Characterization of vinculin´s lipid anchor regionThermodynamic evidence of non-muscle myosin...

Characterization of vinculin´s lipid

anchor region

Der Naturwissenschaftlichen Fakultät

der Friedrich-Alexander-Universität Erlangen-Nürnberg

zur

Erlangung des Doktorgrades

vorgelegt von

Gerold Diez

aus Bad Kissingen

Als Dissertation genehmigt von der Naturwissenschaftlichen

Fakultät der Universität Erlangen-Nürnberg

Tag der müdlichen Prüfung: 18.02.2009

Vorsitzender der

Promotionskommission: Prof. Dr. Eberhard Bänsch

Erstberichterstatter: Prof. Dr. Wolfgang H. Goldmann

Zweitberichterstatter: Prof. Dr. Hojatollah Vali

Partial results from this thesis were published in Biophysical and Biochemical Research

Communications. The following publication and conference contributions contain scientific

results of the thesis presented here.

Publications Mierke CT, Kollmannsberger P, Paranhos Zitterbart D, Raupach C, Diez G, Koch TM, Fabry B, Wolfgang H. Goldmann. Vinculin enhances cell invasion by increasing contractile force generation. Submitted to MCB 2009 Diez G, Kollmannsberger P, Koch TM, Mierke CT, Vali H, Fabry B, Goldmann WH. Anchorage of vinculin to lipid membranes influences cell mechanical behaviour. Submitted to Biophysical Journal (under revision) 2009 Möhl C, Kirchgeßner N, Schäfer C, Küpper K, Diez G, Goldmann WH, Merkel R, Hoffmann B; Modulation of vinculin exchange dynamics regulates adhesion site maturation and adhesion strength. Submitted to Cell Motility and Cytoskeleton (under revision) 2009 Klemm AH, Diez G, Alonso JL, Goldmann WH. Compareing the mechanical influence of vinculin, focal adhesion kinase and p53 in mouse embryonic fibroblasts. BBRC, Vol. 379, Page 799, Feb 2009 Diez G, List F, Smith J, Ziegler WH, Goldmann WH. Direct evidence of vinculin tail - lipid membrane interaction in beta-sheet conformation. BBRC Vol. 373,Page 69, Jun 2008 Schewkunow V, Sharma PS, Diez G, Klemm AH, Sharma PC, Goldmann WH. Thermodynamic evidence of non-muscle myosin II-lipid-membrane interaction. BBRC Vol. 336, Page 500, Feb 2008 Smith J, Diez G, Klemm AH, Schewkunow V, Goldmann WH CapZ-lipid membrane interactions: a computer analysis. Theor. Biol. Med. Model. Vol. 3 Page 30, Aug 2006 Scott DL, Diez G, Goldmann WH. Protein-Lipid Interactions: Correlation of a predictive algorithm for lipid-binding sites with three-dimensional structural data. Theo. Biol. Med. Model. Vol. 3, Page 17, Mar 2006 Conference contributions Diez G, Kollmannsberger P, Koch TM, Zitterbart DP, Mierke CT, Krukiewicz AA, Vali H, Fabry B, Goldmann WH. Anchoring of vinculin to the lipid membrane influences its tyrosine phosphorylation on position 1065. Meeting of the ASCB, San Fransisco, December 2008 Diez G, List F, Smith J, Himmel M, Ziegler WH, Goldmann WH. Characterization of vinculins membrane binding anchor-an in vitro study. 31st Annual Meeting of the DGZ (Marburg), EJCB Vol.87S1, Suppl.58, March 2008

Diez G, Kollmansberger P, Paranhos-Zitterbart D, Mierke CT, Smith J, Goldmann WH. Head and tail interactions of vinculin influence cell mechanical behaviour. 47.th Annual Meeting of the ASCB, Washington, December 2007

Diez G, Kollmansberger P, Goldmann WH. Anchoring of vinculin to lipid membranes influences its binding strength in living cells. 2nd European Meeting on Cell Mechanics, Barcelona September 2007

Diez G., Kollmansberger P., Vali H., Goldmann W.H. Anchoring of Vinculin to the membrane influences its binding strength in living cells. Annual CBB Meeting, McGill University Montreal (Canada), May 2007 Diez G., Smith J., Stiebritz M., Goldmann W.H. Molecular dynamics and secondary structure behaviour of the C-terminus of vinculin that includes a membrane binding anchor. German Physical Society Spring Meeting, Regensburg, Mar 2007 Diez G., Smith J., Goldmann W.H. Secondary structure computer analysis of vinculin’s C-terminus. 51st Biophysical Society Symposium, Mar 2007 Diez G., Scott D.L., Goldmann W.H. Protein-Lipid interactions: Correlation of a predictive algorithm for lipid-binding sites with 3D-structural data. Journal of Biomechanics, Vol. 39 (Sup.) P. 578, Aug 2006

Table of contents

1 Introduction ...................................................................................................................... 1

1.1 The extracellular matrix determines the different tissues .......................................... 1 1.2 Mechanical perturbations regulate cell survival and tissue formation....................... 3 1.3 The focal adhesion complex....................................................................................... 6 1.4 Focal adhesion formation ........................................................................................... 8 1.5 Vinculin a key player of the focal contact................................................................ 10

1.5.1 Lipid binding of vinculin.................................................................................. 12 1.6 Aims ......................................................................................................................... 14

2 Materials and Methods .................................................................................................. 15 2.1 Materials................................................................................................................... 15

2.1.1 Chemicals and enzymes ................................................................................... 15 2.1.2 Cell culture medium and plastic ware .............................................................. 15 2.1.3 Oligonucleotides............................................................................................... 15 2.1.4 Vectors ............................................................................................................. 16 2.1.5 Bacterial cultures and cell lines........................................................................ 16 2.1.6 Antibodies ........................................................................................................ 17 2.2.1 Methods in molecular biology.......................................................................... 17

2.2.1.1 Quantification of Desoxyribonuclein acid (DNA) ....................................... 17 2.2.1.2 DNA electrophoresis .................................................................................... 17 2.2.1.3 Restriction endonuclease digestion of DNA ................................................ 18 2.2.1.4 Ligation ........................................................................................................ 18 2.2.1.5 Preparation of chemical competent DH5α ................................................... 18 2.2.1.6 Transformation of competent cells............................................................... 19 2.2.1.7 Polymerase chain reaction (PCR) ................................................................ 19 2.2.1.8 Site directed mutagenesis ............................................................................. 19 2.2.1.9 Purification of plasmid DNA from bacteria................................................. 20 2.2.1.10 DNA sequencing ...................................................................................... 20 2.2.1.11 DNA extraction from agarose gels........................................................... 20 2.2.1.12 Cloning strategy – EGFP expression vector ............................................ 20 2.2.1.13 Cloning strategy – vinculin full length and tail domain constructs.......... 21

2.2.2 Cell biology methods ....................................................................................... 22 2.2.2.1 Cell culture ................................................................................................... 22 2.2.2.2 Transient transfection of cells ...................................................................... 22 2.2.2.3 Culturing of cells on the cover slips for microscopic analysis..................... 23 2.2.2.4 Fixation and permeabilization...................................................................... 23 2.2.2.5 Immunolabelling, fluorescence microscopy and image processing............. 23 2.2.2.6 Determination of the spreading area ............................................................ 24 2.2.2.7 Fluorescence recovery after photo-bleaching (FRAP) measurements ......... 24 2.2.2.8 Electron microscopy measurements............................................................. 25

2.2.3 Biochemical and biophysical methods............................................................. 26 2.2.3.1 Bead coating................................................................................................. 26 2.2.3.2 Peptide synthesis .......................................................................................... 26 2.2.3.3 Lipid vesicle preparation.............................................................................. 26 2.2.3.4 Differential scanning calorimetry (DSC) measurements ............................. 27 2.2.3.5 Circular dichroism (CD) spectroscopy measurements................................. 29 2.2.3.6 Solid state NMR........................................................................................... 30 2.2.3.7 Cell lysis....................................................................................................... 32 2.2.3.8 Protein concentration determination according to Bradford ........................ 32 2.2.3.9 Western blot analysis ................................................................................... 32

2.2.4 Computational methods.................................................................................... 33

Table of contents

2.2.4.1 Molecular dynamics (MD) simulations........................................................ 33 2.2.4.2 Analysis of molecular dynamics simulations............................................... 34 2.2.4.3 Cluster analysis ............................................................................................ 35 2.2.4.4 Visualization of calculated structures .......................................................... 35

2.2.5 Cell mechanical methods ................................................................................. 35 2.2.5.1 Magnetic tweezer measurements ................................................................. 35 2.2.5.2 2D-traction microscopy measurements........................................................ 37

3 Results ............................................................................................................................. 38 I. In vitro ...................................................................................................................... 38 3.1 Differential scanning calorimetry (DSC) measurements ......................................... 38

3.1.1 DSC measurements of the C-terminal arm peptide.......................................... 38 3.1.2 DSC measurements of the mutated C-terminal arm......................................... 39

3.2 Molecular dynamics simulations.............................................................................. 40 3.2.1 MD Simulations of the C-terminal arm............................................................ 40 3.2.2 MD Simulations of the vinculin-tail................................................................. 46

3.3 CD-spectroscopy measurements of the C-terminal arm .......................................... 47 3.4 Solid state NMR measurements of the C-terminal arm ........................................... 49 II. In vivo ....................................................................................................................... 50 3.5 Cloning and expression of the different vinculin constructs.................................... 50

3.5.1 Cloning and expression of EGFP linked vinculin and vinculinΔC.................. 50 3.5.2 Cloning and expression of EGFP-linked vinculin-tail and vinculin-tailΔC..... 52 3.5.3 Mutagenesis of vinculin´s src dependent phosphorylation site Y1065F ......... 52

3.6 Magnetic tweezer measurements ............................................................................. 53 3.6.1 Magnetic tweezer measurements of MEF-resc and MEF-vinΔC cells ............ 54 3.6.2 Creep measurements of MEF-vtail and MEF-vtailΔC cells ............................ 58

3.7 Determination of the spreading area ........................................................................ 63 3.8 Determination of the FA per cell.............................................................................. 63 3.9 FRAP measurements ................................................................................................ 65 3.10 2D-traction microscopy measurements.................................................................... 66

3.10.1 Strain energy measurements of MEF-vinΔC ................................................... 66 3.10.2 Strain energy measurements of different vinculin mutants.............................. 67

4 Discussion ........................................................................................................................ 70 4.1 Vinculin´s lipid anchor revealed membrane insertion potential .............................. 71 4.2 Vinculin´s lipid anchor is involved in beta-sheet formation .................................... 72 4.3 Vinculin´s lipid anchor regulates cell stiffness ........................................................ 74 4.4 The lipid anchor affects traction generation via phosphotyrosine 1065 .................. 76 4.5 The Model ................................................................................................................ 78 4.6 Outlook..................................................................................................................... 79

5 References ....................................................................................................................... 81 6 Appendix ......................................................................................................................... 88

Abbreviations

A Alanine ADF Actin depolymerization factor AFM Atomic force microscope Amp Ampicillin AmpR Ampicillin-Resistenz Arp2/3 Actin related protein 2/3 ATP Adenosine-triphosphate ATPase Adenosine-triphosphate phosphatase BDM 2,3-Butanedione monoxime Bp Base pair BP320 Blocking peptide 320 BSA Bovine serum albumin C Cysteine Ca2+ Calcium CapZ Capping protein Z CD Circular dichroism cDNA complementary DNA CFP Cyane fluorescent protein CM Calcium/ Manganese CO2 Carbon dioxide CPU Central processing unit D Aspartate dATP Deoxyadenosine-triphosphat dCTP Deoxycytosine-triphosphat dGTP Deoxyguanosine-triphosphat Dia1 Diaphanous 1 peptide DMEM Dulbeccos modified eagle medium DMPC Dimyristoyl-L-α-phosphatidylcholine DMPG Dimyristoyl-L-α-phosphatidylglycerol DMSO Dimethyl sulfoxid DNA Deoxyribonucleic acid dNTP Deoxynucleotide-triphosphat DSC Differential scanning calorimetry DSSP Dictionary of secondary structure prediction DTT Dithiothreitol dTTP Deoxythymidine-triphosphate E Glutamate ECL-reagent Enhanced chemiluminescence reagent ECM Extracellular matrix EDTA Ethylenediaminetetraacetic acid EGFP Enhanced green fluorescent protein ERK-protein Extracellular signal regulated protein kinase F Phenylalanine FA Focal adhesion FAK Focal adhesion kinase FCS Fetal calf serum FERM F for Band 4.1, E for Ezrin, R for Radixin, M for Moesin FRAP Fluorescence recovery after photobleaching FRET Fluorescence resonance energy transfer Fs Femto-second Fx Focal complex

Abbreviations

g Gramm G Glycine Gd2+ Gadolinium GHz Giga-Herz GROMACS Groningen Machine for Chemical Simulations GTP Guanosine-triphosphate GTPase Guanosine-triphosphate phosphatase h Hour H Histidine HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid HMW marker High molecular weight marker HPC High performance computer I Isoleucine ILK Integrin linked kinase IRSp53 Insulin receptor substrate protein 53 K Lysine Kan Kanamycine kb Kilo-base Kcal Kilo-calories kDa Kilo-dalton kJ Kilo-joule kPa Kilo-pascal KT5926 A protein kinase inhibitor l Litre L Leucine LB Lura-Bertoni-medium LRW London resign white M Methionine mA milli-Ampere MCS Multiple Cloning site MD Molecular dynamics MEF Mouse embryonic fibroblast MgCl2 Magnesium-chloride MHZ Mega-Herz min Minute ML-7 Myosin light chain kinase inhibitor No. 7 MLV Multi-lamellar vesicles mM milli-molar N Asparagine Na+ Sodium NaCl Sodium-chlorid nm Nano-meter NMR Nuclear magnetic resonance ns Nano-seconds OD Optical density P Proline p.A. Per analysis p21 Cyclin-dependent kinase inhibitor 1A or CDKN1A p53 Protein or tumor protein of 53 kDa p95PKL Paxillin kinase linker PAA Polyacrylamide

Abbreviations

PAGE Polyacrylamide gel electrophoresis PAK p21 activated protein kinase PBS Phosphor buffered saline PC Phosphatidyl-choline PCR Polymerase chain reaction PDB Protein data base PEG Polyethylene glycol Pfu DNA polymerase from Pyrococcus furiosus pH Potential hydrogenii PH Pleckstrin homologue PI Isoelectric point PIP Phosphatidyl-inositol-phosphate PIP2 Phosphatidyl-inositol 4,5-bisphosphate PMSF Phenylmethanesulphonylfluoride POPC 1-Palmitoyl-2-Oleoyl-sn-Glycero-3-Phosphocholine POPG 1-Palmitoyl-2-Oleoyl-sn-Glycero-3-Phosphoglycerol ppm Parts per million ps Pico-second Q Glutamine R Arginine RAM Random access memory ROCK Rho associated kinase rpm Rotation per minute RPTP Receptor tyrosine phosphatases RT Room temperature s Second S Serine SCAR Suppressor of cAMP receptor SDS Sodiumdodecylsulfat SUV Small unilamellar vesicles T Threonine TAE Tris-Acetate buffer with EDTA Taq DNA polymerase from thermus aquaticus TBS-T Tris-buffered-saline with Tween TE Tris/ EDTA TEM Transmission electron microscope TEMED N,N,N’,N’, Tetraethylendiamin Tet Tetracycline TM Tris/ Magnesiumacetat Tris Tris-(hydroxymethyl)-Aminomethan TRITC Tetramethyl Rhodamine Iso-Thiocyanate V Valine Vinculin-CT EGFP-linked vinculin; RK1060/61 mutated to Q Vinculin-H3 EGFP-linked vinculin; K952, K956, R963, R966 mutated to Q Vinculin-LD Vinculin-CT + Vinculin-H3 VinculinY1065F EGFP-linked vinculin Y at position 1065 mutated to F VinculinΔC EGFP-linked vinculin (residues 1-1052) w.o. the lipid anchor Vol. Volumen Vtail EGFP-linked vinculin-tail (residues 858–1066) VtailΔC EGFP-linked vinculin-tail (residues 858-1052) w.o. the lipid

anchor

Abbreviations

W Tryptophane WASP Wiskott aldrich syndrome protein WAVE Wiskott aldrich Verprolin homologue protein Y Tyrosine YFP Yellow fluorescent protein

Zusammenfassung

Zusammenfassung Die extrazelluläre Matrix (ECM) beeinflusst und kontrolliert die Adhäsion sowie die

Migration von Zellen. Der fokale Adhäsionskontakt (FA) verbindet intrazellulär das Aktin-

Zellskelett über den transmembranen Integrin-Rezeptor mit Komponenten der ECM, wie

Kollagen und Fibronektin (FN). Der Auf- und Umbau dieser Adhäsionskontakte ist unter

anderem nur durch die Wechselwirkung von einigen fokalen Proteinen mit der Zellmembran

möglich. Das Vinkulin-Protein, welches in eine Kopf- (95 kDa) und eine Schwanzgruppe (30

kDa) unterteilt werden kann, zeigt solche Membran-bindende Strukturen. Es konnte

experimentell bestätig werden, dass Helix 3 (Aminosäuren 935-978) sowie die letzten 15

Aminosäuren des C-terminus (Aminosäuren 1052-1066; Lipid Anker) der Vinkulin-Schwanz

Gruppe mit Lipid-Vesikeln interagieren. Sogenannte Pull-down Experimente mit

Lipidvesikeln, die mit der Vinkulin-Schwanz gruppe ohne den Lipid Anker (vtailΔC)

inkubiert wurden, zeigten im Gegensatz zu den Versuchen mit dem gesamten Vinkulin-

Schwanz (vtail) keine Interaktion mit der artifiziellen Membran. Ob allerdings der Lipid-

Anker tatsächlich direkt an der Membran Wechselwirkung beteiligt ist, wurde im Rahmen

dieser Experimente nicht geklärt.

In dieser Arbeit wurde mittels „Differential scanning Calorimetry“ (DSC) versucht, die Frage

der direkten Beteiligung des Lipid-Ankers (Aminosäuren 1052-1066) an der

Membranbindung von Vinkulin zu klären. Diese Messungen haben gezeigt, dass der C-

terminale Arm von Vinkulin (Aminosäuren 1045-1066) mit dem hydrophoben Bereich der

Lipidvesikel in Kontakt tritt. Molekulardynamische Simulationen und Circular Dichroismus

(CD) Messungen lassen vermuten, dass der Lipid-Anker eine für Lipid-Interaktionen günstige

anti-parallele beta-Faltblatt Konformation einnimmt. Nuclear magnetic resonance (NMR)-

Messungen in Anwesenheit von POPC/ POPG Membranen bestätigten dies.

Weiterhin ist bekannt, dass Zellen die Vinkulin exprimieren welches nicht mit Membranen

wechselwirken kann, eine verminderte FA-Umbaurate aufweisen. Dies wirkt sich negativ auf

die Adhäsion und Migration der jeweiligen Zellen aus. Basierend auf diesen Ergebnissen

wurden im Rahmen dieser Arbeit in zusätzlichen in vivo Experimenten der Einfluss des Lipid-

Ankers auf die mechanischen Eigenschaften der Zellen getestet. Zu diesem Zweck wurden

MEF-Vinkulin(-/-) Zellen mit Vinkulin dessen Lipid-Anker fehlte (MEF-vinΔC)

retransfiziert. Diese MEF-vinΔC Zellen wurden mit Fibronectin beschichteten

paramagnetischen „beads“ inkubiert und mit einer „magnetischen Nadel“ untersucht. Im

Vergleich zu wildtyp (MEF-wt) und rescue (MEF-resc) Zellen verhielten sich die MEF-

vinΔC Zellen weniger steif. Gleichzeitig rissen während der Messung von diesen Zellen mehr

Zusammenfassung

„beads“ ab. Dies ist ein Indiz für die verminderte Anbindung an ECM beschichteten

Oberflächen von MEF-vinΔC Zellen. 2D-traction microscopy Experimente zeigten weiter,

dass diese MEF-vinΔC Zellen im Vergleich zu MEF-wt bzw. MEF-resc Zellen während der

Adhäsion geringere Kräfte und weniger fokale Kontakte ausbilden. Dies wird auf die

verminderte Actin-myosin Aktivität zurückgeführt. Messungen der Kraftentwicklung von

Zellen, die weitere Lipidbindedefiziente Varianten von Vinkulin exprimierten, zeigten, dass

lediglich der Knock-out des Lipid-Ankers von Vinkulin zu einer Reduktion der Kräfte führt.

Der Knock-out der src-Phosphorylierungstelle im Lipid-Anker (Y1065F) führt zu ähnlich

verringerten Kräften. Diese lässt den Schluss zu, dass die Zellmechanik über die

Membranbindung des Lipid-Ankers, sowie der src-abhängige Phosphorylierung von Vinkulin

beeinflusst wird.

Summary

Summary The contact of the cell to the extracellular matrix components such as collagen and fibronectin

is important for cell adhesion and migration. Most of the cell matrix contacts are linked to the

actin filaments by the focal adhesions (FA). At the FA, heterodimeric transmembrane integrin

receptors link the ECM to the cytoskeleton via adaptor proteins that are part of a sub

membrane plaque. A variety of those adaptor proteins have been shown to associate with, and

in some cases insert into, lipid bilayers. The focal adhesion protein vinculin (1066 residues),

which can be separated into a 95 kDa head and a 30 kDa tail domain, shows such lipid

binding sites. However, the function of vinculin´s lipid binding is still an enigma. Two

regions on the 30 kDa tail domain have been experimentally identified as candidates for lipid-

binding: Helix 3 (residues 935–978) and the lipid anchor (residues 1052–1066). The alteration

of helix 3 (residues 944-978) and the unstructured C-terminal arm (residues 1052-1066, so-

called lipid anchor) resulted in impaired lipid vesicle interaction of the vinculin-tail. Pull-

down assays with artificial lipid membranes revealed that in contrast to vinculin-tail (vt), a

variant lacking the lipid anchor (VtΔC), does not interact with vesicles. To what extent the

last 15 residues are involved in lipid interaction was not determined.

In this study the lipid-binding ability of the lipid anchor of vinculin as well as the influence to

cell mechanical behavior were determined. Differential scanning calorimetry (DSC)

demonstrated that vinculin´s C-terminal arm, which includes the lipid anchor, is directly

involved in lipid binding. The peptide inserts into the lipid vesicle consisting of DMPC/

DMPG at various molar ratios. The secondary structure of the C-terminal arm was also

explored under different ionic conditions which represent nominal basic, neutral and acidic

pH´s using molecular dynamics simulations. The generated trajectories predicted an

antiparallel beta-sheet followed by an unstructured C-terminal end for the peptide

representing vinculin´s C-terminal arm under “basic” and “neutral” conditions. This

conformational behavior was investigated in more detail in the presence/absence of DMPC/

DMPG vesicles using CD-spectroscopy. The results suggest direct association of vinculin’s

lipid-binding region (residues 1052–1066) with membranes whilst forming a beta-sheet. To

determine the orientation of the lipid anchor during membrane interaction, solid state NMR

measurements were performed using vinculin´s C-terminal arm peptide. Those results imply

that in presence of POPC/ POPG vesicles, the beta-sheet inserts into the lipid membrane.

Furhermore, it was demonstrated that cells expressing vinculin without the lipid anchor

(vinΔC) showed a decreased focal adhesion turnover rate, which results in impaired cell

adhesion and migration. In additional in vivo experiments, the influence of the lipid anchor

Summary

region (residues 1052–1066) in terms of cell mechanical behavior was determined using

vinculin deficient mouse embryonic fibroblasts, retransfected with EGFP-linked vinculin

lacking the lipid anchor (MEF-vinΔC). Magnetic tweezer experiments revealed that MEF-

vinΔC cells, incubated with fibronectin coated paramagnetic beads, were less stiff and more

beads detached during these experiments compared to MEF-resc cells. Cells expressing

vinΔC formed fewer focal contacts as determined by confocal microscopy. 2D-traction

measurements showed that MEF-vinΔC cells generated less force compared to rescue cells.

Attenuated traction forces were also found in cells that expressed vinculin with point

mutations of the lipid anchor that either impaired lipid binding or prevented src-

phosphorylation at site Y1065. However, traction generation was not diminished in cells that

expressed vinculin with impaired lipid binding due to point mutations on helix 3. These

results show that both the lipid binding and the src-phosphorylation of vinculin's C-terminus

are important for cell mechanical behavior, but the lipid binding of helix 3 is not, suggesting

that both the lipid anchor and the src-phosphorylation of Y1065 affect cell mechanical

behavior.

Introduction

1

1 Introduction

1.1 The extracellular matrix determines the different tissues Most of the eucaryotic cells in multicellular organisms are organized as cooperative

assemblies called tissue, which in turn are associated in various combinations to form larger

functional units called organs [1]. In higher organisms such as vertebrates, the major tissue

components are nerves, muscles, blood, lymphoid, epithelial and connective (mesenchymal)

tissue. The cells in tissues are usually in contact with a complex network of extracellular

macromolecules referred to as extracellular matrix (ECM) which is mainly secreted by

fibroblasts. This matrix organizes the contact to neighbouring cells and provides a lattice for

cells to migrate. The matrix also determines the localized function of the tissue. To maintain

the tissue, the cells need mechanical strength, which is often provided by the extracellular

matrix. Connective- and epithelial tissues represent two extremes in which structural roles

played by the matrix and by cell-cell contacts are radically different (figure 1-1). In

connective tissues, the extracellular matrix is abundant and cells are sparsely distributed in it.

The matrix is full of fibrous polymers, such as collagen. Cells as well as the matrix bear most

of the mechanical perturbations to which the tissue is subjected. The cells such as fibroblasts

are attached to the ECM components and exert forces, whilst direct interactions of the

embedded cells are relatively unimportant. In contrast, epithelial tissues are built of tightly

bound cells which form layers. The ECM is here sparsely distributed and consists mainly of a

thin mat called basal lamina that can be found under each cell layer.

Figure 1-1: Simplified cartoon of a cross section through part of the intestine [1]. Here, the cells themselves develop and deal with mechanical stresses by protein filaments

such as the cytoskeleton. The components criss-cross the cytoplasm of each epithelial cell to

Introduction

2

transmit any mechanical perturbations from one cell to the other. The cytoskeleton is directly

or indirectly linked to transmembrane proteins in the plasma membrane where specialized

junctions connect the surface of adjacent cells with the underlying basal lamina. Epithelial

cell sheets line all the cavities and free surfaces, and the specialized junctions between the

cells enable these sheets to form barriers between solutions and cells from one body

compartment to another. Epithelial sheets nearly always rest on a supporting bed of

connective tissue, which may attach them to other tissue types such as muscle [1].

The formation and maintenance of tissue is dependent on the ability to resist surface tension

and shear stress [2]. Mechanical loads exerted at the macroscale affect conformational

changes in connective tissue and ECM. This results in production of stress at the microscale

level that produces global structural rearrangements of the individual molecular components

such as the ECM and the interconnected cytoskeleton of adherent cells. Figure 1-2 shows that

tension-dependent changes in the orientation of collagen bundles, cytoskeletal and nuclear

rearrangements within adherent fibroblasts, as well as distortion of interconnected basement

membrane scaffolds. Those in turn produce similar cytoskeletal and nuclear rearrangements

within associated endothelial cells.

Figure 1-2: Cellular mechano responsiveness and physical connectivity between ECM, cells and the cytoskeletal network. The shear stress and the cells´ own cytoskeletal movement results in changes of the collagen fibers conformation [2].

At the same time, fluid shear stresses applied at the apical surface of certain endothelial and

epithelial cells can also influence the cytoskeletal structure by deflecting the primary cilium

found on the surface of many cells, and by exerting tension on the plasma membrane and

apical cell-cell junctions. Because these shear stresses are also channelled through the

cytoskeleton to basal cell-ECM adhesions, they can alter the structure and function of

underlying connective tissue as well. The entire cellular response to stress may therefore vary

depending on the structural integrity and organization of the whole cytoskeleton-cell-ECM

Introduction

3

lattice, as well as the tension in the network prior to load application. For instance, in the lung

the residual filling pressure that remains after expiration is responsible for tensing and

stiffening the ECMs (basement membranes, collagen fibers, elastin bundles) that surround

each alveolus, and for resisting surface tension forces acting on the epithelium. This force

balance stabilizes the alveoli in an open form [3]. Lung expiration and inspiration influence

this force balance and produce complex micromechanical responses in the lung parenchyma,

including lengthening and shortening (and tension and compression) of alveolar walls

depending on the direction of the applied stress [2]. This is accompanied by extension and

linearization of some collagen fibers on inspiration, as well as buckling of the same fibers on

expiration. Breathing also causes the lateral intercellular spaces between epithelial cells to

reversibly shrink and expand without compromising the structural integrity of the tissue.

Those form of reversible mechanical deformation within surrounding alveolar cells by

altering the local concentration of soluble ligands for epidermal growth factor receptors [4].

1.2 Mechanical perturbations regulate cell survival and tissue formation Throughout their life, cells participate in numerous physical interactions with their

neighbourhood. Endothelial cells sense physical perturbations such as shear stress, produced

by the blood flow [2; 5-6]. In kidney epithelial cells, fluid shear stress produces Ca2+ influx by

deflecting the primary cilium, which acts like a long, microscopic, vertical lever arm on the

apical surface of the cells [2]. In the lung the residual filling pressure that remains after

expiration is responsible for tensing and stiffening the tissue. Cells embedded into the

muscular tissue are dealing with mechanical pressure, generated by their one contractile

machinery such as the actomyosin filaments [5-6]. At cell contact sites, the extracellular

domains of the transmembrane adhesion molecules interact with partners localized on the

surface of the adjacent cells or in the extracellular matrix, whilst the intracellular parts are

attached to components of the cytoskeleton. The connections provide anchor points to mediate

the mechanical stability and integrity of the cells. Furthermore, cell-cell and cell-matrix

contacts are important for the formation and preservation of tissues, cell survival and

proliferation. In order to form and maintain such tissues, eucaryotic cells have elaborated

systems that enable the cell to interpret and amplify signals from other cells. The signalling

systems includes cell surface and intracellular receptor proteins, protein kinases, protein

phosphatases, GTP-binding proteins and many other proteins with which those proteins

interact. This classical process so called signal transduction refers to any process by which a

cell converts one kind of a stimulus into biochemical signal [1]. Additional, an increasing

Introduction

4

body of evidence suggests that besides the classical biochemical signals such as cytokines and

hormones, force dependent mechanical interactions play a central role in regulating cell

behavior [7]. Cell adhesion to other cells or extracellular matrix components (ECM) such as

collagen, fibronectin and laminin trigger biochemical or biomechanical signals inside the cells

that initiate cell survival, proliferation or cell migration by regulating protein synthesis and

cytoskeletal reorganization [2; 7-9]. Most of the cell-cell and cell matrix contacts are linked to the intermediate- or actin filaments.

These transmembrane driven connections are summarized and described in figure 1-3.

Tight junctions are structures between two endothelial or epithelial cells and prevent the

leakage of water-soluble molecules into the tissue. They serve as semi-permeable diffusion

barrier between two cells by separating distinguished areas. These structures are Ca2+

dependent built. Chelating Ca2+ ions lead to tight junction dissolution. In order to transport

high amounts of amino acids and glucose, the tight junctions can be opened. Integral

transmembrane proteins such as claudin, and occluding link apically two neighbouring cell.

These peptide structures are intracellular linked to the actin cytoskeleton by adaptor peptides

form the zonula occludens (ZO) family [10].

Adherens junctions are adhesion belts, commonly found between epithelial cells in the

small intestine. The belt-like junction encircles each of the interacting cells. Its most obvious

feature is a contractile bundle of actin filaments running along the cytoplasmic surface of the

junctional plasma membrane. In this intercellular connection, the actin filaments are joined

from cell to cell by transmembrane adhesion proteins called cadherins. The cadherins form

homodimers in the plasma membrane of each interacting cell. The extracellular domain of one

cadherin dimer binds to the extracellular domain of an identical cadherin dimer on the

adjacent cell. The intracellular tails of the cadherins bind to anchor proteins that tie them to

actin filaments. These anchor proteins include α-catenin, β-catenin, γ-catenin (also called

plakoglobin), α-actinin, and vinculin. [9-10].

Desmosomes are button-like points of intercellular structures that link mainly epithelial and

heart muscle cells. The desmoglein and desmocollin glycoprotein receptors (also cadherin

superfamily) are indirectly connected with intermediate filaments such as keratin and desmin.

Desmoplactine and plakoglobine built the junction between the receptor and keratin-

filaments. Extracellular the receptors are in contact with cadherin receptors of the same

family, exposed by the neighbouring cell [11]. Those structures prevent the leakage of body

fluids into the loosened epithelium.

Introduction

5

Hemidesmosomes or half desmosomes are morphologically similar to Desmosomes.

Instead of joining adjacent epithelial cell membranes, they connect the basal surface of those

cells via integrins to extracellular matrices (ECM) consisting of laminin – a specialized mat of

ECM at the interface between the epithelium and the connective tissue. Intracellular, the

receptors are linked to the intermediate filaments by plectin and BP230 adaptor peptides [12].

Focal adhesions are the structural junction between the ECM via integrins with the

cytoskeleton. Structural proteins such as talin, alpha actinin and vinculin built the linkage

between the receptors and the actin filaments. Focal adhesions serve as the mechanical

linkages to the ECM, and as a biochemical signalling hub that concentrate and directs

numerous signaling proteins. In sessile cells, focal adhesions are quite stable whilst in moving

cells their stability is diminished: this is because in motile cells, focal adhesions are being

constantly assembled and disassembled as the cell establishes new contacts at the leading

edge, and breaks old contacts at the trailing edge of the cell. One example of their important

role is in the immune system, in which macrophages migrate along the connective

endothelium following cellular signals to damaged tissue [2; 5-8].

Figure 1-3: Schematic of cell contacts visualized in epithelial cells [1].

The most prominent mechanosensory devices that can detect forces and respond to them are

adhesion sites to neighbouring cells and to the ECM. However, the function of such “force

receptors” is still poorly understood and characterized in comparison to the classical

biochemical receptors [2, 5-8]. The most prominent and best characterized mechano-receptor

is the focal adhesion complex or focal contact. Ingber and co-workers showed in magnetic

Introduction

6

twisting experiments that the permanent movement of integrin ligand coated beads, attached

to cells, induces a force dependent “stiffening response” in those cells. During these

experiments, cells were incubated with ECM-coated magnetized beads. To the particles a

magnetic field was applied which in turn displaces the beads connected to the cells. Due to the

applied stress, the cells increased their resistance to mechanical deformation, suggesting that

the focal adhesions respond to applied forces [13].

1.3 The focal adhesion complex Structurally defined focal adhesion sites between cultured cells and ECM were described

about 30 years ago using interference reflection, fluorescence and electron microscopy [5; 7;

14-15]. The focal adhesions anchor the cells to the ECM and can be described as streak like

structures which are connected to actin stressfibers (figure 1-4A). Besides anchorage of cells

to the ECM, focal adhesion assembly is also involved in cell migration. Focal adhesion (FA)

complexes are highly dynamic structures composed of structural and signal transduction

proteins. As described in figure 1-3, heterodimeric transmembrane integrin receptors link the

ECM to the cytoskeleton via adaptor proteins that are part of a sub membrane plaque (figure

1-4B). The beta subunits of integrins play a direct role in the establishment of that connection.

The intracellular part of integrin exhibits two conservative NPxY and NPxF motives. With

those sites the receptor can interact with peptides that show phosphotyrosine binding sites

(PTB like modules).

Figure 1-4: Illustration of focal adhesions. A) Visualization of the focal adhesion complexes (green), stained with EGFP-linked vinculin in mouse embryonic fibroblast. The focal adhesions form streak like structures which are in contact with the actin stress-fibers (red). The bar represents 20 µm. B) A schematic diagram of a focal adhesion complex, modified according to Goldmann et al [16]. The integrin receptors link the ECM to the actin cytoskeleton via the sub membrane plaque. This plaque is composed of structural and signalling proteins which sense mechanical perturbations and translate them into biochemical signals. Talin belongs to that group and connects the integrins directly to the actin cytoskeleton [17].

Other actin related peptides such as alpha actinin and filamin also link the transmembrane

Introduction

7

receptor to the cytoskeleton [7; 18]. In addition to these direct linkers, there are structural

components such as vinculin, actin related protein (Arp2/3) or paxillin that reinforce the

linkage to the cytoskeleton and/or recruit signal transduction proteins to the focal adhesions

[19- 21].

Peptides such as small GTPases (Rho, Rho associated kinase (ROCK), Rac etc.),

phosphotyrosine kinases (focal adhesion kinase (FAK), Src, Fyn), Phosphotyrosine

phosphatase, serin threonine kinases (integrin linked kinase (ILK) or p21 activated kinase

(PAK)) and phosphatidyl-inositol kinases are also localized in the focal adhesions [5; 7; 22].

Altogether, these molecular conductors regulate FA formation by integrating the mechanical

perturbations into biochemical signals such as phosphorylation [5]. This implies

reorganization of the actin cytoskeleton and conversion of physical signals, such as contractile

forces or extracellular matrix perturbations into chemical signals that results in peptide

synthesis. In particular the FAK/Src pathway seems to be involved in the regulation of FA

turnover [18]. FAK (-/-) fibroblasts are deficient in both force–induced FA growth and cell

response to substrate rigidity [23]. Recently receptor tyrosine phosphatases alpha (RPTP-

alpha) which is involved in activation of src family kinases is necessary for FA reinforcement

of αv β3 integrin cytoskeleton connections [24].

Tensions are essential for FA maintenance and development [25-27]. This is reached by using

the contractile actomyosin apparatus. The actin filaments are highly dynamic structures

distributed over the whole cell body. Those filaments are in contact with the myosin motor-

proteins. Activating myosin by phosphorylation causes filament sliding and pre-stress

generation. The inhibition of the myosin II-driven contractility causes a reduction in the

formation of new focal adhesions, and the destabilization of the existing ones [25-27]. The

reduction of myosin II-driven tension by substances such as ML-7, BDM, KT5926 or by the

overexpression of peptides like caldesmon (inhibits actin-dependent myosin II ATPases

activity), brings about a rapid decrease of the focal adhesion size. The block of contractility

leads to complete dissolution of focal adhesions [25-27].

The formation and stability of force-dependent focal adhesion are not only due to the cell’s

own contractile system. The stimulus can also be applied from outside [28-29]. Indeed,

external force applied to focal adhesions can effectively substitute cell-generated forces even

in chemically relaxed cells. It was demonstrated that focal adhesions may be stimulated to

grow even in relaxed cells treated by BDM or transfected with caldesmon by pulling on single

cells using a micropipette [29]. Mechanical force can also be applied to micron-sized beads,

coated with fibronectin or other extracellular matrix proteins attached to the dorsal cell

Introduction

8

surface. Without application of external force, these beads move centripetally, driven by

retrograde actin flow [30]. However, trapping of a bead with laser tweezers directs the force

produced by the flow to the integrin-mediated contact at the interface between the bead and

the cell surface. This force was shown to strengthen the transmembrane link between the bead

and the actin cytoskeleton, which further increases the flow-driven force applied to the bead

and eventually allows it to escape from the trap [31]. This process, termed reinforcement,

seems analogous to the process of focal adhesion assembly and similarly includes recruitment

of the new vinculin molecules into the adhesion plaque [32].

An additional important factor is the mechanical nature of the underlying substrate. When

cells are attached to a soft flexible substrate, which can be easily deformed, the tension acting

on the adhesion plaques may be smaller than the force needed to sustain the adhesion site and

the attached stress-fibers. Consequently, the typical dimensions of focal adhesions formed

with such substrates are considerably smaller than those formed following attachment to a

rigid surface [33]. This ability to discriminate between soft and rigid substrate enables cells to

become oriented [34].

1.4 Focal adhesion formation Focal adhesions (FAs) evolve from “dot-like” structures, smaller than 1 µm in diameter

adhesion sites, termed focal complexes (FXs)[5]. They are nascent integrin- mediated

adhesions in protrusive lamellipodia developed in a Rac dependent manner (figure 1-5) [18;

35-36]. It was demonstrated that the Rac-GTPase gets recruited to the ECM attached

lammelipodia [37] and influences focal complex (FX) formation in three possible ways.

GTPases as Rac are membrane linked peptides which get activated by binding the nucleotide

guanosine- tri-phosphate (GTP). The autocatalytic transformation of GTP to Guanosine-di-

phosphate (GDP) deactivates this peptide. Rac activates the phosphatidyl inositol phosphate

kinase (PIP 5-kinase) which generates phosphatidyl inositol 4-5 bisphosphate (PIP2).

The membrane-associated inositol variant has the ability to uncap existing actin filaments

near the membrane and activate FA molecules such as talin and vinculin [38]. It is believed

that talin is involved in sequestering/ activating integrins. Furthermore, the GTPase Rac

initiate the WAVE/SCAR (peptides from the Wiskott Aldrich syndrome protein or WASP

family) peptide complex formation via the insulin receptor substrate p53 (IRSp53) that in turn

activates the actin nucleator Arp2/3 [39-40]. The serine-threonine kinase PAK is also

activated by Rac [41]. PAK promotes actin polymerization via LIM-Kinase, a kinase which is

deactivating ADF/Cofilin, an actin depolymerization factor. This prevents actin filament

Introduction

9

dissolution in the lamellipodia. Further, PAK gets localized to the membrane after the cell

attaches to ECM molecules. Here it associates with integrins, possibly via paxillin and the

paxillin kinase linker p95PKL [42].

Figure 1-5: Schematics of the Rac dependent focal complex formation (A) and the Rho dependent maturation to focal adhesions (B). Rho signalling is responsible for generating the appropriate tension generation which is necessary for recruiting other actin linked integrins for focal adhesion maturation.

The transformation from FX to FA is force dependent. Experiments using EGFP β3 revealed a

myosin-dependent recruitment of integrin αv β3 during FA maturation [43-44]. In addition,

myosin-driven force extracts molecular complexes consisting αv β1 integrins and tensin from

the focal contact [45-46]. These structures known as fibrillar adhesions move centripetally

and transmit the myosin driven forces to fibronectin molecules [18]. Integrin signalling also

activates Rho kinase, but the mechanism is not clear [5]. Rho coordinates the force-dependent

growth and maturation of focal complexes to focal adhesions [35-36]. It is known that the rho

targets ROCK (Rho associated kinase) and the formin homolog peptide Dia 1 (Diaphanous 1

peptide) are necessary for FA formation [5; 18]). ROCK is known to activate myosin-II by the

phosphorylation of the myosin light chain (MLC) [47]. Inhibition of ROCK function by

chemical inhibitors or dominant negative mutants prevents the formation of FA contacts [29;

48], but not of focal complexes. Dia 1 is also involved in actin cytoskeleton regulation.

Experiments suggest Dia 1 promotes actin polymerization by targeting profilin and Arp2/3

activation. It was also demonstrated that Dia 1 targeting microtubules to the FA [49].

Introduction

10

The focal adhesion mechanosensor is probably not a unique molecular machine but a

prototypic device, which can have analogs in other cellular systems such as adherens junction

complexes and the Z-disc-complex in striated muscle. Focal adhesions have the features to act

as mechanosensors, probe the physical environment and activate certain signal cascades. This

capacity to convert physical perturbations into biochemical signals is based on the existence

of structural and signalling components in the adhesion plaque. However, some of the basic

functions of the mechanosensing apparatus are still unknown. It is still an enigma which

molecules in the focal adhesions sense and transmit the force [5-6; 18]. There are several

promising candidates to act as mechanosensor such as vinculin [20]. For example, blocking

the Ca2+ signalling in fibroblasts with Gd2+, which regulates local traction forces, correlates

with the reduction of vinculin at the focal adhesions which resulted in impaired cell migration

[50]. Furthermore, fluid shear measurements with NIH 3T3 cells revealed that adhesion

strengthening is accompanied by a 300 % increase in vinculin and a 90 % increase in

recruitment of talin to adhesion structures, whereas integrin levels were unchanged [51].

1.5 Vinculin, a key player of the focal contact The 116 kDa vinculin with its 95 kDa head- and 30 kDa tail domain is kinetically one of the

first proteins in FA formation [16]. As demonstrated in figure 1-6, it is available in two states,

the open and the closed conformation [20]. In the closed or autoinhibited state the vinculin-

head keeps its tail pincer-like in place. Activating vinculin frees the binding sites on the tail

for paxillin, actin and phospholipids [19-20]. Several experiments revealed that vinculin is

only in the open state localized at the FA [52-53]. For these experiments, a vinculin FRET

construct with a yellow fluorescent protein (YFP) label in the prolin rich domain and a cyan

fluorophor (CFP) at the C-terminus was developed. The FRET ratio of these constructs was

decreased in the FAs in comparison to the cytosol. This suggests that the recruitment of

vinculin to the focal adhesions is linked with a conformational shift which displaces the

vinculin-head from the tail [52]. Additional studies with this FRET construct revealed that

talin together with a second binding partner (actin) have the ability to displace the vinculin-

head from the tail [54]. Talin interacts with the vinculin-head region (residues 1-258) that

lowers the affinity between the head and tail domain [55-57]. The microinjection of these

vinculin binding sites specifically targets vinculin in cells, disrupting its interactions with talin

and disassembling focal adhesions [57]. Besides talin, alpha-actinin shows also vinculin

activation. Alpha-actinin binds at the same position at the vinculin-head (residue 1-258) as

talin [57-58]. It was also reported that acidic phospholipids such as phosphatidyl-inostiol 4-5

Introduction

11

bisphosphat (PIP2) can cause a similar conformational shift and activate vinculin [59-61].

Experiments with acidic phospholipids incubated with the vinculin-head and the vinculin-tail

domain demonstrated that PIP2 vesicles have the ability to displace the head from the tail [59].

Additional in vitro experiments could demonstrate that the 30kDa vinculin-tail binds to

negatively charged lipid vesicles [60]. Johnson and co-workers incubated the 30kDa tail and

the 95kDa head fragment separately with phosphatidyl-inositol (PI) vesicles. Only the

vinculin-tail could be found together with the PI-vesicles under physiological ionic

conditions. The vinculin-head region showed no lipid interaction potential.

Cells deficient of vinculin are still able to form focal contacts but spread poorly on ECM-

coated surfaces [62-65]. They are more motile and less adhesive than wildtype cells [62; 64].

Furthermore vinculin (-/-) cells showed a higher level on tyrosine phosphorylated proteins

localized in the focal adhesions [64]. Stable retransformation of the vinculin-tail in F9 mouse

carcinoma vinculin (-/-) cells lead to improved adhesion and decreased cell motility compared

to F9 vinculin (-/-) cells [66]. Reintroduction of the complete vinculin molecule rescued the

wildtype phenotype.

Figure 1-6: Schematic of the vinculin molecule in the closed/auto inhibited (a) or the open/active (b) conformation according to Critchley [19]. About 8 % of intracellular vinculin are necessary to recover the wildtype phenotype [66].

Magnetic twisting, magnetic tweezer and atomic force microscope (AFM) measurements of

mouse F9 vinculin (-/-) cells showed softer behavior of vinculin knockout cells compared to

Introduction

12

F9 wildtype cells, suggesting that vinculin acts as an intracellular mechano-coupler [63; 65;

67-69]. It is also believed that phosphorylation of vinculin is important for the mechano-

coupling function of vinculin [70-72]. Removing the phosphorylation site on position Y822

causes an up-regulation of p-ERK which resultes in the reduction of cell migration [70].

Furthermore, c-src-dependent vinculin phosphorylation on position Y100 and Y1065 affects

cell spreading and migration, indicating that the phosphorylation of vinculin stabilizes the

active or open conformation [71]. It was further demonstrated, that phospholipid interaction

of vinculin modifies its c-src- dependent phosphorylation [73-74]. In the presence of acidic

phospholipids vesicles such as phosphatidyl-inositol (PI), the phosphorylated fraction of

vinculin was elevated more than 10 fold compared to vinculin in absence of lipid vesicles

[73].

1.5.1 Lipid binding of vinculin

Since it is known that signal transduction, vesicle trafficking, retroviral assembly, and other

central biological processes involve the directed binding of proteins to membranes,

researchers started to try to elucidate their mechanisms and functions [38]. Soluble proteins

may associate with membranes through well-defined structural domains such as amphipathic

helices and/or unstructured motifs that interact through non-specific electrostatic and apolar

interactions. Post-translational modifications, such as myristylation or palmitoylation, may

also play critical roles in regulating membrane association of peptides. Many cytoskeleton-

associated proteins such as alpha-actinin, talin, Arp2/3, CapZ and vinculin interact, at least

transiently, with lipid membranes [38; 75].

Three regions on the 30 kDa tail domain of vinculin have been identified as candidates for

lipid-binding: residues 935–978, 1020–1040 and 1052–1066 (figure 1-7) [76-78]. Residues

935–978 and residues 1020–1040 were identified as amphiphatic alpha-helices by an

algorithm that recognized highly hydrophobic or amphiphatic amino acid segments with

alpha–helical character, whilst discriminating between surface-seeking and transmembrane

configurations [38; 76]. Johnson and co-workers experimentally confirmed the computational

prediction. They incubated several peptides of the vinculin-tail with radioactive-labeled

phosphatidylserine (PS) lipid vesicles. The fragments of residues 916–970 inserted into the

hydrophobic core of lipid vesicles. Crystal structure analysis revealed an alpha-helical

secondary structure for this lipid binding site [55; 78; 79]. The residues 1052–1066, which

also have lipid interaction potential, remained unstructured during crystallization. Differential

scanning calorimetry (DSC) together with circular dichroism (CD) spectroscopy

Introduction

13

measurements showed that this lipid anchor region of vinculin has the potential to insert into

the hydrophobic core of lipid vesicles consisting of dimyristoyl-L-α-phosphatidylcholin

(DMPC) and dimyristoyl-L-α-phosphatidylglycerol (DMPG) whilst forming a beta-sheet [80].

Pull-down assays with artificial lipid membranes demonstrated that the vinculin-tail variant,

lacking the last 15 amino acids (VtailΔC) was not able to interact with acidic

phosphatidylserine (PS) or phosphatidylinositol (PI) vesicles under physiological conditions

compared to full length vinculin-tail (Vtail) [78; 81].

Figure 1-7: The vinculin-tail with its three potential lipid binding sites (colored in gold), obtained from the crystal structure of chicken vinculin (PDB 1ST6) [55]. It was also shown that the lipid interaction of the vinculin-tail is mainly driven by electrostatic

interaction between the lipid vesicle and peptide surfaces. As demonstrated by additional pull-

down experiments, the mutation of six surface exposed basic residues on vinculin-tail to

glutamine (Q) in helix 3 (K952, K956, R963, R966) and the C-terminal lipid anchor

(RK1060/61) also significantly reduces the interaction potential with negatively charged

vesicles [82]. Furthermore, it was reported that cells transfected with a vinculin variant

including this 6 point mutations which cause lipid binding deficiency (vinculin-LD) showed a

reduced focal adhesion turnover rate and impaired cell adhesion on different extracellular

substrates such as collagen, laminin or fibronectin. The cell motility in vinculin-LD cells was

also decreased [82]. Cells expressing vinculin without the lipid anchor (vinculinΔC) showed

Introduction

14

the same reduced FA adhesion turnover rate and decreased cell motility [81]. These results

imply that only the membrane interaction of vinculin´s lipid anchor influences cell

mechanical behavior.

1.6 Aims Lipid binding of vinculin influences the focal adhesion turnover rate, which results in

impaired cell adhesion and migration [82-81]. Pull-down assays with artificial lipid

membranes consisting of phosphatidylserine (PS) or a mixture of 40 % phosphatidylinositol-

4,5-bisphosphate (PIP2) and 60 % phosphatidylcholine (PC) revealed that in contrast to Vtail,

a variant lacking the last 15 amino acids (VtailΔC), does not interact with vesicles of these

compositions [78; 81]. But whether the last 15 residues are directly involved in lipid

interaction was not determined. Crystal structure analysis together with CD spectroscopy

analysis revealed that the integrity of the helical bundle is ensured in both vinculin-tail

variants, even in the absence of the head domain [78; 81]. In this work, we want to clearify

the question wether the lipid anchor is directly or indirectly involved in lipid membrane

interaction. Furthermore the impact of that interaction due to cell mechanical behavior was

investigated. To determine the membrane interaction of the lipid anchor region, differential

scanning calorimetry (DSC) measurements were performed, using a peptide which represents

vinculin´s unstructured C-terminal arm. In additional molecular dynamics simulations we

calculated possible secondary structures for the peptide. CD-spectroscopy as well as NMR-

measurements were performed to verify the calculated secondary structure and calorimetric

characterization of the peptide. Furthermore, the influence of the lipid anchor due to cell

mechanical behavior was determined using a vinculin variant lacking the last 15 residues

(vinculinΔC). Transient with EGFP-labeled vinculin∆C transfected MEF vinculin(-/-) cells

were examined using a magnetic tweezer setup with force feedback control. Forces between

0.5 and 10 nN were applied in a log-step force protocol to investigate the stiffness of the cells

as well as the stability of the FN-integrin-actin linkage. The actomyosin- driven contractile

forces of MEF-vinΔC cells were obtained by 2D-traction microscopy measurements. Further,

the numbers of FA as well as the turnover rate of the different vinculin constructs were

determined.

Materials & Methods

15

2 Materials and Methods 2.1 Materials 2.1.1 Chemicals and enzymes

All chemicals were of per analysis (p.A) quality and purchased from Sigma (Deisenhofen),

Merck (Darmstadt), Fluka (Neu-Ulm), Difco (Hamburg) and Invitrogen (Karlsruhe). All

enzymes were from the following companies unless otherwise specified, New England

Biolabs (Bad Schwalbach), Stratagene (Heidelberg), Boehringer (Mannheim), Amersham

(Freiburg), Invitrogen (Karlsruhe), and Promega (Mannheim).

2.1.2 Cell culture medium and plastic ware

Cell culture media and additives such as fetal bovine serum (FBS) and the antibiotics

Penicillin and Streptomycine were purchased from Gibco (Karlsruhe) and Sigma

(Deisenhofen). Plastic ware was obtained from Corning (Kaiserslautern), Eppendorf

(Hamburg), Becton and Dickenson (BD-Bioscience, Heidelberg), Greiner (Frickenhausen)

and Nunc (Langenselbold).

2.1.3 Oligonucleotides

The Oligonucleotides employed in cloning and sequencing reactions were obtained from

MWG-Biotech (Ebersberg). The primers for sequencing cloning and mutagenesis are listed in

Table 2.1, and primers for cloning and mutagenesis are listed in Table 2.2. The restriction

sites present in the primers are specified.

Number Name Target sequence Sequence 5’ to 3’ Primer P1 Seq_pEGFP_for1 pEGFP, start in front of

EGFP TGGGCGGTAGGCGTGTA

Primer P2 Seq pEGFP rev1 pEGFP, start at the end of EGFP

CAGGTTCAGGGGAGGT

Primer P3 Seq_EGFP_rev pEGFP, start in the middle of EGFP

GAGCAGGATGGGCAC

Primer P4 Seq_pcDNA_rev pcDNA3.1/Hygro, behind MCS

GGTCAAGGAAGGCACG

Primer V1 Seq vinc rev1 mvinculin, start at 703bp-720bp

TTCAGCACTCATCTTTTC

Primer V2 Seq vinc for1 mvinculin, start at 551bp-568bp

ACCAGGAACACCGTGTG

Primer V3 Seq vinc rev2 mvinculin, start at 1953bp-1970bp

GTCGATTTATTGGCAGTA


GATACTACAACTCCTATC


CTCAGGAGCCTGACTTC

Tabelle 2-1: List of sequencing primers used during this work; bp is an abbreviation for “base pair”.

Materials & Methods

16

Number Name Purpose Sequence 5’ to 3’ Primer 1a pEGFP, for, Nhe I

amplification of pcEGFP-N2 vector

GCGCTAGCGCTACCGGTCG

Primer 1b pEGFP, rev, Xho I

amplification of pcEGFP-N2 vector

GCCTCGAGCTTAAGGAGTCCGGACTTGTACAG

Primer 2a mvinc_N, for, Afl II

amplification of vinculin/ vinculinΔC for pcEGFP-N2

GCCTTAAGATGCCGGTGTTTCACACG

Primer 2b mvinc_N, rev, Xba I

amplification of vinculin/ vinculin-tail858-1066 for pcEGFP-N2

GCTCTAGACTACTGGTACCAGGGAGT

Primer 2c mvinc0.5, for

gene SOEing of vinculin AATAGGGAAGAGGTATTTGAT

Primer 2d mvinc0.5, rev

gene SOEing of vinculin ATCAAATACCTCTTCCCTATT

Primer 2e mvincdelC, rev, Xba I

amplification of vinculin ΔC/ vinculin-tail858ΔC for pcEGFP-N2

GCTCTAGACTAATCTGTTCGGATTTTGATTGA

Primer 2f Vinculin-tail, for, XbaI amplification of vinculin-tail858/ vinculin-tail858 ΔC for pcEGFP-N2

GCCTTAAGATGCTGGCTCCTCCTAAGCCA

Primer 3a Vinculin Y1065F, for Mutagenesis of the C-terminal src phos-phorylation site

CAGAAAGACTCCCTGGGAACAGTAGTCTAGAGGG

Primer 3b Vinculin Y1065F, rev Mutagenesis of the C-terminal src phos-phorylation site

GTCTTTCTGAGGGACCCTTGTCCATCAGATCTCCC

Table 2-2: Primers for cloning and mutagenesis; Note, SOEing is an abbreviation for “splicing by overlap extension”

2.1.4 Vectors

The pcDNA 3.1 Hygro/(+) vector (Invitrogene, Karlsruhe) was used for generating an

enhanced green fluorescent protein (EGFP) vector carrying a hygromycin resistance gene.

The EGFP cassette was amplified via a polymerase chain reaction (PCR) using a pEGFP-

actin vector from BD Bioscience (Heidelberg). The vinculin, vinculinΔC, vinculin-tail,

vinculin-tailΔC and vinculinY1065F were subcloned into the generated pcEGFP-vector. The

vinculin-LD, vinculin-CT and vinculin-H3 were cloned into a pEGFP-C2 (Clontech,

Heidelberg) vector and kindly provided by Dr. Wolfgang Ziegler (University of Leipzig).

2.1.5 Bacterial cultures and cell lines

The cloning was performed in the E. coli strain DH5α with the following genotype: F-,

φ80dlacZΔM15, Δ(lacZYA-argF)U169, deoR, recA1, endA1, hsdR17(rk-, mk+), phoA, supE44,

λ-, thi-1, gyrA96, relA1. The strains were grown in LB-medium, and LB-agar was prepared

according to the manufactor´s describtion. Antibiotics such as Ampicillin (100µg/ml) or

Kanamycin (30µg/ml) were used for liquid medium and plates. The cell lines employed in

this study were mouse embryonic fibroblasts (MEF) isolated from vinculin knock out mice

Materials & Methods

17

(MEF-vin(-/-)).The unaffected control fibroblasts (MEF-wt) were kindly provided by Dr. E.D.

Adamson (La Jolla, CA).

2.1.6 Antibodies

Antibodies were mainly employed for immuno detection in Western Blots and in vivo

imaging of cells by immuno fluorescence. The antibodies used in immunoblots and

immunofluorescence are listed in Table 2.3. The secondary antibodies and toxins used in

these approaches are listed in Table 2.4.

Description Antigen Company Organism Clonal type Anti-vinculin Vinculin-head Sigma Mouse Monoclonal

Anti paxillin Paxillin BD Bioscience Mouse Monoclonal

Anti-Actin Beta-actin Sigma Mouse Monoclonal

Tabelle 2-3: List of primary antibodies used in immuno blots and immuno fluorescence.

List of secondary antibodies used in immuno blots and immuno fluorescene. 2.2 Methods 2.2.1 Methods in molecular biology

2.2.1.1 Quantification of Desoxyribonuclein acid (DNA) DNA concentrations were determined photometrically. The absorption maximum of DNA is

at 260 nm. An absorption value of 1 at the given wavelength of 260 nm is equal to 50 µg/ ml

DNA. The concentration of DNA in a diluted solution is calculated as follows:

OD260

× dilution factor × 50 μg/ ml = Concentration of DNA μg/ ml.

2.2.1.2 DNA electrophoresis The electrophoresis of DNA is an essential technique in molecular biology relying on the

negative charge of DNA for size separation in a sieving matrix. Traditionally, electrophoresis

is performed using agarose (an extract of seaweed) and a Tris(hydroxymethyl)aminomethane

(Tris) buffering solution as described in Sambrook et al [83]. Agarose gels of 1-2 % were

Description Antigen Company Organism Clonal type Alexa FluorTM545 phalloidin

Amanita phalloides Molecular Probes - Monoclonal

HRP-linked Antibody Anti-Mouse IgG Amersham From sheep Monoclonal TRITC Anti-Mouse IgG Jackson

ImmunoResearch Donkey Monoclonal

FITC Anti-Mouse IgG Jackson ImmunoResearch

Donkey Monoclonal

Table 2-4:

Materials & Methods

18

used. The gels were prepared using 1 x Tris-acetate (TAE) buffer pH 8 (40 mM Tris, 20 mM

acetate and 1 mM Ethylendiamintetraacetat ((EDTA)). Between 0.5-1 % of Ethidium

Bromide was added to the gel and the running buffer for visualization of the DNA. DNA

sample buffer (0.25 % Bromphenol blue, 0.25 % Xylencanol, 30 % Glycerin) was added to

the DNA. The DNA was separated at 120 mA. As standard, the 1 kb DNA ladder from New

England Biolabs (NEB, Frankfurt/ Main) was used.

2.2.1.3 Restriction endonuclease digestion of DNA Restriction enzymes were purchased from NEB (Frankfurt/ Main), and the reaction was set up

using the appropriate amount of enzyme in units as recommended by the manufacturer. For

preparative digestion, 2.0-5.0 μg DNA was used. For analytical purposes 0.2 – 0.3 µg DNA

was used. The reaction was performed in the suitable 1 × buffer provided by NEB (Frankfurt/

Main). The reaction mix was incubated for 1-2 hrs at 37 °C, if not otherwise indicated, as

recommended by the manufacturer. The digestion assays were separated and monitored on

agarose gels as described under 2.2.1.2.

2.2.1.4 Ligation The ligation reaction covalently links the phosphordiester bonds between two fragments of

DNA using the ligase enzyme. DNA ligase from T4 Bacteriophage (Boehringer, NEB) was

employed in this work to link vector DNA with the insert DNA. Appropriate molar ratios

(usually 1:3) between vector and insert DNA were used in a 15 μl reaction with 1-2 units

ligase and 1 × T4 ligation buffer. The reaction mixture was incubated at room temperature for

1 hour or overnight at 12 °C, respectively.

2.2.1.5 Preparation of chemical competent DH5α The bacteria were prepared according to a modified protocol of Hannahan [84]. A pre-culture

of DH5α was grown over night in Luria-Bertani (LB) medium. 100 µl of that culture was

transferred into 50 ml LB-medium and inoculated at 37 °C and 220 rpm until an OD600 of 0.4-

0.5 was reached. Prior to centrifugation (2000 g, 7 min at 4 °C), the culture was incubated for

10 minutes on ice. After centrifugation the bacteria pellet was resuspended in 15 ml Calcium/

Magnesium (CM) buffer (50 mM CaCl2, 50 mM MgCl2, 10 % Glycerol), followed by another

incubation step on ice for ten minutes. The bacteria were spun down for 5 minutes at 2000 g

and 4 °C. The pellet was dissolved in 3.5 ml CM and incubated again on ice for 10 minutes.

After the addition of 125 µl Dimethylsulfoxid (DMSO; Sigma, Deisenhofen), the cells were

kept for additional 5 minutes on ice. The bacteria solution was aliquoted and stored at -80 °C

after adding a final volume of 125 µl DMSO followed by 5 minutes of incubation on ice.

Materials & Methods

19

2.2.1.6 Transformation of competent cells The ligated DNA was transformed in chemically competent cells. The E. coli strain DH5α

was used for this purpose. Cells were incubated for 30 minutes on ice with the vector DNA. A

heat shock at 42 °C was applied for 90 s followed by 45 minutes incubation in 0.6 ml LB-

medium at 37 °C and low agitation for cell regeneration. The cells were spun down by

centrifugation at 8,500 g, resuspended in 100 μl medium and plated onto LB agar medium

containing appropriate antibiotics.

2.2.1.7 Polymerase chain reaction (PCR) PCR is a technique employed for rapid in vitro amplification of DNA [83]. The amplification

is done using a temperature resistant DNA polymerase, free nucleotides, a primer or the

starter sequence which specifically binds to the template strand. Three major steps are

involved in the reaction: the denaturation of the template DNA at high temperature (95 °C),

annealing or primer binding to the template strand at 55 °C – 60 °C and finally an extension

step at 72 °C for the actual amplification. For exponential increase of amplified DNA, the

steps are repeated for 25 - 30 cycles. DNA polymerase from thermophilic bacterias such as

Thermus aquaticus (Taq) does not have the 3’ to 5’ exonuclease or “proof reading” properties

to prevent mutations. Therefore, this enzyme was only used for the amplification of small

fragments (<1000 bp) such as the EGFP cassette or the vinculin-tail. The DNA polymerase

from Pyrococcus furiosus (Pfu) was used for amplifying whole vinculin and vinculinΔC

cDNA fragments >1000 bp. The standard PCR reaction mixture contained template plasmid

DNA (50 ng concentration), template specific primers (100 pmol), free nucleotides or dNTP’s

(0.3 mM), MgCl2

(0.5 – 2.5 mM), 1 × PCR buffer and 0.5 - 5.0 units of DNA polymerase

(Taq/Pfu) in 25 – 50 μl reaction volume.

2.2.1.8 Site directed mutagenesis The two step site directed mutagenesis protocol according to Wang and co-workers was

employed to introduce the Y1065F mutation in vinculin [85]. Oligonucleotide sequences

(primer) with the respective bases exchanged were used. In a first step, two separate PCR

reactions, one for the coding and one for the non-coding strand, were performed using only

one mutagenesis primer for each reaction. This first step ensures the efficient introduction of

point mutations. After 5 separate cycles, the PCR assays were combined for additional 25

cycles. The PCR reactions contained the same reagents as described in 2.2.1.7. The PCR

products were digested with the restriction enzyme Dpn I at 37 °C for one hour. Dpn I

specifically digests the methylated parental DNA used as template and retains only the newly

Materials & Methods

20

synthesized DNA. The reaction mixture was then transformed in DH5α competent cells and

plated onto LB agar medium with an appropriate antibiotic selection.

2.2.1.9 Purification of plasmid DNA from bacteria Plasmid DNA amplified in DH5α cells was purified by the modified alkaline lysis method

[83]. Plasmid mini-preparations were performed using the Nucleo Spin plasmid extraction kit

from (Macherey-Nagel, Düren); plasmid maxi-preparations were performed using Qiagen

columns (Qiagen) according to the manufacturer’s recommendation.

2.2.1.10 DNA sequencing DNA sequencing reactions were performed by MWG Biotech according to Sanger [86]. For

that purpose, 1 µg of the plasmid DNA for each reaction together with 10 pmol of the

appropriate sequencing oligonucleotide were sent to the company.

2.2.1.11 DNA extraction from agarose gels DNA was recovered from agarose gel slices with the QIAex Gel Extraction Kit (Qiagen) as

suggested by the manufacturer.

2.2.1.12 Cloning strategy – EGFP expression vector The EGFP expression system was generated as described in figure 2-1. For the generation of

an EGFP expression vector with a hygromycin resistance gene, a pcDNA 3.1 Hygro/(+)

vector (Invitrogene, Karlsruhe) was used. The EGFP cassette was amplified from a pEGFP-

Actin Vector (BD Bioscience) by a PCR reaction and subcloned N-terminally to the multiple

cloning site (MCS) of the pcDNA 3.1 Hygro/(+) vector via Nhe I/ Xho I.

Materials & Methods

21

Figure 2-1: Cloning strategy for the generation of the pEGFP expression system.

2.2.1.13 Cloning strategy – vinculin full length and tail domain constructs The full length mouse vinculin cDNA was kindly provided by Dr. E.D. Adamson (La Jolla,

CA) and Dr. W.H. Ziegler (Leipzig). Sequencing analysis identified several mutations in

these clones. There were three mutations which resulted in amino acid replacements in the

vinculin cDNA provided by Dr. Adamson (F633S, V828E, L936P). The vinculin cDNA

obtained from Dr. Ziegler showed two mutated residues in the vinculin-head region (V246A,

A486V). All those mutations were considered critical for the protein function. The following

strategy was developed to obtain a vinculin cDNA without any mutations. The first part of the

vinculin cDNA from Dr. Adamson was fused with the unaffected part of Dr. Ziegler’s cDNA

using a gene SOEing protocol as described in figure 2-2. In a first PCR reaction, vinculin part

I from Dr. Adamsons cDNA and vinculin part II from Dr. Ziegler’s cDNA were amplified

by overlapping the complement ends. In a second PCR, the products of the first two separate

reactions were combined. They aligned on the complement ends which gave the PCR together

with primer 2a and 2b the possibility to synthesize the strain.

Materials & Methods

22

Figure 2-2: Cloning strategy for the vinculin cDNA. Black arrows mark the positions of the mutated codons. The generated vinculin cDNA was cleaved into the constructed pcEGFP-N2 vector (see

2.3.1.10) using the restriction sites of Afl II and Xba I. The fused vinculin cDNA was used as

template for the generation of vinculinΔC (1-1052), vinculin Y1065F, vinculin-tail (858-

1066) and the vinculin-tailΔC (858-1066) cDNA. The endproduct was also verified by

sequencing.

2.2.2 Cell biology methods

2.2.2.1 Cell culture The mouse embryonic fibroblasts (MEF-wt) and vinculin null fibroblasts (MEF-vin(-/-)) were

grown in Dulbecco´s modified eagle medium (DMEM), 1 g/ l glucose (Biochrom, Berlin)

with 10 % FCS (Biochrom, Berlin) and 1 % Penicillin/ Streptomycin antibiotics at 37 °C and

5 % CO2. For liquid nitrogen storage, 80 µl of DMSO was added to 2 x 105 – 5 x 105 cells

dissolved in 1 ml DMEM with additives.

2.2.2.2 Transient transfection of cells Transfections were carried out with LipofectaimeTM (Invitrogen, Karlsruhe) according to the

manufacturer’s protocol. In brief, 4 μg DNA and 10 µl LipofectaimeTM were dissolved in 250

µl pure DMEM, respectively. After 5 min incubation at RT, the dissolved DNA and

Materials & Methods

23

lipofectaimeTM solutions were combined and incubated for additional 20 min at RT. The

reaction assay was pipetted onto 2 x 105 cells plated in a dish of 3.2 cm diameter

approximately 24 h prior to transfection. The cells were incubated for 4 h at 37 °C with a total

volume of 2.5 ml pure DMEM. After 4 h of incubation, the pure DMEM was replaced against

DMEM with additives as described under 2.2.3.1. The constructs were allowed to express for

16-24 hours prior to further procedures.

2.2.2.3 Culturing of cells on the cover slips for microscopic analysis 1 x 105 – 2 x 105 of transfected or non-transfected cells were seeded onto 15 mm glass

coverslips that had been pre-treated with a mixture of 70 % ethanol. The ethanol on the

coverslips was allowed to evaporate under laminar flow prior to coating. Glass coverslips

were coated with 25 μg/ml fibronectin (Roche, Penzberg) for at least 2 hours at room

temperature or overnight at 4 °C. Uncoated coverslips were used for focal contact

determination.

2.2.2.4 Fixation and permeabilization Cells (transfected and non-transfected) were fixed with 3 % para-formaldehyde (PFA) in PBS

for 15 minutes. After fixation, the cells were permeabilized with 0.1 % Triton X-100 for 5

minutes. The cells were thoroughly washed three times with 1 × PBS and stored at 4 °C until

they were stained.

2.2.2.5 Immunolabelling, fluorescence microscopy and image processing To avoid non-specific interactions, cells were blocked prior to antibody staining with 1 %

bovine serum albumin (BSA, Sigma, Deisenhofen) in phosphate buffered saline (PBS) for 20

minutes. Thereafter, the coverslips were incubated with 50 μl of primary antibody solution on

parafilm for 1 - 2 hours at room temperature followed by 3 washes in 1 × PBS. Staining with

secondary antibody with or without the addition of fluorescently coupled phalloidin was

carried out for 1 hour at room temperature. Coverslips were mounted in 30 μl Mowiol on

microscope glass slides, dried and stored in the dark at 4 °C until analysis. Cells were

observed on a LEICA microscope DMI6000 equipped for fluorescence and phase contrast

using 20 x/0.4NA-, 40 x/0.6NA- as well as 63 x/1.3NA objectives. The immersion oil was

obtained from Leica. Data were acquired by a CCD camera (ORCA ER, Hamamatsu) driven

by the WASABI software package. Data were stored and analyzed using the Image J

software. The cells for focal adhesion determination were examined using a Zeiss LSM 510

META confocal laser scanning microscope system with a 25 x/0.8 NA Plan-Neofluar oil

objective. Images were acquired and processed using Zeiss LSM Image browser (Zeiss, Jena).

Materials & Methods

24

2.2.2.6 Determination of the spreading area Prior to fixation (see chapter 2.2.2.3), transfected and non-transfected MEF cells were

cultured on coverslips for 1h (see chapter 2.2.2.2). The actin cytoskeleton at non-transfected

cells was stained as described under 2.2.2.5. Pictures were acquired at 20x magnification

using a fluorescence microscope. The number of cells and their spreading area computed

using a MatLab image analysis program.

2.2.2.7 Fluorescence recovery after photo-bleaching (FRAP) measurements MEF-vin(-/-) cells were transfected with EGFP-labeled vinculin and vinculinΔC constructs as

described under 2.2.3.2. 1 x 105 of the transfected cells were seeded on fibronectin-coated

(25µg/ml) glass bottom wells. The glass bottom wells with the transfected cells were placed

in the chamber on a temperature-regulated 37 °C microscope stage 30 minutes before

measurements. Time-separated confocal 8-bit images were collected using a Zeiss LSM 510

META confocal laser scanning microscope system with a 63 x Plan-Apochromat oil objective

(NA 1.4). GFP was excited with the 488 nm line of the Argon laser and reflected and emitted

light was separated using a 505 nm long pass filter (for viewing GFP emission). During

bleaching, the 488 nm laser line output was set to 100 % using 50 iterations. Prior to

bleaching, the sample cells were scanned for 60 s to determine the stability of focal contacts.

For determination of the background photobleaching caused by the line scans, at least 3

reference focal contacts and one cytosolic region were selected. The line scans were collected

for at least 5 minutes. To eliminate the background fluorescence at the cytoplasm as well as

incomplete bleaching, the offset cs(t) was calculated as follows:

)((t)c 0s tFad= (1)

where Fad represents the fluorescence intensity of the bleach field at the FA and Fcp the

fluorescence intensity of the bleach field of the cytoplasm. Time point t0 determines the start

of bleaching. The mean of the intensity of the reference focal adhesions were averaged (Fref

(t)) and the bleach function was normalized due to following equation:

0n ))()((

)()((t)F

FtctFtctF

sref

sad

−−

= (2)

The pre-bleach intensity F0 was determined over ten timepoints (tp) before starting the

bleaching according to

spref

spad

ctFctF

−

−=

)()(

F0 (3)

Materials & Methods

25

Prior to analysis, the curvatures were fitted to exponential function

)1(f(t) kte−−= α (4)

Using the MATLAB function fminsearch, whereas α represents the saturation value and k

the turnover rate. The recovery halftime was determined according to

k2lnt1/2 = (5)

2.2.2.8 Electron microscopy measurements The focal contacts in MEF cells were investigated using a transmission electron microscope

(TEM). Dependent on the density of cell compartments, electrons go through the sample and

hit the detector or were scattered on the dense structures which result in a picture. The

resolution of such a microscope is around 1Å. 3 x 105 MEF cells were seeded on a positively

charged “Melinex” plastic strip (2.8 cm x 0.5 cm) deposited in a 3.2 cm cell culture dish. The

plastic strip with the overnight attached cells was transferred into a glass dish prior to fixation.

The cells were incubated for 20 minutes with a fixative consisting of 4 % para-formaldehyde,

0.5 % glutaraldehyde dissolved in 1 M sodium-cacodylate buffer. The cells were washed

twice using 1 x PBS. The sample was sequentially incubated in ethanol solutions of 30 %, 40

%, 50 %, 60 % 70 %, 80 % 90 % and 100 %. Each step took 10 min and occurred at RT.

Afterwards the cells were treated with the London resin white (LRW; Sigma, Deisenhofen) by

incubating them sequentially for 30 minutes at room temperature with a mixture of

ethanol/LRW (1 : 1), ethanol/LRW (1 : 3) followed by 2 steps of pure LRW incubation. The

strip with the cells was embedded into a gelatine capsule using LRW. The resin was hardened

by incubating the capsule for 24 h at 60 °C. The gelatine shell was removed by a razor blade,

the LRW block splitted longitudinal to the cells and the plastic strip was ripped of as

illustrated in figure 2.3. The resin block with the cells was re-embedded in a second gelatine

capsule and filled with additional LRW resin to prevent the loss of the cells during sectioning.

After additional incubation for 24 h at 60 °C ultra-thin sections of 70-100 nm thickness were

prepared using a microtom. The sections were deposited on a copper grid. Before immuno-

labelling, the sections were blocked by a 1 % BSA solution dissolved in filtered 0.01 M PBS

(pH 7.4). Thereafter, the samples were incubated with the primary antibody solution for 1 h at

room temperature followed by 4 washes in 1 × PBS. Staining with gold-labeled secondary

antibody was carried out for 30 minutes at room temperature. After washing the samples with

filtered 0.01 M PBS, the sections were jet-washed for 30 seconds with distilled water. Finally,

the sections were incubated for 7 minutes with 4 % uranyl-acetate followed by 4 minutes of

Materials & Methods

26

incubation with lead to improve the contrast during the measurement. The samples were

investigated using a JEOL (JEM – 2000FX) TEM.

Figure 2-3: Sample preparation for TEM measuremnts. (a) Melinex plasic strip bottom, (b) Melinex plastic strip top. After the LRW hardening, the crystal was separate longitudinal to the plastic strip and the plastic strip was ripped off. The crystal part with the cells (A) was re-embedded into another gelatine capsule and after hardening used for preparation of the microsections.

2.2.3 Biochemical and biophysical methods

2.2.3.1 Bead coating

Superparamagnetic 4.5 µm epoxylated beads (Invitrogen, Karlsruhe) were coated with

100µg/ml fibronectin (Roche, Mannheim) in PBS at 4 °C for 24 h. Beads were washed in

PBS and stored at 4 °C. The Integrin receptors bind to the RGD motif, localized in FN.

2.2.3.2 Peptide synthesis

The different vinculin peptides, representing vinculin´s C-terminus were kindly provided by

Dr. Rothemund and Dr. W.H Ziegler (University of Leipzig). The peptides, listed in Table 2.4

were generated according to the method of Marryfield [87]. Peptides were synthesized by

coupling the activated backbone carboxyl group of one amino acid to the backbone amino

group of another residue. The possibility of unintended reactions is obvious, thus protecting

groups are usually necessary.

Vinculin Peptide (residue 1045-1066) Sequence

C-terminal arm (residue 1045–1066) IKIRTDAGFTLRWVRKTPWYQ

C-terminal arm R1060, K1061 to Q IKIRTDAGFTLRWVQQTPWYQ Table 2-5: Vinculin´s C-terminal peptides used in this work

2.2.3.3 Lipid vesicle preparation Multilamellar vesicles (MLVs) were prepared as follows: lipid stock solution consisting of

Dimyristoyl-L-α-phosphatidylcholin (DMPC) and Dimyristoyl-L-α-phosphatidylglycerol

Materials & Methods

27

(DMPG) were purchased from Avanti Polar Lipids (Birmingham, AL). Mixtures of

crystalline DMPG and DMPC were dissolved in chlorophorm/methanol, 2 : 1 (v/v). To form a

dry lipid film on the glass wall of an Erlenmeyer-flask, the solvent was evaporated by

nitrogen followed by a vacuum desiccation step for at least 2 h. The lipid film was suspended

in a lipid buffer containing 20 mM HEPES (pH 7.4), 2 mM EDTA, 5 mM NaCl and 0.2 mM

DTT for generating multilamellar vesicles. The vesicles equilibrated overnight at 35 °C.

For the CD-spectroscopy measurements small unilamellar vesicles (SUVs) were prepared.

For this purpose, the lipid film in the Erlenmeyer-flask was dissolved in 10 mM potassium

phosphate buffer (pH 7.4) and sonified for 10 minutes after the equilibration step.

2.2.3.4 Differential scanning calorimetry (DSC) measurements DSC is a technique to resolve conformational transitions of biological macromolecules by

energy determination. It provides an immediate access to the thermodynamic mechanism that

governs a conformational equilibrium, i.e. between folded and unfolded forms of a protein or

DNA. The theory of DSC and the thermodynamic interpretation of the experiment have been

the subject of excellent reviews {Goldmann, 2008 #953]. In brief, the DSC registers phase

transition energy of biological macromolecules and can be used to determine the heat capacity

calorimetrically. The enthalpy of an isobaric system is defined as

ΔQΔH = (6)

where, H is the enthalpy and, Q the heat energy. Furthermore, under these conditions, the heat

capacity can be described as

ΔTΔHCp = (7).

Integration of the determined heat capacity (Cp) over the change in temperature gives the

phase transitions enthalpy (H):

∫=fluid

gelpdTCΔH (8).

The DSC experiments in this work focused on the phase transition enthalpy of phospholipids

in presence and absence of peptides. The principle is described in figure 2-4. Phospholipids in

membranes can exist in a gel-like (ordered) as well as fluid-like (disordered) phase dependent

on the temperature. At the ordered phase, all acyl-residues of a fatty acid are in the

energetically favourable trans-conformation. Transferring the phospholipids into the fluid-like

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28

phase by heating up the system constantly, leads to a change in conformation and allows the

energetically unfavourable gauche conformation of the acyl residues. This results in a

disturbed package of the acyl-residues in the vesicles which is responsible for the transition to

the liquid state. The phase change is regarded as first order phase transition. The energy,

necessary for that conformational change could be determined by the specific heat capacity

(Cp), also called specific heat.

Figure 2-4: Phase transition thermogram of DMPG/DMPC vesicles. Both pre-transition and transition occur with increased temperature. TV marks the maximum temperature of the pre-transition and, TM represents the melting point for the main phase transition, where the specific heat has the calorimetric maximum. TS and TL mark the start and end points of the transition. As the high energy (gauche) conformations increase, the energetically favourable (trans) conformations decrease during phase transition [88]. Some lipids show a pre-transition phase at which the gel-like membrane changes from a

lamellar (Lβ´) to a ripple phase (Pβ´), before undergoing the transition to the fluid phase (Lα) at

the melting temperature (TM). A polypeptide with the ability to insert into a lipid membrane

interacts non-covalently with some of the acyl-residues and, therefore, prevents phase

transition for these lipids. This results in a reduced TM and a decreased phase transition

enthalpy (H).

The heating of the sample and reference solution is performed at a preset heating rate, ß = ΔT/

Δt. The principle of the DSC requires the temperature of the sample (TP) and the reference

(TR) solution to remain constant (T = TP = TR) as described in figure 2-4. To ensure an

endothermic phase transition, the sample solution must be heated to a higher temperature than

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29

the reference solution to keep both temperatures constant. The heat output for the sample (Pp)

will be higher than for the reference (PR). This is reflected in the heat capacity ΔC (T)

between sample and reference ΔC (T) = Cp - CR =ΔP (T) / ß [88].

Figure 2-5 A schematic of the DSC apparatus with the sample (1), reference (2), heater (3), insulation coat (4) and temperature observation points (5) and PP and PR the probes of the sample (P) and the reference (R) respectively [88].

A differential scanning calorimeter Q100 from TA Instruments was used for all

measurements. The peptide of vinculin’s lipid binding region and insulin (purchased from

Sigma, Deisenhofen) were dissolved in a lipid buffer (see 2.2.3.3). The lipid buffer solution

was placed in the reference cell and lipid peptide solution in the sample cell. MLVs were

used. Under sealed conditions, both solutions were heated at a rate of 0.5 °C/ min and cooled

at a rate of 1 °C/ min. The heat capacity was captured between +7 °C and +30 °C until the

equilibrium of the phase transition enthalpy was reached. A phase transition was observed at

~23 °C depending on the molar ratio of DMPG and DMPC. Data analysis was performed

using the software from Universal Analysis 2000 (TA Instruments) and Origin 7G.

2.2.3.5 Circular dichroism (CD) spectroscopy measurements Circular dichroism (CD) is a spectroscopic technique which measures differences in the

absorption of left-handed and right-handed circularly polarized light of optical active

molecules such as chiral compounds. Amino acids and so peptides are photoactive molecules.

The Cα atoms of amino acids are achiral. This leads to a disproportionated distribution of the

peptide bond electrons, which is responsible for the difference in absorption of right and left

polarized light. The conformation or secondary structure of the peptide influences this

electron distribution. This results in certain spectra for typical secondary structures as alpha

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30

helices and beta sheets. A CD spectrum is measured typically between 180 – 300 nm. The

first part is reflecting the absorption of the peptide bond (180 – 230 nm). At 240 – 300 nm,

the aromatic residues give the signal. The difference in absorbtion of left and right polarized

light is blotted as ellipticity (θ).

The 21 residue peptide of vinculin was dissolved in a 10 mM potassium phosphate buffer (pH

7.5). The concentration of the peptide was determined photometrically using an extinction

coefficient of 12,490 M-1 cm-1 at 280 nm. CD spectra were recorded on a JASCO J-815 CD

Spectrometer at 1 nm intervals over the 180 - 260 nm wavelength range using a 70 µM

protein solution. The spectra of the lipid/protein mixture were recorded at a lipid/protein

molar ratio of 40 : 1. Peptides and lipid vesicles were dissolved in 10 mM potassium

phosphate buffer (pH 7.5) and measured in a cuvette of 0.1 cm path length. Three scans were

recorded for each sample and averaged. The protein spectra were corrected by subtraction of

the spectrum of 10 mM potassium phosphate buffer (pH 7.5) or the spectrum of lipid vesicles,

respectively. The corrected protein spectra were adjusted for protein concentration and path

length to obtain the mean residue molar ellipticity at each wavelength.

CD spectra of the vinculin peptide 1045-1066 were obtained at 30 °C and smoothed with the

Savitzky – Golay algorithm. Secondary structure analysis was performed using the

CONTINLL, CDSSTR algorithms provided by DICHROWEB [89-90]. The quality of the fit

between experimental and back-calculated spectrum corresponding to the derived secondary

structure element fractions was assessed from the normalized root mean square deviation

(NRMSD).

2.2.3.6 Solid state NMR Among the biophysical techniques that allow the investigation of peptides and proteins in

bilayer environments solid-state NMR spectroscopy has proven to be a valuable tool. The

technique resolves the structure, dynamics and topology of membrane-associated

polypeptides [88]. The tilt angles of helices with respect to lipid bilayers have been

determined using static-oriented samples. It has been demonstrated that this approach has also

been shown to be suitable for the complete structure determination of membrane-bound

peptides by measuring a large number of conformational constraints. If the lipid peptide

interactions are oriented, NMR interactions are used to extract angular constraints from such

static-oriented samples. Proton-decoupled 15N solid-state NMR spectroscopy of peptides

labeled at the backbone amides with 15N has been proven particularly convenient as this

method provides the approximate tilt angle of membrane-associated helices in a direct

manner. Whereas transmembrane helical peptides exhibit 15N chemical shifts around 200

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31

ppm, sequences oriented parallel to the surface resonate at frequencies < 100 ppm (Figure 2-

6). Due to fast axial rotation of the phospholipids around their long axis the 31P chemical shift

is characterized by an averaged symmetric tensor. The singular axis (σ||) coincides with the

rotational axis, i.e., the bilayer normal. In the 31P solidstate NMR spectra of pure liquid

crystalline phosphatidylcholine bilayers the signal at 30 ppm is thus indicative of

phosphatidylcholine molecules with their long axis oriented parallel to the magnetic field

direction, whereas a –15 ppm 31P chemical shift is obtained for perpendicular alignments. In

perfectly aligned samples the phospholipid bilayer spectra consists of a single line. Routinely

the 31P NMR spectra of phospholipid bilayers of the peptide carrying samples are recorded to

test for the quality of order and alignment of phospholipid bilayers.

Figure 2-6: Simulated solid-state NMR spectra. (A) and (B) show simulated 15N solid-state NMR spectra of an a-helical polypeptide oriented with the helix long axis perpendicular (A) or parallel (B) relative to the bilayer surface. The membranes are aligned with their normal parallel to the magnetic field of the NMR spectrometer (B0) [88].

NMR measurements were performed under supervision of Dr. Philipe Bertani at the

University of Strassbourg. The vinculin peptide representing the last 21 residues with the

sequence IKIRTDAGFTLRWVRKTPWYQ was used. At the underlined positions the 15N-

labeled analogue of leucine (L) was used (again). Typically, 10 mg of the polypeptide was

resolved in 50µl formic acid. A phospholipids solution consisting of 30 mg 1-Palmitoyl-2-

Oleoyl-sn-Glycero-3-Phosphoglycerol (POPG) and 70 mg 1-Palmitoyl-2-Oleoyl-sn-Glycero-

3-Phosphocholine (POPC) was dissolved in Hexafluor-2-Isopropanol. Both solutions were

mixed and dried onto approximately 30 ultra-thin cover slips (9-22 mm) using a dessicator.

After the complete evaporation of the organic solvents the samples equilibrated at 93 %

relative humidity. The glass plates are then stacked on top of each other, which results in

small brick-shaped samples of 3–4 mm thickness. These are stabilized and sealed with teflon

tape and plastic wrappings. The sample was placed into a coil which was placed that the

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32

membrane was aligned parallel to the magnetic field direction. Cross-polarization or Hahn

echo NMR pulse sequences are typically used to acquire 15N and 31P NMR spectra. The

measurements were performed in a 400MHz Burker Avance solid state NMR spectrometer

using a magnetic field of 9.4 T.

2.2.3.7 Cell lysis The cells were seeded onto fibronectin (FN) coated cell culture treated dishes of 3.2 cm

diameter 24 h prior to lysis. The attached cells were put in 100 µl radio immuno precipitation

assay (RIPA) –buffer (50 mM Tris-Cl pH 7.4, 150 mM NaCl, 1 % NP40, 0.25 %

sodiumdeoxycholate, 1 mM PMSF) containing 1 mM sodiumvanadate and 1 x CompleteTM

protease inhibitor (Roche, Basel, Switzerland). After washing with 1 x PBS, the RIPA treated

cells were removed from the surface using a cell scratcher. Lysis was enhanced by agitating

the dissolved cells for 15 minutes and 4 °C. For separating the cell debris from the cytosolic

components, the lysate was centrifuged at 4 °C and 14,000 g for 15 minutes. The supernatant

was diluted with 6 x SDS sample buffer and boiled at 95 °C for 3 minutes to prevent further

protein degradation.

2.2.3.8 Protein concentration determination according to Bradford The protein determination according to Bradford is based on the interaction of the Coomassie

dye with the peptides [91]. The built peptide-Coomassie complex causes a shift of the dye´s

absorbtion pattern from 465 nm to 595 nm under acidic pH conditions because of the

interaction between the sulfonate groups of the Coomassie dye and the cationic or apolar

peptide residues. The sensitivity of the assay is between 50 - 500 ng/ ml of peptide. The

Bradford assay was performed according to the manufacturer’s instructions. The peptide

concentration was determined using the absorbtion values at 595 nm of a reference

measurement with BSA protein of different known concentrations.

2.2.3.9 Western blot analysis Sodium-dodecylsulphate-polyacrylamide gel electrophoresis (SDS-PAGE) was performed

with a protocol adapted by Laemmli [92]. Separating gels of 10 % were mostly employed in

this work. The protein solutions or cell lysates were diluted in 6× SDS sample buffer, boiled

at 95 °C for 5 minutes and loaded into gels. “High molecular weight marker” (HMW) from

Paclab was used to determine the molecular weight. Proteins were transferred from the gels

onto a nitrocellulose membrane using a semi-dry blotting apparatus from Biometra. The

membrane, gel and Whatman filter papers were equilibrated in the glycine-methanol buffer

(25 mM Tris pH 8.5, 150 mM glycine, 10 % methanol). The proteins were transferred from

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the gel (negative electrode) to the membrane (positive electrode) at 25 V for 40 minutes. The

efficiency of the transfer was monitored by staining the membranes with PonceauS solution

(0.5 % PonceauS, 40 % methanol, 15 % acetic acid; Sigma, Deisenhofen).

Membranes were blocked for 30 minutes to 1 hour at room temperature or at 4 °C overnight

in blocking buffer (5 % dry-milk in TBS-T buffer (20 mM Tris-HCl pH 7.6, 140 mM NaCl,

0.1 % Tween 20)). Afterwards membranes were incubated for 1 h at room temperature or at 4

°C overnight in TBS-T buffer containing the primary antibody. Membranes were washed

three times for 15 minutes in TBS-T and incubated for 1 hour at room temperature in TBS-T

buffer containing the secondary antibody followed by another 4 washes as described above.

Membranes were then incubated for up to 5 minutes with the chemiluminescence substrates

provided by Amersham Biosciences and exposed to Hyperfilm ECL (Amersham Biosciences)

for 10 seconds – 5 min. Films were developed manually using the Kodak developer and fixing

solutions.

2.2.4 Computational methods

2.2.4.1 Molecular dynamics (MD) simulations One of the principle tools in the theoretical study of biological macromolecules is the method

of molecular dynamics simulations. This computational method calculates the fluctuation and

conformational as well as the energetically changes of peptides and nucleic acids in a time-

dependent manner. The theory has been described in detail [93]. In brief, the MD simulation

calculates the accelerated motion of all atoms in a system during a certain time period

according to a defined force field. This force fields represents the energy values of the

covalent bond as the binding-angles and –distances and the non-covalent bond as the van-der-

Waals and the Coulomb interactions. The resulting forces are then applied to each atom

localized in the system using Newton´s law of motion

ma=F (9)

where (F) is the force and (a) the acceleration which can be described as

dtdv

=a (10)

The integration of equation (9) over the time (t) results in the velocity (ν), described in

0)( vattv += (11)

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Further, ν can be expressed as

dtdx

=v (12)

where (x) represents the position. Integrating equation (10) leads to

0)( xvttx += (13)

that give the position of a particle after a certain timepoint. Due to time scales of molecular

movements the integration time-steps are in the range of 1 - 2 femto seconds (fs). Starting

structures on the atomic level are necessary for the MD simulations. Crystal structures of

peptides determined via X-ray and NMR deliver the appropriate coordinates for the start of a

simulation. As starting structure for all molecular dynamic runs for secondary structure

determination of vinculin´s C terminus, the last 20 amino acids were extracted from the

crystal structure of chicken vinculin (PDB 1ST6). For adding the last absent amino acid (Q),

swissprot PDB viewer (http://www.expasy.org/spdbv/) was used. The entire sequence was

IKIRTDAGFTLRWVRKTPWYQ. All simulations were carried out with the GROMACS

3.3.1 package [94-95]. The simulations with the peptide representing vinculin´s last 21

residues were run on a 4 Processor HPC unit with 2.2 GHz for each Processor and 6.2 GB

RAM using Linux. Simulations of the vinculin-tail variants (residues 890-1066 or residue 890

- 1052) were also pre-processed on the 4 Processor HPC system. The final runs were

performed with 16 Xeon 5160 "Woodcrest" CPUs (3 GHz) on a cluster consisting of 217 nods

using Linux. All simulation were performed in an octahedron box filled with approximately

7850 spc216 water molecules [96] by using periodic boundary conditions. The molecule was

centred in the box and the distance to each border was 1.5nm. For keeping the net charge

neutral during the simulations, we added CL- ions to the octahedron box. The GROMACS

Force field ffG53a6 was used [97]. All bonds were constrained using LINCS [98]. Long range

electrostatics were handled using the Particle Mesh Ewald (PME) method. A non bonded cut-

off of 0.9 nm for Lennard - Jones potential was used. The runs were performed at 300 K by

using a pressure coupled Berendsen temperature bath [99]. Energy minimization with a

tolerance of 1000 kJ mol-1 nm-1 was carried out, using the steepest descent method. Position

restraint was used to equilibrate the system for 50 ps. The final MD runs were carried out with

time steps of 2 fs. Snapshots were taken every 100 ps.

2.2.4.2 Analysis of molecular dynamics simulations The dictionary of secondary structure prediction (DSSP) was used for secondary structure

calculation [100]. The program performs its sheet and helix assignments solely on the basis of

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35

backbone-backbone hydrogen bonds from the generated trajectories. The DSSP algorithm

defines a hydrogen bond when the bond energy is below -0.5 kcal/ mol from a Coulomb

approximation of the hydrogen bond energy. The structure assignments are defined in a way

that visually appealing and unbroken structures result. In case of overlaps, alpha-helix is

given first priority, followed by beta-sheet.

2.2.4.3 Cluster analysis The snapshots 1 - 100 of the vinculin peptide representing vinculin´s last 21 residues were

used for a cluster analysis. A pair-wise difference Euclidian distance matrix was generated

using the available sixty dihedral angles (20 peptide bonds x phi, omega and psi). The

difference between dihedrals (Δθn,m) was defined as

Δθn,m = min ( | θn,p - θm,p |, | θn,p - θm,p + 360° |, | θm,p - θn,p - 360° | ) (14)

for any two structures n and m for a given dihedral angle p, where p = 1 to 60. Ward’s

geometric (cluster centered), minimal variance, agglomerative hierarchical clustering was

applied to each pair-wise matrix [101]. The resulting dendogram was partitioned using

Mojena’s Stopping Rule (No.1) to identify significantly different clusters, representing

distinct conformational groups [102].

2.2.4.4 Visualization of calculated structures All the results were visualized using swissprot (http://www.expasy.org/spdbv/) and the NOC

PDB viewer (http://en.bio-soft.net/3d/noc.html).

2.2.5 Cell mechanical methods

2.2.5.1 Magnetic tweezer measurements The principle of the magnetic tweezers device used here was described in Kollmannsberger et

al. [103]. Prior to measurements, fibronectin-coated beads (see 2.2.3.1) were sonicated, added

to cells (1 x 105 beads/dish), and incubated for 30 min at 5 % CO2 and 37 °C. A magnetic

field with a high field gradient was generated using a solenoid (250 turns of ∅ 0.4 mm copper

wire, solenoid length = 2 cm, mean solenoid diameter 1 cm) with a needle-shaped core

(HyMu80 alloy, Carpenter, Reading, PA). The needle tip was placed at a distance of 20 - 30

µm from a bead bound to a cell using a motorized micromanipulator (Injectman NI-2,

Eppendorf, Hamburg, Germany). Bright-field images of the cell, bead, and needle tip were

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36

taken by a CCD camera (ORCA ER, Hamamatsu) at a rate of 40 frames/ s. The bead position

was tracked on-line using an intensity-weighted center-of-mass algorithm. A preset force was

maintained by continuously updating the solenoid current or by moving the solenoid so that

the needle-tip to bead distance was kept constant. Measurements on multiple beads per well

were performed at 37 °C for 1 h, using a heated microscope stage on an inverted microscope

at 40x magnification (NA 0.6) under bright-field illumination. Transfected MEF (-/-) cells had

been identified in the fluorescence mode using an EGFP-filter.

To ensure that cells had not experienced any significant forces resulting from previous

measurements, the needle was moved by at least 0.5 mm between two measurements. The

bead position, the needle tip position and the solenoid current was continuously recorded at a

rate of 40 s-1. Image acquisition was triggered and synchronized with the solenoid current

generator using a custom-made C ++ program run on a PC equipped with an AD-DA board

(NI-6052E, National Instruments).

When a force step with amplitude ΔF was applied to a bead, it moved with a displacement

d(t) towards the needle tip. The ratio d(t)/ΔF defines a creep response J(t) that for all force

amplitudes was well described by a power-law

J(t) = a(t/t0)b (15)

where t0 is a reference time that was arbitrarily set to 1 s. The parameter a (units of µm/nN)

characterizes the elastic cell properties and corresponds to the compliance (i.e. inverse of

stiffness). The exponent b reflects the viscoelasticity of the force-bearing structures of the cell

that are connected to the bead. The numbers of b are between 0 and 1, whereas 0 reflects an

elastic and 1 a viscos behavior [65; 103]. The parameter a changed with the amplitude of the

applied force, indicating a force-dependent non-linearity of the creep modulus: a decreased

with increasing force. The parameter a of the creep response were replaced by an arbitrary

force-dependent functions a(F) and to describe the force-dependence of the creep response,

J(t,F) = a(F) (t/t0)b(F) (16)

The force-dependent differential elastic modulus, 1/a(F), can be easily evaluated at discrete

forces if the force protocol follows a staircase-like pattern. A force protocol with nearly

logarithmically spaced force steps according to 0.5, 1, 1.5, 2, 3, 4, 5, 6, 8, and 10nN was

found to be most versatile, each step lasting for 1 s. In this way, a value for a(F) was obtained

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37

for every force level. The median correlation coefficient between the fit and the data was r2 =

0.99.

2.2.5.2 2D-traction microscopy measurements Gels (6.1 % acrylamide/ 0.24 % bis-acrylamide) for traction experiments were prepared on

rectangular 75 x 25 mm nonelectrostatic silane-coated glass slides according to the procedure

described by Pelham and Wang [33]. The Young’s modulus of the gels was measured with a

magnetically driven plate rheometer and found to be 12.8 kPa. Red fluorescent 1 µm

carboxylated beads (Molecular Probes) were suspended in the gels and centrifuged at 300 g

toward the gel surface during polymerization at 4 °C. These beads served as markers for gel

deformations. The surface of the gel was activated with Sulfo-SANPAH (Pierce

Biotechnology, Rockford, IL) and coated with 50 mg/ml bovine collagen G (Biochrom). The

cell suspension added on the gel was contained within a silicone ring (flexi-perm, in Vitro,

Göttingen, Germany) attached to the glass slide. Cell tractions were computed from an

unconstrained deconvolution of the gel surface displacement field measured before and after

cell detachment with 80 µM CytochalasinD and Trypsin/EDTA (0.25/ 0.02 %) in PBS [104].

During the measurements, the cells were maintained at 37 °C in humidified atmosphere. Gel

deformations were estimated using a Fourier-based difference-with-interpolation image

analysis [105].

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3 Results I. In vitro 3.1 Differential scanning calorimetry (DSC) measurements The membrane interaction of vinculin influences cell adhesion and migration [81-82]. There

are three regions localized at the 30 kDa tail domain (residues 858-1066) that have been

identified as candidates for lipid membrane interaction [38]. Two of them are well

characterized alpha-helical regions. The third one is localized at the C-terminal arm which

remained unstructured during crystallization [78]. Pull-down experiments with artificial lipid

vesicles incubated with a vinculin-tail variant lacking the last 15 residues (vtailΔC), so called

lipid anchor, demonstrated that this variant was not able to interact with acidic lipid

membranes [78; 81]. However, it was not determined to what extent the lipid anchor (residues

1052-1066) is involved in lipid interaction.

In this experiment the lipid-binding ability of the C-terminal arm, which includes the lipid

anchor of vinculin, was explored and characterized using differential scanning calorimetry

(DSC). Multilamellar lipid vesicles (MLVs) consisting of DMPC/ DMPG at various molar

ratios were incubated with peptides representing different variants of vinculin´s C-terminal

arm (refer to 2.2.3.4). Since peptides have the ability to insert into a lipid membrane and

interact with the hydrophobic acyl chains, the specific heat capacity and the phase transition

enthalpy (H) will decrease. Insulin which has approx. the same molecular weight (5,8kDa) as

the synthesized vinculin peptides was used as negative control. It is known that insulin is not

inserting into lipid membranes.

3.1.1 DSC measurements of the C-terminal arm peptide

As demonstrated in figure 3-1, the specific heat was determined from MLVs at a molar ratio

of 70:30 DMPC:DMPG that were incubated with the 21 residue vinculin peptide, representing

the C-terminal arm. With increasing peptide concentration (from 0 to 180 µM), the specific

heat of MLVs decreased compared to MLVs in the absence of vinculin peptide (figure 3-1A).

The relative flattening of the curve together with the shift of Tm to lower temperatures are

indicative for peptide insertion into lipid bilayers [106-107]. As a control, DSC measurements

in the presence of lipid vesicles and insulin at different molar ratios were performed. With

increased insulin concentration, the specific heat did not alter significantly (figure 3-1B). The

decrease in relative transition enthalpy (dH/ dH0) as a function of increasing peptide

concentration was plotted for DMPC/ DMPG vesicles of different molar ratios (figure 3-1C).

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39

Increasing the concentration of negatively charged lipids in vesicles caused a decrease in

peptide-lipid insertion. Insulin (control) showed no changes in relative transition enthalpy.

These results suggest that the C-terminal peptide of vinculin is directly involved in membrane

interaction of vinculin.

Figure 3-1: The peptide, representing vincuin´s C-terminal arm (residue 1045-1066), interacts with MLVs. (A) Thermograms from DSC measurements with lipid vesicles containing DMPC and DMPG at a molar ratio of 70:30. The specific heat decreased with increasing peptide concentration or the peak expanded (n = 4–6 runs). Note, that the integral of the specific heat provides the phase transition enthalpy dH. (B) The phase transition enthalpy of insulin incubated with DMPC/DMPG vesicles of a molar ratio of 70:30. The specific heat remained almost constant with increasing the insulin concentration. (C) Relative phase transition enthalpy changes of DMPC/DMPG vesicles at different molar ratios. With increasing vinculin peptide concentration, the phase transition enthalpy decreased as a consequence of the peptide insertion into the lipid membrane. Note, that there were no detectable changes in enthalpy at all lipid compositions for insulin (data not shown).

3.1.2 DSC measurements of the mutated C-terminal arm

Pull-down experiments with artificial lipid vesicles revealed that the mutation of the basic

amino acids Arginine 1060 and Lysine 1061 to Glutamine (vtail-RK 1060/61 Q) results also

in impaired lipid vesicle interaction [82]. To test the nature of this lipid interaction, the

peptide representing this altered C-terminal arm (RK 1060/61 Q) was also measured using

DSC. With increasing peptide concentration, the specific heat of MLVs incubated with RK

1060/61 Q also decreased in comparison to the measurements performed with MLVs only,

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40

whilst Tm was shifted to lower temperatures (figure 3-2A). The relative transition enthalpy

(dH/dH0) for DMPC/DMPG vesicles at different molar ratios (figure 3-2B) also decreased;

even more, compared to the measurements of the non-mutated peptide (for comparison see

also figure 3-1C). Decreasing the concentration of negatively charged lipids, the insertion of

the peptide was also increased, judged by the relative transition enthalpy (see figure 3-2B).

The results suggest that the RK 1060/61 Q peptide of vinculin showed a better membrane

insertion behavior than the wildtype peptide.

Figure 3-2: Interaction of the mutated vinculin-tail peptide (RK1060/61 Q) with MLVs. (A) Thermograms from DSC measurements with lipid vesicles containing DMPC and DMPG at a molar ratio of 70:30. The specific heat decreased or expanded with increasing peptide concentration (B) Relative phase transition enthalpy changes of DMPC/ DMPG vesicles at different molar ratios of DMPC/ DMPG. The increasing vinculin peptide concentration results in a decreased phase transition enthalpy as a consequence of the peptide insertion into the lipid membrane.

3.2 Molecular dynamics simulations Proteins and peptides are highly dynamic structures. Their function is determined by their

secondary structure. The interaction with substrates or other proteins induces conformational

shifts which are important for their functions. Crystal structure analysis delivers only a static

picture of such molecules. In contrast, molecular dynamic investigations give an indication

for proteins in motion during their function.

3.2.1 MD Simulations of the C-terminal arm

Crystal structure analyses of the vinculin molecule demonstrated that the C-terminal arm

remained unstructured and can be divided into three different regions: a flexible loop

(residues 1047–1052), a beta-clamp (residues 1053–1061) and a hydrophobic hairpin

(residues 1062–1066) [78]. As demonstrated in pull-down experiments, parts of this C-

terminal arm known as the lipid anchor (residues 1052–1066) influence the membrane

binding of vinculin [78; 81]. In order to elucidate the secondary structure of the mostly

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41

unstructured C-terminal lipid binding region of the vinculin molecule, molecular dynamics

simulations were carried out using the GROMACS package (see also chapter 2.2.4.1). The

MD calculations were performed under different ionic conditions representing basic, neutral

and acidic pHs. The trajectories of vinculin´s last 20 residues were extracted from the crystal

structure of chicken vinculin (PDB 1ST6) [55]. Because of the high flexibility of the C-

terminal arm during crystallization, the position of vinculin´s last residue is missing. So, the

last glutamine (Q) residue was added. The N-terminal isoleucine was always considered as –

NH2 (uncharged) and the five basic residues, the single acidic residue and the terminal

carboxyl-group were either protonated or deprotonated, respectively, depending on the

nominal pH conditions. Under “neutral” conditions, the five basic residues were protonated

and both, the single acid residue and the terminal carboxyl group, deprotonated. Under the

“acid” condition both carboxyl groups were protonated. During the “basic” condition the five

basic residues were completely deprotonated. The nominal charges were therefore -1, 0 or +1.

Charge neutrality of the entire system was achieved by adding either Cl- or Na+ counter ions.

Three 10 ns runs of each condition were performed. The calculated conformations were

collected every 100 ps. The program dictionary of secondary structure prediction (DSSP)

[100] was used for identifying the secondary structure elements of each run (refer to 2.2.4.2).

Further, a conformational analysis of all runs was performed under nominal “basic”, “neutral”

and “acidic” conditions to identify a representative secondary structure. Each simulation was

repeated three times to proof the reliability of the secondary structure determination. The

results are plotted in figure 3-3, 3-4 and 3-5. Under nominal “basic” (Figure 3-3A) and

“neutral” (Figure 3-4A) conditions, an anti-parallel beta-sheet emerged after 3.5 ns and

remained constant until the end of the simulation at 10 ns. Residues 2-12 were part of that

secondary structure element (illustrated in red). Residues 13-21 remained unstructured under

those conditions. Under basic conditions, all three runs delivered the same result whilst only

two of three runs delivered the anti-parallel beta-sheet under “neutral” conditions. The

simulation, performed under nominal “acidic” conditions, obtained no defined secondary

structure elements (Figure 3-5A). Ward’s geometric (cluster centered), minimal variance,

agglomerative hierarchical clustering was applied to perform the cluster analysis for the

snapshots 1 to 100 under all nominal pH conditions. The resulting dendograms were

partitioned using Mojena’s Stopping Rule (No.1) to identify significantly different clusters

[102]. The analysis as demonstrated in figure 3-3B revealed 6 statistically significant

conformational groups (p<0.05) under nominal “basic” conditions. The clusters on the

diagram are illustrated in red and black, respectively. The representative geometry for each

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42

cluster is colored in green. The cluster labeled with an asterisk is the most densely packed one

and represents the most energetically favorable cluster. Where the snapshots appear to jump

between clusters, the associated conformations are likely to lie on a plateau, and not a well-

defined local minimum, on the potential energy landscape for the simulation. The cluster

analysis for the run under “neutral” conditions delivered 4 distinct groups (Figure 3-4B)

whilst the simulations under nominal “acidic” (Figure 3-5B) conditions obtained also 6

distinct groups. The structures extracted from the most densely packed groups represent the

most energetically favorable structure of the run and can be seen as the representative

structure of the simulation. Under nominal “basic” (figure 3-3C) and “neutral” (figure 3-4C)

conditions, the extracted structure showed an anti-parallel beta-sheet followed by a mostly

unstructured part. The “acidic” run revealed an undefined secondary structure (figure 3-5C).

These results suggest that the first 6 residues of the lipid anchor (residues 1052-1057) are part

of the turn and the second strand of the beta-sheet. This implies an involvement of vinculin´s

third lipid binding site (residues 1052-1066) in beta-sheet formation. Furthermore, the

simulations indicate a pH-dependent bi-stable behavior. The presence and absence of the beta-

sheet is a passive response of the putative anchor region in vivo, which is controlled by the

charged residues. Such behavior could be influenced by local proton or ion gradients in the

cytosol. To test, if the emerged secondary structure was determined by its trajectories

extracted form the crystal structure, the protonation states during the runs were altered. The

simulations were started under acidic or basic conditions, respectively. After 10 ns, the

trajectories of the last calculated structure was extracted and used for the following run under

the next nominal pH condition for additional 10 ns. As described in figure 3-6, runs from

acidic to neutral to basic and vice versa were performed. There was no alteration in secondary

structure between the acidic and the neutral run. However, after changing the conditions from

neutral to basic, the anti-parallel beta-sheets emerged. The simulations which started with the

basic run delivered a similar result (figure 3-6B). During the first 10 ns the anti-parallel beta-

sheet was visible after ~3.5 ns and remained constant until the end of the basic run at 10 ns.

During the run under “neutral” conditions, the beta-sheet was available until ~18 ns and

vanished at the last 2 ns of that step. Under “acidic” conditions, there was also no beta-sheet

conformation detectable. This result, together with the determined secondary structure under

neutral conditions visualized under 3-3, indicates a bi-stable behavior for the peptide under

nominal “neutral” conditions. The anti-parallel as well as the unstructured configuration was

calculated. Further it suggests that the starting structure does not determine the secondary

structure. The anti-parallel beta-sheet formation depends on the amino acid composition.

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Figure 3-3: (A) Emergent secondary structure of vinculin´s C-terminal 21 residues under charge “basic” conditions according to DSSP. (B) Clustered conformations of 21 residue vinculin-tail peptide and putative lipid anchor. The fusion distance is a scale bar for measuring the distances between compact clusters. The labels on the x-axis represent the snapshots taken during the molecular dynamics simulation. The distinguished clusters are colored black and red, respectively. The representative structure of each cluster is colored in green. Note, that the asterisk marks the most compact cluster which is magnified. (C) The representative backbone geometry of the most compact cluster at time point 7100 ps.

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Figure 3-4: (A) Emergent secondary structure of vinculin´s C-terminal 21 residues under charge “neutral” conditions according to DSSP. (B) Clustered conformations of 21 residue vinculin-tail including the lipid anchor. Note, that the most compact cluster was magnified and labeled by an asterik. (C) The representative backbone geometry of the most compact cluster was at time point 8400 ps.

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45

Figure 3-5: (A) Emergent secondary structure of vinculin´s the C-terminal 21 residues under charge “acidic” conditions according to DSSP. (B) Clustered conformations of 21 residue vinculin-tail including the lipid anchor. Note, that the most compact cluster was magnified and labeled by an asterik. (C) The representative backbone geometry of the most compact cluster at time point 8400 ps

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46

Figure 3-6: MD-simulations of the 21 residue peptide, representing vinculin´s unstructured C-terminal arm. The nominal pH conditions were altered each 10ns from (A) “acidic” to “neutral” to “basic” or (B) from “basic” to “neutral” to “acidic”. The result implies that the secondary structure was not influenced by the starting structure.

3.2.2 MD Simulations of the vinculin-tail

The results determined in 3.2.1 suggest that the lipid anchor of vinculin is involved in beta-

sheet formation. All those simulations were performed out of the context of the vinculin-tail.

To test if the lipid anchor is also involved in beta-sheet formation, molecular dynamics

simulations were performed, using the whole vinculin-tail molecule (residues 890-1066). The

trajectories were also extracted from the crystal structure (PDB 1ST6) of chicken vinculin and

prepared as described under 2.2.4.1. The simulations were performed under charge neutral

conditions. Because of the emerged secondary structure under “neutral” conditions, which

reflects the cytosolic pH, the MD-simulation of the whole vinculin-tail was also performed

under those conditions. As described in figure 3-7, no beta-sheet emerged at the lipid anchor

region in the context of the whole vinculin-tail molecule. The C-terminal arm remained

unstructured during the whole simulation. This may imply that a conformational switch of the

vinculin-tail, induced by local pH shift or interaction with binding partners such as actin, is

needed to set the unstructured C-terminal arm free which could allow for anti-parallel beta-

sheet formation of the lipid anchor.

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47

Figure 3-7: Secondary structure of the vinculin-tail (residue 890-1066). The red square marks the unstructured C-terminal 21 residues (residues 1045-1066) including the lipid anchor (1052-1066). Note that no anti-parallel beta-sheet emerged after 150 ns.

3.3 CD-spectroscopy measurements of the C-terminal arm The MD-Simulations of the vinculin peptide, representing its C-terminal arm, revealed an

involvement of the lipid anchor in beta-sheet formation under nominal “basic” and “neutral”

conditions (refer to 3.2). Furthermore, it was demonstrated by DSC measurements that this

peptide has the ability to insert into lipid vesicles (refer to 3.1). To proof the in silico

determined secondary structure in the presence and absence of lipid vesicles CD-

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48

spectroscopic measurements were performed. As described under 2.2.3.5, SUVs consisting of

DMPC/ DMPG at a molar ratio of 70:30 were used in these experiments. At this composition,

the best insertion behavior was determined according to DSC measurements (refer to 3.1).

With respect to the cytosolic environment, all measurements were performed at physiological

pH. The CD-spectra of the C-terminal peptide in the presence/ absence of lipid vesicles were

plotted in figure 3-8. Measuring between 180 and 260 nm, the spectrum of pure peptide

showed a minimum at 200 nm, whilst in the presence of SUVs this minimum was shifted to

223 nm. This observation suggests a clear change in conformation of the peptide. Analyses of

the spectral data using algorithms (CDSSTR and CONTINLL) that employ different

approaches for deconvoluting the CD data from a database of secondary structure

contributions, show similar structural elements within the vinculin C-terminal peptide. As

described in figure 3-8B, CDSSTR shows 25 % beta-strand 1, 13 % beta-strand 2, 43 %

random coil (NRMSD < 0.016) and CONTINLL 21.9 % beta-strand 1, 13 % beta-strand 2,

42.6 % random coil (NRMSD < 0.134).

Figure 3-8: Results from CD-spectroscopic measurements of the C-terminal peptide in the presence/ absence of SUVs. The CD spectra of 70 µM peptide were obtained in 10mM sodium phosphate at pH 7.4 and 30 °C (n = 3). (A) The plot of the mean residue molar ellipticity, Θ (deg cm2dmol-1) in the presence and absence of DMPC/ DMPG lipid vesicles. Secondary structure prediction according to the algorithms provided by Dichroweb in the (B) absence and (C) presence of lipid vesicles. Note, that the result is in good agreement with the calculated secondary structure using GROMACS.

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49

These analyses indicate a possible anti-parallel beta-sheet conformation. It underlines that the

CD spectrum of the peptide allows an interpretation which agrees with the representative at

8400 ps derived from the MD simulation under “neutral” conditions. In the presence of SUVs,

CD spectroscopy of the vinculin peptide provided similar results (figure 3-8C). In comparison

to the spectra in the absence of lipid vesicles, the two algorithms indicate also two beta-

strands and unstructured parts at a similar distribution (NRMSD< 0.05) for both CDSSTR and

CONTINLL. However, the difference in spectral characteristics of the spectra for the peptide

in the presence/ absence of lipid vesicles suggests distinguishable beta-sheet conformations.

3.4 Solid state NMR measurements of the C-terminal arm The results of the MD-simulations together with the CD-spectroscopic measurements in

presence and absence of lipid vesicles indicate that the lipid anchor is involved in an anti-

parallel beta-sheet formation whilst interacting with lipid membranes. To proof the orientation

of this interaction, solid state NMR measurements were performed as described under 2.2.3.6.

The 15N- label was placed in the leucine residue on position 11 of the C-terminal arm peptide

(IKIRTDAGFTLRWVRKTPWYQ), which was part of the second beta-strand. One single

peak at 100 ppm or 200 ppm would suggest a distinct conformation of the peptide during lipid

membrane interaction. As described in figure 3-9, a plateau around 200 ppm followed by one

prominent single peak at ~100 ppm appeared. This implies a random distribution with a

tendency to one certain conformation during interaction (in detail of the labeled part of the

peptide) with the lipid membrane. The shift to ~100 ppm together with the proven secondary

structure of the beta-sheet (refer to 3.5 and 3.6) implies a surface seeking interaction of the

peptide. The spectra of the lipids (figure 3-9B) demonstrated a well oriented lipid bilayer into

which the peptide inserted. Additional measurements are necessary to elucidate the lipid

interaction behavior and the insertion angle of the peptide in more detail.

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50

Figure 3-9: Result of the NMR measurement of vinculin´s unstructured C-terminal arm. (A) Proton-decoupled 15N solid-state NMR spectra of 10mg peptide reconstituted into 100 mg of 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC)/ palmitoyl-2-oleoyl-sn-glycero-3-phosphoglycine (POPG) membranes at a molar ratio of 70 : 30. (B) The proton-decoupled 31P solid-state NMR spectrum of the sample is shown in panel A. The 31P NMR line shape represents the distribution of alignments of the phospholipid head group in the sample.

II. In vivo 3.5 Cloning and expression of the different vinculin constructs

3.5.1 Cloning and expression of EGFP linked vinculin and vinculinΔC

The structural and functional role of vinculin´s lipid anchor in the regulation of adhesion sites

was studied using recombinant vinculin proteins. Eukaryotic expression constructs were used

for biochemical and cellular characterization of the protein. EGFP linked full length vinculin

and vinculinΔC was synthesized as described under 2.2.1.13. To test if the EGFP labeled

constructs were localized at the FA, MEF-vin(-/-) cells were transfected with the vinculin and

vinculinΔC constructs (Refer to chapter 2.2.2.2), reseeded on coverslips (refer to 2.2.2.3) and

fixed (refer to 2.2.2.4) for microscopic analysis. Furthermore, the lysates of the transfected

cells were used for Western blot analysis (Refer to chapter 2.2.3.9.). Figure 3-10 shows the

images of the MEF-wt (wildtype), MEF-vin(-/-) (vinculin knock out), MEF-resc (rescue) and

MEF-vin C (transfected with vinculin lacking the lipid anchor; residues 1-1052) cells. The

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focal adhesions (green) were determined in MEF-wt and MEF-vin(-/-) cells using antibodies

against vinculin. In MEF-resc and MEF-vin C cells the EGFP molecule connected to the

different vinculin peptides was used. As observed in the pictures, vinculin is localized in the

focal adhesions. The focal adhesions are connected to the actin stress-fibers colored in red.

According to the Western Blot analysis (Figure 3-10B) vinculin and vinculinΔC linked to

EGFP were expressed in the cells. There are no additional bands below 120 kDa visible,

suggesting there was no degradation of the vinculin peptides.

Figure 3-10: Expression of vinculin and vinculinΔC in the different MEF cell lines determined by (A) Immunofluorescence and (B) Western blot analysis. (A) The FA were stained with a monoclonal antibody against vinculin in MEF-wt and MEF-vin(-/-). For visualization of the vinculin constructs in MEF-resc and MEF-vinΔC, the EGFP label was used. The actin cytoskeleton was stained with TRITC-Phalloidin in each cell line. The scale bar represents 50 µm. The Inset shows FA in MEF-vin(-/-) stained with paxillin (B) Western Blot analysis show the cell lysates of MEF-wt, MEF-vin(-/-), MEF-resc and MEF-vinΔC cells. There was no vinculin in the MEF vin(-/-) cells detected. Note, that the EGFP tag on the different vinculin constructs in MEF-resc and MEF-vinΔC cells caused a shift of the molecule in the direction to higher molecular weight.

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3.5.2 Cloning and expression of EGFP-linked vinculin-tail and vinculin-tailΔC

F9 Vinculin (-/-) carcinoma cells showed a decrease in stiffness and develop less force in

comparison to their wildtype cells [65; 69]. Furthermore, it was reported that the

retransfection of the vinculin-tail domain is sufficient to fully restore the wildtype phenotype,

suggesting that the interaction between the actin cytoskeleton and vinculin is responsible for

those effects [65]. To test the influence of the lipid anchor to cell mechanical behavior in

absence of the vinculin-head, an EGFP linked vinculin-tail (vtail, residues 858-1066) and

vinculin-tailΔC (vtailΔC, residue vtail 858-1052) construct was prepared as described under

2.2.1.13, using the primer pair 2b/ 2f or 2e/ 2f, listed under 2.1.3. MEF-vin(-/-) cells

transfected with EGFP linked vtail and vtailΔC were used to obtain the difference to cell

mechanics of this two constructs (refer to chapter 2.2.2.2). As described in figure 3-11, both

constructs were localized at the FA. However, both vinculin-tail peptides also masked the

actin stress-fibers. The cytosolic background was higher in comparison to MEF-vin(-/-) cells

transfected with the full length vinculin constructs (see figure 3-11). The cells expressing the

vtail constructs did not spread well, suggesting that the vtail and vtailΔC constructs show an

impaired FA localization. An expression analysis of the different vtail constructs via Western

Blot analysis was not possible with the used antibody because the epitope of recognition is

localized in the vinculin-head region.

Figure 3-11: Expression of EGFP linked vinculin-tail and vinculin-tailΔC in MEF-vin(-/-) cells determined by fluorescence microscopy. The scale bar represents 20 µm.

3.5.3 Mutagenesis of vinculin´s src dependent phosphorylation site Y1065F

The phosphorylation of vinculin also influences the mechanical properties of the cells. It was

reported that the mutation of the c-src dependent phosphorylation sites on vinculin results in

impaired cell spreading and migration [71-72]. One of this phosphorylation sites, Tyrosine

1065, is localized at the lipid anchor. To determine the influence of this site to cell mechanics,

the tyrosine residue on position 1065 was mutated to a phenylalanine by site-directed

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mutagenesis (refer to 2.2.1.8). This construct was transfected into MEF-vin(-/-) cells and

transferred onto FN-coated glass coverslips (refer to chapter 2.2.2.3). After fixation the actin

cytoskeleton was stained with TRITC-Phalloidin. According to figure 3-12 the

vinculinY1065F construct was properly localized to the FA. Further, no visible difference in

focal adhesion size was determined in comparison to non-mutated vinculin.

Figure 3-12: Expression of vinculin and vinculinY1065F in MEF-vin(-/-) cell lines (green). The actin cytoskeleton was stained using TRITC-Phalloidin (red). The scale bar represents 20 µm.

3.6 Magnetic tweezer measurements Control of cell mechanics and shape is crucial for many cellular functions, including growth,

differentiation, migration, and gene expression. The mechanical properties of the cell, in turn,

are established through a balance of mechanical forces. Tensions, generated by actin filaments

are counteracted by compression resistant extracellular matrix (ECM) anchors as well as

internal microtubule structures [5-6]. To measure the intracellular prestress a magnetic

tweezer was used for the quantification of mechanical properties of individual living cells [69;

103]. The magnetic tweezer measurements were performed as described in chapter 2.2.5.1. In

brief, transfected and non-transfected cells were incubated with fibronectin-coated para-

magnetic beads 30 min prior to measurements. During the measurements, a solenoid with a

needle-shaped core was placed in front of a FN-coated bead connected to a single cell. After

the application of a magnetic field, the bead moved towards the needle. The displacement of

the bead (d) follows a power law

btta )/(d 0= (15)

where parameter a (units of µm/nN) characterizes the elastic cell properties and corresponds

to a compliance whilst the exponent b reflects the viscoelasticity. The inverse of compliance

is defined as stiffness 1/a (units of nN/µm).

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3.6.1 Magnetic tweezer measurements of MEF-resc and MEF-vinΔC cells

Cell attachment to the ECM via focal adhesions is prerequisite for cell survival and mainly

driven by transmembrane receptors, like intergin [22]. The co-localization of the focal

adhesions and the beads were tested by confocal microscopy. MEF-vin(-/-) cells transfected

with EGFP linked vinculin or vinculinΔC, were seeded on coverslips (refer to 2.2.2.3),

incubated with FN-coated beads and fixed (refer to 2.2.2.4). As control, the focal adhesions of

MEF-wt and MEF-vin(-/-) cells, seeded and fixed on cover slips, were stained using an

antibody against the FA protein paxillin (refer to 2.2.2.5). The images in figure 3-13 show

MEF-wt, MEF-vin(-/-), MEF-resc and MEF-vin C cells.

Figure 3-13: Co-localization of the auto-fluorescent beads (red) with the focal adhesions (green). Transfected and non-transfected MEF vin(-/-) cells were seeded on glass slides. The cells were allowed to adhere overnight. Prior to fixation, the cells were incubated with FN-coated beads for 1 h. The FAs in MEF-wt and MEF-vin(-/-) cells were stained with antibodies against paxillin; in MEF-vin(-/-) cells transfected with vinculin (MEF-resc) and vinculinΔC (MEF-vin C) focal adhesion were determined through visualization of the N-terminal EGFP label. The arrows mark the FN-coated beads which are in close proximity to the focal adhesions.

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The focal adhesions (green) were determined in MEF-wt and MEF-vin(-/-) cells using

antibodies against paxillin, and in MEF-resc and MEF-vin C cells using the EGFP molecule

connected to the different vinculin peptides. The position of the 4.5 µm FN-coated beads (red,

arrow) was determined using its auto-fluorescent properties. In close proximity to the beads,

an accumulation of the FA components such as paxillin or vinculin was detected, suggesting

that beads were linked to focal contacts. These pictures represent this common phenomenon.

Note, that there was no visual difference in FA size in all of these cells. Cells expressing

vinculin lacking the lipid anchor region showed a reduced FA turnover rate and impaired cell

migration [81]. This implies that the lipid anchor influences cell mechanical behavior. To

determine the influence of vinculin´s lipid anchor on cell stiffness MEF-wt, MEF-vin(-/-),

MEF-resc and MEF-vinΔC were measured using the magnetic tweezer device. Transfected

and non-transfected cells were measured as described under 2.2.5.1. The results are shown in

figure 3-14A, where we applied forces of 1 nN to FN-coated beads attached to MEF-wt and

MEF-resc cells. These cells indicated similar stiffness, whilst MEF-vin(-/-) and MEF-vin C

cells showed significantly reduced stiffness values of ~ 48 % and ~26 %, respectively,

compared to MEF-wt cells. This suggests that the lipid anchor is important for cell stiffness.

To test, whether the binding strength of the different MEF-cells to fibronection-coated

surfaces holds over a wide range of forces, the externally applied force was increased over 10

s in a staircase-like manner from 0.5 – 10 nN. With increasing forces (0.5 – 10 nN), the bead

detachment also increased (figure 3-14B). At 10 nN force, ~20 % and ~14 % beads detached

from MEF-wt and MEF-resc, respectively, whilst 37 % of the beads detached during force

application on MEF-vin(-/-) cells. MEF cells transfected with vinculin C showed at low

forces (0.5 - 5 nN) similar behavior to MEF-wt and rescue cells. Only at higher (5 – 10 nN)

force nearly the same level of beads detached in comparison to MEF-vin(-/-) cells. The

fraction of beads that detached at a given force level can be regarded as a direct measure of

the adhesion strength (yielding force) of the force-transmitting elements and bonds between

the fibronectin-coated beads, integrins, focal adhesion proteins and the cytoskeleton. These

results may imply that the lipid anchor of vinculin influences the adhesion strength between

the cytoskeleton and ECM-coated surfaces.

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Figure 3-14: Result of magnetic tweezer measurements. A) Stiffness (geometric mean) of the transfected and non-transfected MEF cells determined at 1 nN force. The error bars represent the standard error (SEM). B) Percentage of detached beads vs. forces in MEF-wt and vinculin mutant cells averaged for all experiments. Besides the increased bead detachment level at high forces, the differential stiffness of all cell

lines increased also with the rising force. In figure 3-15 the stiffness of the different MEF-cell

lines was plotted against the applied force. The stiffness calculation in figure 3-15A includes

the displacements of the detached beads. In figure 3-15B, the stiffness was calculated only

with the beads that remained attached until the end of the measurement at 10 nN force. MEF-

wt and MEF-resc cells displayed at all force steps significantly higher stiffness values in

comparison to MEF-vin(-/-) cells, independent from the level of disrupted beads. In contrast,

the stiffness values for MEF-vin C cells were over a wide range significantly less stiff in

comparison to MEF-wt and MEF-resc performing the calculation with all measured beads

(figure 3-15A). Only at high forces (6 – 10 nN) they reached almost wildtype and rescue

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level. Calculating the stiffness of MEF-vin C cells excluding the detached beads (figure 3-

15B) demonstrated that MEF-vin C cells showed similar stiffness levels than MEF-wt and

MEF-resc cells over the whole force range. In contrast, the stiffness of MEF-vin(-/-) cells was

always ~50 % decreased in comparison to MEF-wt and rescue, independent from the

disrupted beads. This suggests that the reduced binding strength in vinculinΔC cells is

influencing also its stiffness.

Figure 3-15: Differential cell stiffness of transfected and non-transfected MEF cells at given forces between 0.5 to 10 nN visualized in a log-log plot (p<0.05). The cell stiffness was determined including (A) or excluding (B) the detached beads. The stiffness increased with force nearly parallel for MEF-wt, MEF-resc, MEF-vin(-/-) cell lines independent from the detachment level. However, calculating the stiffness of MEF-vinΔC cells, including the disrupted beads (A), the values were significantly different to MEF-wt and MEF-resc for forces between 0.5 – 4 nN. The application of higher forces resulted in stiffness values near to MEF-wt and -resc cells. Excluding the detached beads (B) resulted in similar stiffness than MEF-wt and MEF-resc cells. The error bars represent the standard error (SEM). To determine the viscoelasticity of the transfected and non-transfected cells, the power-law

exponent b was plotted against the applied force (figure 3-16). The exponent b determines the

visco-elastic behavior of the cells [65; 103]. The values are between 0 and 1, whereas 0

reflects elastic and 1 viscous behavior. In figure 3-16A the b-Factor was calculated using all

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measured beads, whilst in figure 3-16B the disrupted ones were excluded. The exponent b of

the MEF-vin(-/-) cells was significantly higher in comparison to MEF-wt, MEF-resc and

MEF-vinΔC over the applied force range (force 1-10 nN), independent from the level of

disrupted beads. This indicates a more viscous behavior for MEF-vin(-/-) cells in comparison

to MEF-wt and MEF-resc cells. No significant difference was detected between MEF-wt,

MEF-resc and MEF-vinΔC cells, suggesting that the lipid anchor is not affecting the

viscoelasticity of the cell.

Figure 3-16: Geometric mean of the power-law exponent b over a range of applied forces from 0.5-10 nN, calculated with all (A) and without the disrupted beads (B). There was no significant difference between MEF-wt, MEF-resc and MEF-vinΔC cells. The error bars represent the standard error (SEM).

3.6.2 Creep measurements of MEF-vtail and MEF-vtailΔC cells

F9 mouse carcinoma cells expressing only the vinculin-tail showed nearly the same

mechanical properties as the F9 wildtype or rescue cells [65]. It is assumed that this is due to

vinculin´s ability to interact with the actin cytoskeleton [65]. Generally, the access of the

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vinculin-tail is controlled by the vinculin-head domain which masks the cryptic binding-sites

for actin, paxillin and phospholipids localized at the tail [52]. Talin- and also lipid interaction

displace the vinculin-head domain from the tail and free those interacting sites [20]. To

reveal, whether the influence on cell mechanics of the lipid anchor is due to the activation or

the anchorage of vinculin to the membrane itself, magnetic tweezer measurements (refer to

2.2.5.1) were also performed using MEF-vtail (residue 858-1066) and MEF-vtailΔC (residue

858 – 1052) cells. The results are plotted in figure 3-17. Forces of 1 nN were applied to FN-

coated beads attached to the MEF cells, transfected with the different EGFP-linked vinculin-

tail constructs. MEF-vtail and MEF-vtail C cells showed a stiffness reduction to

approximately 40 % in comparison to MEF-wt. There was no difference in stiffness between

MEF-vtail and MEF-vtail C, suggesting that the lipid anchor only influence stiffness in cells

expressing full length vinculin constructs. Furthermore, measuring the binding strength

between the beads and MEF-vtail and MEF-vtailΔC cells revealed higher detachment values

in comparison to MEF-vin(-/-) cells (figure 3-17; B). At 10 nN force, ~41 % and ~52 % of

beads detached from MEF-vtailΔC and MEF-vtail, respectively. In contrast, only 37 % of the

beads detached from MEF-vin(-/-) cells. The account of detached beads did not move

between the values determined by MEF-vin(-/-) and MEF-wt, suggesting that the expression

of the EGFP linked vinculin-tail variants negatively affects cell mechanical behavior.

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Figure 3-17: Magnetic tweezer measurements of MEF-vtail and MEF-vtailΔC in comparison to MEF-wt and MEF-vin(-/-) cells. A) Stiffness (geometric mean) of transfected and non-transfected MEF cells determined at 1nN force. There was no detectable difference between MEF-vtail and MEF-vtailΔC. The error bars represent the standard error (SEM). B) Percentage of detached beads vs. forces, averaged for all experiments. Note, that the detachement levels of MEF-vtail and MEF-vtailΔC were not in between those values, determined by MEF-wt and MEF-vin(-/-) cells. In figure 3-18 the stiffness of MEF-wt, MEF-vin(-/-), MEF-vtail and MEF-vtailΔC was

plotted against the applied force ramp. Figure 3-18A represents the stiffness calculation

including the displacements of the detached beads. The stiffness values of the cells transfected

with the different vinculin-tail constructs were between the values of MEF-wt and MEF-vin(-

/-) cells. No difference in stiffness was determined for MEF-vtailΔC and MEF-vtail over the

whole force range. In figure 3-18B, the stiffness was calculated only with the beads that

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remained attached until the end of the measurement. There was also no difference in stiffness

between MEF-vtailΔC and MEF-vtail. Furthermore, MEF-vtailΔC and MEF-vtail were as

stiff as MEF-wt over the whole force range, suggesting the vinculin-tail hosts the mechano-

coupling properties.

Figure 3-18: Differential cell stiffness of MEF-vin(-/-) cells transfected with vtail and vtailΔC at given forces between 0.5 to 10 nN visualized in a log-log plot. The cell stiffness was determined (A) with all (A) as well as without the disrupted beads (B). The stiffness increased with force nearly parallel for all measured cell lines independent from the bead detachement level and the values. The error bars represent the standard error (SEM). In figure 3-19, the viscoelasticity (determined by the power-law exponent b) of cells

expressing the vinculin-tail constructs, was plotted against the applied force. Including all

beads into the calculation, the power-law exponent b of the MEF-vtail and MEF-vtailΔC cells

jumps between the values of MEF-wt and MEF-vin(-/-). The evaluation of the b-values with

the beads that remained attached until the end of the measurements delivered a clearer picture

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(figure 3-19B). The MEF-vtail and MEF-vtailΔC are almost as elastic as the MEF-wt cells.

No difference between MEF-vtail and MEF-vtailΔC was detected. This could be seen as

additional evidence that the lipid anchor is not affecting the viscoelasticity of the cell.

Figure 3-19: The mean of the power-law exponent b over a range of applied force from 0.5-10 nN, determined including (A) or excluding (B) the detached beads. There is no significant difference between MEF-wt, MEF-vtail and MEF-vtailΔC, suggesting no effect of the lipid anchor due to the cellular viscoelastic behavior. The error bars represent the standard error (SEM).

During the creep measurements, no significant differences in stiffness, binding strength and

viscoelasticity between MEF-vtail and MEF-vtailΔC could be detected, independent if the

calculation was performed with all beads or only with the beads which remained attached

until the end of the measurement at 10 nN. The result suggests that the lipid anchor influence

stiffness only in cells expressing full length vinculin constructs. This could be indicative for

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an involvement of the lipid anchor in regulating vinculin´s head/ tail interaction. In the

following experiments only full length vinculin constructs were used.

3.7 Determination of the spreading area Cell stiffness and force generation are important for cell adhesion and migration [18]. To test

the influence of the lipid anchor on cell mechanic properties, the spreading area was

determined. As described in chapter 2.2.2.6, transfected and non-transfected cells were seeded

on FN-coated coverslips. After 1h incubation, the samples were fixed and images were taken

(refer to chapter 2.2.2.6). As demonstrated in figure 3-20, MEF-vin(-/-) cells were half the

size of MEF-wt. In contrast, Saunders and co-workers demonstrated that MEF-wt cells are

double the size in comparison to MEF-vin(-/-) cells [81]. No significant difference was

detected between MEF-resc and MEF-vinΔC cells. Furthermore, the retransformation of

vinculin in MEF-vin(-/-) cells did not fully restore the wildtype phenotype, suggesting the

vinculin knock out caused subsequent alterations which determines the spreading area of the

cell. Thus the data are not consistent, meaning the spreading area is no criteria for evaluating

any difference between MEF-wt, MEF-resc and MEF-vinΔC cells.

n=108

n=96

n=566

n=498

0

500

1000

1500

2000

2500

3000

3500

4000

MEF-wt MEF-vin(-/-) MEF-resc MEF-vin∆C

Spre

adin

g ar

ea (µ

m2 )

Figure 3-20: Spreading area of transfected and non-tranfected MEF cells. Note, that the data were inconsistent because they were not in the given parameters, determined by the MEF-wt and MEF-vin(-/-) cells. The error bars represent the standard error (SEM).

3.8 Determination of the FA per cell The FAs are the linkers between the ECM and the actin cytoskeleton. The connection to the

extracellular matrix determines cell size and shape as well as controls its survival. It was

reported that the vinculin molecule influences focal adhesion enlargement and its maturation

[108]. It is also known that the number of focal adhesions depends on the internal tension of

the cell [109]. Because of the inconsistent spreading area (see also chapter 3.7), the number of

focal adhesions in the different MEF cell lines was determined using cells of a similar

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spreading area (Figure 3-21). MEF-wt and MEF-resc cells showed a similar number of focal

adhesions, whilst MEF-vin(-/-) cells transfected with vinculinΔC displayed 23 % and MEF-

vin(-/-) cells showed 40 % less focal adhesions compared to MEF wildtype and rescue cells.

This result suggests an involvement of the lipid anchor in focal adhesion formation.

Figure 3-21: Determination of focal adhesions (FA) per cell line. (A) MEF-wt and MEF-resc show a similar number of FA. MEF-/- cells transfected with vinculin∆C show sligthly more FAs in comparison to wt and rescue cells. To stain the FA in MEF-wt and MEF-vin(-/-) cells, an antibody against paxillin (green) was used. In MEF-resc and MEF-vinΔC cells, the EGFP-molecule linked to the vinculin proteins was used (green). (B) Numbers of the FA plotted for each cell line. The spreading area of the cells used for FA determination was ~4000 µm2 in all cell lines (inset). The error bars represent the standard error (SEM).

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3.9 FRAP measurements MEF-cells expressing vinculinΔC showed a reduced focal adhesion turnover rate whilst

there was no detectable difference in the exchange rate of vinculin and vinculinΔC molecules

localized in FA, judged by FRAP measurements [81]. However, in this article they did not

report wether they determined the recovery half time in the presence or absence of

endogenous vinculin.

Figure 3-22: Turn over rate of EGFP-vinculin and vinculinΔC transiently expressed in MEF-vin(-/-) cells. The red circle marks the measured FA before and after bleaching. There was no significant difference in recovery half time (t ½) between MEF-resc and MEF-vinΔC cells. The error bars represent the standard error (SEM).

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To test the exchange rates of EGFP-linked vinculin and vinculinΔC molecules in individual

adhesion sites expressed in MEF-vin(-/-) cells, fluorescence recovery after photo bleaching

(FRAP) experiments were carried out. The technique calculates the rate of re-incorporation of

GFP-tagged vinculin variants after bleaching them with 100% laser intensity. Transfected

MEF-vin(-/-) cells expressing GFP vinculin and vinculinΔC were seeded on glass bottom

wells and the measurements were performed as described in 2.2.2.7. As described in figure 3-

22, individual adhesions were bleached by increasing the laser intensity and subjected to time

lapse video microscopy. The halftime of fluorescence recovery after photobleaching was

about 96,91s for vinculin and 119,86s for vinculinΔC molecules. According to the standard

error, there was no detectable difference in exchange rates of the protein in individual

adhesion sites, suggesting the anchor does not influence the exchange rate of individual

vinculin molecules.

3.10 2D-traction microscopy measurements During their life cells exert tractions on their surroundings including contraction, spreading,

crawling as well as invasion. These functions are associated with complex mechanical

interactions between the extracellular substrates, focal adhesion molecules such as vinculin,

cytoskeletal elements, and molecular motors. Dembo and co-workers recently demonstrated

that the traction field of a cell exerted on its surroundings can be mapped from knowledge of

the displacement field in a flexible substrate on which cells are adherent [110]. The

measurement of the displacement field is accomplished by tracking small beads, embedded on

the surface of polyacrylamide gels of a known elasticity module. The raw displacements

themselves can be seen as a qualitative map of the local tractions, generated by a single cell.

3.10.1 Strain energy measurements of MEF-vinΔC

The distribution, the size and the numbers of FAs are dependent on internal and external

forces [25; 29]. The contractile forces of the actomyosin motor activity, transferred via the

focal adhesions to the ECM-coated PAA gel, was tested in these experiments using various

MEF cell lines. Figure 3-23A shows images of MEF-wt, MEF-vin(-/-), MEF-resc and MEF-

vinΔC together with the traction-fields generated by those cells. MEF-vin(-/-) cells displayed

about 80 % and MEF-vin(-/-) cells transfected with the vinculinΔC mutant showed about 41

% less traction force during adhesion compared to the MEF-wt and rescue cells (figure 23B).

These results suggest that vinculin´s lipid anchor is important for generating the appropriate

force during cell adhesion.

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Figure 3-23: Results of the 2D-traction microscopy measurements for MEF-wt, MEF-vin(-/-), MEF-resc and MEF-vinΔC. (A) Pictures of the traction maps and the corresponding cells, generating the deformations. The pseudo-color range in the pictures of the strain fields represents the different strain field values. To visualize the MEF-resc and MEF-vinΔC, the EGFP label was used. The scale bar represents 50 µm. (B) Plot of the elastic strain energy (mean) stored in the extracellular matrix due to cell tractions. The error bars represent the standard error (SEM).

3.10.2 Strain energy measurements of different vinculin mutants

Previously, it was demonstrated by Chandrasekar et al. that the mutation of six surface

exposed basic residues on vinculin-tail to glutamine (Q) in helix 3 ( H3; K952, K956, R963,

R966) and the C-terminal lipid anchor (CT; R1060, K1061) result in impaired lipid membrane

interaction [82]. To test, whether the difference in tractions is due to the lipid binding of

vinculin in general, we performed additional 2D-traction microscopic measurements with

those vinculin mutants deficient in lipid binding. MEF-vin(-/-) cells were transfected (refer to

chapter 2.2.2.2) with EGFP-linked vinculin-H3 (K952, K956, R963, R966 to Q), vinculin-CT

(R1060, K1061 to Q) and vinculin-LD (K952, K956, R963, R966, R1060, K1061 to Q; equals

CT + H3) constructs. As described in figure 3-24A, all constructs were localized in the focal

adhesions. There was no detectable difference in FA size and distribution. The 2D traction

measurements were performed as described in chapter 2.2.5.2. The results from these

experiments were plotted as relative strain energy compared to MEF-resc cells (figure 3-24B).

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MEF-vin(-/-) cells transfected with the lipid binding deficiency variant of vinculin displaying

all six point mutations (MEF-LD) showed a reduction in strain energy of about 50 %

compared to MEF-resc cells. MEF-CT cells, expressing a vinculin variant with two point-

mutations in the lipid anchor region which also impairs lipid binding displayed the same

reduced strain energy level than MEF-LD. However, MEF-vin(-/-) cells carrying vinculin

with the mutated H3 domain, which also influence the lipid binding [82], showed a similar

strain energy level as MEF-resc cells, suggesting that only the change of vinculin´s lipid

anchor localized at the C-terminus affects force generation in MEF cells.

Figure 3-24: Results of the strain energy measurements of MEF-LD, MEF-CT and MEF-H3 cells. (A) Pictures of MEF-vin(-/-) cells transfected with EGFP labeled vinculin-LD (MEF-LD), vinculin-CT (MEF-CT) and vincunlin-H3 (MEF-H3). The actin cytoskeleton was stained with TRITC-Phalloidin in each cell line. The scale bar represents 20 µm. (B) Relative strain energy (mean) stored in the PAA gel, coated with the extracellular matrix protein collagen. The energy values of LD (K952, K956, R963, R966 and R1060, K1061 transferred to Q), CT (R1060, K1061 transferred to Q) and H3 (K952, K956, R963, R966 transferred to Q) were correlated to the rescue (resc) values. The absolute strain energy values of MEF-LD, -H3 and CT were ~1 pJ. The energy values of Y1065F were ~10 pJ (Note, the higher value was due to different gel coating). The error bars represent the standard error (SEM).

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Furthermore, a src-dependent phosphorylation site is also localized at the lipid anchor [71-

72]. The mutation of this tyrosine phosphorylation site on position 1065 to phenylalanine

(Y1065F) results in impaired cell spreading and migration. This suggests that the src

phosphorylation affects cell mechanical behavior. As demonstrated in Figure 3-24, the

vinculinY1065F mutant also showed a decreased force generation in comparison to the rescue

cells. The lipid binding abilities of vinculinY1065F are not impaired. This implies that the

interaction of the lipid anchor with the cell membrane influences cell mechanical behavior via

src-phosphorylation.

In summary, vinculin´s membrane binding of the lipid anchor influences cell stiffness and is

necessary for traction generation. 2D-traction measurements with MEF-Y1065F as well as

MEF-CT imply that the anchorage of the C-terminal arm to the cell membrane provides

vinculin for src phosphorylation. This in turn may alter the affinity of vinculin for other

binding partners such as the actin crosslinker Arp2/3 [72].

Discussion

70

4 Discussion Signal transduction, vesicle trafficking, retroviral assembly, focal adhesion formation and

other central biological processes directly involve membrane-embedded protein complexes.

Over the last decades, several structural elements have been identified that provide proteins

for transiently lipid membrane interaction [38; 111-112]. There are proteins such as spectrin,

an actin associated protein, which show pleckstrin homology (PH) domains. This binding

motif recognizes different phosphatidylinositol (PIP) lipids localized in cell membranes [111].

Another PIP binding motif is the FERM domain, named after the four proteins in which this

domain was originally described: F for Band 4.1, E for Ezrin, R for Radixin, M for Moesin

[112]. Post-translational modifications, such as myristylation or palmitoylation, may also play

critical roles in regulating membrane associations [113]. There are also secondary structure

elements such as amphiphatic helices and hydrophobic beta-sheets that interact transiently

with lipid membranes [38; 75; 80; 112]. The application of biophysical techniques including

Fourier-transformed infrared spectroscopy (FTIR), neutron reflection, electron spin resonance

(ESR), nuclear magnetic resonance (NMR) and X-ray crystallography has been helpful in

characterizing these protein-membrane interaction in vitro [114]. Unfortunately, the

mechanism and structural consequences of membrane association in cells are still poorly

understood [38; 112].

The cytoskeleton and its associated proteins such as spectrin, alpha-actinin, Arp2/3, CapZ,

talin and vinculin are important ubiquitous cellular components that determine cell shape,

locomotion and adhesion [38; 75; 112]. To influence these processes, the cytoskeleton must

be connected reversibly to transmembrane receptors in a manner that is regulated by

signalling events. In the most cases, these receptors are linked via adaptor proteins, such as

talin, alpha-actinin and vinculin, to the cytoskeleton [19-20]. A variety of these cytoskeletal

interacting proteins have been shown to associate with, and in some cases insert into, lipid

bilayers [38; 112]. Indeed, in vitro evidence for specific membrane interactions has been

provided for several proteins and their interaction partners [38; 106; 112; 115-118]. In a few

studies, the existence of such interactions was also shown in intact cells [112; 119].

Experiments using a hydrophobic photolabel localized within the membrane of intact chicken

fibroblasts demonstrated the existence of such an insertion potenial for vinculin [119]. Further

experiments identified three lipid binding regions on the 30 kDa tail domain of vinculin [76-

78]. Whilst residues 935–978 and 1020–1040 were identified as two amphiphatic alpha-

helices [76-77], residues 1052–1066 (so-called lipid anchor) are involved in beta-sheet

formation [80]. Recent studies have shown that adhesion site dynamics and turnover as well

Discussion

71

as cell motility are directly affected by vinculin-tail binding to phospholipids [78; 81-82].

Two different strategies generating lipid-binding deficient vinculin mutants were examined:

one approach was based on a variant lacking the last 15 residues of vinculin’s C-terminus [78;

81]. In another experiment six point-mutations on surface-exposed basic residues (K952,

K956, R963, R966 and RK1060/61 to Q) were introduced to achieve similar impaired lipid

membrane binding [82]. However, the in vivo relevance of vinculin´s membrane interaction is

still far from being resolved [38; 112]. It was demonstrated that the interaction of vinculin

with acidic phospholipids regulates focal adhesion disassembly [82]. Other scientists propose

membrane binding triggers vinculin´s activation and FA recruitment [59-60; 74]. However,

this work revealed clear evidence for the involvement of vinculin´s lipid anchor in the

regulation of cell mechanical properties.

4.1 Vinculin´s lipid anchor revealed membrane insertion potential It was reported that expressed vinculin deficient in lipid binding still locates at focal

adhesions, but that the FA turnover was impaired [81-82]. Moreover, the same studies showed

that binding of the vinculin-tail to lipid membranes is dependent on the capacity of vinculin to

interact with acidic phospholipids, such as phosphatidylserine (PS) and phosphatidylinositol-

4,5-bisphosphate (PIP2). Work, focussing on the possible involvement of acidic phospholipids

in adhesion site turnover left the question unanswered of whether or not the vinculin lipid

anchor interacts directly with phospholipid membranes. To this end, an investigation into the

function of the C-terminal arm (residues 1045-1066), which includes the lipid anchor, was

required as many studies had previously indicated that the lipid anchor of the vinculin-tail

interacted with phospholipids such as phosphatidylcholine, phosphatidyl-serine as well as

PIP2 [78; 81]. To gain information about the thermodynamic behavior of artificial lipids in the

presence/ absence of the 21-residue C-terminal peptide of the tail domain, we applied

differential scanning calorimetry (DSC). Using multilemellar lipid vesicles (MLVs) allowed

the detection of subtle perturbations during the lipid melting process and gave direct evidence

of peptide– lipid interaction. As described in chapter 3.1.1, the C-terminal peptide of vinculin

showed membrane insertion behavior. Since membrane–protein interactions are known to

depend on the negative surface charge of membranes, we increased the charge of lipid

vesicles to mimic the inner leaflet of the cell membrane. The results showed that the phase

transition enthalpy increased, suggesting a reduced binding of vinculin’s C-terminal region

with the membrane. This result demonstrates that the lipid interaction of the C-terminal site is

driven by the peptide’s hydrophobic potential and not by the acidic head groups of the

Discussion

72

phospholipids. It is assumed that in vivo the C-terminal arm is exposed to the cytosolic

environment and available even in the closed protein conformation. Therefore, the C-terminal

arm may tether vinculin to the cell membrane and/ or facilitate its interaction with other focal

adhesion components such as talin. Site-directed mutagenesis revealed that intermolecular

interactions between the vinculin-tail and lipids are controlled by surface exposed, basic

residues [82]. Furthermore, a change of two residues (RK 1060/61 Q) located in the C-

terminal lipid-binding site significantly impaired the interaction of the vinculin-tail with

vesicles containing acidic phospholipids. DSC measurements using a 21-residue peptide

carrying this mutation indicated, however, a higher lipid insertion potential for DMPC/

DMPG vesicles (see chapter 3.1.2). Thus, changing residues from positively charged (R, K) to

uncharged (Q) increased the hydrophobic character of this peptide and resulted in increased

interactions with moderately charged lipid vesicles. The in comparison to the unmutated C-

terminal arm increased hydrophobicity may also explain why the lipid binding of vinculin-tail

(RK 1060/61 Q) was suppressed [82]. A conformational change in vinculin-tail triggered by

the mutations may bury the C-terminal arm inside the helical bundle, which results in the loss

of the lipid anchor as membrane interaction site and reduces lipid binding.

4.2 Vinculin´s lipid anchor is involved in beta-sheet formation The protein crystals revealed no well defined secondary structure for the C-terminal arm

which hosts the lipid anchor [55; 78-79]. Protein crystals are often obtained under non-

physiological pH and the proteins are compressed due to crystal packing with minimal solvent

(water) molecules around their surface. Furthermore, a protein crystal displays only one

possible protein conformation which gives no information about flexibility and

conformational changes of the peptide during their function. In order to determine a

secondary structure of the C-terminal arm, the local conformational flexibility of this part was

explored using molecular dynamics simulations (see Chapter 3.2). The observations indicated

for nominally “basic” and “neutral” conditions an anti-parallel beta-sheet followed by an

unstructured C-terminus. In contrast, for an “acidic” environment no stable secondary

structural elements emerged and the C-terminus remained unstructured. This result indicates

that the secondary structure of the C-terminal arm depends on the ratio of positively to

negatively charged residues. As seen in repeated simulations, changing the ionic conditions

after each 10ns, the anti-parallel beta-sheet secondary structure was not influenced by its

previous starting conformation. However, the MD-study exploring the conformational

behavior of the full length vinculin-tail domain under nominal “neutral” settings,

Discussion

73

demonstrated that no anti-parallel beta-sheet emerged at the C-terminal arm (refer to chapter

3.2.2). The vinculin-tail simulation shows that the C-terminus is hindered more sterically by

the rest of protein. This implies that a conformational shift of the vinculin-tail, induced by a

local pH-shift or other interaction partners such as paxillin and actin, is necessary to free the

C-terminal arm before beta-sheet formation. Experimental support for the secondary structure

formation of the 21 residue peptide comes from CD spectroscopy (refer to chapter 3.3). An

alignment using the DICHROWEB database, indicates also two beta-strands and unstructured

parts of the C-terminal region suggesting that at a pH of 7.5 an beta-sheet conformation is

preferred. Together with the conformations observed from the MD-simulations, the following

summary of the C-terminal region secondary structure can be made: (i) residues 2–6 form an

initial beta-strand that represents the first part (residues 1045–1051) and (ii) residues 7–12

complete the beta-strand of the C-terminus of the tail domain (residues 1052–1057). The CD-

spectrum measured in presence of lipid vesicles revealed a totally different absorption curve

in comparison to the measurement without the lipid vesicles. However, the database analysis

obtained also an anti-parallel beta-sheet and mostly unstructured parts for the peptide

representing vinculin´s last 21 residues. This suggests a conformational shift of the peptide,

representing vinculin´s C-terminal arm, during lipid membrane interaction. The difference in

spectral characteristics of the 21-residue peptide during CD spectroscopy in the presence/

absence of lipid vesicles can be explained by the high variance of beta-sheet structures. This

could be due to parallel and anti-parallel orientations and different twists which are reflected

in different backbone angles [89; 120]. In order to determine the orientation of the C-terminal

peptide during lipid membrane interaction, solid state NMR-measurements were performed

(see chapter 3.4). The 15N label was placed in leucine at position 11, which is part of the beta

strand. The result may indicate membrane insertion for the beta-sheet secondary structure

element. However, to verify those results, further NMR measurements are needed with

radioactive labeled residues on different positions. A different solvent for the peptide is also

necessary. Because of the poor solubility in polar solvent at high concentrations, vinculin´s C-

terminal peptide was dissolved in formic acid. However, formic acid has the ability to

denature and degrade peptides. So it is not known if the peptide still exists and has the same

secondary structure, as determined under the conditions of CD-spectroscopy measurement. In

conclusion, this study indicates that the C-terminal 21 residues of the vinculin-tail have the

capability to associate with, or indeed insert into, artificial lipid membranes probably by

forming an anti-parallel beta-sheet.

Discussion

74

4.3 Vinculin´s lipid anchor regulates cell stiffness Mechanical tension between the interconnected assemblies of the extracellular matrix and the

cytoskeleton play a critical role in determining cell structure and function. It was

demonstrated that forces generated by and in a cell regulate many biological functions. This

implies the involvement of key molecules which recognize and generate tension for specific

cellular features [13]. Changes in cell tension influence the molecular structure and

biochemical activity of the cell. To determine the effect of the lipid anchor in respect to cell

mechanics, several EGFP-linked vinculin and vinculin-tail constructs, deficient in lipid

binding, were designed and transiently transfected in MEF-vin(-/-) cells. As demonstrated in

chapter 3.5, the N-terminal EGFP label has no obvious effect on FA localization of full length

vinculin (residues 1-1066) and vinculinΔC (residues 1-1052) molecules in MEF cells. In

contrast, MEF-vin(-/-) cells transfected with the EGFP-vtail (residues 858-1066) and -vtailΔC

(residues 858-1052) constructs showed no well-defined localization to the focal adhesions

(see chapter 3.5.2). Besides focal adhesion localization, the used EGFP-vtail and EGFP–

vtailΔC constructs also covered the actin stress-fibers outside of the focal adhesions. Because

of the lack of the vinculin-head domain, all the binding sites localized at the tail region,

including the actin binding sites, were freely accessible. This exposure of the sites could

explain the interaction of the vinculin-tail constructs with the actin filaments. In the inactive

or closed conformation, the vinculin-head masks the cryptic actin-binding sites and prevents

actin binding [55]. Only in its open or activated form, vinculin is available at the focal

adhesions [52]. This localization ensures that vinculin is connected with actin filaments only

in the focal adhesions. The covered actin filaments in both MEF-vtail and MEF-vtailΔC cells

may explain the magnetic tweezer results. The determination of the binding strength between

ECM-coated surfaces and cells displayed a higher bead detachment for MEF-vtail and MEF-

vtailΔC in comparison to MEF-vin(-/-) (see chapter 3.6.2). The vtail or vtailΔC which were

localized on the actin-filaments outside the focal adhesions may impair the actin turnover.

This disturbs the integrin clustering and FA formation around the bead. However, the stiffness

of MEF-vtail and MEF-vtailΔC was increased by ~20 % in comparison to MEF-vin(-/-) cells.

The stiffness calculation, excluding the detached beads, obtained the same stiffness values for

the MEF-vtail and MEF-vtailΔC cells as for MEF-wt cells, suggesting that the vinculin-tail is

responsible for vinculin´s mechano-coupling properties. This is consistent with results

obtained from magnetic tweezer using F9-vinculin(-/-) cells, stably expressing the vinculin-

tail domain. It was reported that the F9-vtail cells restore the wildtype phenotype

mechanically [65-66]. In contrast to MEF cells, expressing vinculin (residues 1-1066) and

Discussion

75

vinculinΔC (residues 1-1052) (see chapter 3.6.1), no difference could be detected between

MEF-vtail and MEF-vtailΔC, suggesting vinculin´s lipid anchor is involved in the regulation

of vinculin´s accessibility for other binding partners. The so-called MEF-resc cells showed

almost identical mechanical properties to their wildtype counterparts. However, MEF-vin(-/-)

cells transiently transfected with EGFP-vinculinΔC were less stiff compared to MEF-resc

cells. It is known that the vinculin-tail lacking the lipid anchor showed impaired interaction

with acidic phospholipid vesicles [78; 81]. This indicates that the difference in stiffness of

MEF-vinΔC compared to MEF-resc and MEF-wt cell might be due to the linkage of the cell

membrane with the lipid anchor of vinculin. Support for this assumption also comes from the

DSC measurements (see chapter 3.1.1) [80]. The results of the binding strength measurements

are also indicative for an involvement of the lipid anchor in controlling cell mechanical

properties. With increased pulling force, more beads detached from vinculinΔC cells than

from MEF-wt or rescue cells during the measurements. At high forces, they displayed almost

the same level of disrupted fibronectin-coated beads as the vinculin knock-out cells,

suggesting that the lipid anchor is important for the binding strength to ECM-coated surfaces.

Furthermore, the reduced binding strength of MEF-vinΔC cells to the beads may also explain

the reduced stiffness in MEF-vinΔC cells. As demonstrated in figure 4-1, the relative stiffness

values of MEF-vinΔC cells, determined at 1 nN force, were significantly reduced in

comparison to MEF-wt and MEF-resc cells. In contrast, the values of MEF-vinΔC cells

obtained at 10nN were as stiff as MEF-wt and MEF-resc cells. For the stiffness calculation at

1nN all measured beads were used, including those which could not sustain high forces and

detach during the measurement. In general, such weakly bound beads show increased

displacements during tweezers measurements in comparison to tightly bound beads [69; 65].

After these weakly bound beads had been removed from the analysis, as demonstrated for the

measurement at 10 nN force, MEF-vinΔC cells were indistinguishable from MEF-wt and

MEF-resc cells in terms of average stiffness (see chapter 3.6.1). This indicates that vinculin

must be anchored to the cell membrane with its C-terminal arm to guarantee a well

established connection between the bead and the FA, suggesting that MEF-vinΔC cells are

less stiff because of the reduced binding strength to ECM-coated surfaces. This connection

between binding strength and stiffness leads to the conclusion that there are two

subpopulations of beads in MEF-vinΔC cells. The well attached and the weakly bound ones.

The well attached beads show the same stiffness than the beads on MEF-wt and MEF-resc

cells. This suggests a difference in the turnover rate for vinculin and vinculinΔC molecules at

the focal adhesions. However, FRAP measurements in transfected MEF-vin(-/-) cells revealed

Discussion

76

no significant differences between the turnover rate of EGFP vinculin and vinculinΔC (refer

to chapter 3.9), indicating that the lipid anchor influences the recruitment of other FA proteins

which in turn control the turnover rate of the whole focal adhesion. Indeed, it was

demonstrated that the FA turnover rate in cells expressing vinculinΔC was decreased in

comparison to cells expressing intact vinculin [81]. However, FRAP measurements performed

in Keratoncytes revealed that the vinculin turnover rate in nascent FA is higher than in mature

adhesions (In: Modulation of vinculin exchange dynamics regulates adhesion site maturation

and adhesion strength; by Christoph Möhl, Norbert Kirchgeßner, Claudia Schäfer, Kevin

Küpper, Gerold Diez, Wolfgang H. Goldmann, Rudolf Merkel, Bernd Hoffmann; submitted to

Cell Motility and Cytoskeleton). During the FRAP-measurements, performed in MEF-cells

(see chapter 3.9), it could not be distinguished between nascent and mature focal adhesions.

This might also explain the similar turnover rate for vinculin and vinculinΔC, measured in

MEF-vin(-/-) cells.

0

0,2

0,4

0,6

0,8

1

1,2

1,4

11 nN 10 nN

Rel

ativ

e st

iffne

ss

MEF-wtMEF-rescMEF-vin∆CMEF-vin(-/-)

Figure 4-1: Relative stiffness of the different MEF-cells at 1nN and 10nN, respectively. The stiffness calculation at 1nN force includes also beads that detached during the measurement, whilst at 10nN only the beads could be used which remained attached. The error bars represent the standard error (SEM).

4.4 The lipid anchor affects traction generation via phosphotyrosine 1065 Additional evidence that the lipid anchor is important for regulating the mechanical properties

in cells came from 2D-traction microscopy measurements. The strain energy for MEF-wt and

rescue cells was about 8-fold higher compared to MEF-vin(-/-). In contrast, MEF-vinΔC cells

generate only 2 times more strain energy in comparison to MEF-vin(-/-) cells (refer to chapter

3.10.1). This indicates that the lipid anchor is important for internal force generation which in

turn controls FA growth and maintenance [109]. Cell adhesive forces are believed to be due to

Discussion

77

the actin ECM connection and the myosin-II driven force development [5; 7; 15]. It was

demonstrated when blocking the contractility with substances such as ML-7, BDM, KT5926

or by the over-expression of peptides like caldesmon (inhibits actin-dependent myosin-II

ATPases activity), that it leads to dissolution of focal contacts [27; 109]. This could explain

the decreased number of focal adhesions in MEF-vinΔC in comparison to MEF-wt and rescue

cells (refer to chapter 3.8).

Chandrasekar and co-workers demonstrated that the introduction of point mutations in Helix 3

(residues 935 – 978) and the C-terminal arm (residues 1045-1066) of the vinculin-tail result

also in impaired lipid membrane interaction [82]. To test the influence of each lipid binding

site to force generation, additional 2D-traction experiments were performed using MEF-vin(-

/-) cells, transfected with vinculin-H3 (mutated residues: K952, K956, R963, R966 to Q),

vinculin-CT (mutated residues: RK1060/61 to Q) and vinculin-LD (vinculin-H3 + CT). MEF-

LD as well as MEF–CT cells developed ~ 50 % less tractions in comparison to MEF-resc

cells. This was in the same range as MEF-vinΔC cells. However, MEF-H3 cells displayed

almost the same strain energy than MEF-resc cells (see chapter 3.10.2). It suggests that only

the membrane association of the lipid anchor (residue 1052-1066) is important for force

generation, whilst the lipid binding of vinculin-H3 has no influence on cell mechanical

behavior. Recently, it was demonstrated, that src phosphorylates vinculin on tyrosine 100 and

1065 [71]. It was reported when preventing c-src phosphorylation, the interaction of vinculin

with the Arp2/3 subunit p34Arc is inhibited [72]. This results in impaired cell spreading and

migration [71-72]. One of those src phosphorylation sites, tyrosine 1065, is localized in the

lipid anchor. To test the influence of Y1065 due to traction generation, 2D-traction

microscopy experiments of MEF cells transfected with vinculinY1065F were performed.

Indeed, MEF-Y1065 cells generated the same reduced strain energy compared to MEF-resc.

Those reduced values of vinculinY1065F cells were comparable to MEF-LD, MEF-CT and

MEF-vinΔC cells (refer to chapter 3.10.2). In vitro experiments demonstrated that acidic

phospholipid vesicles stimulate the phosphorylation of vinculin by src kinase [73-74]. In the

presence of divalent cations and acidic phospholipids such as phosphatidyl-inositol (PI) or

phosphatidyl-glycerol (PG), the phosphorylated fraction of vinculin was elevated more than

10-fold in comparison to vinculin in the absence of lipid vesicles. However, the

phosphorylation levels of other src targets such as alpha-actinin and casein were not affected

by artificial membranes, which implies a conformational shift of vinculin prior to

phosphorylation [73]. Indeed, protease digestion assays of vinculin in the presence and

absence of such lipid vesicles demonstrated a higher degradation of the membrane bound

Discussion

78

vinculin peptide in comparison to unbound vinculin, suggesting that the interaction of

vinculin with the lipid membrane induces a conformational shift [74]. This implies that the

membrane interaction of the lipid anchor provides vinculin´s src-dependent phosphorylation

which in turn is necessary for traction generation.

4.5 The Model Previous studies indicate that the loss of the lipid anchor as well as the introduction of six

surface-exposed basic residues on the vinculin-tail result in impaired membrane interaction

[78; 81-82]. Cells transfected with vinculin variants containing those alterations displayed

reduced cell migration and adhesion, because of their decreased FA turnover rate [81-82]. The

function of vinculin´s lipid binding is still a controversy. Chandrasekar and co-workers

demonstrated a link between the interaction of vinculin with acidic phospholipids and focal

adhesion disassembly [82]. An increase of PIP2, caused by an over-expression of PIP 5-kinase

alpha, resulted in vinculin delocalization from the focal adhesion whilst vinculin deficient in

lipid binding remained in the focal adhesion complex. In contrast, it was proposed that

membrane binding triggers the displacement of vinculin´s head domain from the tail which

could lead to its activation and FA recruitment [59-60; 74]. There is also evidence for other

proteins to activate vinculin. It is conceivable that there are several intermediate

conformations of vinculin, dependent on its binding partners. In vitro and in vivo experiments

demonstrated that talin, alpha-actinin as well as actin alter also vinculin´s conformation and

activate the molecule [52-54]. Indeed, no difference in FA localization between vinculin and

vinculinΔC cells could be detected. The shape and size of adhesions were also similar. FRAP

measurements in transfected MEF-vin(-/-) cells revealed also no significant differences

between the turnover rate of EGFP vinculin and vinculinΔC (refer to chapter 3.9), indicating

the lipid anchor influences the recruitment of other FA components which in turn controls the

turnover rate of the entire adhesion plaque.

Furthermore, an involvement in cell mechanical regulation was only determined for the C-

terminal lipid anchor. In contrast, the lipid binding site localized on helix 3 (residues 935 –

978) had no influence on cell mechanical behavior. This may imply that each lipid binding

site of vinculin has its own purpose and function, independent from each other. Crystal

structure analysis of vinculin indicates that the lipid anchor region is not buried inside the

vinculin molecule in the closed conformation [55; 79]. This allows the lipid anchor to interact

with the hydrophobic core of lipid membranes even in the closed or auto-inhibited state of

vinculin (current work) [80]. In summary, it is possible that the lipid anchor of vinculin

Discussion

79

guides the molecule to the cell membrane where it becomes prepared for FA recruitment.

Localized at the membrane, vinculin is in close proximity to other focal components like talin

and actin, which have also the ability to activate vinculin for focal adhesion recruitment. The

impaired membrane localization of vinculin lacking the lipid anchor may result in an

incomplete conformational switch that could result in an impaired connection to the actin

cytoskeleton in MEF-vinΔC cells.

The following model was drafted from the described results. As demonstrated in figure 4-2,

the vinculin molecule becomes activated by talin, actin or phospholipids. Further, the

anchorage of vinculin´s C-terminal to the cell membrane prepares the molecule for src-

dependent phosphorylation [73-74]. Identified c-src targets of vinculin are tyrosine 100 and

tyrosine 1065 [71]. The last tyrosine is part of the lipid anchor. The phosphorylation of

vinculin by c-src enables the molecule to bind Arp2/3, an actin nucleator and crosslinker [72;

121]. This interaction may reinforce the connection of the focal adhesion complex to the actin

cytoskeleton, which enables the cells to generate the appropriate pre-stress and strain energy

for adequate cell adhesion and migration. Therefore the lipid anchor regulates the mechano-

coupler vinculin via membrane binding dependent src-phosphorylation, followed by Arp2/3

recruitment. Thus, additional experiments are necessary to prove this mechanism.

Figure 4-2: Lipid binding of vinculin via its lipid anchor provides the molecule for src-dependent phosphorylation.

4.6 Outlook The lipid anchor regulates cell adhesion and migration by src dependent phosphorylation. The

MEF-cells are a model system for further investigations. In the next step, the phosphorylation

Discussion

80

level of tyrosine 1065 should be determined by Western blotting for all those mutant variants

used in this study. Since it is known that tyrosine 100 is also phosphorylated by src, it is also

necessary to characterize its influence on cell mechanical behavior, using the magnetic

tweezer and 2D-traction microscopy. Additional FRAP measurement with vinculin and

vinculin deficient in lipid binding as well as deficient in tyrosine 100/ 1065 phosphorylation

in nascent focal adhesions should reveal a difference in the turnover rate. It is also necessary

to determine the influence of phosphorylation to membrane binding. Molecular dynamics

simulations of the phosphorylated and unphosphorylated vinculin-tail together with lipid

membranes are another possibility to obtain further informations. Additional pull-down

experiments with the phosphorylated vinculin-tail, incubated with lipid vesicles, would also

be helpful to clarify these questions.

References

81

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6 Appendix

6.1 Visualization of focal contacts using the TEM The focal adhesion sites connect the actin cytoskeleton via the integrin receptor to the

extracellular matrix. To investigate the density of focal adhesion plaques, we characterized

the FA distribution of paxillin proteins in mouse embryonic fibroblasts. The cells were

cultured and prepared for TEM characterization as described under 2.2.2.7. The gold labeled

paxillin molecules are near a longitudinal structure of 200-400 nm length and 10-20nm

diameter, which could represent actin fibers. Judged by the density of the beads around the

actin fiber, it is believed to observe a focal contact. The next step clearly is to extend the

investigations to MEF-vin(-/-) cells, for visualizing any differences between MEF-wt and

knock out cells to obtain differences in the density of paxillin at the focal adhesions.

Figure 6-1: Electronmicroscopic images of MEF-wt cells at 50.000x (A) and 130.000x (B) magnification. B represents the magnified spot of image A, labeled with the red circle. The arrows in picture B mark the gold labeled secondary antibodies which represent the paxillin molecules. Note, that the accumulation of the paxillin molecules may represent a focal adhesion complex.

Appendix

89

6.2 pcEGFP-N2 expression vector

MCS: NheI 851 GCTTATCGAA ATTAATACGA CTCACTATAG GGAGACCCAA GCTGGCTAGC 901 GCTACCGGTC GCCACCATGG TGAGCAAGGG CGAGGAGCTG TTCACCGGGG M V S K G E E L F T G V Frame 2 951 TGGTGCCCAT CCTGGTCGAG CTGGACGGCG ACGTAAACGG CCACAAGTTC V P I L V E L D G D V N G H K F Frame 2 1001 AGCGTGTCCG GCGAGGGCGA GGGCGATGCC ACCTACGGCA AGCTGACCCT S V S G E G E G D A T Y G K L T L Frame 2 1051 GAAGTTCATC TGCACCACCG GCAAGCTGCC CGTGCCCTGG CCCACCCTCG K F I C T T G K L P V P W P T L V Frame 2 1101 TGACCACCCT GACCTACGGC GTGCAGTGCT TCAGCCGCTA CCCCGACCAC T T L T Y G V Q C F S R Y P D H Frame 2 1151 ATGAAGCAGC ACGACTTCTT CAAGTCCGCC ATGCCCGAAG GCTACGTCCA M K Q H D F F K S A M P E G Y V Q Frame 2 1201 GGAGCGCACC ATCTTCTTCA AGGACGACGG CAACTACAAG ACCCGCGCCG E R T I F F K D D G N Y K T R A E Frame 2 1251 AGGTGAAGTT CGAGGGCGAC ACCCTGGTGA ACCGCATCGA GCTGAAGGGC V K F E G D T L V N R I E L K G Frame 2

Appendix

90

1301 ATCGACTTCA AGGAGGACGG CAACATCCTG GGGCACAAGC TGGAGTACAA I D F K E D G N I L G H K L E Y N Frame 2 1351 CTACAACAGC CACAACGTCT ATATCATGGC CGACAAGCAG AAGAACGGCA Y N S H N V Y I M A D K Q K N G I Frame 2 1401 TCAAGGTGAA CTTCAAGATC CGCCACAACA TCGAGGACGG CAGCGTGCAG K V N F K I R H N I E D G S V Q Frame 2 1451 CTCGCCGACC ACTACCAGCA GAACACCCCC ATCGGCGACG GCCCCGTGCT L A D H Y Q Q N T P I G D G P V L Frame 2 1501 GCTGCCCGAC AACCACTACC TGAGCACCCA GTCCGCCCTG AGCAAAGACC L P D N H Y L S T Q S A L S K D P Frame 2 1551 CCAACGAGAA GCGCGATCAC ATGGTCCTGC TGGAGTTCGT GACCGCCGCC N E K R D H M V L L E F V T A A Frame 2 BsrGI AflII XhoI 1601 GGGATCACTC TCGGCATGGA CGAGCTGTAC AAGTCCGGAC TCCTTAAGCT G I T L G M D E L Y K S G L L K L Frame 2 XbaI ApaI PmeI 1651 CGAGTCTAGA GGGCCCGTTT AAACCCGCTG ATCAGCCTCG ACTGTGCCTT E S R G P V * Frame 2

Appendix

91

6.3 cDNA of vinculin mouse BsiWI MluI PvuI AflIII BsiEI PfoI BsrFI 1 ATGCCAGTGT TTCATACGCG TACGATCGAG AGCATCCTGG AGCCGGTGGC TACGGTCACA AAGTATGCGC ATGCTAGCTC TCGTAGGACC TCGGCCACCG +H3N- M P V F H T R T I E S I L E P V A Frame 1 1% 6 % % % SexAI DraIII BssSI 51 GCAGCAGATC TCGCACCTGG TGATTATGCA CGAGGAGGGC GAGGTGGACG CGTCGTCTAG AGCGTGGACC ACTAATACGT GCTCCTCCCG CTCCACCTGC Q Q I S H L V I M H E E G E V D G Frame 1 ? 20 a ? a a % % % BsgI EagI BglI BsiEI 101 GCAAAGCCAT TCCTGACCTC ACCGCGCCCG TAGCCGCCGT GCAGGCGGCC CGTTTCGGTA AGGACTGGAG TGGCGCGGGC ATCGGCGGCA CGTCCGCCGG K A I P D L T A P V A A V Q A A Frame 1 40 NciI 151 GTCAGCAACC TCGTCCGGGT TGGAAAAGAG ACTGTTCAGA CCACTGAGGA CAGTCGTTGG AGCAGGCCCA ACCTTTTCTC TGACAAGTCT GGTGACTCCT V S N L V R V G K E T V Q T T E D Frame 1 60 201 TCAGATTCTG AAGAGAGATA TGCCACCAGC CTTTATTAAG GTTGAAAATG AGTCTAAGAC TTCTCTCTAT ACGGTGGTCG GAAATAATTC CAACTTTTAC Q I L K R D M P P A F I K V E N A Frame 1 80 HindIII 251 CTTGCACCAA GCTTGTTCAG GCAGCCCAGA TGCTTCAGTC AGACCCATAC GAACGTGGTT CGAACAAGTC CGTCGGGTCT ACGAAGTCAG TCTGGGTATG C T K L V Q A A Q M L Q S D P Y Frame 1 100 301 TCGGTTCCTG CGCGGGATTA CCTCATTGAC GGCTCTAGGG GAATCCTTTC AGCCAAGGAC GCGCCCTAAT GGAGTAACTG CCGAGATCCC CTTAGGAAAG S V P A R D Y L I D G S R G I L S Frame 1 V-head D1 (6-252) % % BbvCI 351 TGGCACATCT GACCTACTGC TTACCTTCGA TGAGGCTGAG GTTCGTAAAA ACCGTGTAGA CTGGATGACG AATGGAAGCT ACTCCGACTC CAAGCATTTT G T S D L L L T F D E A E V R K I Frame 1 120 % ApoI 401 TTATTAGGGT TTGCAAAGGA ATTTTGGAAT ATCTTACAGT GGCAGAGGTA AATAATCCCA AACGTTTCCT TAAAACCTTA TAGAATGTCA CCGTCTCCAT I R V C K G I L E Y L T V A E V Frame 1 140

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Tth111I 451 GTGGAAACTA TGGAAGACTT GGTCACTTAC ACAAAGAATC TTGGGCCAGG CACCTTTGAT ACCTTCTGAA CCAGTGAATG TGTTTCTTAG AACCCGGTCC V E T M E D L V T Y T K N L G P G Frame 1 160 HincII 501 AATGACTAAG ATGGCCAAAA TGATTGATGA GAGACAGCAG GAGTTGACTC TTACTGATTC TACCGGTTTT ACTAACTACT CTCTGTCGTC CTCAACTGAG M T K M A K M I D E R Q Q E L T H Frame 1 180 % % ? For1 DraIII FalI 551 ACCAGGAACA CCGTGTGATG TTGGTGAACT CAATGAACAC TGTCAAAGAG TGGTCCTTGT GGCACACTAC AACCACTTGA GTTACTTGTG ACAGTTTCTC Q E H R V M L V N S M N T V K E Frame 1 ? 200 601 CTGCTTCCTG TTCTCATTTC AGCTATGAAG ATTTTTGTTA CAACCAAAAA GACGAAGGAC AAGAGTAAAG TCGATACTTC TAAAAACAAT GTTGGTTTTT L L P V L I S A M K I F V T T K N Frame 1 HindIII ApoI 651 CTCAAAAAAC CAAGGAATAG AAGAAGCTTT GAAAAATCGA AATTTTACTG GAGTTTTTTG GTTCCTTATC TTCTTCGAAA CTTTTTAGCT TTAAAATGAC S K N Q G I E E A L K N R N F T V Frame 1 220 Rev1 701 TAGAAAAGAT GAGTGCTGAA ATTAACGAGA TCATTCGTGT GTTACAACTC ATCTTTTCTA CTCACGACTT TAATTGCTCT AGTAAGCACA CAATGTTGAG E K M S A E I N E I I R V L Q L Frame 1 240 A 751 ACTTCCTGGG ATGAAGATGC CTGGGCCAGC AAGGACACTG AAGCCATGAA TGAAGGACCC TACTTCTACG GACCCGGTCG TTCCTGTGAC TTCGGTACTT T S W D E D A W A S K D T E A M K Frame 1 ? a 260 HaeII HgaI AfeI BsaHI PflMI 801 GAGAGCGCTG GCGTCCATAG ACTCCAAATT GAACCAGGCC AAAGGTTGGC CTCTCGCGAC CGCAGGTATC TGAGGTTTAA CTTGGTCCGG TTTCCAACCG R A L A S I D S K L N Q A K G W L Frame 1 280 TspGWI PasI 851 TCCGTGACCC CAATGCCTCC CCAGGGGATG CTGGAGAGCA GGCCATCAGG AGGCACTGGG GTTACGGAGG GGTCCCCTAC GACCTCTCGT CCGGTAGTCC R D P N A S P G D A G E Q A I R Frame 1 300 BsgI 901 CAGATCTTAG ATGAAGCTGG AAAAGTTGGT GAACTTTGTG CAGGCAAGGA GTCTAGAATC TACTTCGACC TTTTCAACCA CTTGAAACAC GTCCGTTCCT Q I L D E A G K V G E L C A G K E Frame 1 AvrII 951 ACGCAGGGAG ATCCTAGGAA CCTGCAAAAT GCTAGGGCAG ATGACTGACC TGCGTCCCTC TAGGATCCTT GGACGTTTTA CGATCCCGTC TACTGACTGG R R E I L G T C K M L G Q M T D Q Frame 1 320

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BmrI 1001 AAGTGGCTGA CCTCCGAGCC AGAGGACAAG GAGCTTCCCC AGTGGCCATG TTCACCGACT GGAGGCTCGG TCTCCTGTTC CTCGAAGGGG TCACCGGTAC V A D L R A R G Q G A S P V A M Frame 1 340

TaiI HpyCH4IV BanII BmgBI 1051 CAGAAGGCCC AGCAAGTGTC TCAGGGGCTC GACGTGCTTA CCGCCAAAGT GTCTTCCGGG TCGTTCACAG AGTCCCCGAG CTGCACGAAT GGCGGTTTCA Q K A Q Q V S Q G L D V L T A K V Frame 1 360 BsmI AlfI 1101 GGAGAATGCA GCTCGGAAGC TGGAAGCCAT GACGAACTCA AAGCAGAGCA CCTCTTACGT CGAGCCTTCG ACCTTCGGTA CTGCTTGAGT TTCGTCTCGT E N A A R K L E A M T N S K Q S I Frame 1 380 BamHI EciI PflMI 1151 TTGCAAAGAA GATTGATGCT GCCCAGAATT GGCTGGCGGA TCCAAATGGT AACGTTTCTT CTAACTACGA CGGGTCTTAA CCGACCGCCT AGGTTTACCA A K K I D A A Q N W L A D P N G Frame 1 V-head D2 (253-485) 400 1201 GGACCTGAGG GAGAAGAACA GATTCGAGGG GCTTTGGCTG AAGCTCGGAA CCTGGACTCC CTCTTCTTGT CTAAGCTCCC CGAAACCGAC TTCGAGCCTT G P E G E E Q I R G A L A E A R K Frame 1 BsrBI 1251 GATTGCAGAA TTATGTGATG ATCCTAAGGA GAGAGATGAC ATCCTCCGCT CTAACGTCTT AATACACTAC TAGGATTCCT CTCTCTACTG TAGGAGGCGA I A E L C D D P K E R D D I L R S Frame 1 420 1301 CCCTTGGAGA GATAGCTGCT CTGACCTCTA AACTAGGAGA CTTGCGAAGA GGGAACCTCT CTATCGACGA GACTGGAGAT TTGATCCTCT GAACGCTTCT L G E I A A L T S K L G D L R R Frame 1 440 XhoI PspXI 1351 CAGGGGAAAG GAGACTCGCC AGAGGCTCGA GCCTTGGCTA AACAAGTGGC GTCCCCTTTC CTCTGAGCGG TCTCCGAGCT CGGAACCGAT TTGTTCACCG Q G K G D S P E A R A L A K Q V A Frame 1 460 PstI 1401 GACGGCACTA CAGAACCTGC AGACCAAAAC CAACAGGGCC GTGGCCAACA CTGCCGTGAT GTCTTGGACG TCTGGTTTTG GTTGTCCCGG CACCGGTTGT T A L Q N L Q T K T N R A V A N S Frame 1 % % 480

Appendix

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XhoI PvuII PspXI 1451 GCAGACCTGC CAAAGCAGCT GTCCACCTCG AGGGCAAGAT TGAACAGGCG CGTCTGGACG GTTTCGTCGA CAGGTGGAGC TCCCGTTCTA ACTTGTCCGC R P A K A A V H L E G K I E Q A Frame 1 V % % K 500 DrdI TaqII 1501 CAGCGGTGGA TTGATAACCC CACAGTAGAT GACCGTGGAG TCGGTCAGGC GTCGCCACCT AACTATTGGG GTGTCATCTA CTGGCACCTC AGCCAGTCCG Q R W I D N P T V D D R G V G Q A Frame 1 % % TspGWI BbvCI BspHI 1551 TGCCATCCGT GGACTTGTGG CTGAGGGGCA TCGGCTGGCC AATGTCATGA ACGGTAGGCA CCTGAACACC GACTCCCCGT AGCCGACCGG TTACAGTACT A I R G L V A E G H R L A N V M M Frame 1 520 PpuMI BsmFI BslFI BslFI FalI AccI 1601 TGGGACCTTA TCGCCAAGAT CTTCTTGCCA AATGTGACCG TGTAGACCAG ACCCTGGAAT AGCGGTTCTA GAAGAACGGT TTACACTGGC ACATCTGGTC G P Y R Q D L L A K C D R V D Q Frame 1 540 V-head D3 (493-717) PvuII BlpI 1651 CTAACAGCTC AGCTGGCTGA CCTGGCTGCC CGAGGGGAGG GGGAGAGTCC GATTGTCGAG TCGACCGACT GGACCGACGG GCTCCCCTCC CCCTCTCAGG L T A Q L A D L A A R G E G E S P Frame 1 560 1701 TCAGGCGAGA GCACTTGCAT CCCAGCTTCA GGACTCCTTA AAGGATCTTA AGTCCGCTCT CGTGAACGTA GGGTCGAAGT CCTGAGGAAT TTCCTAGAAT Q A R A L A S Q L Q D S L K D L K Frame 1 580 BpuEI BciVI 1751 AAGCCCAGAT GCAGGAAGCT ATGACTCAAG AGGTATCCGA TGTTTTCAGC TTCGGGTCTA CGTCCTTCGA TACTGAGTTC TCCATAGGCT ACAAAAGTCG A Q M Q E A M T Q E V S D V F S Frame 1 600 BsaXI BsaXI BsaXI for2 BsaXI 1801 GATACTACAA CTCCTATCAA GCTGTTGGCA GTAGCCGCCA CTGCTCCTCC CTATGATGTT GAGGATAGTT CGACAACCGT CATCGGCGGT GACGAGGAGG D T T T P I K L L A V A A T A P P Frame 1 1851 TGATGCACCC AATAGGGAAG AGGTATTTGA TGAAAGGGCA GCCAATTTTG ACTACGTGGG TTATCCCTTC TCCATAAACT ACTTTCCCGT CGGTTAAAAC D A P N R E E V F D E R A A N F E Frame 1 620 S 1901 AAAACCATTC AGGAAGGCTT GGAGCCACAG CAGAGAAGGC GGCTGCTGTT TTTTGGTAAG TCCTTCCGAA CCTCGGTGTC GTCTCTTCCG CCGACGACAA N H S G R L G A T A E K A A A V Frame 1 640

Appendix

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Rev2 BsmI 1951 GGTACTGCCA ATAAATCGAC AGTGGAAGGC ATTCAGGCAT CTGTGAAGAC CCATGACGGT TATTTAGCTG TCACCTTCCG TAAGTCCGTA GACACTTCTG G T A N K S T V E G I Q A S V K T Frame 1 ? a % % a 660 2001 AGCCCGAGAA CTCACTCCCC AGGTCATCTC CGCTGCTCGG ATCTTACTGA TCGGGCTCTT GAGTGAGGGG TCCAGTAGAG GCGACGAGCC TAGAATGACT A R E L T P Q V I S A A R I L L R Frame 1 680 BstEII BsaI 2051 GGAACCCTGG TAACCAGGCT GCTTATGAAC ATTTTGAGAC CATGAAGAAC CCTTGGGACC ATTGGTCCGA CGAATACTTG TAAAACTCTG GTACTTCTTG N P G N Q A A Y E H F E T M K N Frame 1 700 2101 CAGTGGATTG ATAATGTTGA AAAAATGACA GGGCTGGTGG ACGAGGCTAT GTCACCTAAC TATTACAACT TTTTTACTGT CCCGACCACC TGCTCCGATA Q W I D N V E K M T G L V D E A I Frame 1 % 2151 TGATACCAAG TCTCTGTTGG ATGCTTCTGA AGAAGCAATT AAAAAAGACC ACTATGGTTC AGAGACAACC TACGAAGACT TCTTCGTTAA TTTTTTCTGG D T K S L L D A S E E A I K K D L Frame 1 ? 720 NcoI SspI 2201 TGGACAAGTG TAAGGTAGCC ATGGCCAATA TTCAGCCTCA GATGCTGGTC ACCTGTTCAC ATTCCATCGG TACCGGTTAT AAGTCGGAGT CTACGACCAG D K C K V A M A N I Q P Q M L V Frame 1 740 BsaWI 2251 GCTGGAGCAA CCAGTATTGC TCGTCGGGCC AACCGGATTC TGCTGGTTGC CGACCTCGTT GGTCATAACG AGCAGCCCGG TTGGCCTAAG ACGACCAACG A G A T S I A R R A N R I L L V A Frame 1 V-head D4 (719-835) 760 AloI AloI PpuMI 2301 TAAGAGGGAG GTAGAGAACT CTGAGGACCC GAAGTTCCGA GAGGCTGTGA ATTCTCCCTC CATCTCTTGA GACTCCTGGG CTTCAAGGCT CTCCGACACT K R E V E N S E D P K F R E A V K Frame 1 % % % a % % % 780 NcoI 2351 AAGCTGCCTC TGATGAACTG AGCAAAACAA TCTCCCCCAT GGTGATGGAT TTCGACGGAG ACTACTTGAC TCGTTTTGTT AGAGGGGGTA CCACTACCTA A A S D E L S K T I S P M V M D Frame 1 K 800 PfoI 2401 GCCAAGGCTG TGGCTGGAAA CATCTCTGAC CCTGGCCTGC AAAAGAGCTT CGGTTCCGAC ACCGACCTTT GTAGAGACTG GGACCGGACG TTTTCTCGAA A K A V A G N I S D P G L Q K S F Frame 1 810 D EcoRV BamHI 2451 CCTGGACTCA GGATATCGGA TCCTGGGAGC TGTGGCCAAG GTCAGAGAAG GGACCTGAGT CCTATAGCCT AGGACCCTCG ACACCGGTTC CAGTCTCTTC L D S G Y R I L G A V A K V R E A Frame 1 820 Q For3 2501 CCTTCCAACC TCAGGAGCCT GACTTCCCGC CTCCTCCACC AGACCTTGAA GGAAGGTTGG AGTCCTCGGA CTGAAGGGCG GAGGAGGTGG TCTGGAACTT F Q P Q E P D F P P P P P D L E Frame 1

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% 840% % SacI EcoICRI 2551 CAGCTACGAC TAACTGATGA GCTCGCTCCT CCTAAGCCAC CTCTGCCTGA GTCGATGCTG ATTGACTACT CGAGCGAGGA GGATTCGGTG GAGACGGACT Q L R L T D E L A P P K P P L P E Frame 1 860 BsmFI BslFI AloI BslFI AloI 2601 GGGTGAAGTC CCTCCACCCA GGCCCCCACC ACCAGAAGAG AAGGATGAAG CCCACTTCAG GGAGGTGGGT CCGGGGGTGG TGGTCTTCTC TTCCTACTTC G E V P P P R P P P P E E K D E E Frame 1 879 880 2651 AGTTCCCTGA GCAGAAAGCT GGTGAGGTGA TTAACCAGCC AATGATGATG TCAAGGGACT CGTCTTTCGA CCACTCCACT AATTGGTCGG TTACTACTAC F P E Q K A G E V I N Q P M M M Frame 1 896 900 2701 GCCGCCAGGC AGCTCCACGA TGAAGCTCGG AAATGGTCTA GCAAGGGCAA CGGCGGTCCG TCGAGGTGCT ACTTCGAGCC TTTACCAGAT CGTTCCCGTT A A R Q L H D E A R K W S S K G N Frame 1 % ? Vt Helix I ? 913 AlfI 2751 TGACATCATT GCAGCAGCCA AGCGCATGGC TCTGCTGATG GCAGAGATGT ACTGTAGTAA CGTCGTCGGT TCGCGTACCG AGACGACTAC CGTCTCTACA D I I A A A K R M A L L M A E M S Frame 1 920 Vt Helix II KpnI BanI Acc65I 2801 CTCGGCTGGT AAGAGGGGGC AGTGGTACCA AGCGGGCACT TATTCAGTGT GAGCCGACCA TTCTCCCCCG TCACCATGGT TCGCCCGTGA ATAAGTCACA R L V R G G S G T K R A L I Q C Frame 1 P 938 940 %944 % % EcoRV StuI 2851 GCCAAGGATA TCGCCAAGGC CTCTGATGAG GTGACGAGGT TGGCCAAGGA CGGTTCCTAT AGCGGTTCCG GAGACTACTC CACTGCTCCA ACCGGTTCCT A K D I A K A S D E V T R L A K E Frame 1 Vt Helix III 960 2901 GGTTGCCAAG CAGTGCACAG ATAAGCGGAT TAGAACCAAT CTCTTACAGG CCAACGGTTC GTCACGTGTC TATTCGCCTA ATCTTGGTTA GAGAATGTCC V A K Q C T D K R I R T N L L Q V Frame 1 972 % % %975% % 980% % % % 2951 TATGCGAGCG AATCCCAACT ATAAGCACCC AGCTCAAAAT CCTATCCACA ATACGCTCGC TTAGGGTTGA TATTCGTGGG TCGAGTTTTA GGATAGGTGT C E R I P T I S T Q L K I L S T Frame 1 a % % a a % 1000 3001 GTGAAGGCCA CTATGCTGGG CCGGACCAAC ATCAGTGATG AGGAGTCTGA CACTTCCGGT GATACGACCC GGCCTGGTTG TAGTCACTAC TCCTCAGACT V K A T M L G R T N I S D E E S E Frame 1 Vt Helix IV 1005 % 1013%% % % 3051 GCAGGCCACA GAGATGCTGG TTCATAATGC CCAGAACCTC ATGCAGTCTG CGTCCGGTGT CTCTACGACC AAGTATTACG GGTCTTGGAG TACGTCAGAC Q A T E M L V H N A Q N L M Q S V Frame 1 1020 % ? ? Vt Helix V

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3101 TGAAGGAGAC TGTGCGAGAG GCTGAAGCTG CTTCAATCAA AATCCGAACA ACTTCCTCTG ACACGCTCTC CGACTTCGAC GAAGTTAGTT TTAGGCTTGT K E T V R E A E A A S I K I R T Frame 1 % 1040 T 1046% % KpnI BanI Acc65I 3151 GATGCTGGCT TTACTCTGCG CTGGGTCAGA AAGACTCCCT GGTACCAGTA CTACGACCGA AATGAGACGC GACCCAGTCT TTCTGAGGGA CCATGGTCAT D A G F T L R W V R K T P W Y Q * Frame 1 1052 Flexible C-tail 1060 1066 3201 G C

D1: residues 1-252 D2: residues 253-485 D1-D4 Vinculin-head D3: residues 493-717 D4: residues 719-835 Poly-Proline: residues 838-878 D5: residue 898-1066 → also called vinculin-tail

Appendix

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6.4 List of Figures Front-page: AFM-image of a MEF-cell (kindly provided by Tilman Schäffer) Figure 1-1: Simplified cartoon of a cross section through part of the intestine ......................... 1 Figure 1-2: Cellular mechano responsiveness ........................................................................... 2 Figure 1-3: Schematic of cell contacts visualized in epithelial cells.......................................... 5 Figure 1-4: Illustration of focal adhesions. ................................................................................ 6 Figure 1-5: Schematics of the rac/ rho dependent focal adhesion formation............................. 9 Figure 1-6: Schematic of the vinculin molecule ...................................................................... 11 Figure 1-7: The vinculin-tail with its three potential lipid binding sites.................................. 13 Figure 2-1: Cloning strategy for the generation of the pEGFP expression system.................. 21 Figure 2-2: Cloning strategy for the vinculin cDNA. .............................................................. 22 Figure 2-3: Sample preparation for TEM measuremnts........................................................... 26 Figure 2-4: Phase transition thermogram of DMPG/DMPC vesicles.. .................................... 28 Figure 2-5 A schematic of the DSC apparatus. ........................................................................ 29 Figure 2-6: Simulated solid-state NMR spectra....................................................................... 31 Figure 3-1: Vinculin peptide representing vincuin´s C-terminal arm. ..................................... 39 Figure 3-2: Interaction of the mutated vinculin-tail peptide (RK1060/61 Q) with MLVs. ..... 40 Figure 3-3: Secondary structure of vinculin´s C-terminal arm under “basic” conditions........ 43 Figure 3-4: Secondary structure of vinculin´s C-terminal arm under “neutral” conditions..... 44 Figure 3-5: Secondary structure of vinculin´s the C-terminal arm under “acidic” conditions 45 Figure 3-6: MD-simulations of the 21 residue peptide ............................................................ 46 Figure 3-7: Secondary structure of the vinculin-tail (residue 890-1066)................................. 47 Figure 3-8: Results from CD-spectroscopic measurements. .................................................... 48 Figure 3-9: Result of the NMR measurement. ........................................................................ 50 Figure 3-10: Expression of vinculin and vinculinΔC in MEF cells. ........................................ 51 Figure 3-11: Expression of EGFP linked vinculin-tail and vinculin-tailΔC. ........................... 52 Figure 3-12: Expression of vinculin and vinculinY1065F in MEF-vin(-/-) cells. ................... 53 Figure 3-13: Co-localization of the auto-fluorescent beads with the focal adhesions. ............ 54 Figure 3-14: Result of magnetic tweezer measurements ......................................................... 56 Figure 3-15: Differential cell stiffness of transfected and non-transfected MEF cells ............ 57 Figure 3-16: Geometric mean of the power-law exponent b .................................................. 58 Figure 3-17: Magnetic tweezer measurements of MEF-vtail and MEF-vtailΔC..................... 60 Figure 3-18: Differential cell stiffness of MEF-vtail and vtailΔC cells................................... 61 Figure 3-19: The mean of the power-law exponent b for MEF-vtail and MEF-vtailΔC. ........ 62 Figure 3-20: Spreading area of transfected and non-tranfected MEF cells.............................. 63 Figure 3-21: Determination of focal adhesions (FA) per cell line ........................................... 64 Figure 3-22: Turn over rate of EGFP-vinculin and vinculinΔC .............................................. 65 Figure 3-23: Results of the 2D-traction microscopy measurements ........................................ 67 Figure 3-24: Strain energy measurements of MEF-LD, MEF-CT, MEF-H3, MEF-Y1065F.. 68 Figure 4-1: Relative stiffness of the different MEF-cells at 1 nN and 10 nN .......................... 76 Figure 4-2: Model..................................................................................................................... 79 Figure 6-1: Electronmicroscopic images of MEF-wt cells. ..................................................... 88

Acknowledgements

99

Acknowledgements I extend my sincere thanks to Prof. Dr. Wolfgang H. Goldmann for giving me this great

opportunity, to write my PhD thesis under his guidance. He was always very encouraging and

supportive to my work.

I am grateful to Prof. Dr. Hojatollah Vali for his wonderful supervision, encouragement, and

support during my term at McGill University. I enjoyed working in his group and I am sure

this experience will help me in my future career.

I am thankful for Prof. Dr. Ben Fabry´s financial support and valued discussions throughout

this work. My special thanks go to Dr. James Smith, for his supervision and guidance.

I am grateful to my dear colleague Anna H. Klemm. Without her help and value advices this

work would not have been possible.

I greatly acknowledge the help from Philip Kollmannsberger, Thorsten Koch, Daniel

Paranhos Zitterbart, Carina Raupach and Stefan Münster with useful discussions and

suggestions.

Many thanks to Dr. Fereshte Azzari, University of McGill, for her kind help and support to

this work. I would like to include Aleksandra A. Krukiewicz, Barbara Reischel, Ulrike Scholz

and Christine Albert for her good technical support in this regard.

I am thankful to Dr. Wolfgang H. Ziegler, IZKF Leipzig, for the collaboration, help and

support.

I thank Prof. Dr. Hannappel and Prof. Dr. Brandstätter for their acceptance to be involved in

the thesis committee. Thank you to Prof. Dr. Tilman Schäffer for the AFM-image of a MEF-

cell.

My sincere thanks go to Nicole Dietzel for helping me with my thesis. Many thanks also to

her husband Reinhard Dietzel for the technical assistance.

I thank Dr. Bernd Hoffmann and Christoph Möhl, Forschungszentrum Jülich, as well as Prof.

Dr. Burkard Bechinger (University of Strassbourg) for the useful discussions and support. I

also thank Felix List, University of Regensburg and Volker Wirth for their help and good

chats.

My sincere thanks go to all my good friends (Barbara, Bruno, Michi, Kerstin, Martin, Fariba,

Christian, Ralf and Uwe). All of you deserve a PhD degree.

I am grateful to my family for their constant encouragement, support and trust on me which

helped me to get to this stage.

I am very happy to have Martina in my life; I thank her for her unconditional love, patience,

encouragement and support.

Acknowledgements

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101

Curriculum Vitae Personal Datas: Name: Gerold Diez Adress: Friedrich-Ebert-Str 26

97318 Kitzingen Germany

Date of Birth: 22nd January 1978 Telephone: 049131/8525602 E-mail: [email protected] Education: 09/1984 - 06/1993 Qualifizierter Hauptschulabschluss: Hauptschule Nüdlingen 09/1993 - 06/1995 Staatlich geprüfter Sozialpfleger: Berufsschule Schweinfurt 09/1995 - 06/1996 Realschulabschluss: Berufsaufbauschule Münnerstadt 09/1996 - 06/1999 Abitur: Bayernkolleg Schweinfurt 07/1999 - 07/2000 Wehrersatzdienst 10/2000 - 08/2005 Study of Biochemistry

08/2005 Diploma in Biochemistry: Department of Biochemistry I, University of Regensburg (Germany)

09/2005 - 12/2008 PhD-Student at the Department of Medical Physics and

Technology, Friedrich-Alexander-Universität Erlangen/ Nuremberg (Germany)

12/2008 PhD in Biophysics

Characterization of vinculin´s lipid anchor regionThermodynamic evidence of non-muscle myosin...

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