nascent FAs(Reinhard et al. 1992). VASP contains two conserved domains (EVH1 and EVH2) in theNH2 the and COOH-terminus, respectively, and a proline-rich region between them(Haffner et al. 1995). The proline cluster region is involved in the binding of profilin(Tanaka & Shibata 1985, Reinhard et al. 1995). Thus, VASP could serve as a mediator ofprofilin’s regulatory function against actin. EVH1 is responsible for binding to vinculinand zyxin, while EVH2 mediates tetramerization of VASP, F-actin binding and actin fila-ment bundling (Bachmann et al. 1999, Calderwood et al. 2000).

Several other novel proteins have been documented to modulate actin filaments.b-nexilin, for instance, is an actin-crosslinking protein, which is located at FAs of ratfibroblasts (Ohtsuka et al. 1998). Parvin/actopaxin modulates cytoskeletal organization atFAs by mediating functions of several proteins, such as ILK, paxillin, vinculin, nck2 andguanidine nucleotide exchange factors (reviewed by Brakebusch & Fässler 2003).

2.2.3.2 Dynamics of focal adhesions

During the dynamic cellular processes, such as cell migration and adhesion, FAs arequickly assembled and disassembled. Their dynamics, assembly in the forward-movingend and disassembly at the dorsal surface, allow cells to migrate. Contact of ECM com-ponents with their transmembrane receptors, integrins, or stimulation of a cell by growthfactors results in integrin clustering (Schmidt et al. 1993, Felsenfeld et al. 1996) and sub-sequent association of the cytoskeletal FA elements, such as focal adhesion kinase, tensin,talin, vinculin, α-actinin and paxillin with the cytoplasmic domains of the integrins. Thesmall GTPase Rho is a major regulator of the process (Hall 1994, Hotchin & Hall 1995,Nobes & Hall 1995, Takai et al. 1995, Ren et al. 1999). Of the other regulatory mole-cules Src kinase, Rac, Ras, MAPK cascade proteins, and cortactin are needed for assem-bly (Miyamoto et al. 1995a, Miyamoto et al. 1995b). Specific tyrosine phosphorylationsare prerequisites for FA assembly (Kornberg et al. 1991, Burridge et al. 1992, Miyamotoet al. 1995a, Miyamoto et al. 1995b, Nobes et al. 1995, Craig & Johnson 1996).

Disassembly of FAs includes weakening of the interactions between FA components,severing of portions of FAs and the associating membranes, dispersion of integrins alongthe plane of the membrane, or moving them to new adhesion sites (Chen et al. 1981,Regen & Horwitz 1992, Palecek et al. 1996). Also extracellular anti-adhesive compo-nents, such as thrombospondin, tenascin and SPARC, are involved in cell migration, andin the formation and disassembly of FAs (Murphy-Ullrich et al. 1991, Sage & Bornstein1991, Murphy-Ullrich et al. 1996, Greenwood & Murphy-Ullrich 1998). Stimulation withepidermal growth factor (EGF), for instance, promotes cell migration via regulated disas-sembly of FAs (Xie et al. 1998).

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2.3 Filamin

2.3.1 Splice variants, domain structure and tissue expression

Filamins are a family of homodimeric, high molecular-weight proteins, which areinvolved in organizing actin filaments into a loose network of non-parallel fibers or tightparallel stress fibers. In addition to actin orchestration, filamins link various transmem-brane proteins to the actin cytoskeleton and serve as scaffolds for many cytoplasmic sig-naling molecules (van der Flier & Sonnenberg 2001).

Filamin shows filamentous staining along actin fibers, hence the name (Wang et al.1975). Three filamin genes have been identified in humans: filamin A (filamin-1,α-filamin, ABP-280, 2647 aa; Gorlin et al. 1990), filamin B (filamin-3, β-filamin,ABP-278/276, 2602 aa; Takafuta et al. 1998, Xu et al. 1998) and filamin C (filamin-2,γ-filamin, ABPL, 2705; Xie et al. 1998). They display a high (~70%) sequence similarity,except for the hinge regions H1 and H1 (see below), which have about 45% similarity.Filamin C has an additional 81-aa insert in the repeat 20. Vertebrates seem to have threefilamin genes. In worms and insects, two long and two shorter filamin proteins have beenfound, probably originating from alternatively spliced transcripts (databank search byBLAST; van der Flier & Sonnenberg 2001).

The NH2-terminal 267-aa region of human filamins represents a typical α-actinin-likeABD (Matsudaira 1991), which contains two calponin homology domains (CH1 andCH2). ABD is followed by a rod-like region, which is formed of 24 approximately 100aa-long repetitive segments (filamin repeats), each consisting of immunoglobulin-likeβ-sheet sandwiches (Tyler et al. 1980, Gorlin et al. 1990). Filamin contains two flexiblehinge regions, H1 and H2, between the repeats 15 and 16, and 23 and 24, respectively(Gorlin et al. 1990, Hock et al. 1990). Filamin forms a V-shaped dimer where the repeats24 of the two polypeptides interact noncovalently with each other (Gorlin et al. 1990).

The expression pattern of filamin is complex due to alternative gene promoters andalternative splicing. Splice variants for all human filamins have been detected (Gorlin etal. 1990, Takafuta et al. 1998, Xie et al. 1998). Variants that lack the repeat 15 of filaminA, H1 of filamins B and C, the 41-aa region between the repeats 19 and 20 of filamins Aand B, and the four COOH-terminal repeats of filamin B have been identified (for domainarchitecture, see below). Alternative splicing affects the properties of filamins. Thus, thesplice variant of filamin B with deletions of H1 and the 41-aa region[filamin-Bvar-1(∆H1)] binds more strongly to integrins and localizes to focal adhesions(van der Flier et al. 2002). Furthermore, filamin-Bvar-1(∆H1), but not the other variants offilamin B, accelerates the differentiation of myoblasts into muscle cells in vitro (van derFlier et al. 2002).

Taking together all variants, filamins A and B are expressed in most human tissues. Atthe level of variants, on the basis of RT-PCR, filamin-Avar-1 and filamin-Bvar-1 are weaklyexpressed in all tissues examined (van der Flier et al. 2002). Regarding the presence ofH1, on the basis of RT-PCR, the expression level of wild type filamin B is the predomi-nant form in the prostate, uterus, lung, liver, thyroid, stomach, lymph node, small intestineand spleen, while filamin-B(∆H1) dominates in the spinal cord and in Daudi lymphoma

40

cells. Both forms are seen in the placenta, bone marrow, brain, umbilical vein endothelialcells, retina and skeletal muscle (Xu et al. 1998). A variant lacking four COOH-terminalrepeats is expressed in cardiac muscle (van der Flier et al. 2002). The expression of fil-amin C, mostly as a filamin-C(∆H1) splice variant, is restricted to skeletal and cardiacmuscle (Xie et al. 1998), in which it is enriched in Z-lines and in myotendinous junctionsof skeletal muscle, Z-lines and intercalated discs of heart muscle. It is also present indense plaques and dense bodies of smooth muscle. In cultured cells, filamin A has beenlocalized to stress fibers, cortical actin network, membrane ruffles and cleavage furrow(van der Flier & Sonnenberg 2001).

2.3.2 Functions of filamin

2.3.2.1 Actin cross-linking and organization

Filamin organizes actin filaments into parallel bundles or different types of network. Inthe networks, filamin is located at the crossroads/intersections of the actin filaments, or atthe membrane contact points (Hartwig & Shevlin 1986). Filaments from different sourcesdiffer in their actin cross-linking activity. For instance, macrophage filamin forms tighternetworks than chicken gizzard filamin (Brotschi et al. 1978). The type of the actin net-work depends on the filamin-to-actin ratio. At high filamin excess, parallel bundles areformed, whereas smaller amounts result in loose, orthogonal networks (Brotschi et al.1978, Niederman et al. 1983, Dabrowska et al. 1985).

In vitro, the type of filamin-based network is also influenced by the presence of otheractin cross-linking proteins. When pure filamin and actin are mixed, an orthogonal net-work is produced, while the presence of α-actinin results in the formation of dense cables(Schollmeyer et al. 1978). Lack of the H1 hinge region, which contributes to a less flexi-ble filamin dimer, seems to favor formation of parallel bundles (van der Flier & Sonnen-berg 2001).

2.3.2.2 Filamin as an anchoring protein for transmembrane receptors

Filamins A and B bind to the cytoplasmic tail of the glycoprotein (GP)-Ibα subunit of theheteropentameric, platelet-specific von Willebrand factor (vWF) receptor (Ezzell et al.1988, Andrews & Fox 1991, Meyer et al. 1997, Takafuta et al. 1998, Xu et al. 1998). Inplatelets, joining of GP-Ibα to the receptor complex triggers rearrangement of the cytosk-eleton, as well as platelet aggregation. The repeats 17–19 mediate the binding (Andrews& Fox 1991). Expression of filamin in filamin-A-deficient melanoma cells (M2 cells),which have been stably transfected with the vWF receptor, leads to an increase of thelevel of vWF receptor on the plasma membrane (Meyer et al. 1998).

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Via its four and a half COOH-terminal repeats, filamin interacts with the cytoplasmictails of integrins β1A (Loo et al. 1998, Pfaff et al. 1998, Zent et al. 2000), β2 (Sharma etal. 1995), β3 and β7 (Liu et al. 2000). Filamin-Bvar-1, the variant with a 41-aa deletionbetween the repeats 19 and 20, also binds to integrin β1D and shows increased bindingaffinity to integrin β1A (van der Flier et al. 2002). Similarly to the case of GB-Ibα, theexpression of filamin in M2 cells elevates the level of integrin β1 in the plasma mem-brane (Meyer et al. 1998), suggesting that filamin has a role in the retainment of trans-membrane receptors (van der Flier et al. 2002).

Filamin also interacts with the cytoplasmic tails of other transmembrane proteins suchas γ- and δ-sarcoglycans (Thompson et al. 2000), presenilins (Zhang et al. 1998, Guo etal. 2000), furin (Liu et al. 1997), FcγRI (Ohta et al. 1991), tissue factor (Ott et al. 1998),and probably the acetyl choline receptor as well (Shadiack & Nitkin, 1991). Furthermore,filamin binds and regulates the function of dopamine D2 receptor (Li et al. 2000) and theandrogen receptor (Ozanne et al. 2000).

2.3.2.3 Filamin as a scaffolding protein for signaling molecules

Filamin has been suggested to serve as docking site for signaling molecules and, thus, todirect them to proper localization and orientation for actin filament nucleation, actindynamics and vesicle transport (van der Flier & Sonnenberg 2001).

Several members of the Ras superfamily of GTPases bind to the COOH-terminalrepeats of filamin (Ueda et al. 1992). Among them, the binding of RalA is GTP-depen-dent, while that of the Rho-like GTPases Cdc42 and Rac1 is not (Ohta et al. 1999). AlsoTrio, a guanidine nucleotide exchange factor for RhoG, Rac and RhoA, binds, via itspleckstrin homology domain, to filamin (Bellanger et al. 2000). Transfection and micro-injection experiments with Swiss 3T3 fibroblasts and filamin A-deficient human mela-noma M2 cells show that the formation of filopodia, induced by RalA and Cdc42, unlikethat induced by RhoA and Rac1, is dependent on filamin (Ohta et al. 1999).

Filamin A also binds SEK-1 (MKK-4, JNKK), a kinase that activates severalstress-activated protein kinases (SAPKs) (Marti et al. 1997). In filamin A-deficient M2cells, TNFα- and lysophosphatidic acid (LPA)-induced activation of SAPKs is reduced,suggesting that appropriate TNFα- and LPA-mediated SAPK activation is dependent onthe presence of filamin. Expression of dimerization-deficient mutant filamin fails to nor-malize LPA-activation of SAPK, whereas it corrects the defective SAPK response toTNFα. Moreover, interaction of filamin-A with tumor necrosis receptor-associatedfactor-2 is essential for TNFα-mediated activation of SAPKs and of the transcription fac-tor NF-κB (Marti et al. 1997, Leonardi et al. 2000). Filamin’s role in TNF-receptor sig-naling has been supported by studies in Drosophila. In Drosophila, filamin binds to Toll,a TNF receptor superfamily member, which regulates the development of dorsal-ventralpolarity of the fruit fly, and to Tube, a signaling protein downstream of Toll (Edwards etal. 1997).

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2.3.2.4 Regulational aspects of filamin function

The regulation of filamin function is poorly known. However, some regulatory mecha-nisms have been suggested (Fig. 4). One mechanism could be the occupancy of the trans-membrane receptors that are/become coupled to filamin. Thus, for instance, binding ofimmunoglobulin to FcγRI receptor causes the dissociation of filamin from FcγRI and,thus, release of FcγRI receptor from the cortical actin cytoskeleton (Ohta et al. 1991).FcγRI is a receptor in the plasma membrane of hematopoietic cells, in which it binds tothe Fc domain of the antigen-bound IgG (Strzelecka et al. 1997). In contrast, in the caseof tissue factor, the cellular receptor for Factor VII, occupancy of the receptor is neededfor filamin-A binding (Ott et al. 1998). Binding of Factor VII to tissue factor upon tissueinjury triggers the blood coagulation process (McVey 1999).

In cells with high activity of Ras-related GTPases, filamin is found to be stronglyphosphorylated (Yada et al. 1990, Ueda et al. 1992). Phosphorylation, on the other hand,has an influence on its actin binding and actin cross-linking activities (Zhuang et al. 1984,Ohta & Hartwig 1995). At least EGF, platelet-derived growth factor and LPA induceserine/threonine phosphorylation of filamin (van der Flier & Sonnenberg 2001). Theknown kinases responsible for filamin phosphorylation are ribosomal S6 protein kinase-2(Ohta & Hartwig 1996), PKA, PKC and CaM-kinase II (Wallach et al. 1978, Kawamoto& Hidaka 1984, Chen & Stracher 1989, Ohta & Hartwig 1995).

The hinge regions H1 and H2 contain calpain cleavage sites. Cleavage in H2 results inthe formation of filamin that is unable to dimerize. Thus, it lacks the ability to cross-linkactin filaments (Davies et al. 1978, Gorlin et al. 1990). Cleavage by calpain is probablyregulated by phosphorylation of filamin (Chen & Stracher 1989, Gorlin et al. 1990, Wu etal. 1994, Jay et al. 2000). Upon platelet activation, GP-Ibα is released from filamin aftercleavage of filamin by calcium-dependent protease (Fox 1985). In addition, cleavage offilamin by m-calpain and granzyme is associated with myotube differentiation process(Kwak et al. 1993a, b) and caspase-independent apoptosis (Browne et al. 2000), respec-tively.

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Fig. 4. Suggestions for the regulation of the function of filamin. A. Binding of a ligand to thefilamin-bound receptor causes the dissociation of filamin from the receptor. B. Phosphorylationof filamin enhances the capacity of filamin to bind and cross-link F-actin. C. Cleavage of therepeat 24 by calpain protease cuts the dimerization domain loose, which causes the loss of theactin cross-linking capacity of filamin.

2.4 Endocytosis and the cytoskeleton

Endocytosis is a complex process that includes the endocytotic machinery itself as well asits regulatory system. Currently, the regulatory mechanisms are largely unrevealed. Dur-ing the past few years, a linkage between endocytosis and the actin cytoskeleton has beenestablished. Disturbance of normal actin cytoskeleton by actin-perturbing drugs or mutant

44

Rho-family proteins disrupts endocytosis in some, but not in all assays (Gottlieb et al.1993, Jackman et al. 1994, Lamaze et al. 1997, Fujimoto et al. 2000, da Costa et al.2003). Recent studies have revealed several proteins that are associated both withendocytosis and actin dynamics (Qualmann & Kessels 2002).

2.4.1 Actin and endocytosis

Currently, the regulatory role of actin cytoskeleton cannot be conclusively assigned toany specific stage of endocytosis. Also the molecular mechanisms are largely unknown.Building upon variable models the following modes of involvement of the cytoskeletonin the specific steps of endocytosis have been presented (Qualmann et al. 2000):

First, the network of actin filaments could localize the endocytotic machinery to spe-cific compartments by setting physical barriers or by directly associating with endocytoticproteins. This is supported by the observations that coated pit formation in a defined loca-tion was altered and led to diffusion of clathrin-coated pits from the place of their originupon disrupting the actin architecture by monomer-sequestering drugs (Gaidarov et al.1999).

Another possibility is that actin filaments facilitate invagination of the planar mem-brane, and, in this way, make it more available for endocytotic machinery to attach.

In a third model, actin filaments play a role in the membrane scission event uponwhich the newly formed vesicle is liberated from the plasma membrane. In experimentsutilizing multicolor real-time fluorescence microscopy, rapid actin polymerization hasbeen seen to take place at the sites of vesicle pinching (Kaksonen et al. 2003). On theother hand, inhibition of actin polymerization arrests the endocytotic process to the stageof invaginated coated pit (Lamaze et al. 1997).

Finally, actin polymerization might serve as a motor, which moves the newly formedvesicle through the cytoplasm. This theory is supported by the observation that differentvesicles are associated with actin comet tails (Frischknecht et al. 1999, Merrifield et al.1999, Rozelle et al. 2000, Taunton et al. 2000).

As opposed to these views of an intact actin network as a facilitator of endocytosis,there is also evidence that actin could suppress it. Some assays suggest that the corticalactin cytoskeleton has to be removed prior to initialization of membrane trafficking. Arigid cortical cytoskeleton has been observed to inhibit endocytosis (Trifaró & Vitale1993). Electron microscopy studies of Cos-7 cells have shown that the immediate vicinityof the coated pits is almost free of fibrous actin (Fujimoto et al. 2000).

2.4.2 Proteins acting at the interface between endocytosis and the actin cytoskeleton

Dynamin is a GTPase that regulates the fission stage of endocytosis (for a review, seeSever et al. 2000). Some studies suggest dynamin to be coupled to actin dynamics. For

45

instance, overexpression of the mutant dynamin, which is unable to bind GTP, causes notonly inhibition of endocytosis, but also a redistribution of actin stress fibers and alter-ation of cell shape (Damke et al. 1994). On the other hand, downregulation of dynamin Iin hippocampal cultures led to inhibition of the actin-associated, neurite outgrowth (Torreet al. 1994). Low concentrations of overexpressed dynamin 2 were shown to enhanceactin nucleation, while higher concentrations had an inhibitory effect. Recently, expres-sion of mutant Vps1p, the yeast dynamin-related protein, has been shown to result in bothactin abnormalities and impaired vesicular protein sorting in Saccharomyces cerevisiae(Yu et al. 2004). Vps1p binds to the actin-regulating protein Sla1p (Yu et al. 2004). More-over, dynamin has been found to regulate the dynamics of actin-based comet tails, whichprovide mechanical force for cellular organelles and Listeria monocytogenes to moveinside the cells (Lee & Camilli 2002, Orth et al. 2002).

PACSINs represent novel linkers between endocytosis and actin cytoskeleton. Theybind vesicle-trafficking proteins dynamin I, synapsin I and synaptojanin, as well asN-WASP, a nucleator of actin polymerization (Qualmann & Kelly 2000, Modregger et al.2000). Exogenous expression of the SH3 domain of rat PACSINs in cultured cells leads toa block in receptor-mediated endocytosis. It affects the dynamin-regulated step, in whichthe vesicles are pinched off, and leads to accumulation of clathrin coated pits and vesi-cles at the apical plasma membrane of the cells (Simpson et al. 1999, da Costa et al.2003). Co-overexpression of the VCA domain of N-WASP, the crucial region for the initi-ation of actin polymerization, cancels the disruption. On the other hand, overexpressionof PACSINs induces Arp2/3 complex activation, probably via interaction with N-WASP,resulting in the reorganization of actin cytoskeleton and the formation of filopodia (Qual-mann & Kelly 2000, Kessels & Qualmann 2002). PACSIN 1 also binds to mSos, a gua-nine nucleotide exchange factor for the cytoskeletal regulators Ras and Rac, suggestinganother link between the cytoskeleton and endocytosis (Wasiak et al. 2001). Overex-pressed rat PACSIN locates at the sites of high actin turnover, such as lamellipodia andfilopodia. Chicken FAP52 has been localized to focal adhesions, which also represent thesites of dynamic actin at the plasma membrane (Meriläinen et al. 1997). Thus, FPS pro-teins have been suggested to couple actin polymerization to dynamin function in the fis-sion reaction (Qualmann et al. 1999). The putative mechanisms by which FPS family pro-teins affect endocytosis are presented in Fig. 5.

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Fig. 5. FPS proteins and endocytosis. The mechanism of the FPS proteins in the regulation ofendocytosis may be mediated by its interaction partners, such as N-WASP, dynamin I, synap-tojanin, or huntingtin and its interactor Hip1.

Mammalian Abp1 binds to F-actin by its NH2-terminal binding domains and colocal-izes with highly dynamic actin in lamellipodia (Kessels et al. 2000, 2001). Its involve-ment in actin dynamics is dependent on the GTPase Rac1. In yeast, Abp1 binds both tothe Arp2/3 complex and F-actin (Goode et al. 2001). It has been suggested to recruitArp2/3 complex to sides of actin filaments. Recently, Abp1 has been described to interact,via its COOH-terminal SH3 domain, with dynamin (Kessels et al. 2001). Its overexpres-sion leads to reduction of receptor-mediated endocytosis (Kessels et al. 2001). Thus,Abp1 might serve as a physical link between endocytosis and the actin cytoskeleton.

Similarly to Abp1, the Src kinase substrate cortactin (Wu et al. 1991) binds to F-actin(Wu & Parsons 1993), Arp2/3 complex (Uruno et al. 2001), and dynamin (McNiven et al.2000). Cortactin and dynamin colocalize with F-actin in lamellipodia (Kaksonen et al.2000, McNiven et al. 2000), and promote actin polymerization activity of Arp2/3 and sta-bilize the actin filament network (Weaver et al. 2001)l. Cortactin has been suggested toregulate actin assembly, which propels the movement of endosomes through the cyto-plasm (Kaksonen et al. 2000).

WASP family members enhance the ability of the Arp2/3 complex to nucleate actinpolymerization by binding via its COOH-terminus to Arp2/3 (Machesky & Insall 1998,

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Welch 1999). Platelets and lymphocytes of Wiskott-Aldrich syndrome patients show adecreased number of microspikes and abnormal actin organization (Snapper & Rosen1999). In addition to regulation of actin organization, WASPs have been shown to beinvolved in endocytosis. Upon EGF treatment of Cos-7 cells, neuronal N-WASP forms acomplex with the EGF receptor, probably via the adaptor protein Grb2 (Miki et al. 1996).WASP-knockout lymphocytes exhibited, in addition to impaired actin polymerization,reduction in T cell receptor endocytosis (Zhang et al. 1999). Moreover, N-WASP binds toPACSINs, which are also implicated in both actin dynamics and endocytosis (Simpson etal. 1999, Qualmann & Kelly 2000). Recently, endosomal and lysosomal vesicles havebeen observed to move in cell-free extracts via actin propulsion, an event that exhibitedN-WASP staining in the interface of the vesicle and the actin tail (Taunton et al. 2000).

Synaptojanin is a phosphoinositide phosphatase (McPherson et al. 1996, Woscholskiet al. 1997), which is also implicated in both endocytosis and actin dynamics. Synaptoja-nin binds several SH3 domain-containing proteins, such as PACSINs (Qualmann et al.1999, Qualmann & Kelly 2000), which are involved in endocytosis. It plays a role inuncoating of coated vesicles (Cremona et al. 1999). In vitro, synaptojanin is able tohydrolyze PIP2, which is bound to profilin, cofilin and α-actinin. Overexpression of syn-aptojanin causes rearrangement of actin stress fibers (Sakisaka et al. 1997). This is well inline with the observation that a decrease in the level of PIP2 causes actin depolymeriza-tion.

Sla2 of budding yeast is a component of the cytoskeleton and participates in the inter-nalization step of endocytosis (Wesp et al. 1997). In cells lacking Sla2, non-motileendocytotic vesicles with actin-comet tails are seen (Kaksonen et al. 2003), suggesting itsregulatory role in endocytosis-associated actin events. Similarly to Sla2, lowering thelevel of its mammalian homolog HIP1R by RNAi causes the stable association betweenthe actin assembly and the endocytotic machineries (Engqvist-Goldstein et al. 2004). Viadirect binding to clathrin and α-adaptin, HIP1R closely associates with clathrin andclathrin-coated pits and vesicles (Engqvist-Goldstein et al. 1999, Gaidarov et al. 1999,Waelter et al. 2001). The talin-like COOH-terminus of HIP1R binds F-actin. The bindingsite is necessary for its colocalization with cortical actin (Engqvist-Goldstein et al. 1999).HIP1R promotes clathrin cage assembly and links it to F-actin in vitro(Engqvist-Goldstein et al. 2001). The dual function of HIP1R may be needed for spatialorganization of endocytosis or actin-dependent movement of newly formed vesicles(Engqvist-Goldstein et al. 1999).

3 The aims of the present study

A novel SH3 domain-containing protein, FAP52, was originally characterized by Meri-läinen et al. (1997). In cultured fibroblasts, FAP52 is localized in focal adhesions. InNorthern blotting, its transcript has been detected in most tissues. On the basis of compu-tational analyses of its primary structure, the NH2-terminus of FAP52 is highly α-helical,and it has a tendency to fold into α-helical coiled-coil and form higher-order oligomericcomplexes. The specific aims of the present study were:1. to further characterize FAP52 in structural terms,2. to describe novel binding partners of FAP52,3. to determine the function of FAP52,4. and to determine the distribution of FAP52 in different tissues.

4 Materials and methods

The detailed methods are described in the original articles I–IV.

4.1 General procedures and computer programs (I–IV)

The standard solutions, buffers and procedures for the purification and precipitation ofDNA, restriction enzyme digestions, ligation reactions, SDS-PAGE runs, immunohis-tochemical and histochemical reactions, and stainings were as described in Sambrook etal. (1989). DNA sequencing was carried out on an automated ABI Prism 377XL DNASequencer (PerkinElmer, Branchburg, NJ). For sequence alignment, the program ClustalX, version 1.8 (Thompson et al. 1997), and for sequence comparisons, the programBLAST at the network server http://www.ncbi.nlm.nih.gov/BLAST/ were utilized. Forthe prediction of coiled-coil arrangements, the programs PairCoil (Berger et al. 1995) andMultiCoil (Wolf et al. 1997) were employed at the network servers http://night-gale.lcs.mit.edu/cgi-bin/score and http://nightingale.lcs.mit.edu/cgi-bin/multicoil, respec-tively.

4.2 Materials (I–IV)

The plasmid DNA purification kits were from Qiagen (Hilden, Germany) and Promega(Madison, WI), the) and the DNA gel extraction kit from Qiagen. E. coli strainBL21(DE3) cells, expression vectors pGEX-2T, pGEX-2TK and pGEX-4T-1,glutathione-Sepharose 4B, thrombin, Rainbow High Molecular Weight Markers, andsynthetic oligonucleotides were purchased from Amersham Pharmacia Biotech (Uppsala,Sweden), isopropyl-β-D-thiogalactopyranoside (IPTG) from MBI Fermentas (Vilnius,Lithuania), Site-Directed Mutagenesis kit, pfu polymerase and T4 DNA ligase fromStratagene (La Jolla, CA), restriction enzymes from MBI Fermentas and Promega, andchicken gizzard filamin from Progen Biotechnik (Heidelberg, Germany). For blot overlay

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assay, chicken gizzard filamin was purified according to the method described byFeramisco and Burridge (1980). Fugene 6-transfection reagent, reduced glutathione,protease inhibitors aprotinin, leupeptin and phenylmethylsulphonyl fluoride wereobtained from Boehringer Mannheim (Mannheim, Germany), and ampicillin, bovineserum albumin (BSA) and benzamidine, disuccinimidyl suberate, Nondenatured ProteinMolecular Weight Marker kit, urease, and chicken egg albumin from Sigma Chemicals(St. Louis, MO). Triton X-100 was from Merck (Darmstadt, Germany), γ-globulin fromCalbiochem-Novabiochem (Darmstadt, Germany), and nitrocellulose membranes fromSchleicher & Schuell (Keene, NH). HeLa cells were purchased from the American TypeCulture Collection (Rockville, MD), Immu-Mount mounting medium from Shandon(Pittsburgh, PA), and cell culture media and fetal calf serum from Hyclone Laboratories(Logan, UT).

4.3 Antibodies (I, II, IV)

Rabbit polyclonal antibody to bacterially produced FAP52, denoted Affi-K7, was pro-duced and affinity-purified as described (Meriläinen et al. 1997). Rabbit polyclonal anti-hemagglutinin (HA) antibody (anti-HA) was from Santa Cruz Biotechnology (SantaCruz, CA), mouse monoclonal anti-GST antibody from Biacore (Uppsala, Sweden),mouse monoclonal anti-filamin antibody from Chemicon International (Temecula, CA),and mouse monoclonal anti-paxillin antibody from Zymed Laboratories (San Francisco,CA). Horseradish peroxidase (HRP)-conjugated goat anti-rabbit and anti-mouse immuno-globulins, and goat anti-rabbit and anti-mouse IgG-agarose were purchased from SigmaChemicals, and rhodamine-phalloidin, Alexa Fluor 488 goat anti-rabbit IgG and AlexaFluor 546 goat anti-mouse IgG from Molecular Probes (Eugene, OR). For immunoelec-tron microscopy, Protein A-gold particles (∅ 5 nm and 10 nm), prepared by Slot andGeuze (1985), were used.

4.4 cDNA constructs (I–III)

All the cDNA constructs and corresponding polypeptides were designated as “FAP52” or“Fil” for chicken FAP52 and human filamin A, respectively, followed by subscriptednumbers indicating the first and the last aa residues represented.

To express the recombinant full-length FAP52 and its fragments, the correspondingcDNAs were generated by polymerase chain reaction (PCR) utilizing the oligonucleotideprimers with BamHI and EcoRI restriction sites. The resulting cDNAs were inserted intothe corresponding restriction sites of pGEX-2T or pGEX-2TK vectors. COOH-terminalfragments of filamin were expressed by pGEX-4T-1 vectors with the cDNAs encoding forthe fragments Fil1524–2283, Fil2283–2490 and Fil2495–2647 (a kind gift from Dr. T.P. Stossel,Division of Hematology, Brigham and Women’s Hospital, Boston, USA; Ohta et al.1999). Further COOH-terminal truncation mutants were generated from the construct

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encoding for the fragment Fil1524–2283 by using Site-Directed Mutagenesis kit and oligo-nucleotide primers to generate stop codons at appropriate sites of the cDNA.

To express FAP52 or its fragments in cultured eukaryotic cells, the correspondingcDNA-constructs were produced by PCR with engineered restriction sites. They were cutwith EcoRI and EcoRV and subcloned into EcoRI/EcoRV cloning site of pRK5-vector (akind gift from Dr. Joseph Schlessinger, New York University, Medical Center, New York,NY). An HA-tag was engineered to the COOH-termini of the constructs by primerdesign. The following constructs were generated: FAP52-HA (FAP521–448 plus HA-tag);FAP52Nt-HA (FAP521–293 plus HA-tag); FAP52SH3-HA (FAP52390–448 plus HA-tag).

4.5 Expression and purification of the GST fusion proteins (I–III)

GST fusion proteins were expressed in E. coli BL21(DE3) cells according to standardprotocols, purified from the cell lysates on glutathione-Sepharose 4B beads, and liberat-ed from the beads by incubation with reduced glutathione. For some experiments, FAP52or its mutant forms were released from their fusion partners by incubation with thrombin.For crystallization, FAP52 released from GST was further purified by using a Mono Qanion-exchange chromatography column (Amersham-Pharmacia Biotech) in 20 mM Bis-Tris, pH 6.0, connected to an ÄKTA fast protein liquid chromatography (FPLC) system(Amersham Pharmacia Biotech), and eluted with increasing NaCl concentration. For thecontrol experiments, GST was expressed from the plasmid pGEX-2TK and purified asabove.

4.6 Cell cultures, transfections and immunofluorescence microscopy (I, II, IV)

Chicken embryo heart fibroblasts (CEHF) were prepared and grown as described previ-ously (Meriläinen et al. 1997). HeLa-cells were grown according to the same protocolwith the exception that Dulbecco’s modified Eagle’s medium was used as a growth medi-um.

Transfections were carried out with the cells grown on glass coverslips and by using aFugene 6-transfection reagent and following the manufacturer’s instructions.

For immunofluorescence microscopy, the cells were fixed, stained and viewed asdescribed in the original articles I and II. Briefly, the cells grown on coverslips werewashed with Hank’s salt solution, fixed with paraformaldehyde solution, and post-fixedwith methanol or ethanol. After immersing fetal calf serum, the cells were incubated withthe primary antibodies, washed, incubated with appropriate fluorochrome-conjugated sec-ondary antibodies, washed again, and mounted on an object glass. Viewing was under aZeiss LSM510 laser scanning microscope equipped with a Zeiss Axiovert 110M invertedmicroscope (Carl Zeiss Microscopy, Oberkochen, Germany). The images were processedby using LSM 3D software, version 5.2. (Carl Zeiss Microscopy).

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4.7 Immunoelectron microscopy (I, IV)

Immunoelectron microscopy was carried out as described in the original article I. Briefly,the cells were washed with Hank’s salt solution, fixed with paraformaldehyde-sucrosesolution, harvested by scraping, and pelleted. The pellet was then resuspended in asucrose solution and frozen in liquid nitrogen. Cryosections were cut on a microtome andplaced on Butvar-coated nickel grids. They were then washed and blocked with 5% BSAwith 0.1% ColdWaterFishSkin gelatin.

In the double labeling experiments, the sections were first incubated with monoclonalantibodies targeted to a protein of interest, followed by incubation with rabbit anti-mouseIgG, and then with Protein A-gold (∅ 10 nm). After this, the sections were fixed in glut-araldehyde solution and blocked with BSA. They were then incubated with Affi-K7 fol-lowed by Protein A-gold (∅ 5 nm). The sections were then post-fixed in glutaraldehyde,washed, counterstained with uranyl acetate, and coated with 1.8% methyl cellulose with2% uranyl acetate. The sections were viewed using a Philips 410 LS transmission electronmicroscope (Philips, Eindhoven, Netherlands).

4.8 Cell lysates and immunoprecipitation (I, IV)

The cells grown to confluency were harvested by a scraper, suspended in the lysis bufferand incubated on ice bath for 20 min. The lysates were clarified by centrifugation at12,000 x g.

The whole cell (unfractionated) lysates were incubated with the primary antibodies(Affi-K7 or mouse anti-filamin antibodies), followed by incubation with agarose-coupledanti-rabbit or anti-mouse IgG antibodies, respectively. The control reactions were carriedout with the same protocol, but in the absence of the primary antibodies. The precipitateswere washed and subjected to SDS-PAGE, followed by immunoblotting.

4.9 Affinity chromatography and pull-down experiments (I)

Affinity chromatography was carried out with CEHF lysates, which were incubated withthe recombinant GST-FAP52 or GST alone, followed by incubation with the glutathione-Sepharose 4B beads. The beads were then washed, and the proteins were liberated fromthe beads by adding the SDS-PAGE loading buffer and boiling. The proteins were thensubjected to SDS-PAGE and coomassie brilliant blue staining.

For pull-down experiments, GST fusions constructs of the fragments of FAP52 or fil-amin were immobilized on glutathione-Sepharose 4B beads and incubated with chickengizzard filamin or recombinant FAP52, respectively. For the controls, GST was immobi-lized on the beads instead of the FAP52 or filamin fusion proteins. The proteins werereleased from the beads by adding the SDS-PAGE loading buffer and boiling. They werethen subjected to SDS-PAGE and immunoblotting.

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4.10 Immunoblotting and blot overlay assay (I, II, IV)

For immunoblotting, the electrophoretically separated proteins were transferred to nitro-celllose membranes, which were then blocked in nonfat dried milk powder in tris-buff-ered saline. Thereafter, the filters were incubated with Affi-K7 or mouse anti-filamin.After a 3-hour or overnight washing, the filters were incubated with either HRP-conjugated goat anti-rabbit or anti-mouse IgG, respectively. After washing, the blot wasdeveloped by the enhanced chemiluminescence (ECL) detection system.

Blot overlay assays were carried out with bacterially produced FAP52 and GST, orwith GST-fusions of filamin mutants. They were separated on SDS-PAGE and trans-ferred to nitrocellulose membranes. The filters were incubated with nonfat dried milkpowder in tris-buffered saline, and then overlaid with the purified filamin or recombinantFAP52, respectively. For the visualization of the bound proteins, the overlays werewashed and then incubated with either the mouse anti-filamin antibodies or Affi-K7,respectively, as above, followed by HRP-conjugated goat anti-mouse or anti-rabbit IgG,respectively, and ECL method.

4.11 Electrospray ionization mass spectrometry (I)

For mass-spectrometry, pieces of polyacrylamide gels corresponding to the protein bandsof interest were excised from the coomassie brilliant blue-stained gels and processed asdescribed by Shevchenko et al. (1996). Briefly, the proteins were in-gel reduced, alkylat-ed, and digested with trypsin. The supernatant obtained was acidified with formic acid,loaded onto a Poros R2 microcolumn (Perseptive Biosystems, Fromingham, MA), anddesalted according to Gobom et al. (1999). The peptides eluted with methanol/formicacid solution were introduced into a nanoelectrospray needle (Protona, Odense, Den-mark). Nanoelectrospray tandem mass spectrometry was performed on an API III massspectrometer (PerkinElmer Instruments, Shelton, CT) equipped with a nanoelectrospraysource, as described elsewhere (Wilm et al. 1996).

4.12 Surface plasmon resonance analysis (I, II)

Surface plasmon resonance (SPR) measurements were performed on a BIACORE 3000analyzer under control of BIACORE control software, version 3.1.1. (Biacore). A car-boxymethyl-coated CM5 sensor chip was prepared by immobilization of anti-GST anti-body on the surface of the chip by utilizing the GST Capture and the Amine Couplingkits (Biacore). As ligands, GST-FAP52 or GST-Fil1524–2283, or mutants thereof were cou-pled to the immobilized anti-GST. In order to keep GST-FAP52 a monomer, mildly acid-ic buffer (Bis-Tris, pH 6.0), a condition in which FAP52 exists mainly as monomer (datanot shown), was utilized during the coupling procedure. For the binding assays, chickengizzard filamin or the recombinant FAP52 were then passed over the chip at varying con-

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centrations. The binding of the analyte to the ligand was detected from the change in thesensorgram. In order to correct the sensorgram for background binding and bulk refrac-tive index changes, a different flow cell without the GST-fusion protein was used as a ref-erence. As controls, GST-coated chips were used, over which filamin or FAP52 waspassed. Kinetics was analyzed by BIAevaluation software, version 3.1 (Biacore).

4.13 Chemical cross-linking (II)

For chemical cross-linking, the bacterially produced protein or the CEHF lysate clarifiedby centrifugation was incubated in the presence or absence of disuccinimidyl suberate.The reaction was terminated by adding SDS-PAGE loading buffer into the reaction mix-ture. The samples were run on SDS-PAGE and visualized by coomassie brilliant bluestaining or immunoblotting.

4.14 Gel filtration chromatography (I, II)

Bacterially produced FAP52 or CEHF lysate, clarified by centrifugation, was applied on aSuperdex 200 gel filtration column (Amersham Pharmacia Biotech) connected to anÄKTA FPLC system. The column was pre-equilibrated with PBS and calibrated with themolecular weight markers urease (hexamer 545 kDa, trimer 272 kDa), γ-globulin (150kDa), BSA (dimer 132 kDa, monomer 66 kDa) and chicken egg albumin (45 kDa). Thesample was eluted with PBS and the run was monitored by absorbance at 280 nm. Thepeak fractions were collected and analyzed by SDS-PAGE and immunoblotting.

4.15 Far UV circular dichroism (II)

Determination of the circular dichroism (CD)-spectra was carried out by using Jasco J-715 spectropolarimeter (Jasco Research, Brentwood Bay, B.C., Canada) under control ofJ-700 Hardware Manager, version 1.50.00 (Jasco Research). For the experiments, the pro-tein was dialyzed against 50 mM potassium phosphate buffer, pH 7.4. The CD valuescans were measured from 190 to 260 nm, at a step resolution of 2 nm, and with a scan-ning rate of 20 nm/min. The CD spectra were the average of 10 scans. The data were pro-cessed with Standard Analysis Program, version 1.50.01 (Jasco Research). The content ofα-helicity was calculated by Protein Secondary Structure Program, Model SSE338 (Jas-co Research; Yang et al. 1986).

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4.16 Crystallization of FAP52 (III)

For crystallization, bacterially produced GST-FAP52 was purified by affinity and anion-exchange chromatography as described above. Purified FAP52 was then subjected tobuffer change and desalting by performing three cycles of concentration-dilution in 20mM Tris, pH 8.0 and by using Centriprep concentrating column (Millipore Corp, Bed-ford, MA). Finally, FAP52 was adjusted to 13.5 mg/ml.

The crystals were grown by the hanging drop vapor diffusion technique (HamptonResearch, Laguna Niguel, CA). An ammonium sulphate grid screen was utilized to findproper crystallization conditions. The drops were prepared by mixing 1 µl of the proteinwith 1 µl of the reservoir solution over the 1-ml volume of the reservoir. After one-day ofequilibration, small stick-like crystals quickly appeared. Subsequently, polyethylene gly-col PEG2kMME in concentrations ranging from 10 to 30% was combined with ammo-nium sulphate ranging from 100 to 400 mM. The pH was buffered to 8.0 by 100 mMHEPES. One condition, 20–22% PEG2kMME, 350 mM ammonium sulphate, 100 mMHEPES, pH 8.0, yielded crystals of two different morphologies. Form I displayed a shapeof thick needles or columns, and the form II appeared as prism/plate-like crystals.

5 Results

5.1 Interaction of FAP52 with filamin (I)

To identify new binding partners for FAP52, coimmunoprecipitation and pull-down(affinity binding) experiments were carried out by using the lysates of cultured CEHFs asa target. For co-immunoprecipitation experiments, the lysates were incubated with theAffi-K7 antibodies, followed by agarose-bound anti-rabbit IgG antibodies. The beadsalong with their lysate-derived bound materials were then collected by centrifugation, andsubjected to SDS-PAGE and silver staining. For pull-down experiments, GST-FAP52 wascoupled to the reduced glutathione-conjugated sepharose beads in a column, and the celllysates were passed over the column. The attached proteins were then eluted from thebeads and subjected to SDS-PAGE and coomassie brilliant blue staining. The immuno-precipitation and the pull-down experiments both revealed a 280-kDa polypeptide band,which was present in the experimental but not in the control assay [Fig. 1 in the originalarticle I (Fig. 1/I)]. In pull-down experiments, another prominent band of about 220 kDawas seen.

The 280-kDa band from the pull-down experiments was excised from the gel and sub-jected to in-gel trypsin digestion. The extracted peptides then underwent analysis bynanoelectrospray tandem mass spectrometry. The partial primary structures of severalpeptides in the mass spectrum of the 280-kDa band were compared with the sequences inthe databases. In five cases there was a match with GST, in four cases with FAP52, in onecase with myosin, and in one case with filamin. We concluded that the myosin sequenceamong the peptides derived from the 280-kDa band originated from the more abundantmaterial of the 220-kDa band corresponding to the MW of myocin heavy chain.

5.1.1 Association between FAP52 and filamin in cells in vivo

The association of FAP52 with filamin was further explored by immunoprecipitation ofFAP52 from CEHF-lysates, followed by immunoblotting for coprecipitation of filamin.

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Reciprocal experiments were carried out by immunoprecipitation of filamin fromFAP52+HA –transfected HeLa cells and immunoblotting to probe for the coprecipitationof FAP52+HA. Filamin was present, as judged by immunoblotting, in the Affi-K7 precip-itates. Reciprocally, FAP52+HA was found in the precipitates obtained by immunoprecip-itation with anti-filamin. Thus, FAP52 associates with filamin in vivo. These experimentsdid not resolve whether the interaction is a direct one or mediated by some other compo-nent (Fig. 2/I).

5.1.2 Direct binding of FAP52 to filamin

In order to explore whether the association between FAP52 and filamin is due to directinteraction, blot overlay assays were carried out. Bacterially produced FAP52 or theCOOH-terminal fragments of filamin were subjected to SDS-PAGE and transferred to anitrocellulose filter, which was then incubated with chicken gizzard filamin or recombi-nant FAP52, respectively. Binding of filamin to FAP52 was assayed by using anti-filaminantibody or Affi-K7, respectively, followed by the ECL method. Indeed, filamin was seento directly bind FAP52 (Fig. 3a,b/I).

Surface plasmon resonance is a technique that permits determination of the dynamicsof the interactions between various biomolecules. In the experiments, one interactor isanchored to the solid phase, the sensor chip. The other interactor is then passed over itsimmobilized binding partner. The dynamic association and dissociation can be detected aschanges in the response in the sensorgram. The dissociation and association rates aredetermined from the response profile, and the dissociation constant (KD) can then bederived from their binding equations.

We utilized SPR to further demonstrate the interaction between FAP52 and filamin.Bacterially produced GST-FAP52 was immobilized to the sensor chip, and chicken giz-zard filamin was passed over the chip. In the sensorgram, a distinct change was seen uponfilamin injection. On the other hand, a return to base level was seen upon washing thechip after the injection of filamin (Fig. 3c/I). This indicates a direct and reversible bindingof FAP52 to filamin. The KD was determined to be 9.64 x 10-9 M, suggesting a high-affinity binding between FAP52 and filamin.

5.1.3 Filamin-binding site in FAP52

Next we wanted to map the filamin-binding site in FAP52. For that purpose, the sequen-tially truncated NH2- and COOH-terminal mutant forms of FAP52 in GST fusion wereutilized in SPR and blot overlay experiments and assayed for the binding of filamin. Bothmethods showed a binding of filamin to FAP52 polypeptides corresponding to residues1–184, 1–219, 1–359 and 1–448, but not to 1–145, 146–448, 185–448, 220–448, 281–448, 360–448 and 390–448 (Fig. 4/I). These results point to the region 146–184 of FAP52as a filamin-binding site. However, no binding was seen for the isolated polypeptide cor-responding to the aa 146–184 (GST-FAP52146-184), indicating that the region 146–184 is

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necessary, but not sufficient for filamin binding. For binding in vivo, the residues in theNH2-terminal region 1–145 are probably also needed.

5.1.4 FAP52-binding site in filamin

Similar but reciprocal experiments were then performed in order to map the FAP52-bind-ing site(s) in filamin. Both SPR and blot overlay assay affirmatively indicated thepolypeptides corresponding to the aa 1524–1858, 1524–1956, 1524–2064, 1524–2132,1524–2172 and 1524–2283 of filamin as being capable of binding FAP52 (Fig. 5/I). SPRbut not overlay assay also showed binding to the fragment 1524–1780 and weakly tofragment 1524–1664. This disaccord could very well be due to a higher sensitivity of theSPR method. From these results it can be concluded that the residues 1664–1858, corre-sponding to the repeats 15–16 of filamin, are responsible for binding of FAP52.

5.1.5 Colocalization of FAP52 with filamin in cultured cells

The localization of FAP52 in naive CEHFs in relation to filamin distribution wasexplored by immunofluorescence microscopy and immunoelectron microscopy by usingdouble immunolabeling technique. In immunofluorescence microscopy (Fig. 6/I), fil-amin showed mostly a distinct filamentous staining along actin stress fibers. FAP52, onthe other hand, was seen to reside in FAs (Fig. 6a/I). At the sites where the actin fibersabut the FAs there was a distinct overlap of these otherwise distinct staining patterns.More closely, the overlap was seen in a distinct region corresponding to the hinge regioncomprising the more central (proximal) part of the FA (Fig. 6a/I). Colocalization was alsoseen in small punctates along the very edges of the cell periphery especially at sites corre-sponding to the dynamic, fast-moving, forward-pushing edge of spreading fibroblasts(Fig. 6b/I).

For immunoelectron microscopy, cryosections of cultured fibroblasts were double-labeled with antibodies to FAP52 and filamin, which were then visualized by using appro-priate secondary antibody conjugated with small and large gold particles, respectively(Fig. 7/I). Labeling indicating the presence of filamin was seen in a linear fashion, mostprominently in the cell periphery. FAP52, on the other hand, was only localized to periph-eral parts of the cell, at the elongated structures, which, on the basis of their morphologyand localization, represent FAs (Fig. 7a/I). At a higher magnification, colocalization ofFAP52 and filamin was seen (Fig. 7b,c/I).

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5.2 Overexpression of FAP52 in cultured cells (I, II)

Overexpression of protein by using an exogenously introduced vector has become a val-ued means to explore a role of the given protein in cellular structure and function. Weoverexpressed FAP52 and its truncated forms as HA-tagged fusion proteins, a techniquethat allow easy visualization of the fusion products in the cell by using anti-HA antibod-ies. Due to strong overexpression, FAP52 was seen in most cytosolic compartments.There was, however, also a distinct accumulation in surface protrusions and along theedge of the cell (Fig. 8/I). Moreover, fibroblasts overexpressing FAP52-HA displayedseveral types of abnormal morphologies. A normal, nontransfected cell has an elongatedor polygonal, smooth-contoured morphology (Fig. 8a/I). Some cells transfected withFAP52-HA displayed an elongated appearance with several surface protrusion represent-ing filopodia (Figs 8a/I; 9a/II). More typical were roundish transfectants with abnormal orarrested spreading (Fig. 8b/I). In some cells, ruffling edge-typed formations were seen(Fig. 8c,d/I). Actin, as visualized by phalloidin, and FAP52-HA were seen accumulated insuch membrane ruffles. The cells with a slender appearance displayed attenuated actinstaining (Figs 8c/I; 9b/II), indicating the disturbance of actin filament network. As a con-clusion from these studies, it can be said that FAP52 colocalizes with filamentous actin-based structures, and, upon overexpression, FAP52 modulates the actin cytoskeleton.FAP52 may disturb normal actin orchestration by stimulation of actin polymerization inthe cell periphery, resulting in filopod formation, and, on the other hand, by disruption ofthe actin filament network, probably via accelerated actin depolymerization or inhibitedactin polymerization.

We also wanted to see how the overexpression of FAP52 would affect the distributionof filamin. For that purpose, FAP52-HA was transiently expressed in CEHFs, and anti-HA and anti-filamin antibodies were utilized for the visualization of FAP52 and filamin,respectively. The FAP52-HA transfectants displayed an abnormal filamin distribution(Fig. 8a/I). Instead of the normal linear organization along actin fibers, dot-like densitiesclose to the cell surface were seen. Also FAP52 and actin were present in the dots. Therewere only thin actin filaments connecting the dots to each other. This suggests that FAP52associates with filamin in vivo, and that the modulation of the actin cytoskeleton uponoverexpression of FAP52 may be mediated via filamin.

In order to try to resolve what functions the specific domains of FAP52 could serve incells we also carried out transfections by using FAP52Nt-HA and FAP52SH3-HA con-structs. The cells overexpressing FAP52Nt-HA mostly showed a similar distribution of theexogenously expressed FAP52 as seen in the cells overexpressing the full-length protein.However, also some very long filopodia were seen in FAP52Nt-HA-transfectants (Figs8e,f/I; 9a,c/II). Overexpressed FAP52SH3-HA was mostly located in the perinuclearregion and was not associated with any major detectable changes in the overall cell mor-phology, as verified by visualization of actin filaments (data not shown).

Since FAP52 and filamin are closely associated with FAs, we paid special attention tothe number and morphology of FAs in transfected cells. Double labeling experimentsrevealed that in the cells overexpressing FAP52-HA or FAP52Nt-HA, the FAs were smalland randomly oriented, while in naive cells they were radially oriented and regularlyspaced in the periphery of the cell (Figs 8g/I; 9d/II ). On average, FAs in transfected cells

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were also fewer in number than in naive cells (40 vs. 61). Thus, overexpression of FAP52distorts the structure and reduces the number of FAs in cultured fibroblasts.

5.3 Self-association of FAP52 (II)

5.3.1 Computational analysis and circular dichroism

Secondary structure prediction of FAP52, based on its primary structure, indicated that ithas in its NH2-terminus a long stretch with an α-helical arrangement with a high-degreeof propensity to coiled-coil arrangements. This could be fulfilled either by intra- or inter-molecular interactions. In order to distinguish between these possibilities we started toanalyze the capacity of FAP52 to self-associate. The PAIRCOIL program predicted thatFAP52 includes three regions comprising the residues 146–179, 185–219 and 248–280with the probability of 53%, 100%, and 9%, respectively, to form coiled-coils (Fig. 2/II).Another program, MULTICOIL, further discriminates the propensity for dimeric andhigher oligomeric arrangements, giving the likelihoods of dimer formation of 3%, 74%and 0%, and of trimer formation of 4%, 10% and 0.5% for the same regions, respectively(I). These analyses suggest a high propensity for FAP52 to self-associate and for theregion spanning the aa 146–280 to represent the coiled-coil domain.

The content of α-helicity of the wild-type FAP52 and FAP52Nt was also estimated bycircular dichroism, and found to be 25.8% and 57.7%, respectively. The results are well inline with the computational analyses that predict the α-helicity to reside in the NH2-termi-nus of FAP52.

5.3.2 Self-association in vitro and in vivo

In order to resolve whether FAP52 forms dimers, trimers or higher order oligomers in vit-ro we decided to apply cross-linking of the bacterially produced FAP52, followed bySDS-PAGE and coomassie brilliant blue staining. After covalent cross-linking, FAP52migrated as two separate bands with the estimated molecular weight of 170 kDa and 63kDa. They correspond roughly to the trimultiple and the unit weight of FAP52. The non-crosslinked FAP52 was seen as a single band of 63 kDa. This suggests that FAP52 has thecapacity to self-associate in vitro and, most probably, to arrange trimers.

In order to study whether FAP52 occurs as oligomers also in vivo, we utilized chemicalcross-linking, and, on the other hand, gel filtration chromatography, of the lysates of cul-tured CEHFs. For chemical cross-linking experiments, the cross-linked cell lysates weresubjected to SDS-PAGE and coomassie brilliant blue staining. FAP52 migrated as a sin-gle band of 170 kDa (Fig. 5/II). For gel filtration chromatography, the cell lysates wereapplied in the calibrated gel filtration column, and, upon elution, fractions were col-lected. In order to determine the elution profile of the endogenous FAP52, the fractions

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were then subjected to SDS-PAGE and immunoblotting. The majority of FAP52 eluted ina fraction corresponding to the molecular weight of 170 kDa, whereas the minorityshowed a mass below 60 kDa (Fig. 6/II). Taken together, FAP52 predominantly occurs inoligomeric, most probably in trimeric form in vivo.

5.3.3 Kinetics of self-association

We utilized SPR to look at the dynamics of the self-association of FAP52. For that pur-pose, series of concentrations ranging from 20 nM to 400 nM of recombinant FAP52were passed over GST-FAP52 coupled to the sensor chip, and the binding profile wasdetected as changes in the RU values in the sensorgram. The association and dissociationrates were estimated to be 1.15 x 105 M-1 s-1 and 5.47 x 10-4 s-1, respectively, whichgives a KD of 4.7 x 10-9 M, indicating high affinity between the subunits.

5.3.4 Mapping of the self-association site

Secondary structure prediction strongly implicated the NH2-terminal α-helical domain asthe self-association site of FAP52. The verification for the self-association site was exper-imentally pursued by carrying out SPR experiments with GST-FAP52 immobilized on thesensor chip. Use of the sequentially truncated 5´- and 3´-deletion mutants of FAP52 in themobile phase also allowed a more accurate mapping of the association site(s) within thisdefined region. The NH2-terminal fragments encompassing the residues 1–184, 1–219,1–359, but not the one representing the residues 1–145 were seen to bind to the chip. Adistinct binding was also seen with the COOH-terminal fragments with the residues 146–448, 185–448 and 220–448. These results indicate that several distinct regions within aspan of 146–280 contribute to self-association.

5.4 Crystallization and phasing of FAP52 (III)

In order to determine the 3-D structure of FAP52, we decided to resort to crystallizationof FAP52 and its X-ray diffraction analysis. For crystallization, the hanging drop tech-nique was used. Two crystal forms were detected. The space group of the crystal form Iwas P212121, having the cell dimensions a=101.4, b=105.9, c=126.3 Å. The best crystaldiffracted to a resolution of 2.8 Å. The crystals were column-shaped and their averagevolume was 500x60x60 µm3. The unit cell volume was 1.36 x 106 Å3. The asymmetricunit contains two or three protein molecules, suggesting VM of 3.3 Å3Da-1 or 2.2 Å3Da-1,respectively.

The crystals of the form II were prisms or rhomboidal plates and they represent the C2space group. The unit cell dimensions were a = 164.7, b = 102.2, c = 107.3 Å, β = 131.2o.

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The best crystal diffracted to a resolution of 2.1 Å. In addition, a 3.27 Å low-resolutiondataset of 54 frames with 3 degrees oscillation angle was collected to supplement themissing low-resolution reflections lost due to overload. The average dimension of thecrystals was 250x250x100 µm3. The unit cell volume was 1.36 x 106 Å3, consistent againwith of 2 or 3 molecules per asymmetric unit. These crystals were fragile and broke easilyduring mounting and pressurization, and the trials to obtain xenon-derivatized crystalsfailed utterly, or the datasets showed radiation damage. Therefore 30 frames at the end ofthe dataset were omitted from the processing.

The datasets of two different xenon-derivatized crystals were collected using wave-lengths of 1.5 and 1.9 Å, and the maximum resolution of 3 Å was reached. Multiwave-length anomalous dispersion phasing was successfully performed. Preliminary structuralanalysis revealed a dimer in the asymmetric unit.

5.5 FAP52 in bile canaliculi

The distribution of FAP52 in chicken tissues was studied by using the Affi-K7 antibodiesand immunofluorescence microscopy. A positive staining reaction was seen in several tis-sues. Intriguingly, a very distinct staining pattern was seen in the liver, where FAP52 wasseen almost exclusively in close proximity to bile canaliculi (Fig. 1/IV). Due to this asso-ciation of FAP52 with a highly differentiated and specific structure, we decided to lookmore closely at the role of FAP52 in the structure and function of bile canaliculi.

For closer scrutiny, cultured human hepatoma HepG2 cells, Affi-K7 antibodies, andimmunofluorescence and immunoelectron microscopy were utilized. In immunofluores-cence microscopy, vacuolar-like structures resembling bile canaliculi were seen when theactin microfilaments were visualized by rhodamine-phalloidin. Their authenticity wasjudged by labeling for the canaliculus-specific component, CEA (not shown). FAP52localized along the basal domains of the cells and, more specifically, in subdomains cor-responding to CEA-positive bile canaliculi (Fig. 3/IV). FAP52 was especially stronglypresent in the bases of microvilli, which were visualized in double-labeling experimentsby rhodamine-phalloidin. In immunoelectron microscopy, FAP52 was present under theplasma membrane along the canaliculi and, occasionally, in microvillar projections. Theauthenticity of the bile canaliculi was again judged by labeling for CEA (not shown).

6 Discussion

6.1 FAP52 as a multidomain protein

FAP52 was originally described as a novel SH3 domain-containing protein (Meriläinen etal.. 1997). Analysis of the primary structure revealed the presence of a unique SH3domain in the COOH-terminus of the protein and a very highly α-helical stretch with ahigh propensity to coiled-coil arrangement in the NH2-terminus. In the middle, anunstructured “linker” region was predicted.

Currently, more than 400 SH3 domain-containing proteins are known (search fromhuman protein database at the network server http://pfam.wustl.edu). Therein, the SH3domain serves the function of binding various proteins with a proline-rich consensus rec-ognition motifs (Cesareni et al.. 2002). In many cases, the SH3 proteins also contain otherprotein-protein interaction motifs, such as the SH2 domain (Pawson et al.. 2001).

Sequences with a propensity to coiled-coil arrangements are known to mediate bothhomo- and hetero-oligomerization of proteins. Thus, we set out to study whether FAP52and especially its NH2-terminal part was capable of forming dimers or higher order oligo-mers. First, we resorted to computational analysis in order to determine the nature of α-helicity in FAP52. For that purpose we utilized the programs PAIRCOIL and MULTI-COIL. The results showed the presence of three distinct regions with a high tendency tofold in a coiled-coil arrangement and probably in higher order oligomeric complexes.These regions, spanning the aa residues 146–179, 185–220 and 248–280, reside in theNH2-terminus. Dimer was a preferred form in the calculations based on MULTICOIL,which estimates probabilities for various degrees of oligomerization.

The content of α-helicity was verified also empirically by utilizing circular dichroismanalysis. In full-length FAP52 and in its NH2-terminal 359 aa it was estimated to be25.8% and 57.7%, respectively, which is well in line with the prediction by the PHD pro-gram (Meriläinen et al.. 1997) and with the results of the PAIRCOIL and MULTICOILprograms.

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6.2 FAP52 self-associates

The capacity of FAP52 to self-associate was tested by utilizing chemical cross-linking,gel filtration chromatography and SPR analyses. All methods concertedly showed FAP52to have a capacity to oligomerize. In the cross-linking and gel filtration experiments,FAP52 appeared as species of 63 kDa and 170 kDa, corresponding roughly to the mono-meric and trimeric forms of the protein. The kinetics of the self-association was estimat-ed by SPR analysis. The results showed the dissociation constant KD to be 4.7 x 10-9 M,which indicates high affinity between the subunits. The observation that, despite suchhigh affinity, FAP52 also exists as a monomer in cell extract, may be due to the interfer-ence of the self-association by other proteins of the extract, or to the putative effect ofphosphorylation on self-association.

The region in FAP52 that is responsible for the self-association was also determined byutilizing SPR with the sequentially truncated mutants of FAP52. The results revealedthree distinct regions, encompassing the aa residues 146–179, 185–220 and 248–280 mostlikely responsible for the self-association. Thus, the empirical data completely match thepredictions based on the PAIRCOIL and MULTICOIL programs.

In order to determine the detailed 3-D structure of FAP52, we utilized X-ray diffrac-tion analysis. The preliminary structural analysis revealed an antiparallel dimer in theasymmetric unit. The discrepancy between the results of X-ray crystallography and chem-ical cross-linking/gel filtration, which suggested the presence of trimers rather dimers,could be due to the abnormal migration of FAP52 in gel electrophoresis and size exclu-sion chromatography. The calculated molecular mass of FAP52 is 52 kDa, whereas itsmigration in SDS-PAGE corresponds to the molecular mass of 63 kDa. The difference inthe calculated mass and the molecular weight determined by migration in gel may be dueto an exceptionally rigid secondary structure of FAP52.

The NH2-terminal region is highly conserved among the members of the FPS proteinfamily. In a comparison with the BLAST program at the web server http://www.ncbi.nlm.nih.gov/BLAST, their similarity with FAP52Nt ranges from 72% to 91%(data not shown). Among PACSIN 2 proteins, the human, rat and mouse orthologs are91% identical with the NH2-terminal region of chicken FAP52. In addition to FAP52, allthree PACSINs have a tendency to self-associate, but also to form heteroaggregates withthe other PACSINs (Modregger et al.. 2000).

Several FA components, such as α-actinin (Blanchard et al.. 1989) and talin (Gold-mann et al., 1994) have been shown to exist as antiparallel dimers. The rod-shaped coreof native α-actinin is made up of two α-actinin subunits, in which the coiled-coil regionsmediate dimerization (Djinovic-Carugo et al.. 1999). Due to self-association, proteinsmay have a capacity to cross-link actin filaments (α-actinin) or to serve as a docking sitefor various structural or signaling molecules to FAs (talin). In the case of FAP52, dimer-ization could bring proteins that bind to the SH3 of FAP52 to close proximity of eachother. The SH3 domain of FPS proteins binds to a number of proteins involved in vesicu-lar trafficking and actin dynamics, such as synapsin I, dynamin I, synaptojanin, N-WASPand mSos (Qualmann et al.. 1999, Modregger et al.. 2000, Wasiak et al.. 2001). Thus,FAP52 may serve as a coordinator of their proper spatial placement in the cellular pro-cesses.

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6.3 FAP52 as a filamin-binding protein

Elucidation of the function of FAP52 was pursued by determining its binding partners,and by in vivo studies, in which FAP52 or its fragments were overexpressed in culturedcells. By using affinity chromatography, immunoprecipitation, blot overlay and SPR anal-yses, FAP52 was here found to bind to filamin, a dimeric, high-molecular weight protein,which cross-links actin filaments and organizes them into parallel bundles or nonparallel-ly to form orthogonal networks. The COOH-terminus of filamin is composed of 24repeats, which form a rod-like appearance. The 24th repeat forms the self-associationdomain. In the NH2-terminus, there is an actin-binding region. The FAP52-binding site infilamin was mapped to the repeats 15–16. This region of filamin also serves as the bind-ing site for e.g. TNF receptor-associated factor 2 (TRAF2; Leonardi et al. 2000), dopam-ine D2-receptor (Li et al. 2000), and calcium-sensing receptor (CaR) (Awata et al. 2001).The filamin-binding site in FAP52 overlaps with the self-association site. The question ofhow the self-association of FAP52 and the interaction with filamin are related is the sub-ject of further studies.

Filamin has been suggested to have a crucial role in mediating inflammatory stimulivia the TNF receptor for the activation of SAPKs and NF-κB (Leonardi et al. 2000). Fil-amin binds to TRAF2, an intracellular adaptor in TNF receptor signaling (Leonardi et al.2000). Overexpression of filamin blocks the TNF-induced SAPK and NF-κB activation.On the other hand, in M2 melanoma cells, which lack filamin, TNF fails to activateSAPKs and NF-κB (Leonardi et al. 2000). Filamin is also needed in the signaling fromCaR to MAPK cascade activation. By experiments in which CaR was transientlyexpressed in M2 cells and in A7 melanoma cells, which stably express filamin, it wasshown that CaR activates ERK only in the presence of filamin (Awata et al. 2001, Pi et al.2002). Filamin also enhances dopamine D2-receptor signaling. Dopamine D2-receptormediates signaling in a multitude of neurobiological and endocrine processes, e.g. byinhibiting adenylate cyclase (Li et al. 2000). Cells with mutant filamin display attenuatedinhibition of adenylate cyclase (Li et al. 2000).

In the studies discussed above, filamin is suggested to serve as a scaffold and dockingmolecule for the signaling proteins. Filamin also binds same small-molecular weightmodulators of the actin cytoskeleton, such as RhoA, Rac1 and Cdc42, and actin itself(Ueda et al. 1992, Ohta et al. 1999). Due to a multitude of binding sites of signaling mol-ecules in filamin, it may have the capacity to “trap” the entire repertoire of signaling mol-ecules needed for a given cellular process within narrow confines and to guarantee theirproper orientation. It may also bring about a high local concentration of signaling mole-cules involved in a specific cellular process at a focal point.

Filamin also binds to integrins β1A, β1D, β2, β3 and β7 (Pfaff et al. 1998, Takafuta etal. 1998, Liu et al. 2000). Integrin β1, the receptor for the ECM component fibronectin,has been shown to fix the cytoskeleton to cell surface, and, on the other hand, the cell sur-face to ECM in stationary or less motile cells (Critchley 2000). β2, on the other hand,plays a similar role in more motile cells, such as leukocytes (Arnaout 1990, Sharma et al.1995). Tight integrin-filamin interaction has been shown to inhibit cell migration (Calder-wood & Ginsberg 2003). Thus, by linking actin microfilaments to integrins, filamin regu-lates the physical properties of cells. The lymphocyte-specific integrin β7 mediates thecontact of the cell with ECM and the vascular wall. In these cells filamin is suggested to

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contribute the targeting of lymphocytes (Pfaff et al. 1998). In platelets, filamin binds toβ3 integrin (Goldmann 2000). The significance of the interaction is not known.

6.4 FAP52 as a regulator of the cytoskeleton

Overexpression of FAP52 or its filamin-binding region in cultured fibroblasts inducesreorganization of the actin cytoskeleton and the formation on filopodia to the cell sur-face. Similar effects were recently demonstrated also for rat PACSIN 2 (Qualmann &Kelly 2000).

Formation of filopodia is mediated by the Arp2/3 complex, a nucleator of actin fila-ment polymerization (Miki et al. 1998, Qualmann & Kelly 2000). Its activation leads toformation of either filopodia or lamellipodia, depending on the nature of the activatormolecule. Activation by N-WASP leads to formation of filopodia. PACSINs (Qualmann& Kelly 2000, Modregger et al. 2000) bind N-WASP. Thus, the formation of filopodia byFAP52 is probably mediated by N-WASP.

The basis of the formation of the filopodia is the reorganization of the dendritic actinfilaments into parallel bundles facing the plasma membrane (Svitkina et al. 2003). Ourhypothesis is that FAP52, via binding to filamin, controls the actin-filamin interaction in away that enhances the bundling event. This could happen e.g. by bringing the two “arms”of the filamin dimer into a more parallel orientation.

In fibroblasts, overexpression of FAP52 or its NH2-terminal fragment, which containsthe filamin-binding site, led to a distinct reduction in the number of FAs. Whereas thenaive cells expressed on average 61 FAs per cell, the transfected fibroblasts had only 40FAs. Also the morphology and orientation of FAs in FAP52-overexpressing cells wasabnormal.

Currently, no interactions of FAP52 with any FA-specific proteins have beendescribed. Thus, the mechanism responsible for the localization at FAs is not clear. Weare currently working to identify putative interactions of FAP52 with FA-proteins to elu-cidate the molecular mechanisms of its targeting.

Recently, a novel, FA-specific splice variant for filamin B [Filamin Bvar-1(∆H1)] wasdescribed (van der Flier et al. 2002). It lacks 41 aa between the repeats 19 and 20, and hasa deletion between the repeats 15 and 16, corresponding to the hinge region H1. Interest-ingly, the FAP52-binding site in filamin A spans from the repeat 15 to repeat 16, and,thus, overlaps with the deletion site in filamin B. Whether the reduction of the number ofFAs is due to an interference of the excess FAP52 with the normal molecular architectureis a subject of additional research.

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6.5 FAP52 as a component of the pericanalicular web of the bile canaliculi

To expand our insight of the significance of FAP52, we screened its expression in vari-ous tissues by immunohistochemistry and FAP52-specific antibodies (Affi-K7). Apartfrom a fairly wide-spread expression in many tissues we also found a distinct and promi-nent expression present in bile canaliculi.

The liver produces and secretes bile, which is a mixture of various end products of themetabolism of the organism. Bile is secreted by hepatocytes to bile canaliculi, which forman anastamosing network within plates of hepatocytes (Young & Heath 2000). Bile canal-iculi are formed of the apical regions of the plasma membranes of the adjacent hepato-cytes, which are highly specialized, polarized cells. Apical membrane extends smallmicrovilli to the lumens of canaliculi (Young & Heath 2000).

Human hepatoma HepG2 cells are an ideal model for studies on bile canaliculi. Theygrow to islets and spontaneously form short, round bile canalicular structures. In cultures,they retain the biosynthetic capability and the specialized functions of normal liver paren-chymal cells (Chiu et al. 1990, Sormunen et al. 1993).

In immunofluorescence microscopy of cultured HepG2 cells, bile canaliculi, whichwere visualized by rhodamine-phalloidin, were seen as circular structures with themicrovilli directed towards their lumens. FAP52 was seen to localize in the walls of thebile canaliculi. FAP52 was especially strongly present in the bases of microvilli. In elec-tron microscopy studies, FAP52 was localized just below the lateral plasma membrane.Moreover, it was also present in microvilli.

The canaliculus-associated cytoskeleton consists of three distinct zones of actinmicrofilaments (Tsukada et al. 1995; see Fig. 6). First, the parallel bundles of actinmicrofilaments form the core of the microvilli. Actin filaments are there associated withvillin, an actin-splicing protein (Craig & Powell 1980). Second, the membrane-associ-ated actin microfilament band is attached to the membrane. It is associated with myosinII, and is probably involved in the maintenance of the elasticity and the structural integ-rity of the membrane, and in the regulation of the vesicle transport of the canaliculus(Tsukada et al. 1995). Third, the contractile pericanalicular actin filament band is a cir-cumferential belt around the canaliculus. The contraction and relaxation, which is due tothe organization of actin microfilaments with myosin II and tropomyosin, allows the nar-rowing and dilatation, respectively, of bile canaliculi.

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Fig. 6. Actin microfilament architecture in pericanalicular web. Actin filaments are organizedinto three distinct zones. 1, cores of microvilli; 2, membrane-associated microfilaments; and 3,circumferential microfilament band. BC, bile canaliculus; C, cytoplasm; APM, apical plasmamembrane; M, microvillus. (Modified from Tsukada et al. 1995).

As judged by immunofluorescence microscopy and immunoelectron microscopy,FAP52 is located in the regions where the actin filaments of microvilli intersect the per-pendicular pericanalicular actin filament band. The capacity to bind filamin, an actin-orchestrating protein, and the ability to modulate actin organization on cultured cells sug-gests the involvement of FAP52 in the regulation of actin architecture in these nodes ofthe actin network.

7 Conclusions

FAP52 binds directly to filamin. The binding site was mapped to the α-helical NH2-ter-minus of FAP52, and to repeats 15–16 of filamin.

FAP52 self-associates in vitro and in vivo. The site responsible for the self-associationresides in the α-helical region spanning the aa residues 146–280, which also contains thefilamin-binding site. The preliminary X-ray analysis suggests FAP52 to exist as dimer.

Cultured fibroblasts overexpressing FAP52 or its filamin-binding NH2-terminusundergo dramatic morphological alterations. These cells typically extend numerousfilopodia on their surfaces. The distribution of filamin was also changed to the filopodia.In addition, instead of normal, filamentous organization in naive cells, numerous dot-likedensities were seen along the actin fibers. It seems that FAP52 is important in organizingthe actin filaments to parallel stress fibers.

FAP52 has a specific pattern of localization in the liver. Both chicken liver and cul-tured human hepatocytes showed an accumulation of FAP52 under the plasma membraneof bile canaliculi and, especially, at the intersection of the microvillous cores and the peri-canalicular actin microfilaments. Thus, FAP52 may have an important role in organizingthe actin architecture of the pericanalicular web.

Conclusively, FAP52 participates in the organization of the actin cytoskeleton, proba-bly via its direct interaction with filamin. FAP52 self-associates in vivo, which may pro-vide FAP52 the ability to cross-link filamin molecules or regulate the flexibility of the fil-amin dimer, or to serve as a coordinator of the cytoskeleton-associated proteins.

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