The Effect of Thymosin Beta 4 on Cell Mechanics and Motility

Die Wirkung von Thymosin Beta 4 auf Zellmechanik und Motilität

Der Naturwissenschaftlichen Fakultät/dem Fachbereich Physik der Friedrich-Alexander-Universität Erlangen-Nürnberg zur Erlangung des Doktorgrades Dr. rer. nat.

vorgelegt von Leila Minori Jaafar aus Kuala Lumpur, Malaysia

Als Dissertation genehmigt von der Naturwissenschaftlichen Fakultät/vom Fachbereich Physik der Friedrich-Alexander-Universität Erlangen-Nürnberg

Tag der mündlichen Prüfung: 6. Juni, 2014

Vorsitzender des Promotionsorgans: Prof. Dr. Johannes Barth Gutachter: Prof. Dr. Ben Fabry Prof. Dr. Ewald Hannappel

Table of Contents

Zusammenfassung ...... 1 Summary ...... 3 1 Introduction...... 5 1.1 Cell motility: a biochemical and biophysical process ...... 6 1.1.1 The biochemical regulation of cell migration 6 1.1.2 Cellular mechanics and dynamics: biophysical processes regulating cell migration 9

1.2 Thymosin β4: structure and function ...... 12

1.2.1 The molecular structure of Tβ4 12

1.2.2 Tβ4: a multi-functional protein in health and pathogenesis 13 1.3 Open questions ...... 18 2 Material and Methods ...... 20 2.1 Biological and biochemical methods ...... 21 2.1.1 Cells and cell culture 21 2.1.2 Cloning cassettes of EGFP-Actin for generating the GSP 22 2.1.3 Ligation 22 2.1.4 Cloning of EGFP-Actin into the pIRESpuro3 selection vector 24 2.1.5 Transfection of the EGFP-Actin vector construct 24 2.1.6 Stable transfection into mammalian cell lines 24 2.1.7 Immunofluorescence and fluorescence microscopy 25 2.1.8 Synthesis of lipid vesicles 27

2.1.9 Synthesis of Tβ4-Alexa488 labeled peptide 27 2.2 Biophysical methods ...... 29 2.2.1 Three-dimensional collagen cell invasion assay 29 2.2.2 Two-dimensional cell motility assay 29 2.2.3 Cell-spreading assay 30 2.2.4 Two-dimensional cellular traction microscopy 30 2.2.5 Magnetic tweezer microrheology 32 2.2.6 Nano-scale particle tracking 34 2.2.7 Differential scanning calorimetry (DSC) 37 2.2.8 Fluorescence correlation spectroscopy 39

iii

3 Results ...... 43

3.1 The effect of extracellular Thymosin β4 on the motility of SW480 cells ...... 44 3.1.1 Three-dimensional cell migration 44 3.1.2 Two-dimensional cell motility 46

3.2 The effect of extracellular Tβ4 on cell morphology ...... 49 3.2.1 Analyses of SW480 cell morphology in 3D 49 3.2.2 Analyses of SW480 cell morphology in 2D 51

3.3 Effect of extracellular Tβ4 on cell compliance, dynamics and contractility ...... 54 3.3.1 Creep response and cell mechanics 54 3.3.2 Two-dimensional cellular tractions 57 3.3.3 Nanoscale particle tracking of cytoskeletal-bound beads: studying the time- dependant activity of Tβ4 on cytoskeletal dynamics 59

3.4 Tβ4 and ILK: The Biphasic Switch in 3D Cell Invasion Assays ...... 64

3.4.1 Role of ILK as a molecular switch involved in the Tβ4 biphasic response ...... 64

3.4.2 Bypassing RhoA inhibition: the effect of calyculin-A on Tβ4-induced cell invasion 66

3.5 Tβ4 – membrane interaction ...... 69

3.5.1 Determining the binding potential of Tβ4 to lipid membranes using differential scanning calorimetry (DSC) 69 3.5.2 Fluorescence correlation spectroscopy (FCS) of membrane-bound fluorescent Tβ4 71 4 Discussion ...... 75

4.1 The effects of Tβ4 on Cellular Mechanics and Motility ...... 76

4.1.1 Tβ4 regulates the stiffness and contractility of SW480 cells 76

4.1.2 Tβ4 regulation of acto-myosin contractility is involved in the biphasic response of SW480 cell morphology and motility 78

4.2 Tβ4 and the cell lipid membrane: a novel mechanism for outside-in signaling ...... 82

4.3 The Tβ4-ILK Molecular Switch ...... 84 Bibliography...... 88

List of Figures

Figure 1.1 Two-dimensional cell migration...... 7 Figure 1.2 The dynamic turnover of the actin cytoskeleton at the leading edge of a cell...... 8

Figure 1.3 Molecular structure of Tβ4 ...... 13 Figure 1.4 Extracellular application and genetic overexpression: the multi-faceted role of

Tβ4 in health and disease...... 14 Figure 2.1 Cell lines...... 21 Figure 2.2 Stable transfection of NIH 3T3 cells with an EGFP-Actin plasmid vector...... 23 Figure 2.3 Close-up of an NIH 3T3 mouse fibroblast stably transfected with the pIRES- EGFP-Actin construct...... 25 Figure 2.4 Actin fluorescent staining with Alexa Fluor 546-phalloidin...... 26 Figure 2.5 Fluorescent staining of vinculin...... 27

Figure 2.6 Linking the Alexa Fluor 488 fluorophore to Tβ4 molecules...... 28 Figure 2.7 Two-dimensional traction microscopy...... 31 Figure 2.8 2D traction microscopy images...... 31 Figure 2.9 Magnetic tweezer measurements...... 33 Figure 2.10 Nano-scale particle tracking of fibronectin coated superparamagnetic beads ...... 36 Figure 2.11 Analysis of nano-scale particle tracking...... 36 Figure 2.12 Phase transition of synthetic lipid vesicles...... 38 Figure 2.13 Differential Scanning Calorimetry (DSC): experimental setup...... 39 Figure 2.14 Schematic diagram of the Fluorescence Correlation Spectroscopy (FCS) detection volume...... 40 Figure 2.15 Principle of Fluorescence Correlation Spectroscopy (FCS)...... 42 Figure 3.1 Analysis of SW480 cell migration in three-dimensional collagen gels...... 45 Figure 3.2 Analysis of two dimensional SW480 cell motility...... 47

Figure 3.3 Biphasic response of SW480 cell morphology to Tβ4 concentration in 3- dimensional collagen gels...... 50

Figure 3.4 Biphasic response of SW480 cell morphology to Tβ4 concentration on two- dimensional surfaces...... 53

Figure 3.5 Stiffness of Tβ4 induced SW480 cells...... 55

Figure 3.6 Viscoelasticity of Tβ4 induced SW480 cells...... 56

Figure 3.7 Traction force maps of SW480 cells: contractility of Tβ4 stimulated cells on 2D substrates...... 58

Figure 3.8 The effect of exogenous Tβ4 on the contractility of SW480 cells on 2D substrates...... 59

Figure 3.9 Short term effects of extracellular Tβ4 on cytoskeletal remodeling...... 61

Figure 3.10 Long term effects of extracellular Tβ4 on cytoskeletal remodeling...... 62

Figure 3.11 Tβ4-ILK molecular switch...... 65

Figure 3.12 The combined effect of calyculin-A and Tβ4 on the 3D invasion of SW480 cells into collagen gels...... 67

Figure 3.13 Lipid-binding qualities of Tβ4 to DMPG/DMPC lipids analyzed using differential scanning calorimetry (DSC)...... 70

Figure 3.14 Lipid membrane binding of Tβ4 measured through fluorescence correlation spectroscopy...... 72

Figure 3.15 The effect of acidic lipid head groups on the binding of Tβ4 -Alexa488 on membranes...... 74

Figure 4.1 3D stucture of Tβ4...... 83

Figure 4.2 The regulation of SW480 cellular mechanics in response to extracellular Tβ4 concentration – a biphasic molecular switch mechanism...... 86

Abbreviations

2D two dimensional 3D three dimensional ADF actin depolymerizing factors ADP adenosine diphosphate AKT protein kinase b Arp2/3 actin-related protein 2/3 ATP adenosine triphosphate BSA bovine serum albumin CCD charged-coupled device Cdc42 cell division control protein 42 cDNA complementary deoxyribonucleic acid dH20 distilled water DMEM Dulbecccco’s modified Eagle’s medium DMPC 1,2-Dimyristoyl-sn-Glycero-3-Phosphocholine DMPG Dimyristoyl-L- -phosphatidylglycerol DMPS 1,2-Dimyristoyl-sn-glycero-3-phospho-L-serine DNA deoxyribonucleicα acid dNTPs deoxynucleotide triphosphate DSC differential scanning calorimetry DTT Dithiothreitol EDTA ethylenediaminetetraacetic acid EGFP enhanced green fluorescent protein EMT epithelial-mesenchymal transition FAK focal adhesion kinase FCS fluorescence correlation spectroscopy FCS fetal calf serum FITC fluorescein isothiocyanate GSP green stoic puppies (EGFP-actin stably transfected NIH 3T3 cells) GTP guanosine triphosphate HEPES (Hydroxyethyl-Piperazine Ethanesulafonic Acid)-Saline-Albumin-Gelatin HIV-1 human immunodeficiency virus type 1 HRP horseradish peroxidase ILK integrin-linked kinase IRES viral internal ribosome entry site KC keratinocyte-derived chemokine

Kd binding constant kDa kilo Dalton MAPK Mitogen-activated protein kinase MARCKS myristoylated alanine-rich C kinase substrate MCS multiple cloning site MIP-2 macrophage inflammatory protein 2 MLC myosin light chain MLCK myosin light chain kinase MLVs Multilamellar lipid vesicles

vii

MMP matrix metalloproteinase MSD mean square displacement MT6-MMP membrane-type 6 metalloproteinases NF-kB PAA polyacrylamide PBS phosphateNuclear factor buffer κ B saline PCR polymerase chain reaction pCum cumulative probability PINCH particularly interesting new cysteine-histidine rich protein PMN polymorphonuclear macrophage Rac related to A and C protein kinase Rho ras homolog RhoA ras homolog A ROCK Rho-kinase SD standard deviation SE standard error SEM scanning electron microscope Shc SH2-containing collagen-related proteins Src Rous sarcoma oncogene cellular homolog SUVs small unilamellar vesicles

Tb10 Thymosin beta-10

Tb4 Thymosin beta-4

Tb5 Thymosin beta-5 TGA transglutaminase

Tm annealing temperature

TM melting temperature Tris HCl tris(hydroxymethyl)aminomethane hydrochloric acid buffer VEGF vascular endothelial growth factor WASP Wiskott-Aldrich syndrome protein

viii

Zusammenfassung

Die Zellmotilität oder die -migration sind fundamentale Prozesse in der Entwicklung und Funktionsweise von multizellulären Organismen im gesunden wie pathologischen Zustand, beispielsweise bei der embryonalen Entwicklung und bei der Metastase von Krebszellen. Zellen bewegen sich dadurch, dass deren Zellgerüste stetig neu aufgebaut werden und sich dadurch verformen. Die koordinierte Polymerisation von G-Actin- Monomeren zu F-Actin-Filamenten und Stressfasern findet dabei am vorangehenden Rand der Zelle statt. Aufgrund der Polymerisation wirkt dort eine Kraft, die eine Nettobewegung der Zelle verursacht. Der Zusammen- und Abbau dieser F-Actin- Fasern werden durch eine Vielzahl Actin-bindender Proteine gesteuert. Um diese Actin-Filamente bei Bedarf sehr schnell zu produzieren, benötigen Zellen einen großen Pool an nicht-polymerisiertem G-Actin, das für die Polymerisation zur Verfügung steht.

Thymosin beta-4 (Tβ4) ist ein kleines, ca. 5 kDa großes Peptid, das einen 1:1 Komplex mit G-Actin formt und die spontane Polymerisation von G-Actin zu F-Actin hemmt. Größere Konzentrationen von Tβ4, etwa bis 500 µM, sind in den Zellen aller Säugetiere vorhanden, Spuren des Peptids können auch im Blutserum nachgewiesen werden. Kürzlich wurde beobachtet, dass eine zelluläre Überexpression bzw. eine extrazelluläre Behandlung mit Tβ4 mit dem bösartigen Verlauf und der Migration von Krebszellen mehrerer Linien korreliert. Darunter auch mit den sonst nichtinvasiven kolorektalen Krebszellen SW480. Da der Prozess der Zellmigration und die Verformung des Zellgerüsts essentiell zusammenhängen, untersucht diese Arbeit die Auswirkung von extrazellulärem Tβ4 auf die Mechanik des Zellgerüsts, die Kontraktilität und die Motalität von Zellen im Experiment.

Die Untersuchungen der Steifheit und Kontraktilität von Zellen werden mit magnetic tweezer und 2D-tractions microscopy durchgeführt. Die Ergebnisse zeigen eine deutliche biphasische Reaktion mit steigenden Konzentrationen an extrazellulären Tβ4. Der höchste Wert an Steifheit und Kontraktilität wird mit SW480-Zellen und einer Konzentration an Tβ4 von ca. 0,2 µM erreicht. Es wird nachgewiesen, dass die biphasische Reaktion in der Steifheit und Kontraktilität der Zellen mit Änderungen in der Morphologie und Migrationsverhalten derselben einhergeht. Zellen die mit 0,2 µM Tβ4 behandelt wurden, zeigen eine betont längliche, mesenchymale Morphologie. Ihr Migrationsverhalten auf einer flachen Oberfläche ist bei dieser Konzentration ausgeprägt räumlich gerichtet. In Experimenten mit 3D-Kollagengelen verhalten sich die Zellen bei dieser Konzentration stark invasiv. Eine längliche Zellmorphologie zusammen mit einer erhöhten Zellkontraktilität bei Behandlung mit 0,2 µM Tβ4 sind zwei starke Indikatoren für eine verstärkte Zellmigration in einer 3D Kollagenmatrix. Die Messungen zeigen jedoch keine Auffälligkeiten bei der Behandlung mit Konzentrationen von Tβ4 größer als 1 µM.

Es bleibt zunächst unerklärt, warum durch eine spezifische Konzentration an extrazellulären Tβ4 eine so starke Reaktion hervorgerufen wird, zumal eine 100mal höhere Konzentration des Peptids bereits in der Zelle vorhanden ist. Außerdem ist für Tβ4 derzeit auch kein Zelloberflächenrezeptor oder eine andere Möglichkeit für Signalübertragung von außerhalb der Zelle nach innen bekannt. Die Aminosäureanalyse von Tβ4 zeigt jedoch, dass ca. 60% des Peptids mit Clustern von basischen und hydrophoben Aminosäuren besetzt ist. Es ist bekannt, dass basische Aminosäuren mit sauren Bereichen auf einer Lipidmembran wechselwirken und dass hydrophobe Aminosäuren als Anker wirken können. Diese beiden Eigenschaften würden es dem Peptid erlauben an Zellmembrane zu binden und in einer Konfiguration darin einzudringen, die es dem Peptid erlaubt mit weiteren membranassoziierten Partnern zu interagieren, um weitere intrazelluläre Signale zu veranlassen. Die Bindung von Tβ4 an synthetische Lipidmembranen wird in dieser Studie mittels differential scanning calorimetry (DSC) und fluorescence correlation spectroscopy (FCS) nachgewiesen.

Mit der Einbettung und Bindung von extrazellulärem Tβ4 an die Zellplasmamembran wird somit ein möglicher Mechanismus aufgezeigt, wie die lokale Konzentration des Peptids stark erhöht werden kann. Gleichzeitig wird beobachtet, dass die Diffusiongeschwindigkeit des Peptids stark abnimmt und somit die Reaktionswahrscheinlichkeit mit anderen membranassoziierten Signalproteinen wie beispielsweise integrin-linked kinase (ILK) und particularly interesting new cysteine- histidine rich protein (PINCH) erhöht wird.

Die Messergebnisse aus dieser Studie und die oben beschriebenen Beobachtungen zeigen, dass membrangebundenes Tβ4 mit Membran- und Zellgerüsten-assoziierter Proteine (ILK und PINCH) wechselwirken und dabei ein funktionales Komplex ausbilden kann. Bei geringer Konzentration fördert dieses Komplex die Zellkontraktilität. Hohe Konzentrationen des Komplexes hemmen die Kontraktilität hingegen, was zu einer biphasischen Reaktion bei ansteigender Konzentration an extrazellulären Tβ4 führt. Diese Arbeit zeigt somit auf, dass eine Stimulierung mit extrazellulären Tβ4 als biphasischer molekularer Schalter wirken kann, um beispielsweise die Motilität von SW480 Zellen zu steuern.

Summary

Cell motility or migration is a process crucial for the development and function of a multi-cellular organism in a normal or pathological state, for example, during embryonal development and cancer metastasis. In a moving cell, the cytoskeleton is remodeled dynamically. Coordinated polymerization of G-actin monomers into F-actin filaments and stress fibers at the leading edge of the cell provides force for the net movement of the cell. The assembly and disassembly of these F-actin fibers are regulated by a myriad of actin-binding proteins. To assemble actin filaments rapidly when needed, cells keep a large pool of non-polymerized G-actin ready for polymerization.

Thymosin beta-4 (Tβ4), a small 5 kDa peptide, forms a 1:1 complex with G-actin and

inhibits the spontaneous polymerization of G-actin to F-actin. Tβ4 is found in trace amounts in serum and in abundant concentrations of up to 500 µM in virtually every mammalian cell. Recently, the cellular overexpression and extracellular

administration of Tβ4 has been correlated with the malignant progression and migration of several tumor cell lines including the otherwise non-invasive SW480 colorectal cancer cells. As the process of cell migration involves substantial changes in

the cytoskeleton, the effect of extracellular Tβ4 on cytoskeletal mechanics, contractility, and motility was measured in this study.

Cell stiffness and contractility measurements using the magnetic tweezer and 2D tractions microscopy showed a distinct biphasic dose response with increasing

concentrations of extracellular Tβ4; the highest level of stiffness and contractility was

recorded in SW480 cells stimulated with 0.2 µM Tβ4. The biphasic response in cell stiffness and contractility were accompanied accordingly with changes in cell

morphology and migration. Cells treated with 0.2 µM Tβ4 displayed a pronounced elongated, mesenchymal morphology, migrated with higher directional persistence on flat 2D surfaces, and became highly invasive in a 3D collagen invasion assay. An elongated cell morphology, together with an increased contractility, are two potent factors for enhanced cell migration in a 3D collagen matrix in response to stimulation

with 0.2 µM Tβ4. All parameters return, however, to baseline levels at concentrations larger than 1 µM.

It is puzzling how minute amounts of extracellular Tβ4 can elicit such a massive response, considering that a more than 100 fold higher concentration of the peptide is already present in the cell. Furthermore, a cell surface receptor or another way of

outside-in signaling for Tβ4 has yet to be found. Amino acid analysis of Tβ4 showed that clusters of basic and hydrophobic residues make up about 60% of the entire

peptide. Basic amino acids are known to interact with acidic domains on a lipid membrane and hydrophobic residues may function as an anchor. Both factors would enable the peptide to bind and insert into the cell membrane in a configuration allowing it to interact with membrane-associated partners to trigger downstream signaling events. Differential scanning calorimetry (DSC) and fluorescence correlation spectroscopy (FCS) measurements using fluorescently labeled Tβ4 and synthetic lipid membranes were used to demonstrate the binding of Tβ4 to the synthetic lipid membranes. The insertion and binding of extracellular Tβ4 to the cell plasma membrane offers a mechanism of how the local concentration of the peptide can be greatly increased. At the same time its diffusion speed is greatly decreased so as to increase the reaction probability with other membrane-associated signaling proteins such as integrin-linked kinase (ILK) and particularly interesting new cysteine- histidine rich protein (PINCH).

Data from this study together with previous observations suggest that membrane- bound Tβ4 interacts with membrane and cytoskeleton associated proteins, ILK and PINCH, to form a functional complex that promotes cell contractility at lower concentrations, and suppresses contractility at higher concentrations resulting in a biphasic response to increasing concentrations of applied Tβ4. As a conclusion, this study postulates that extracellularly applied Tβ4 may act as a biphasic molecular switch to regulate cell motility in SW480 cells.

1 Introduction

1.1 Cell motility: a biochemical and biophysical process Cell motility or migration is a process crucial for the development and function of a multi- cellular organism in a normal or pathological state, for example, during embryonal development and cancer metastasis. One of the main processes involved in cell motility functions is the regulation of the actin filament assembly and disassembly in the cytoskeleton. Although three types of cytoskeletal filaments are involved in cell motility, namely actin filaments, microtubules and intermediate filaments, actin filaments play the leading role [1-2]. In this chapter, the biochemical and biophysical aspects of the cytoskeleton and its associated constituents in cell motility are described.

1.1.1 The biochemical regulation of cell migration The movement of a cell is controlled by coordination of several complex biochemical and biomechanical mechanisms. At the start of the migration process on two dimensional surfaces, a cell develops protrusions at its leading edge (see Figure 1.1). These structures, which can be sheet-like (lamellipodia) or finger-like (filapodia), are driven to protrude forth by the polymerization of actin monomers (G-actin) into actin filaments (F-actin). New focal adhesions, multi-protein assemblies through which the mechanical force and regulatory signals of a cell are transmitted, are formed at the front of the cell. As the cell contracts, focal adhesions are detached at the rear and the cell body is translocated forward. Cycles of cell contraction and relaxation occur through the interaction of head domains of myosin, an ATP dependent motor protein, and actin filaments. This acto-myosin contraction is regulated by the phosphorylation of the light chain domain of myosin (MLC: myosin light chain) [1].

Each phase during cell movement, as described above, is modulated by biochemical signaling mechanisms controlling actin polymerization and acto-myosin contraction. Responsible for this regulation are mainly the Rho family of GTPases (signal transducers), which include Cdc42, Rac and Rho [1, 3-4]. Cdc42 is involved with the formation of filapodia, whereas Rac with that of lamellipodia. The activation of Rho increases actin polymerization, the development of stress fibers and acto-myosin contraction. Cdc42 and Rac can both activate the Wiskott-Aldrich syndrome protein (WASP), which leads to the recruitment of Arp2/3 and actin polymerization. The activation of LIM-kinase 1 by Cdc42 and LIM-Kinase 2 by Rac leads to the increase in cofilin phosphorylation which inhibits the disassembly of actin filaments. Rho, however, stimulates the formation of stress fibers and focal adhesions through the activation of Rho-kinase (ROCK). Not only does ROCK phosphorylate MLC, which is responsible for acto-myosin contraction, it can inhibit the dephosphorylation and deactivation of MLC by inhibiting MLC phosphatase.

Introduction 1

Figure 1.1 Two-dimensional cell migration. Four phases can be distinguished during the process of a cell moving on a two-dimensional surface (extracellular substrate seen in blue). (1) Protrusion: at the leading edge of a cell, actin polymerization stimulated through Rac and Cdc42 leads to the formation membrane protrusions in the shape of the finger-like filapodia or sheet-like lamellipodia. (2) Adhesion: new focal adhesions (light yellow) are formed. These connect the actin cytoskeleton to the extracellular substrate via cell surface integrins. (3) Translocation: Forces generated by acto-myosin (green rods) contraction modulated by Rho cause the forward translocation of the cell body and nucleus. (4) Retraction: Lastly, disassembly of focal adhesions at the rear end of the cell causes the retraction of the trailing edge.

1 Introduction

The dynamic turnover of the cytoskeleton from filamentous F-actin to G-actin monomers and back can be examined largely in the cell lamellopodia, the major organ in responsible for moving the cell forward. Proteins taking part in this dynamic process are localized in the leading edge of the lamellipodia. The spatial orientation of key proteins allows for the highly specific compartmentalization of actin-filament assembly at the front and disassembly at the rear of the lamellopodia, which together with myosin motors generates a directional force for forward movement of the cell [2].

Figure 1.2 The dynamic turnover of the actin cytoskeleton at the leading edge of a cell. Shown above is a schematic diagram of the dynamic process of the assembly of G-actin monomers into F-actin and disassembly again into G-actin monomers. This cycle occurs typically at the leading edge of a moving cell. (Diagram taken from [2])

Introduction 1

As seen in Figure 1.2, starting at the leading edge of the lamellipodia of a moving cell, the actin fibers form a dense branched network with barbed ends of F-actin filaments facing the edge of the cell and pointed end of the filaments forming Y-shaped, ~70° angle junctions with other filaments. These filaments are joined together by the Arp2/s complex. Upon the activation by the WASP (1), the Arp 2/3 complex can nucleate G-actin assembly into F-actin [5] (2) and caps the free pointed ends of the filaments, starts filament growth at the side of another filament and/or catch the pointed end of an existing filament (3) [6]. Actin-filaments at the leading edge are made of mostly ATP-actin and ADP-actin and do not bind ADF/cofilin. Filament growth at the leading edge of a lamellipodia is a rapid process when compared to depolymerization of filaments. Further elongation of the actin fibers is prevented by capping proteins (4). The ADF/cofilin complex promotes the depolymerization of ADP-actin at the distal end of the lamellipodia (5). ADF-cofilin and ADP-Actin complexes that break away from the F-actin filament ends are in equilibrium with the ADF-cofilin and ADP-actin monomers (6). The slow process of nucleotide exchange on the G-actin monomer is inhibited by ADF-cofilin but is increased by profilin (7) [7-8]. Dissociated ADP-actin then undergoes nucleotide exchange with ATP to form ATP-

actin. At this end, Thymosin beta 4 (Tβ4) binds to the free G-actin monomers and sequesters them to make a ready pool of intracellular G-actin to be polymerized into

F-actin when needed (8) [9-10]. The interaction of G-actin and Tβ4 is, however, dependent on the binding of nucleotides ADP and ATP to G-actin. The binding affinity

of Tβ4 to ATP-G-actin is about 50 times higher than to ADP-G-actin [11]. The G-actin binding protein profilin promotes the nucleotide exchange of ADP-G-actin to ATP-G-

actin, whereas Tβ4 inhibits this process.

During cell migration, mechano-chemical signaling occurs in an outside-in and inside- out manner through the interaction of the cytoskeleton, the focal adhesion complex proteins and integrins, cell surface extracellular matrix receptors. Upon binding ligands in the extracellular matrix, integrins transmit these extracellular signals into the cell by modulating the activity of Rho GTPases and focal adhesion proteins such as focal adhesion kinase (FAK), Src, integrin-linked kinase (ILK) and Shc [12-15]. These proteins in turn transmit the signals to the cytoskeleton.

1.1.2 Cellular mechanics and dynamics: biophysical processes regulating cell migration In the past, the phenomenon of a moving cell has been explained through the various molecular and biochemical signaling pathways. However, the process of cell motility has distinct physical aspects which have remained undiscovered until recently. For example, cell traction and adhesion forces are involved cell morphology and anchoring of the cell onto extracellular substrates during migration. The cell also 9

1 Introduction

generates contractile forces to propel itself forward on two-dimensional surfaces as well as to overcome physical barriers within a three-dimensional matrix. In a moving cell, the cytoskeleton and focal adhesions are force-transmitting structures which undergo constant dynamic reorganization.

Recently, efforts have been made to elucidate the purely physical and mechanical processes governing cell motility. The mechanics and dynamics of the cytoskeleton and its constituents, parameters both of which influence the rate of cell migration, have been studied extensively using high-force magnetic tweezers. With this method, the application of forces can be precisely applied to the cytoskeleton. This method can be used to apply up to 100 nN force onto Ø 4.5 fibronectin-coated beads which bind to the cytoskeleton via cell surface integrins [16-17]. Experiments with the magnetic tweezer have been used to uncover the non-linear material properties and viscoelastic characteristics of the cytoskeleton of various cell types. Through these experiments, it was discovered that cells control their mechanical properties by generating an internal cytoskeletal prestress powered actively by motor proteins. Further studies were carried out using the magnetic tweezer to examine the force transducing function of focal adhesion proteins including vinculin [18-19] and FAK [20] Recent studies also using this method have shown that the loss of filamin A leads to a more viscous-like cytoskeleton in M2 human melanoma cells [21].

Increased cell stiffness has been correlated with increased cell contractility, characteristic for motile cells [19]. Using two-dimensional traction microscopy, a method developed initially by Pelham and Wang et al. [22] , contractile forces produced by individual cells can be measured. In this method, cells are cultivated on collagen-coated polyacrylamide (PAA) gel with a known Young’s modulus. The contractility of each cell can be characterized as the elastic strain energy stored in the extracellular matrix because of cell tractions. For example, studies with vinculin wild- type and vinculin deficient mouse embryonal fibroblasts have shown that vinculin- expressing cells generated higher traction forces when compared with vinculin knockout cells. As a result, vinculin expressing fibroblasts are able to invade through a three-dimensional collagen matrix by utilizing traction forces to actively overcome steric barriers in the extra-cellular matrix [23]. Recent advances in three-dimensional traction microscopy can be used to extend studies of cellular forces in cell invasion into a three dimensional environment [24-25].

As mentioned previously, the dynamic reorganization of the cytoskeleton is one of the major processes occurring in a migrating cell. Cytoskeletal dynamics can be examined by tracking the spontaneous motion of cytoskeletal-bound markers such as fibronectin coated beads [26]. This technique enables the measurement of stress fluctuations within the cytoskeleton, which is constantly transmitting stresses to the

Introduction 1

extra-cellular matrix, especially during the process of migration. The fibronectin coated beads bound to the cytoskeletal network of acto-myosin stress fibers and were observed to move, powered by ATP, in a directionally persistent super-diffusive manner in virtually all cell types. Through these biophysical measurements, a physical model of a tensed and continuously reorganizing system has complemented the widely established knowledge on the biochemical process of the constant assembly and disassembly of actin stress fibers in the cytoskeleton.

In the past, simple methods have been used to qualitatively examine cell migration such as the Boyden-Chamber assay and in vitro scratch/wound assays. However, these methods do not precisely quantify the speed and persistence of cell movement. Recent advances in microscopy techniques have overcome these barriers. By using nanoscale particle tracking methods similar to studying stress fluctuations in the cytoskeleton, bright cell-internal structures such as the nucleus can be used as an indicator for the entire cell and can be tracked. With this, the mean square displacement of a moving cell can be obtained, which gives information on the speed and persistence of its movement. The latter is particularly interesting for studies using chemoattractants. Great improvement in three dimensional microscopy and automated methods of tracking cells in collagen gel has enabled the accurate quantification of cell invasion [23, 27].

1 Introduction

1.2 Thymosin β4: structure and function

Among other cytoskeleton-associated proteins, Tβ4 has been determined as a key peptide in the regulation of the cytoskeleton. In this section, a detailed description of the structure and function of this peptide is given. Furthermore, the role of Tβ4, as described in past literature, in health and pathological development is discussed.

1.2.1 The molecular structure of Tβ4

Thymosin β4 (Tβ4) is a small 5 kDa peptide belonging to the family of thymosins originally isolated from the calf thymus. The 43-amino acid polypeptide has its isoelectric point at a pH of 5.1 (slightly acidic) and is water soluble [28]. Although

largely unstructured in water, NMR spectroscopic analysis shows that Tβ4 tends to form α-helices in aqueous solutions containing fluorinated alcohols. The two helices formed involve amino acid residues 4 – 16 and 30 – 40 [29-30]. This biologically active peptide is present in high concentrations (of up to 500 µM) in virtually every mammalian cell line [31-33], being highly conserved across all species. Up to 1% of

the whole Tβ4 blood level has been found in serum largely due to its release from

damaged cells. Erythrocytes remain the exception; no Tβ4 has been found in these cells [28, 34].

Tβ4 is known as the main intracellular G-actin sequestering protein, as established by

Safer et al. initially in human platelets [35-36]. Tβ4 was found to form a 1:1 complex with G-actin and, thereby, inhibit the spontaneous polymerization of G-actin through steric hindrance [28, 37]. On the molecular level, the sequence motif 17LKKTETQEK25

in Tβ4 has been established by previous studies to be directly involved with G-actin binding. Further NMR analysis has come to the conclusion that for the binding and

sequestration of G-actin, the N-terminus of Tβ4 must form an α-helix [29-30]. Additionally, the three hydrophobic residues (6M, 9I and 12F) and the first three residues of the actin-binding motif (17L, 18K and 19K) are essential for the interaction.

Introduction 1

Figure 1.3 Molecular structure of Tβ4 Adapted from Safer et al [37], Kabsch et al. [38] and Huff et al [28]). Tβ4 is a 5 kDa peptide with 43 amino acid residues. Shown above is a schematic list of the three main domains, the two α-helices spanning from residues 4 – 16 and 30 – 40 and the G-actin binding ’LKKTETQEK’ motif. Tβ4 binds to the G-actin subdomains 3, 1 and 2 through the interaction of G-actin residues E167, D1-E4 and Q41 with the Tβ4 residues K3, K18 and K38. Through steric hindrance, Tβ4 inhibits the spontaneous polymerization of G-actin monomers.

1.2.2 Tβ4: a multi-functional protein in health and pathogenesis

Tβ4 in dermal wound closure The course of wound healing involves stages of regulated process such as inflammation, cell proliferation, the formation of blood vessels, migration of

fibroblasts and keratinocytes, and deposition of tissue collagen. Tβ4 has been shown in different in vitro and in vivo animal studies to play a major role in wound healing. The

concentration of Tβ4 is highest in platelets and polymorphonuclear leukocytes. These cells are responsible for releasing factors which attract other cells involved in closing

the wound. Wound serum is known to have a Tβ4 concentration of up to 13 µg/ml, largely due to release from damaged platelets[39].

1 Introduction

Figure 1.4 Extracellular application and genetic overexpression: the multi-faceted role of Tβ4 in health and disease. (adapted from [34]) Aside from its classical role as an actin sequestering protein, Tβ4 has been documented to induce other cellular processes. Here, a distinction between the extracellular induction and the genetic overexpression of Tβ4 has been made. In situ animal studies have shown that extracellular application of Tβ4 has lead to cardio-protective effects following a myocardial infarct [40] and enhanced wound healing in the dermis and cornea [41-46]. In these studies, it has been reported that Tβ4 interacts with PINCH which results in the activation of integrin-linked kinase (ILK). In addition to its enhancing effects on cell motility, ILK modulates the activity of AKT (protein kinase B) in cell survival processes. Furthermore, the release of inflammatory cytokines, chemokines and proteases are regulated. Genetic overexpression of Tβ4 in SW480 cells, however, has led to the epithelial-mesenchymal transition (EMT) of SW480 cells. Through the simultaneous down-regulation of E-cadherin, up-regulation of b-catenin and MMP-7, the increase in the invasiveness of SW480 cells and the subsequent distant metastases of human colorectal cells in mouse models have been documented [47-48].

In the wound, Tβ4 has been documented to promote several processes of wound healing. Firstly, it increases the formation of new blood vessels from the pre-existing vasculature, a process known as angiogenesis, which is essential for tissue growth 14

Introduction 1

[41-42]. Both keratinocyte and endothelial cell migration into the wound are then increased by Tβ4 stimulation as shown in in vitro and in vivo studies [49]. The actin- binding domain of the peptide, 17LKKTETQEK25, was found to be responsible for the increase in cell migration activity and vessel sprouting [50]. Tβ4 has been shown to increase matrix metalloproteinase (MMP) production in vitro, which is essential for migrating endothelial cells, fibroblasts, keratinocytes, chondrocytes and monocytes [51-52].

Usual symptoms of a chronic wound are prolonged inflammation, dysfunctional cell proliferation and collagen deposition. Thus, the process of wound healing is also enhanced by the anti-inflammatory effects of Tβ4. Here, the N-terminus of the molecule is believed to be responsible for the inhibition of inflammatory cell activity in wounds [34].

Tβ4 in cardiac repair Heart disease including myocardial infarction, stroke and peripheral vascular disorders has been shown to be one of the leading causes of death. Myocardial infarction is the acute loss of myocardial function due to vascular occlusion of a coronary artery. Bock-Marquette et al.[40] have shown that with the administration of

Tβ4 immediately following an induced myocardial infarct in mice, Tβ4 can prevent cell death of heart tissue. It also increases the motility and survival of myocytes after hypoxia [53-54].

The postulated mechanism of action is that exogenously administered Tβ4 upregulates the expression of PINCH, a LIM domain protein. Tβ4 binds to PINCH and forms a functional complex with integrin-linked kinase (ILK), a scaffolding protein [40]. It has also been shown that Tβ4 interacts with the kinase region of ILK [55]. Through the kinase activity of ILK, Akt2 is phosphorylated, which increases the production of MMP-2, a metalloproteinase responsible for matrix degradation. The process of breaking down extracellular matrix, a hindrance to movement, is necessary during the migration of myocardial cells. In addition to myocardial cell migration, the effect of exogenous Tβ4 on the increased migration of endothelial cells in the embryonic mouse heart was also recorded. All these effects, taken together, contribute to an enhanced myocyte survival after hypoxia and an improved cardiac function.

Tβ4 in corneal repair and anti-inflammatory processes

Tβ4 has been implicated with corneal repair in several animal models [45-46] through several mechanisms. Upon external application in the eye, Tβ4 promotes cell-cell and cell-matrix contacts and clarity of the cornea after injury. It also increases epithelial cell migration of the conjunctiva. Unlike in the heart and dermis, Tβ4 speeds up the healing of the cornea by down regulating the expression of metalloproteinases, chiefly 15

1 Introduction

those which break down gelatin (MMP-2 and MMP-9), as well as membrane-type

metalloproteinases (MT6-MMP) [56]. Tβ4 is also found to inhibit injury-induced apoptosis.

Prolonged and massive inflammation usually exacerbates wounds in the eye caused by chemical burns and has an influence on healing. Uncontrolled inflammation is mainly caused by the infiltration of polymorphonuclear macrophages (PMNs) in the eye wound and surrounding stroma. Studies have shown that topical administration

of Tβ4 inhibits the gross migration of PMNs into damaged tissue by decreasing the expression of chemokines, keratinocyte-derived chemokine (KC) and macrophage inflammatory protein 2 (MIP-2), which act as main chemotactic agents for PMNs [45].

Tβ4 in tumor metastases The process of tumor metastasis involves several stages: tumor cell migration and invasion, angiogenesis and, finally, tumor cell growth and proliferation [57-58]. The

overexpression of thymosins (Tβ4, Tβ10 and Tβ15) has been recorded in tumors including thyroid and breast tumors and in melanoma and fibrosarcoma [59-60]. Cha et al. [61] have previously undertaken in vitro and in vivo studies to elucidate the

effect of Tβ4 on the highly malignant B16-F10 cell line cultured from lung tumors. In

these studies, Tβ4 cDNA was introduced into B16-F10 cells through a genetically

engineered adenoviral infection. The resulting overexpression of Tβ4 was correlated with the increase in levels of secreted vascular endothelial growth factor (VEGF), a factor essential for tumor growth and angiogenesis. Further in vitro studies of B16-

F10 cells showed an increase in cell invasion with Tβ4 overexpression although no change in levels of MMPs (MMP-2 and MMP-9).

SW480 colon carcinoma: epithelial-mesenchymal transition

The cellular overexpression of Tβ4 has been implicated with the malignant progression of SW480 colon carcinoma cells. Wang et al. have shown that SW480 cells

overexpressing Tβ4 demonstrate increased levels of growth and invasivity in in vitro and in vivo mouse models. Here, invasivity of cells is directly connected with the elevated levels and activity of MMP-7, b-catenin and c-myc, all well recognized agents

in the process of cell invasion. Furthermore, SW480 cells overexpressing Tβ4 exhibited lower levels of E-cadherin, a protein responsible for the integrity of cell-cell junctions [62].

The overexpression Tβ4 has been linked with the epithelial-mesenchymal transition of SW480 cells [47, 62]. In this process, cells exhibit a loss of cell adhesion through the loss of E-cadherin activity. These cells undergo a change of morphology into a more mesenchymal phenotype typical for migratory cells in normal embryonic development and the malignant progression of non-invasive cells. In these studies, 16

Introduction 1

increased levels of Tβ4 expression in SW480 cells correlate with the upregulation of ILK. Together with the loss of E-cadherin, the increase in activity of ILK and its downstream effectors, including Akt, is postulated to induce EMT and enhanced invasivity of SW480 cells.

1 Introduction

1.3 Open questions What are the effects of Tβ4 on cytoskeletal mechanics and dynamics? Extensive studies have been undertaken to elucidate the molecular and biochemical

role of Tβ4 in the cellular signaling pathway. These studies have shown that not only

does Tβ4 play a major role in the regulation of the cytoskeleton as a G-actin sequestration protein; it also interacts with other cytoskeletal associated proteins

such as ILK and PINCH. Through these interactions, Tβ4 can elicit various cellular reactions involved in healing and repair processes as well as in cancer metastases. The underlying mechanism behind these processes is the increase in the motility of the cells. The molecular pathways governing these processes have only been partially understood and the results of previous studies show only a correlation between the

extracellular application of Tβ4 and the endogenous intracellular overexpression of the protein with cell motility and invasion. No quantitative studies have yet been

made to assess how Tβ4 affects cell motility. The effects of Tβ4 on cell motility imply a distinct change in the mechanics and dynamics of the cytoskeleton. Until now, these aspects have not been examined in any of the cellular models.

Exogenous application vs. intracellular over-expression?

In the past, studies concerning Tβ4 have been made with the external addition of the peptide as well as with the genetic overexpression in cells and often the observed effects of both types of studies, such as increased cell motility, have been interpreted

as originating from the same mechanism. In investigating how Tβ4 can affect cytoskeletal mechanics and dynamics, these two aspects; exogenous application and endogenous genetic overexpression, must be examined separately. This is especially true when considering the simple fact that the cytoskeleton and its constituents which

interact with Tβ4 cannot be accessed in the same manner from inside and from outside the cell. The following studies are concerned mainly with the effect of

exogenous Tβ4 on cellular mechanics and dynamics.

Exogenous Tβ4: How can so little do so much?

Tβ4 is present at a high concentration of up to 500 µM in virtually every cell in

humans and mammals [63-64]. Up to 1% of the total Tβ4 concentration is circulating in blood serum found in wound fluid [28, 65] and is believed to originate from damaged platelets. Although this may be the case, studies have shown that even nanomolar concentrations of the peptide can induce a major cellular response. The

molecular mechanism of Tβ4 entry into the cell is currently unknown. Until now, no

known cell surface receptor has been found for Tβ4 cellular outside-in signaling.

Introduction 1

Clinical studies: a biphasic dose response – a biophysical phenomenon?

As recorded in previous studies, the role of Tβ4 as an agent in wound healing, cardiac repair, anti-inflammatory pathways and in cancer metastases has made it a target for clinical application. Clinical trials of using Tβ4 have been carried out for the treatment of a number of diseases including venous stasis ulcers [66], myocardial infarction [67], chronic non-healing corneal ulcers [68], epidermolysis bullosa and for dermal wounds

[69]. In these clinical trials, the efficacy of the drug, synthetic Tβ4, was examined in a dose dependant manner. In the clinical application for the treatment of venous stasis ulcers and in the healing of dermal wound, separate and independent clinical trials were carried on patients with increasing concentrations of Tβ4 (0.01%, 0.03% and 0.1% topical peptide concentration) with a placebo used as a control [66, 70]. Remarkably, the most rapid healing in both cases was observed at the mid-dose level of applied Tβ4; a clear biphasic response to increasing Tβ4 doses. Here the question arises of why the mid-dose elicited the highest efficacy and how this phenomenon is correlated to cell mechanical responses.

2 Material and Methods

Material and Methods 2

2.1 Biological and biochemical methods

2.1.1 Cells and cell culture The SW480 cell line was established from a primary human colorectal adenocarcinoma and can be acquired commercially at America Type Culture Collection (cat. No. CCL-228 ATCC, Manassas, VA). NIH 3T3 cells (cat. No. CRL-1658 ATCC, Manassas, VA) originate from primary mouse embryonic fibroblast cells. Immortalized cells with a stable growth rate were established from the original culture after 20–30 generations. Both established cell lines were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 1 mg/mL D- Glucose, 100 U/ml Penicillin/Streptomycin and 10% fetal calf serum (FCS) (v/v)

(Biochrom) at a constant temperature of 37°C and 5% CO2 on cell culture surface treated flasks (Nunclon Surface, Nunc). The GSP-cell line is a product of an NIH 3T3 cell stably transfected with a plasmid vector (pIRESpuro3) carrying a gene for EGFP- Actin. This cell line was maintained in DMEM supplemented with 1 mg/mL D-Glucose, 100 U/ml Penicillin/Streptomycin, 10% FCS (v/v) (Biochrom) with 1 µg/ml

Puromycin (Sigma) at a constant temperature of 37°C and 5% CO2. Cells were passaged at 80% confluency.

Figure 2.1 Cell lines. (A) SW480 human colorectal carcinoma cells, Hoffmann modulation contrast image with 20x objective (B) NIH 3T3 mouse fibroblasts, field contrast image with 10x objective

2 Material and Methods

For routine passaging, 5 ml Trypsin (Biochrom AG) was used to detach cells from T- flasks. Prior to measurements, cells were suspended in 10 ml Accutase (PAA Laboratories GmbH) to ensure gentle detachment. Cells were incubated at 37°C during detachment. When using non-treated culture surfaces (glass-bottom wells), 10 g/ml fibronectin in PBS (phosphate buffered saline) was applied to the surfaces and incubated overnight at 4°C prior to seeding.

2.1.2 Cloning cassettes of EGFP-Actin for generating the GSP The cloning cassette with the EGFP-Actin (Enhanced GFP) target DNA was produced through PCR (polymerase chain reaction). The PCR reaction mixture contained 100 ng of template DNA 0.5 µM of each primer, 200 µM dNTPs (Nucleotide Solution Mix, New England Biolabs) and 1.0 U polymerase (Phusion ® Hot Start DNA-Polymerase,

Finnzymes). The mixture was added to a final volume of 50 µL with dH2O. The PCR program cycle was set in four steps. Initial denaturation of DNA was done at 95°C for 30 s. This was followed by 34 cycles of denaturation at 98°C, annealing at gradually rising temperatures of between 67°C and 71.1°C and extension at 72°C. A final

extension step at 72°C was then done for 10 minutes. The Tm (annealing temperature)

was varied to account for uncertainties resulting in differences between the Tm of the 2 primers used (see 2.1.4). After amplification, the PCR mixtures were purified using a PCR purification kit (Qiagen). The DNA products were checked by gel-electrophoresis on 1% Agarose gel before purification and used subsequently in digestion reactions with the appropriate restriction enzymes (New England Biolabs) according to the manufacturer’s protocol. DNA fragments were purified again after digestion.

2.1.3 Ligation Ligation of the DNA target insert and the vector backbone was done by incubating the insert with the vector (in this case both digested with NheI and BamHI restriction enzymes) at a vector:insert mol ratio of 3:1 and 5:1. 20 mM ATP, 5x ligase buffer and 2 U ligase (T4 DNA-Ligase, Invitrogen) was added to the reaction mixture and topped

off with dH20 to a final volume of 10 µL. Reactions were incubated at either 14°C overnight or at room temperature for 3 h before heat-inactivation at 65°C for 15 min. A third of the mixture was used to be transformed into DH5 E. coli strain by heat shock according to standard protocols [71]. α

Material and Methods 2

Figure 2.2 Stable transfection of NIH 3T3 cells with an EGFP-Actin plasmid vector. (1) The EGFP-Actin cassette was cloned with 5’-NheI and BamHI-3’ restriction sites through polymerase chain reaction. Template DNA for the EGFP-Actin cassette was obtained from a pEGFP-Actin vector (Clontech). The EGFP-Actin DNA cassette was ligated into the pIRESpuro3 backbone at the multiple cloning site (MCS) resulting in a pIRES-EGFP-Actin vector, (2). The pIRES-EGFP-Actin plasmid vector was transformed into the DH5-α E. coli strain (3) to multiply the plasmid. The plasmid vector was purified using a conventional purifying kit, EndoFree Maxi Kit (Qiagen) (4). NIH 3T3 mouse fibroblasts were transfected with the resulting plasmid vector using the Effectane (Qiagen) transfection method (5). Puromycin (1 µg/ml) was used as a selective agent for further cultivation. Limited dilution of transfected cells (1 cell in every 4 of 96x wells) enables the growth of positive clones from a single cell. Clones with EGFP-actin are maintained in DMEM with 100 U/ml Penicillin/Streptomycin, 10% FCS and 1 µg/ml Puromycin.

2 Material and Methods

2.1.4 Cloning of EGFP-Actin into the pIRESpuro3 selection vector The pIRESpuro3 vector (Clontech) contains a viral internal ribosome entry site (IRES) which allows for the bicistronic expression of the resistance gene and the target gene of interest. This enhances the translation of both the Puromycin resistance gene in combination with the target DNA. A cloning cassette of EGFP-Actin with restriction sites for NheI (5’) and BamHI was created through PCR using the primers (NheI-EGFP- Actin forward) 5’-GTC AGC GCT ACC GGT CGC C-3’and EGFP-Actin-BamHI reverse) 5’- ATC CCT AGA AGC ATT TGC GGT GG-3’ using the EGFP-Actin vector (Clontech) as a template (see 2.1.2). Digestion and ligation were performed as described previously. The cloned DNA product was transformed into DH5 E. coli by heat-shock according to standard protocols and subsequently purified using the EndoFree Plasmid Maxi Kit (Qiagen). α

2.1.5 Transfection of the EGFP-Actin vector construct Transfection of the NIH 3T3 cells was done by incubating 1 µg of the pIRES-puro3- EGFP-Actin vector construct with the 10 µl Effectane Transfection Reagent (Qiagen) and 8 µl Enhancer according to the manufacturer’s protocol. The DNA-Effectane complexes were added drop-wise with constant gentle mixing onto 80% confluent cells grown on Ø 35 mm cell-culture treated wells in 2 ml normal growth medium (DMEM with 100 U/ml Penicillin/Streptomycin and 10% FCS). Transfected cells were cultivated further for 24 h before further observation.

2.1.6 Stable transfection into mammalian cell lines Prior to stable transfection, Puromycin resistance of NIH 3T3 and SW480 cells were examined. 3 x 105 cells in Ø 35 mm wells were incubated with concentrations of Puromycin ranging from 1 to 10 µg/ml over a period of three weeks. As a control, no Puromycin was added to the cells. Cell medium (DMEM with 100 U/ml Penicillin/Strepromycin and 10% FCS) containing the antibiotic was replaced every 2 days. Throughout this period, cell growth was observed. An optimum Puromycin concentration of 1 µg/ml was chosen for both cells lines.

Material and Methods 2

Figure 2.3 Close-up of an NIH 3T3 mouse fibroblast stably transfected with the pIRES- EGFP-Actin construct. Cells were seeded on glass bottom wells. The live image was made using a 63x objective.

24 h after transfection (see 2.1.5), growth medium was replaced. 48 h after transfection, 1.0 µg/ml Puromycin was added to the growth medium for selection. Puromycin-containing medium was replaced every 2 days regularly until small colonies of resistant cells were visible. Limited dilution of the cells was performed in 96-well plates at a calculated concentration of 1 cell in every 4 wells to ensure single- cell populations. These cells were grown with enriched DMEM (4.5 mg/ml D-glucose, 20% FCS) containing 1.0 µg/ml Puromycin.

2.1.7 Immunofluorescence and fluorescence microscopy Up to 5 x 104 cells were seeded on 10 µg/ml fibronectin-coated glass cover slips

(Menzel, Braunschweig, Germany) and incubated overnight at 37°C and 5% CO2. Adherent cells were fixed with 3% paraformaldehyde for 20 min prior to permeation with 0.2% Triton-X-100 for 5 min. Cells were blocked using 0.5% BSA (bovine serum albumin) in PBS for 1 h at room temperature. After washing, primary antibodies of anti-actin and anti-vinculin (Sigma) were diluted 1:8,000 and 1:1,000,000, respectively, with 0.5% BSA in PBS and incubated with the fixed cells at room temperature for 1 h. Secondary HRP (Amersham) or FITC (Jackson ImmunoReaserch) labeled anti-mouse antibodies were then diluted at 1:1,000 and 1:200, respectively, and incubated for another hour with the fixed cells. For actin staining, 1 µg/ml Alexa 25

2 Material and Methods

Fluor 546-phalloidin (Molecular Probes, Eugene, Oregon) was added during incubation with the secondary antibody. Between each step, cells were washed with 0.5% BSA in PBS. The cell samples were mounted on microscope slides using Mowiol (Sigma) mounting solution. All fluorescent microscopy was carried out on a Leica microscope DM16000 with both phase contrast and fluorescence modi. The objectives for magnification used in the imaging and other experiments were 20x/0.4NA-, 40x/0.6NA- and 63x/1.3NA. Data acquisition was performed by a CCD (charged- coupled device) camera (ORCA ER, Hamamatsu).

Figure 2.4 Actin fluorescent staining with Alexa Fluor 546-phalloidin. (A) The diffuse actin fibers in SW480 cells are made visible using a 10x objective. (B) The cortical actin network of an NIH 3T3 cell seen through a 63x objective.

Material and Methods 2

Figure 2.5 Fluorescent staining of vinculin. Focal adhesions of an NIH 3T3 cell are made visible with vinculin fluorescent staining. Images were taken with a 63x objective.

2.1.8 Synthesis of lipid vesicles Multilamellar lipid vesicles (MLVs) were synthesized by dissolving Dimyristoyl-L- - phosphatidylcholin (DMPC), Dimyristoyl-L- -phosphatidylglycerol (DMPG), or 1,2- dimyristoyl-sn-glycero-3-phospho-L-serine (sodium salt) (DMPS) (Avanti Polarα Lipids, Birmingham, Alabama) in a 2:1 (v/v)α mixture of chloroform/methanol in an Erlenmeyer flask. Through nitrogen evaporation and subsequent vacuum desiccation for 2 h, a dry lipid film was produced on the glass wall of the flask. The lipid film was then resuspended in a buffer solution containing 20 mM HEPES (pH 7.4), 2 mM EDTA, 5 mM NaCl and 0.2 mM DTT. The resulting MLVs were left to equilibrate overnight at 35°C. Fluorescence correlation spectroscopy measurements require the use of small unilamellar vesicles (SUVs) produced by dissolving the lipid film in 10 mM potassium phosphate buffer (pH 7.4) followed by equilibration and finally sonication for 10 min before use.

2.1.9 Synthesis of Tβ4-Alexa488 labeled peptide

Tβ4 and scrambled Tβ4 (used as a negative control) were obtained courtesy of RegenerRex Biopharmaceuticals (J.J. Finkelstein, 15245 Shady Grove Rd, 470

Rockville, MD 20850-6243). The amino acid sequence of scrambled Tβ4 synthetic peptide is: acetyl-EIPKETEKFDKETMEQKSIDKSQLEKPEAQNKELPTSKSGDKA. The 27

2 Material and Methods

labeling of Tβ4 with Alexa-488 was done according to methods as described in [72]. Using this procedure, 500 µg Alexa Fluor488 cadaverine sodium salt (Invitrogen) and

250 µg Tβ4 was incubated with 0.2 U guinea pig transglutaminase in a reaction

mixture containing 50 mM Tris HCl, pH 7.4; 15 mM CaCl2; 3 mM DTT while adding

0.3 mM TGA. This transamidation reaction labels the Tβ4 peptide with Alexa Fluor488 at glutamine residues. The labeled product was characterized by using reverse phase- HPLC and MALDI-TOF analysis.

Figure 2.6 Linking the Alexa Fluor 488 fluorophore to Tβ4 molecules. Using the transglutaminase enzyme as a transamidation agent, Alexa Fluor 488 cadaverine sodium salt reacts with the Tβ4 peptide in which the fluorophore is linked to glutamine residues of the peptide.

Material and Methods 2

2.2 Biophysical methods

2.2.1 Three-dimensional collagen cell invasion assay Collagen gel were prepared by mixing 775 µl collagen R (2 mg/ml rat collagen type I; Serva, Heidelberg, Germany) and 775 µl collagen G (4 mg/ml bovine collagen type I;

Biochrom). The mixture was then neutralized by adding 135 µl 26.5 mM NaHCO3 buffer, 43 µl 1 mM NaOH and 135 µl 10x DMEM. 1.2 ml of the preparation was

pipetted into Ø 35 mm cell culture dishes left to polymerize for 2 h at 37°C, 5% CO2 and 95% humidity. 2 ml DMEM was layered on top of the resulting gel to prevent

dehydration of the matrices before overnight incubation at 37°C, 5% CO2 and 95% humidity.

After equilibration, the DMEM layer was replaced with a 2 ml suspension of 1 x 105 cells in DMEM (with 1 mg/ml D-Glucose, 2 mM L glutamine, 100 U/ml

Penicillin/Streptomycin). As Tβ4 can be found in serum, cells were cultured without

FCS. Tβ4 was added to the medium upon cell seeding. Cells were left to migrate within the gel for 72 h. In experiments using calyculin-A (Calbiochem), cells were left to adhere to the gel for 24 h before 10 nM or 50 nM of calyculin-A was added for 2 h. Afterwards, the culture dish was washed with pre-warmed (37°C) DMEM, and cells were allowed to migrate for another 48 h. Gels were then fixed with 2.5% glutaraldehyde. Nuclei of individual cells were stained with Hoechst 33342. The number of invaded cells and their invasion depths were determined in 25 randomly selected fields of view.

2.2.2 Two-dimensional cell motility assay 1 x 104 cells were seeded in DMEM medium in a 35 mm dish and stimulated with

various concentrations of Tβ4 (0.05 – 1.0 µM) 24 h before measurements. Phase contrast images of cells (x10 magnification) in 10 random fields of view were taken every 60 s for a period of 3.5 h. Cells were kept in a stable environment throughout the entire measurement in a heated chamber mounted onto the microscope set at

37°C with a constant supply of CO2. Cell movement was calculated as a measure of mean square displacement (MSD) through a Fourier-based difference-with- interpolation image analysis [73]. The MSD followed the power law as below:

= ( / ) + 훽 0 where t0 is the time interval푀푆퐷 between퐷 ∙ recorded∆푡 푡 images푐 (60 s), the pre-factor D represents the apparent diffusivity of movement, and the exponent β is a measure of movement persistence [74]. Values of β ~1 are seen with cells migrating randomly, and β ~2 for cells moving ballistically. The additive constant c represents 29

2 Material and Methods

measurement noise. In our case, because of the large time interval between successive images, this constant is negligible.

2.2.3 Cell-spreading assay 5 x 104 cells were seeded on 10 µg/ml fibronectin-coated glass cover slips in a 35 mm

dish. After overnight incubation at 37°C and 5% CO2, adherent cells were fixed with 3% paraformaldehyde for 15 min at room temperature. After washing with 0.5 % BSA in PBS solution, cells were stained for 30 min with 1 µg/ml Alexa Fluor546-phalloidin (Molecular Probes, Eugene, Oregon) at room temperature. Cell nuclei were stained with 1 mg/ml Hoechst 33342 dye (Pierce) for 5 min at room temperature. Fluorescent and bright field images were taken from 10 – 20 random fields of view. Each cell nucleus was counted to represent a single cell. The projected area of a cell was calculated as the area bounded by the stained actin cytoskeleton measured with a custom image processing program written in MATLAB.

2.2.4 Two-dimensional cellular traction microscopy Polyacrylamide (PAA) gels (6.1% acrylamide/0.31% bis-acrylaminde) used for traction microscopy measurements were cast on 75 mm x 25 mm silane-coated microscope glass slides (Menzel) as described in [22]. For silanization the glass slides were immersed first in 0.1 N NaOH for 5 min and, after drying, in 2.0% (3- Aminopropyl)-trimethoxysilane for another 5 min. The slides were then washed in

dH2O twice before further immersion in 2.5% glutaraldehyde for 30 min. The slides

were ready for use after washing twice again in dH2O and drying at room temperature. Green fluorescent carboxylated beads were suspended in the PAA gel before casting into 10 x 10 mm frames (Abgene, Thermo) on the silanized glass slides. The gel suspension was centrifuged upside down at 288 g and 4°C for 3 min to ensure a uniform distribution of the fluorescent beads in a thin layer on the upper surface of the gel. After polymerization, the gels were cross linked with Sulfo-SANPAH (Pierce Biotechnology) and activated under UV-light for 5 min at room temperature. Following two wash intervals with 50 mM HEPES buffer, the PAA gel were coated with 50 µg bovine Collagen G (Biochrom AG) in 5 mM HEPES buffer overnight at 4°C. The Young’s modulus of the gels was 12.8 kPa ± 0.8 kPa as measured by a magnetically driven plate rheometer.

Material and Methods 2

Figure 2.7 Two-dimensional traction microscopy. Serum starved SW480 cells are left to adhere overnight onto collagen-coated PAA gel with a Young’s modulus of 12.8± 0.8 kPa. Upon seeding, concentrations of Tβ4 were added to the cells. Fluorescence and bright field images at 20x are taken before and after detaching the cells with a mixture of Cytochalasin D/Trypsin. The images were analyzed to determine the displacement of the fluorescent markers.

Figure 2.8 2D traction microscopy images. (A) Bright field image of SW480 cells on the surface of PAA gel. (B) Fluorescence image of the underlying fluorescent beads in the same field of interest as in (A).

1.5 ml DMEM suspension (without FCS) containing 2 x 104 cells was pipetted onto the gel. The cell suspension was contained within a silicone ring (In Vitro, Göttingen,

Germany) attached to the glass slide. For measurements with Tβ4, concentrations of

2 Material and Methods

the peptide were added to the cell suspension while seeding. Cells were left to adhere

on the gel overnight at 37°C, 5% CO2 and 95% humidity. During tractions measurements, cells were detached from the gel using a mixture of 80 µM cytochalasin D and 0.25% trypsin. The displacement of fluorescent markers below the surface of the gel was determined using a Fourier-based difference-with-interpolation image analysis [74]. Cell tractions were then calculated from an unconstrained deconvolution of the gel surface displacement field measured before and after detachment of the cells [75].

2.2.5 Magnetic tweezer microrheology The magnetic tweezer device is described in [17]. Fibronectin beads used in this method were produced by incubating 4x108 superparamagnetic epoxylated beads (Ø 4.5 µm, Dynabeads, Invitrogen) with fibronectin (5 µg per 1x107 beads; Roche) for 24 h at 2°C. After incubation, beads were washed with 0.1% BSA in PBS. Fibronectin coated beads were kept in this buffer at 4°C. Beads were sonicated directly each time before adding to cells to avoid clumping. 1.5 x 105 cells were seeded in DMEM medium without FCS on Ø 35 mm cell-culture treated dishes (Nunclon) 24 h before tweezing

measurements. Cells were stimulated with different concentrations of Tβ4 upon seeding. On the day of the measurements, 2 x 105 fibronectin coated beads were

added to each dish with adherent cells and incubated for 30 min at 37°C with 5% CO2. Directly before measurements, culture dishes were washed with fresh, pre-warmed medium to remove unbound beads.

Material and Methods 2

Figure 2.9 Magnetic tweezer measurements. (A) Fibronectin-coated beads are bound to the cytoskeleton through cell surface integrins. Force directed toward the tip of the needle pulls at the bead resulting in its displacement. Inset: bright-filed image of the needle tip pulling at a fibronectin-coated bead. (B) Staircase-like protocol: forces exerted onto the bead are raised sequentially (inset). Bead displacement over time is then recorded over forces from 0.5 nN to 10 nN. Images are modified from [76].

2 Material and Methods

A fibronectin coated bead is bound to the cytoskeleton via integrins spanning the cell membrane. The magnetic tweezer needle (HyMu80 alloy, Carpenter) was aimed by a micromanipulator (Injectiman NI-2, Eppendorf) 20-30 µm away from a bound bead on a cell. A solenoid coil around the needle generated a magnetic field with a high field gradient at the tip of the needle. The resulting force pulled at the fibronectin-coated bead attached to the cell. Bright-field images of the cell, bead and needle were taken continuously during the measurements with a charged-coupled device (CCD) camera (ORCA ER, Hamamatsu) at a rate of 40 frames/s. Bead positions were tracked using an intensity-weighted centre of mass algorithm. A preset force was kept constant by updating the current flowing through the solenoid or by moving the solenoid to keep the distance between the needle-tip and the bead constant. Acquisition of images was triggered and synchronized with the solenoid current generator. All measurements were performed on a heated microscope stage.

With each force step with an amplitude of ∆F applied onto a fibronectin-coated bead, the bead moves with the displacement d(t) towards the tip of the needle. The creep response J(t) , which is the ratio of d(t)/ ∆F, followed a power law:

( ) = ( / ) 푏 0 for all amplitudes of force where퐽 t푡0 is a푎 reference∆푡 푡 time arbitrarily set to 1 s. The inverse of the prefactor a (in units of nN/µm) is a measure of cell stiffness and the exponent b describes the visco-elastic properties of the cytoskeleton. b ~ 1 indicates a Newtonian fluid-like behavior, whereas b ~ 0 an elastic solid-like behaviour. A stepwise increasing force protocol was performed with steps of 0.5 nN, 1.0 nN, 1.5 nN, 2 nN, 3 nN, 4 nN, 5 nN, 6 nN, 8 nN and 10 nN, with each force-step lasting for 1 s. Values of 1/a and b were determined at each force step.

2.2.6 Nano-scale particle tracking Nano-scale tracking of cytoskeleton-bound beads was described by [73]. 4 x 104 cells were seeded in DMEM medium without FCS on flat bottomed Ø 6.4 mm cell culture

wells (Corning® Sigma) and left to adhere overnight at 37°C and 5% CO2. Prior to measurements approx. 2 x 104 pre-sonicated fibronectin-coated beads were added to the culture wells. The same Ø 4.5 µm fibronectin-coated beads were used as described previously in the magnetic tweezers method (see 2.2.5). The beads were incubated

with the cells for 30 min at 37°C and 5% CO2. For overnight measurements,

concentrations of Tβ4 were added during seeding. Short term stimulation was done by

adding Tβ4 during measurements. Bright field images were recorded at a rate of 8.3 images/s with a CCD camera (ORCA ER, Hamamatsu) on an inverted microscope with a 10x, 0.3 NA objective (Leica) placed on a vibration isolated table (Newport, Irvine CA). 100-200 bead positions were tracked continuously for 5 min using an 34

Material and Methods 2

intensity-weighted center of mass algorithm with an accuracy of 10 nm (rms). All measurements were performed in a cell chamber mounted onto the microscope stage maintained at 37°C with a constant supply of CO2. The stage drift, which was estimated from the mean motion of all beads in the field of view, was accounted for in order to calculate the accurate bead positions.

The mean-squared displacement of the beads was calculated as:

( ) = ( + )– ( ) 2 2 where is the position of〈∆ the푟 bead∆푡 〉 and〈� →푟 is푡 the∆ 푡time→푟 lag.푡 The� 〉 brackets … is the time average. The MSD of bead movement is fitted to the power law using a least-squares →푟 ∆푡 〈 〉 fit and, thus, the equation can be rewritten as:

= ( / ) + 훽 The power-law cofactor D (in 푀푆퐷the unit퐷 of∙ nm∆푡2)푡 0is a measure푐 of diffusivity of the bead motion influenced by the stability of the cytoskeleton and the binding quality of the bead to the cytoskeleton via cell-surface integrins. The persistence of bead movement is reflected in the β exponent. Values of β = 1 indicates Brownian/diffusive, β < 1 subdiffusive, β > 1 superdiffusive and β = 1 ballistic motion of the bound bead.

2 Material and Methods

Figure 2.10 Nano-scale particle tracking of fibronectin coated superparamagnetic beads Beads are attached to the cytoskeletal network through integrins. Bead displacement due to movement of the cytoskeleton is recorded. Inset: an SEM image of a cell with attached beads (taken by C. Mierke, bar indicates 10 µm).

Figure 2.11 Analysis of nano-scale particle tracking. A) The trajectory of cytoskeletal bound beads is shown as a series of dots. Bead positions are recorded every 120 ms. Measurements are taken for 5 minutes.(B) The mean square displacement (MSD) of a bead in correlation with ∆t following the power law. This particular analysis is from a recording of trajectories of a bead bound to an NIH 3T3 cell.

Material and Methods 2

2.2.7 Differential scanning calorimetry (DSC)

Preliminary measurements of Tβ4 – lipid interaction were made using this method. Basically, DSC was used to measure the melting phase transition temperature and the

phase transition enthalpy of lipid vesicles in the presence of Tβ4. In membranes, phospholipids exist in a gel-like (ordered) or fluid-like phase (disordered) depending on the temperature. In the ordered arrangement, acyl residues of the fatty acids are in the energetically favorable trans-conformation. A transition into the more disorderly, fluid phase requires energy, leading to the change into the more energetically unfavorable gauche fatty acid conformation. The energy needed for this conformational change can be determined by the specific heat capacity of the lipids

(Cp). Some lipids go from the gel-phase through intermediate phase changes, the

lamellar (Lβ) then the ripple phase (Pβ) before reaching the fluid phase at the melting temperature (TM) (see Figure 2.12). With the non-covalent interaction of a peptide with the acyl-residues, a disruption of the lipid arrangement occurs. As a result, less energy in the form of transition enthalpy is needed to cause a phase transition, thus, a

reduction in the TM.

In DSC measurements, the sample and reference solution is heated at a preset heating rate of:

of 0.5°C/min to +30°C, and cooled 훽at a ∆rate푇⁄∆ 푡of 1°C/min to +7°C until the phase transition enthalpy reached equilibrium (Q100, TA Instruments). The difference in the heat capacity ∆C(T) of the sample and reference is:

( ) = = ( )

where and are the∆ 퐶specific푇 퐶heat푃 − 퐶capacities푅 ∆푃 푇 ⁄of훽 the probe and reference, respectively. 퐶푃 퐶푅 The integration of the heat capacity, ∆C, over the change in temperature gives the phase transitions enthalpy, ∆H. A phase transition was observed at around 23°C, depending on the molar ratio of DMPG and DMPC lipids used. Analysis of the raw data recorded during DSC measurements was performed using custom-programmed software in MATLAB. Small unilamellar vesicles (SUVs) used in these experiments were synthesized as described in 2.1.

2 Material and Methods

Figure 2.12 Phase transition of synthetic lipid vesicles. Shown above is a thermogram of the transition phases of DMPG/DMPC lipid vesicles with increasing temperature. A peak

temperature of TV is reached between the lamellar (Lβ) and the ripple phase (Pβ). The melting point, TM, marks the maximum specific heat which is reached before the lipid bilayer shifts from the ripple phase into the fluid phase (Lα). The onset of the rise in specific heat is marked by Ts .TL marks the end of the main phase transition. Diagram taken from [77].

Material and Methods 2

Figure 2.13 Differential Scanning Calorimetry (DSC): experimental setup. The temperatures of the sample (1) and reference (2) are increased with the heater (3). Loss of heat is prevented by the insulation coat (4). Temperatures are recorded by the temperature observation point (5). PP and PR are heat outputs of the sample and reference, respectively.

2.2.8 Fluorescence correlation spectroscopy

The binding of Tβ4 to lipid membranes was examined further with the use of fluorescence correlation spectroscopy (FCS). This method was used to measure the

diffusion of Tβ4-Alexa488 labeled peptide in the presence of various concentrations of lipid vesicles. A custom FCS system was built as described in [78]. The setup consisted of an excitation laser (20 mW, 488 nm; Changchun Dragon Lasers) with a beam that was spatially filtered and expanded by a lens (f1 = 30 mm), a 50 µm pinhole, and a second lens (f2 = 75 mm) before being passed through an aperture. The laser power was reduced to ~100 µW with a neutral density filter. The excitation beam was then passed through a beam-splitter and reflected by a dichroic mirror into an oil immersion objective (NA 1.25, 100x). Low-fluorescence immersion oil was used to decrease background fluorescence. Fluorescence emitted from the samples was passed through the same objective and reflected by the dichroic mirror into an avalanche photodiode (APD) with an active area of 625 µm². Scattered laser light at 488 nm was eliminated by a blocking filter. Fluctuations in the excitation laser intensity were monitored by a photodiode. Longer correlation times of fluorescence

2 Material and Methods

intensity fluctuations were expected for fluorescent peptides bound to larger lipid vesicles compared to free fluorescent peptides.

Figure 2.14 Schematic diagram of the Fluorescence Correlation Spectroscopy (FCS) detection volume. Free fluorescent peptide (Tβ4-A488) and fluorescent peptide bound to much large lipid vesicles diffuse in and out of the illuminated volume, causing characteristic fluctuations in the detected fluorescence intensity. A detection volume of ca. 1 µm3 was used in the setup.

DMPC and DMPS SUVs were synthesized as described in 2.1.8 and sonicated again directly before measurements. The focal volume was determined to be 1 µm3 through calibration with Alexa-488 (D = 435 µm2 s-1 and Rhodamine 6G solution (D = 414 µm2 s-1) [79]. Measurements were performed at 25 kHz for 10 s. Data analysis of fluorescence auto-correlation was done using a custom program developed in MATLAB.

The principle of fluorescence correlation spectroscopy has its basis in the analyses of correlations in fluorescence fluctuation intensities. According to this principle, longer correlation times are expected of fluorescent peptides bound to larger lipid vesicles than free fluorescent peptides travelling in and out of the detection volume. The normalized autocorrelation function of the fluorescence fluctuations, ( ), calculated from the fluorescence intensity, ( ), detected in the sample volume is as follows: 퐺 휏 퐹 푡 ( ) . ( + ) ( ) = ( ) 〈훿퐹 푡 훿퐹 푡 휏 〉 퐺 휏 2 40 〈퐹 푡 〉

Material and Methods 2

The autocorrelation signal was fitted to

1 1 1 ( ) = < > (1 + ) 1 + ( )² 휏 퐺 휏 ∙ ∙ 0 푉푒푓푓 퐶 휏퐷 푟 휏 � 푧0 ∙ 휏퐷 where Veff is the effective observation volume; is the average concentration of particles; τD is the correlation time for translational diffusion; r0 and z0 are the radial and axial dimensions of the observation volume.

Dissociation constants for the binding for Tβ4 to SUVs, Kd, were obtained by fitting τD versus lipid concentration [L] to the Michaelis-Menten equation

[ ] = , 퐷 푚푎푥+ [ ] 퐷 휏 퐿 휏 푑 where τD,max is the maximum correlation퐾 time 퐿when all fluorophores are bound to ( ) SUVs. τD,max was obtained from = [78], and D is the diffusion constant of SUVs 2 푟0 (8.08x10-12 m2/s) calculated from the Stokes-Einstein equation, assuming an average 휏퐷 4퐷 radius of 30 nm [80].

2 Material and Methods

Figure 2.15 Principle of Fluorescence Correlation Spectroscopy (FCS). (A) A fluorescent peptide moving in Brownian motion in and out of the detection volume has a much shorter

correlation time (τfree) than a fluorescent peptide bound to a slower-moving lipid vesicle. This produces characteristic fluorescence intensity patterns, with longer bursts of intensities for bound peptide recorded over time. Autocorrelation analysis of intensity fluctuations results in an autocorrelation function against correlation time. (B) In an ideal two-component

system, the autocorrelation function for τfree is much smaller that of τbound. When the fraction of free and bound fluorescent peptide are the same, however, the molar partition coefficient can be calculated as = 1 [ ].

퐾 ⁄ 푙푖푝푖푑 42

3 Results

3.1 The effect of extracellular Thymosin β4 on the motility of SW480 cells

Previous studies have shown that Tβ4 is directly involved in regulating cell motility. In the metastases of human colorectal carcinoma, the over-expression of Tβ4 is believed to be responsible for the increased invasion of malignant cells [48]. In vivo and in vitro studies on embryonal cardiomyocytes and endothelial cells have shown that induction with extracellular Tβ4 has resulted in increased cell migration [40]. Increased migration of endothelial progenitor cells (EPCs) in response to stimulation with extracellular Tβ4 has also been recorded [81]. Although this phenomenon has been qualitatively discussed, the biophysical mechanisms underlying the dose response of extracellular Tβ4 induction on cell migration is still not known. In this chapter, the dose-dependent correlation of increasing Tβ4 concentrations to the migration of SW480 colon carcinoma cells is quantitatively examined.

3.1.1 Three-dimensional cell migration

The migration of SW480 cells in response to increasing extracellular doses of Tβ4 was characterized using a three-dimensional collagen gel assay. The gels used in this assay had an average shear modulus of 58 Pa and a thickness of 500 ± 100 µm (mean ± SD). The lattice of collagen fibers within each gel had an average pore size of 1.3 ± 0.2 µm

(mean ± SE)[27]. The SW480 cells were treated with Tβ4 directly upon seeding onto

the gels. The cells were induced with a range of Tβ4 concentrations (0.05 µM – 1 µM).

0.2 µM scrambled Tβ4 peptide was used as a negative control. The cells were cultured for 3 days before labeling their nuclei with Hoechst 33342 vital stain for cell counting and measuring the invasion depth.

The invasivity of the cells was quantified by obtaining the invasion depths of the cells in multiple randomly chosen fields of view. The cumulative probability of finding a cell (pCum) at or below a given invasion depth is expressed in the invasion profiles (see Figure 3.1 (A)). As a robust measure of cell invasiveness, a characteristic invasion depth was defined as the depth that a given percentile of cells have reached or exceeded. A threshold was chosen at pCum = 0.1; the characteristic invasion depth used in this study is the depth that 10% of all counted cells had reached. This approach was selected to accurately demonstrate the differences in migration at

different Tβ4 concentrations while at the same time remaining conservative, ensuring a high number of cell counts for sampling.

The SW480 colon cancer cell line is known to be non-invasive, and these cells

normally do not invade collagen gels. However, cells induced with Tβ4 at concentrations between 0.05 µM and 0.5 µM were able to migrate into the collagen gel. The cells showed a biphasic migratory response to increasing concentrations of

Tβ4; SW480 cells were most invasive at an applied Tβ4 concentration of 0.2 µM. Cells

induced with 1 µM Tβ4 exhibited a marked decrease in invasiveness, with a migration

profile almost equivalent to that of the control (0.2 µM scrambled Tβ4). 44

Results 3

Figure 3.1 Analysis of SW480 cell migration in three-dimensional collagen gels. (A) A Tβ4 dose dependant cellular invasion profile; pCum is the cumulative probability of finding a cell at a given depth and lower. (B) Invasion depth of cells at pCum of 10; 10% of cells are seen at this depth or lower.

3 Results

Closer analysis of the invasion profile shows that 10% of SW480 cells treated with the control have migrated into the gel to a depth of 99 µm or below. Compared to the

control, cells induced with Tβ4 at concentrations of 0.1 µM and 0.2 µM have migrated

over threefold deeper into the collagen gels. Tβ4 concentrations used above and below

these levels resulted in a reduced cell migration. At an induction with 1 µM Tβ4, however, cells migrate in the same manner as the control; 10% of cells are seen at a depth of only 110 µm and below. These results demonstrate a clear biphasic response

in cell migration to the dose of exogenous Tβ4 applied to SW480 cells (see Figure 3.1 (B)).

3.1.2 Two-dimensional cell motility Two-dimensional cell motility of SW480 cell was analyzed to acquire more

information on the effect of Tβ4 on the dynamics of cell movement. Sub-confluent

SW480 cells were induced with 0.2 µM Tβ4, the peak working concentration as seen in the 3D-invasion assays, and at the maximum dose of 1 µM where cell motility had

receded. Scrambled Tβ4 was used as a negative control. 24 h after induction, bright field time lapse images from multiple fields of view were taken every 60 seconds over three and a half hours. Using the images, the movement of each single cell was recorded by tracking bright nuclear structures with an intensity-weighted center-of- mass algorithm [26, 82]. From the recorded trajectory of these nuclear structures, the mean square displacement (MSD) of an entire cell was obtained.

The MSD is a measure of the Euclidean distance the cell has moved within a given time span. This was calculated for each cell using the following equation

( ) = [( ( + ) ( )] t 2 2 where is the two-dimensional∆푟 ∆ 푡position〈 푟 of푡 the∆ cell푡 − and푟 푡 〉 the time lag. The brackets indicate an average over absolute times . The MSD follows the power-law relationship푟 as a function of the time lag. Thus, the MSD ∆can푡 be fitted according to the power-law as below 푡

= ( / ) 훽 where the coefficient, bearing a푀푆퐷 unit of퐷 µm∙ 2∆, is푡 a푡 0 measure of the apparent diffusivity. The exponent describes the movement as diffusive (Brownian motion) when = 1, sub-diffusive퐷 when < 1, super-diffusive when > 1 and ballistic when = 2. The persistence훽 of cell movement can be obtained by assessing the turning angles between훽 two successive trajectories훽 of the cell within a time훽 lag segment. Since this 훽is strongly coupled to the exponent, the exponent is used as an indicator for the persistence of movement [26]. 훽 훽 46

Results 3

Figure 3.2 Analysis of two dimensional SW480 cell motility. Cells were seeded sub- confluently on Ø 35 mm plastic dishes and were stimulated with Tβ4 directly upon seeding. Time lapse measurements were taken after 24 h of induction. (A) MSD plots of cells after induction with Tβ4 concentrations of 0.2 µM (seen in red), 1 µM (blue) and the control using 0.2 µM scrambled Tβ4 peptide (green). Black lines indicate the power-law fit according to ( / ) + . The values of the coefficient are given in (C) and in (D). (B) Examples of cell trajectories훽 tracked in a measurement lasting 3.5 h. Dots show successive cell positions tracked퐷 ∆푡 푡 0 every 60푐 seconds. Seen in red and blue퐷 are trajectories of cells훽 treated with 0.2 and 1.0 µM Tβ4. The control is shown in green. Cells treated with 0.2 µM Tβ4 moved faster on short time scales (higher apparent diffusivity values) and showed highly persistent movement (MSD exponent β = 1.40). In contrast, control cells and cells treated with 1.0 µM

Tβ4 displayed sub-diffusive to diffusive movement (β = 0.90 and β= 0.98, respectively) and 3-fold lower apparent diffusivity values.

3 Results

As shown in Figure 3.2 (A), the log average of the MSDs of all SW480 cells from each

concentration of Tβ4 used and the control was plotted as a function of the time lag. In all cases, the MSDs increased with the time lag, , in accordance with a power law. The solid lines represent the least squares fit to the data. From this line, the coefficient and the exponent were obtained. A marked∆푡 difference can be seen in the 퐷 MSDs slopes of cells treated with 0.2 µM Tβ4 and cells treated with the control or 훽 1.0 µM Tβ4. This is reflected in the coefficients and the exponents as shown in

Figure 3.2 (C) and (D). The coefficient of cells treated with 0.2 µM Tβ4 was more 퐷 훽 than three-fold higher than in cells treated with 1 µM Tβ4 and the control. Analysis of 퐷 the exponents show a highly persistent (super-diffusive; = 1.40) movement of

SW480 cells treated with 0.2 µM Tβ4. Cells induced with 1.0 Tβ4 and the control 훽 훽 demonstrate sub-diffusive to diffusive movement ( = 0.90 and = 0.98, respectively). Taken together, these results show clearly that the persistent directional movement and average velocity of an SW480훽 cell stimulated with훽 0.2 µM

Tβ4 allows it to travel much larger distances. As seen with the results obtained from the 3D invasion assays, these results also demonstrate a clear biphasic dose response

of Tβ4 concentration to 2D cell motility.

Results 3

3.2 The effect of extracellular Tβ4 on cell morphology Cells employ a myriad of strategies to overcome tissue barriers in the process of invasion. One of these is a distinct change in cell morphology called the epithelial-mesenchymal transition (EMT) which occurs during malignant progression of metastatic carcinomas. As the name suggests, EMT is marked by a change from an epithelial to a mesenchymal phenotype following primarily the loss of cell-cell adhesion mediated by the down- regulation of E-cadherin expression. This results in solitary cells bearing a more elongated, fibroblastic morphology [83]. Overexpression of the Tβ4 gene has been shown to trigger EMT in SW480 colorectal carcinoma cells [62]. Until now, the effects of extracellular Tβ4 stimulation of SW480 on cell morphology have not been investigated.

3.2.1 Analyses of SW480 cell morphology in 3D The morphology of SW480 cells was observed using three dimensional collagen gel. 5 x 104 cells were seeded onto the gels in Ø 35 mm dishes and were incubated for 3 days before image stacks through the entire gel were taken. As in the previous

experiments, the cells were treated with 0.2, 1.0 µM Tβ4 and 0.2 µM scrambled Tβ4 peptide (control) upon seeding.

Wild-type SW480 cells exhibit a round, pancake-like phenotype and, when seeded sub-confluently (5 – 10 x 104 cells), tend to group together into large monolayer clusters of between 5 – 10 cells. This is seen clearly with wild-type SW480 cells lying

on the surface of 3D collagen gels (Figure 3.3; 0 µM Tβ4 at a depth of 0 µm). After

incubation with 0.2 µM Tβ4, SW480 cells undergo a drastic change in their morphology and are found to have invaded into the collagen gel (Figure 3.3; 0.2 µM

Tβ4 at a depth of 100 µm). On the surface of the gel, induced cells assume an elongated, spindle-like morphology, forming long filapodia stretching throughout the gel (see white arrows in Figure 3.3) and are found largely as single cells within the collagen gel. These fibroblastic cells which have invaded as far as 100 µm into the gel and beyond can be identified with the same outer characteristics. SW480 cells treated

overnight with 1 µM Tβ4 were observed to be in mixed states on the surface of collagen gels, with populations of induced cells with a spindle-shaped, fibroblastic morphology (see red arrows in Figure 3.3) as well as round cells similar to the wild- type gathered in clusters. Although some cells are elongated, they do not invade into the gel and remain largely on the surface. The phenomenon of mixed phenotypes was

not seen in either the control group or with cells induced with 0.2 µM Tβ4.

3 Results

Figure 3.3 Biphasic response of SW480 cell morphology to Tβ4 concentration in 3- dimensional collagen gels. SW480 cells were seeded onto collagen gel in Ø 35 mm dishes. 0.2 µM, 1 µM Tβ4 and 0.2 µM scrambled Tβ4 (control) were added upon seeding. Cells were incubated for a total of 3 days before images of 25 random fields of view through the entire depth of the gels. Here, sample images are shown of the surface of the gels (0 µm depth) and at a depth of 100 µm. On the surface of the gel, large clusters of round cells are seen after induction with the control. Cells induced with 0.2 µM Tβ4 have undergone EMT and exhibit long filapodia (white arrows). At this Tβ4 concentration, cells have migrated up 100 µm into the gel and beyond. Cells incubated with 1 µM Tβ4 are seen to be in mixed phenotypes; round cells in clusters as well as spindle-like, fibroblastic cells (red arrows) are found on the surface of the gel. Cells treated with the control and 1 µM Tβ4 are not found at depths beyond 100 µm. Bars measure 20 µm.

Results 3

3.2.2 Analyses of SW480 cell morphology in 2D Cell morphology and spreading area on 2D surfaces were examined to accurately

quantify the changes in phenotype as a response to exogenously applied Tβ4. Cells cultured on two-dimensional glass slides were incubated for 24 h with PBS (control)

and 0.2 µM and 1 µM doses of Tβ4. Fixed cells were then stained with phalloidin to visualize actin fibers. From 25 random fields of view, fluorescent images were taken to calculate the cell area with the actin staining serving as a cell area boundary.

As seen in Figure 3.4 (A), wild-type SW480 cells are typically round and tend to form clusters in a monolayer as seen previously on 3D gel surfaces. Cells treated overnight

with 0.2 µM Tβ4 undergo distinct morphological changes, being much larger and better spread. These cells have a polarized morphology, with a clear leading (white arrow) and trailing edge (red arrow), while forming long filapodia. Cells induced with

1 µM Tβ4 have also formed filapodia but tend to cluster and revert to a shape similar to that of the control, with no clear cellular polarization.

An analysis of the calculated cell area in Figure 3.4 (B) shows that SW480 cells

treated with 0.2 µM Tβ4 are 40% larger than the control (PBS induced; average

spreading area= 905 µm2). The spreading area of cells induced with 1 µM Tβ4 is 26%

larger when compared with the control. These results show a clear biphasic Tβ4 dose response in cell spreading and are consistent with all measurements discussed previously. A cell count of each concentration group as seen in Figure 3.4 (C) revealed that SW480 cells treated with 0.2 µM showed more than a 3-fold higher incidence of single cells (13%) within a field of view when compared with the PBS control (4.5%). Consistent with the biphasic dose response, a reduction to 7% single cells were found

in the population treated with 1.0 µM Tβ4.

Results from 3D and 2D analyses of cell morphology have shown that stimulation with

extracellular Tβ4 has induced EMT similar to the overexpression of Tβ4 in SW480 cells. A major factor involved in this transition may be the cell-cell disassociation caused by the loss of E-cadherin and a massive cytoskeletal reorganization [84]. The transition from cell-cluster to single cells is seen with SW480 cells induced with

extracellular 0.2 µM Tβ4 and intracellular genetic overexpression of the peptide [48, 62]. The event of cell polarization into a leading and a trailing edge indirectly causes the reorientation of traction-generation of the cell and the formation of protrusions on one side of the cell and, thus, an increase in directional motility [85]. The increase in

cell area as seen with 0.2 µM Tβ4 treated cells indicates higher cellular traction forces generated for this directional movement. However, unlike the effects of genetic

3 Results

intracellular overexpression, the effect of extracellular Tβ4 is distinctly biphasic with respect to increasing peptide concentration. This implies that the molecular

mechanism with which extracellular Tβ4 regulates cell morphology and motility in

SW480 cells may involve other mechanisms to that of intracellular Tβ4. This point will be examined further in the proceeding sections.

Results 3

Figure 3.4 Biphasic response of SW480 cell morphology to Tβ4 concentration on two- dimensional surfaces. (A) SW480 cells were stained with fluorescently labeled phalloidin to visualize actin fibers. Cells were treated overnight with either PBS (control), 0.2 µM Tβ4 or 1 µM Tβ4. Twenty-five random fields of views were taken to quantitatively measure the cell spreading area marked by the actin boundary. The cells induced with 0.2 µM Tβ4 showed a marked polarization into a leading (white arrow) and trailing edge (red arrow). Cells of the control group showed the round wild-type morphology, forming clusters. Cells treated with 1 µM Tβ4 have formed filapodia but do not exhibit any signs of polarization. Scale bars are 20 µm. (B) Analysis of cell spreading area. Cell of the control (PBS) have an average cell spreading of 905 µm2 ± 10 µm2 (mean ± SE). 0.2 µM Tβ4 induced cells are 40% larger than the control (1257 ± 17 µm2) and 1 µM Tβ4 induced cells are 26% larger (1135 ± 12 µm2). Bars indicate standard errors. (C) Percentage of single cells found on 2D culture surfaces. 4.6% ± 0.5% of cells treated with PBS are single cells. 12.9 % ± 0.8% of cells treated with 0.2 µM Tβ4 and 7.1 % ± 0.4% of cells treated with 1 µM Tβ4 are single cells. Bars indicate standard errors.

3 Results

3.3 Effect of extracellular Tβ4 on cell compliance, dynamics and contractility The process of directional migration involves the polarization of a cell into a leading and trailing edge. At the leading edge of the cell, protrusions are generated by the polymerization of actin filaments which are stabilized by adhesions to the substrate lying under the cell [86-87]. Generation of traction forces by actin-myosin contraction enables the formation of adhesions at the leading edge and the disassembly of adhesions at the rear to allow for the forward locomotion of the cell [88]. This is a dynamic process which requires constant remodeling of the cytoskeleton and turnover of adhesions. In this chapter, the effects of extracellular Tβ4 on the cytoskeletal mechanics, dynamics and cellular tractions of SW480 cells are examined.

3.3.1 Creep response and cell mechanics The magnetic tweezer method was employed to examine the effect of exogenously

applied Tβ4 on cytoskeletal stiffness and viscoelasticity. Measurements were performed on SW480 cells incubated with fibronectin-coated Ø 4.5 µm superparamagnetic beads. The fibronectin coating links the beads to the cytoskeleton and acto-myosin fibers through cell-surface integrins [89]. The magnetic tweezer exerts an external force onto the bead, thereby actively deforming the cytoskeleton in the process. This method allows for the quantification of several physical properties of the cytoskeleton. Firstly, the stiffness of the cell, which is a measure of the ability of the cytoskeleton to withstand the forces, and secondly its hysteresivity or loss tangent, which is a measure the turnover rate of cytoskeletal bonds in response to the active external force [17, 90]. A staircase-like sequence of increasing force steps from 0.5 nN to 10 nN was applied on fibronectin coated super-paramagnetic beads bound to the cytoskeleton. The displacements of the beads were tracked online followed by statistical analysis. Separate measurements on SW480 cells were made to observe the

response of 24 h induction with either the control (PBS) or Tβ4 concentrations of 0.2 µM and 1 µM as done in previous measurements.

The displacement ( ) of the fibronectin coated bead in response to a force step of magnitude Fstep follows the power law [90] as below: 퐽 푡 ( ) = ( / ) 푏 The coefficient is the compliance퐽 푡 of the퐹푠푡푒푝 cytoskeleton푎 푡 푡0 equal to the inverse stiffness. This coefficient indicates the extent of cytoskeletal deformation caused by a unit force. The power푎 -law exponent describes the viscoelasticity of the cytoskeleton. A value of = 0 is obtained from a purely elastic solid whereas a value of = 1 is obtained from a purely viscous fluid. A 푏 exponent value of between 0 and 1 is expected for 훽 푏 viscoelastic systems such as the cytoskeleton. b is related to the loss tangent, or 푏 hysteresivity η of the system by η=tan(b π/2). Within each group of measurements, a

Results 3

log normal distribution of stiffness values (1/ ) and a normal distribution of the power-law exponent were recorded. Due to this, the geometric mean of stiffness and the arithmetic mean of the power-law exponent푎 were calculated [17]. The average response of the cells was taken as the value averaged over all cells within each concentration group.

Figure 3.5 Stiffness of Tβ4 induced SW480 cells. Cells stimulated overnight with the PBS control, 0.2 µM and 1.0 µM Tβ4 were incubated with fibronectin-coated super-paramagnetic beads. External force applied to the bead were increased step-wise from 0.5 – 10 nN. (A) Displacement of the bead following the direction of the force followed the power law, ( ) = ( / ) . The stiffness of the cell is given by the inverse of the power-law coefficient , a, in units of nN/µm푏 at all force steps. (B) A biphasic response in stiffness was recorded at 0 퐽external푡 푎 forces푡 푡 of 0.5 nN; cells stimulated with 0.2 µM Tβ4 were more than twofold stiffer (13.3 nN/µm) than the control (5.8 nN/µm) and cells stimulated with 1.0 µM Tβ4 (6.2 nN/µm). Bars indicate the standard error.

3 Results

A non-linear response of bead displacement ( ) to increasing force was seen with cells in all groups as shown in Figure 3.5 (A). Bead displacement decreased with the increase of force less than proportional as a result퐽 푡 of cellular stress stiffening. Non- linear stress stiffening is a behavior typical of cross-linked biomaterial such as extracellular matrix and the actin cytoskeleton [91-92]. Stress stiffening of cells

treated with 1 µM Tβ4 and the control were similar; both populations showed a twofold increase in stiffness at 10 nN compared to at 0.5 nN. However, cells induced

with 0.2 µM Tβ4 displayed less stress-stiffening; only a one and a half fold increase in stiffness was recorded over the force range. These results suggest that cells treated

with 0.2 µM Tβ4 have an inherently more pre-stressed cytoskeleton and as a result have a smaller increase in stress-stiffening in response to higher external force exerted onto the cytoskeleton. Closer analysis of cell stiffness at an external force of

0.5 nN revealed that cells treated with 0.2 µM Tβ4 showed a slightly more than twofold increase in stiffness compared to the PBS control. No statistically significant difference was found between the stiffness values of cells induced with 1 µM and that of the control.

Figure 3.6 Viscoelasticity of Tβ4 induced SW480 cells. (A) Power law exponent b represents the viscoelastic properties of the cells as a function of increasing force. (B) Power law exponent b at an external force of 0.5 nN; no changes in viscoelasticity was seen at all concentrations. Bars indicate the standard error.

Results 3

The dynamic turnover rate of cytoskeletal bonds was examined by analyses of the viscoelastic exponent. The exponent of this power-law creep response tended to

decrease at a Tβ4 concentration of 0.2 µM especially at higher force values, as 푏 expected for stiffer cells, although the differences compared to control conditions or

Tβ4 concentrations of 1 µM were not statistically significant (see Figure 3.6).

3.3.2 Two-dimensional cellular tractions As previously recorded in literature, cell stiffness correlates directly with prestress and internal contractility [93]. Cell contractility is caused by the ATP-dependent action of myosin, which cross-link actin filaments to form stress fibers. This is a crucial step required in cell adhesion, cell movement and cell division [94]. The contractile force a cell exerts on a two dimensional collagen substrate was examined through 2D traction microscopy. Polyacrylamide (PAA) gels with a known stiffness of Young’s modulus = 12.5 kPa were coated with collagen to enable cell adhesion. Due to cell tractions, elastic strain energy is stored in the gel [75]. The total strain energy of each cell, , is obtained as the sum of local tractions causing gel deformation following the equation given below: 푈 = 1 푈 �2� �→푇 �→푟 � ∙ →푢 �→푟 � 푑푥푑푦 Here, is the traction vector and the displacement vector. The resulting value of elastic strain energy is given in units of pico-Joules per cell. As cellular →푇 �→푟 � →푢 �→푟 � tractions are dependent on the spreading area of the cells, the calculated strain energy was normalized to the cell area giving a value in strain energy density with a unit of pJ/µm2.

Marked differences in cell contractility were measured between Tβ4 induced cells and the controls. Compared to the PBS control (0.108 pJ/cell ± SE 0.028), cells treated

overnight with 0.2 µM Tβ4 exhibited a threefold increase (0.430 pJ/cell ± SE 0.063)

and cells induced with 1 µM Tβ4 showed an 85% increase (0.235 pJ/cell ± SE 0.03) in elastic strain energy. The differences became more pronounced when the cell spreading area is taken into consideration as shown in Figure 3.8 (B). SW480 cells

show an apparent biphasic response to the increasing dose of Tβ4 in contractility, with

the highest contractility at stimulation with 0.2 µM Tβ4. These results are consistent with those recorded with cell stiffness measurements as described previously in section 3.3.1.

3 Results

Figure 3.7 Traction force maps of SW480 cells: contractility of Tβ4 stimulated cells on 2D substrates. Cells were seeded on collagen coated PAA gels (Young’s modulus = 12.5 kPa) and immediately treated with exogenous Tβ4. 2D- traction microscopy was performed after 24 h incubation with the peptide. On the left are bright field images of the cells, scale bars indicate 20 µm. Traction field maps of cells are given on the right. Color bars indicate differences in local tractions in units of Pa. Cells were sampled from random fields of view.

Results 3

Figure 3.8 The effect of exogenous Tβ4 on the contractility of SW480 cells on 2D substrates. (A) Strain energy of SW480 cells was calculated from cellular tractions integrated over the spreading area of each cell. The highest level of strain energy per cell (0.430 pJ/cell ± SE 0.063) was recorded on cells treated with 0.2 µM Tβ4 compared with that of cells induced with the PBS control (0.108 pJ/cell ± SE 0.028) and 1.0 µM Tβ4 (0.235 pJ/cell ± SE 0.028). (B) The cell spreading area was taken into consideration when calculating the strain energy density. Cells treated overnight with 0.2 µM Tβ4 have a strain energy density (343.8 µN/m ± SE 34.3) significantly more than that of the control (144.4 µN/m ± SE 14.4) and cells treated with 1.0 µM Tβ4 (207.3 µN/m ± SE 20.7). Bars indicate the standard error.

3.3.3 Nanoscale particle tracking of cytoskeletal-bound beads: studying the time- dependant activity of Tβ4 on cytoskeletal dynamics Cytoskeletal dynamics can be quantified by directly tracking the spontaneous movement of fibronectin coated beads (Ø 4.5 µm), which are directly connected to the cytoskeleton through cell-surface integrins. In comparison with methods that actively employ external force such as the magnetic tweezer, this method detects the internal cellular forces that cause bead movement, mainly the formation of stress fibers and the reorganization of the cytoskeleton, both of which involve actin in filamentous and monomeric form as well as motor proteins such as myosins. To analyze the

differences between the short term and long term effects of Tβ4 on the cytoskeletal remodeling of SW480 cells, this method was used on cells stimulated for 30 min

(short term) and 24 h (long term) with exogenous Tβ4.

3 Results

Cytoskeletal dynamics was measured by tracking the position of bound beads at 10 times per second over a period of 5 min. The mean square displacement (MSD) of the moving bead is fitted according to the power law equation, = ( / ) + as seen previously in section 3.1.2 (2D cell motility measurements). Here,훽 the fit parameter, D, carries a unit of µm2 and measures the 푀푆퐷 퐷 ∙ ∆푡 푡 0 푐 diffusivity of bead motion, correlating directly to the amplitude of its movement. D values in each of the scenarios (PBS control and induction with either 0.2 µM or 1 µM

Tβ4) were found to be distributed over a wide range in a log-normal manner. Thus, an accurate method of obtaining the average MSD for a given concentration would be to use geometric mean of all MSD values [26]. The power-law exponent, β, describes the persistence of bead motion.

Bead diffusion measurements of SW480 cells after 30 min incubation (short term) revealed a sharp decrease in diffusivity (pre-factor D) values with cells stimulated with both 0.2 µM and 1 µM compared with the control (see Figure 3.9 (A)). D values of

cells incubated with 0.2 µM Tβ4 (1.94 x 10-4 µm2 ± 0.13 x 10-4) (mean ± SE) were 42% smaller and with 1 µM (1.62 x 10-4 µm2 ± 0.15 x 10-4) 51% lower than that of the control (3.33 x 10-4 µm2 ± 0.27 x 10-4). β coefficient values of cells treated with 0.2 µM

Tβ4 (1.47 ± 0.015) and 1 µM Tβ4 (1.44 ± 0.017) were significantly higher than that of the PBS control (1.39 ± 0.013). No statistically significant differences could be seen

between β coefficient values of cells treated with 0.2 µM or 1 µM Tβ4.

Diffusivity values are influenced by a network of tension-carrying stress fibers spanning the entire cell, which are constantly being restructured. Following short term stimulation, lower bead diffusivity recorded in cells induced with both 0.2 µM

and 1 µM Tβ4 is a result of the restricted movement of beads bound tightly to the cytoskeleton. This phenomenon is indicative of the higher amounts of polymerized actin fibers compared to the total amount of actin in these cells, causing rigidity of the cytoskeleton. On the other hand, we assume the persistence (β coefficient) values to reflect the directional persistence of cytoskeletal remodeling events. The higher bead

persistence values of cells stimulated with 0.2 µM and 1 µM Tβ4 indicate that the actin fibers of these cells are more aligned than those of the control. Taken together, these

results show that short term induction of cells with Tβ4 causes a pronounced remodeling of the cytoskeleton due to the formation of stress fibers.

Results 3

Figure 3.9 Short term effects of extracellular Tβ4 on cytoskeletal remodeling. The spontaneous motion of fibronectin coated Ø 4.5 µm beads followed the power-law, = ( / ) + . (A) After 30 min of induction, power-law cofactor, D, has decreased with increasing휷 concentrations of Tβ4. This indicates the immediate increase in actin푴푺푫 polymerization푫 ∙ ∆풕 풕 ퟎ into풄 stress fibers causing less diffusivity in bead movement due to the effect of Tβ4 application. (B) The increase in the power-law exponent, β, values indicate an alignment of cytoskeletal components. No statistically significant differences were recorded with β values between cells induced with 0.2 µM and 1.0 µM Tβ4. Bars indicate one standard error.

Long-term stimulation (>24 h) of SW480 cells reveal a biphasic response to increasing concentrations of Tβ4. A decrease in bead diffusivity values of cells incubated with

0.2 µM Tβ4 (2.45 x 10-4 µm2 ± 0.16 x 10-4) (mean ± SE) compared with the PBS control (3.22 x 10-4 µm2 ± 0.24 x 10-4) was recorded. Diffusivity values of cells treated overnight with 1 µM Tβ4 (2.96 x 10-4 µm2 ± 0.30 x 10-4) are not significantly lower than that of the control. Analyses of bead persistence, power-law factor β, show distinct changes in the turnover rate of the cytoskeleton. Cells treated overnight with

0.2 µM Tβ4 have significantly lower β values Tβ4 (1.33 ± 0.012) compared with the PBS control (1.39 ± 0.012). However, compared to the control, cells induced overnight with 1 µM Tβ4 show a significant increase in bead persistence (1.47 ± 0.015) (see

Figure 3.10). Thus, long term stimulation with Tβ4 shows an apparent biphasic response in cytoskeletal dynamics; cells treated overnight with 0.2 µM Tβ4 still show increased formation of stress fibers resulting in lower diffusivity in the motion of

3 Results

bound fibronectin-coated beads compared to the control and 1.0 µM Tβ4. These results are consistent with increase in stiffness and contractility seen in preceding measurements in magnetic tweezer and 2D traction microscopy. The biphasic

response in the power-law exponent show cells treated with 0.2 µM Tβ4 have a more dynamic turnover rate of the actin cytoskeleton than that of cells treated with the β control or 1 µM Tβ4.

Figure 3.10 Long term effects of extracellular Tβ4 on cytoskeletal remodeling. (A) Significantly lower bead diffusivity, power-law cofactor, D, values were seen with 0.2 µM Tβ4 treated cells after 24 h when compared with cells treated with 1.0 µM Tβ4 and the control. No significant differences were seen between the control and 1.0 µM induced cells. Here, only cells treated with 0.2 µM Tβ4 have retained increased levels of actin polymerization. (B) The lowest β values were recorded on cells treated with 0.2 µM Tβ4 is indicative of the significantly higher turnover rate of the cytoskeleton at this concentration. Cells induced with 1.0 µM Tβ4 show a higher β value compared with the control. After 24 h of induction, a biphasic response to increasing Tβ4 concentrations can be seen in SW480 cytoskeletal dynamics and production of actin stress fibers. Bars indicate one standard error.

When comparing the effects of long term and short term Tβ4 stimulation of SW480 cells, it becomes evident that the initial increase in the polymerization of actin fibers, reflected by the drop in bead diffusivity (power law D coefficient values) after 30 min

stimulation with Tβ4, is transient. After 24 h, less stress fibers are formed in

stimulated cells, as seen with the slight rise in D values after 0.2 µM Tβ4 stimulation and a rise to the same as the control with 1.0 µM shown in Figure 3.10 (A).

Results 3

Considering the dynamic reorganization of the cytoskeleton, Tβ4 induces a short term decrease in the turnover rate of actin polymerization, seen with the increase in the power-law exponent β. However, after 24 h, only cells treated with 0.2 µM exhibit a more dynamic cytoskeleton. Taken together, the biphasic cellular response to Tβ4, with a combination of higher formation of actin stress fibers and a more dynamic turnover can be seen only on longer timescales with a peak at 0.2 µM stimulation.

3 Results

3.4 Tβ4 and ILK: The Biphasic Switch in 3D Cell Invasion Assays Biphasic cellular regulation is by no means an isolated phenomenon. This type of regulation has been found in the MAPK [95-97] and NF-kB signaling pathways [98]. In both examples, two components in the signaling cascade are required to induce a biphasic response. The first is involved in triggering the system’s activation whereas the second regulates the concentrations of the members in the system to elicit the biphasic response. Thus, an over-expression or reduced expression of a protein within the system will lead to inhibition of the signaling cascade. This mechanism, also known as combinatorial inhibition, may be involved in the biphasic response seen in cellular mechanics and dynamics to Tβ4. Combinatorial inhibition usually occurs in the presence of a molecule capable of binding and interacting with multiple molecules such as in scaffolding and adapter proteins. In this chapter, we will examine this hypothesis closely and present a model to explain the biphasic phenomenon involving Tβ4 and ILK.

3.4.1 Role of ILK as a molecular switch involved in the Tβ4 biphasic response Several processes are involved in cell migration. As discussed previously, a major factor in this event is the generation of contractile forces within the cell and the transmission of these forces to the substrate [99]. Cell contractility is caused by actin- myosin contraction regulated by the phosphorylation of the myosin light chain (MLC) domain. With its role as a Ca2+ independent myosin light chain kinase, studies have linked ILK with cell contractility through its ability of phosphorylating MLC [100].

Recently, however, a contradictory role of ILK in the regulation of MLC has been documented. Elevated activity of ILK has been known to negatively regulate RhoA. RhoA plays an indirect role in the MLC pathway [101-102]. Through its effector, Rho Kinase (ROCK), MLC phosphatase is phosphorylated and remains inactive, thus, unable to dephosphorylate and inactivate myosin light chain. However, through the inhibition of RhoA by ILK, MLC phosphatase is maintained in the active state and leads to the inhibition of actin-myosin contraction.

In the cellular biphasic response seen throughout all our experiments, it is assumed

that exogenous Tβ4 activates ILK which in turn regulates actin-myosin contractility via the direct phosphorylation of MLC or the negative regulation of RhoA. At this point, these opposing pathways are not necessarily mutually exclusive events. Initially,

increasing concentrations Tβ4 triggers higher levels of active ILK which function as a Ca2+ independent MLC kinase. This in turn leads to increasing levels of actin-myosin

contractility directly related to increased cell motility. At levels of Tβ4 exceeding 0.2 µM, increasing levels of RhoA inhibition overrides the ability of ILK to phosphorylate MLC leading to a decrease in cell motility (see Figure 3.11).

Results 3

Figure 3.11 Tβ4-ILK molecular switch. Upon initial activation by Tβ4, ILK assumes the role of a Ca2+ independent [103] MLC kinase and phophorylates MLC at Thr18/Ser-19 which leads to increased actin-myosin mediated contraction and increased cell motility. At higher levels of exogenously applied Tβ4, the activity of ILK as a regulator of RhoA increases which leads to increasing inhibition of ROCK. As a result, higher levels of active MLC phosphatase dephosphorylate MLC, causing less actin-myosin contraction and a decrease in cell motility. In this model, ILK acts as a molecular switch in the biphasic cellular response to increasing levels of extracellular Tβ4.

3 Results

3.4.2 Bypassing RhoA inhibition: the effect of calyculin-A on Tβ4-induced cell invasion

Tβ4 induced SW480 cells were treated with calyculin-A, an MLC phosphatase inhibitor [104-105] to study the effect of bypassing ILK-induced RhoA inhibition. Through MLC phosphatase inhibition, phosphorylated MLCs are kept at higher levels, thereby, causing further stimulation of actin-myosin contraction [106]. Invasion assays were

carried out on SW480 cells incubated with Tβ4 concentrations of 0.2 µM and 1.0 µM

and 0.2 µM scrambled Tβ4 as a control. At each Tβ4 concentration level, invasion assays were carried out with increasing levels of calyculin-A (0 nM, 10 nM and 50 nM). Cells were allowed to invade into the collagen gel overnight before stimulation with calyculin-A.

The general effect of MLC phosphatase inhibition is demonstrated clearly in the

control measurements with SW480 cells incubated with 0.2 µM scrambled Tβ4 and 10 nM and 50 nM calyculin-A stimulation. At pCum 10%, cells stimulated with calyculin-A have invaded almost twofold deeper into the gel than the control. No significant difference was seen between cells stimulated with 10 nM or 50 nM calyculin-A. This trend is also observed in further invasion assays involving

stimulation with Tβ4. Cells stimulated with calyculin-A and 0.2 µM Tβ4 show a 20% increase in invasion depth when compared with non-stimulated cells. However, when

compared with cells stimulated with only 0.2 µM Tβ4, cells induced with both

calyculin-A and 0.2 µM Tβ4 show up to 30% lesser invasion depth. A 28% increase in

cell invasivity is seen further with cells stimulated with both 1.0 µM Tβ4 and calyculin- A when compared with non-stimulated cells. A dramatic increase of more than

twofold in invasivity of cells stimulated with both calyculin-A and 1.0 µM Tβ4 is seen,

however, when compared to cells stimulated with only 1.0 µM Tβ4.

Taken together, these results show that the motility of SW480 cells can be enhanced through maintaining cell contractility by sustaining MLC phosphorylation. A steady increase in three-dimensional cell motility is seen as a result of both the effects of

calyculin-A and Tβ4 stimulation. Experiments with both 1.0 µM Tβ4 and calyculin-A show clearly that the role of ILK as a regulator of RhoA, which is assumed to be the

cause of the decrease in the motility of cells stimulated with only 1.0 µM Tβ4, is compensated by enhancing MLC phosphorylation through calyculin-A. On the other hand, the finite number of MLC available for phosphorylation is a limiting factor leading possibly to a plateau in the combined effect of the two stimuli which can be

expected when applying higher concentrations of Tβ4 and calyculin-A.

The peak level of SW480 invasion seen with cells stimulated with 0.2 µM Tβ4 was not

observed with cells treated with both calyculin-A and 0.2 µM Tβ4. At this point, we can

Results 3

assume that the overall synergistic effect of Tβ4 and calyculin-A on MLC phosphorylation is limited. MLC phosphorylation is not the only pathway determining the increase in cell invasion in connection with Tβ4 stimulation of SW480 cells.

Figure 3.12 The combined effect of calyculin-A and Tβ4 on the 3D invasion of SW480 cells into collagen gels. 3D invasion profiles of SW480 cells were analyzed after incubation with both calyculin-A (at concentrations of 10 and 50 nM) and Tβ4. (A) Influence of only calyculin-A on 3D cell invasion. Due to sustained cell contractility as a result of continued phosphorylation of MLC, cells treated with 10 and 50 nM Calyculin invade twofold deeper

3 Results

into collagen gels. (B) Influence of calyculin-A combined with 0.2 µM Tβ4 on 3D cell invasion. Cells treated calyculin-A and 0.2 µM Tβ4 invade 30% less than cells treated with only 0.2 µM Tβ4. (C) Influence of calyculin-A combined with 1.0 µM Tβ4 on 3D cell invasion. Cells treated with both calyculin-A and 1.0 µM Tβ4 were able to invade 28% deeper into gels than when treated with only 1.0 µM Tβ4. (D) Summary: A steady linear increase in cell invasion depth was recorded with the addition of calyculin-A to increasing concentrations of Tβ4. At a Tβ4 concentration of 0.2 µM Tβ4, however, cells treated with only Tβ4 were able invade 30% further into collagen gel than with the addition of calyculin-A. This shows that prolonged cell contractility due to the phosphorylation of MLC alone is not the only factor determining the invasivity of SW480 cells after stimulation with Tβ4.

Results 3

3.5 Tβ4 – membrane interaction

Until now, the effect and pathway of exogenously applied Tβ4 on cells is not known. The mechanisms of entry or a cell surface receptor for this peptide have not yet been discovered. So far, previous studies have given no explanation as to why minute concentrations of Tβ4, as low as 0.1% of intracellular concentrations, are able to elicit pronounced changes in cellular mechanics resulting in altered morphology and motility. Due to its molecular properties, which will be examined further in this chapter, Tβ4 presents itself as a plausible candidate for membrane binding.

3.5.1 Determining the binding potential of Tβ4 to lipid membranes using differential scanning calorimetry (DSC)

Amino acid analysis of Tβ4 showed that clusters of basic and hydrophobic residues make up about 60% of the entire peptide. Basic amino acids are known to interact with acidic domains on a lipid membrane. In addition, hydrophobic residues in the peptide may enable its insertion into a lipid membrane and function as an anchor

[107]. NMR spectroscopic analysis of Tβ4 in solution has shown that the peptide is largely unstructured in water and consists of two α helices in fluorinated alcohols [30], the first helix extending from residues 4-16, and a second helix from residues 30- 40. Among the 43 amino-acids of the entire peptide, 9 residues are basic and 1 residue

is aromatic [108]. These characteristics combined make Tβ4 a good candidate for membrane binding. Similar modes of membrane association have been reported for other proteins such as myristoylated alanine-rich C kinase substrate, MARCKS [107] and Human Immunodeficiency Virus Type 1, HIV-1 [109].

Using differential scanning calorimetry, studies were carried out to explore the lipid-

binding ability of Tβ4. Theoretically, the insertion of the peptide into the lipid membrane and interaction with the hydrophobic acyl chains causes the expansion, realignment and destabilization of the phospholipid membrane. This effect lowers the transition temperature of the lipid membrane from the gel to the fluid phase. Evidence of this phenomenon is given as the decrease in the heat capacity and phase transition enthalpy.

3 Results

Figure 3.13 Lipid-binding qualities of Tβ4 to DMPG/DMPC lipids analyzed using differential scanning calorimetry (DSC). (A) DSC thermogram of 175 µM Tβ4 and the control in a lipid mixture with a molar ration of 70:30 DMPC to DMPG. The distinct reduction in the Tm of the Tβ4 lipid vesicle mixture indicates the insertion of the peptide into the lipid bilayers. (B) Relative transition enthalpy (∆H/∆H0) as a function of increasing peptide concentration. Raising the negative charge of the lipids caused only a slight decrease in peptide insertion into the membrane. As expected, no changes in relative transition enthalpy were recorded with increasing concentrations of insulin used as the control.

In this experiment, small unilamellar vesicles (SUVs) with different ratios of

DMPC/DMPG were incubated with increasing concentrations of Tβ4 (from 60 µM to 175 µM peptide). In these experiments, insulin, a molecule confirmed to having no membrane-binding properties, was used as a control. An example of a thermogram

comparing the specific heat of 175 µM Tβ4 in lipid with a molar ratio of 70:30 DMPC to

DMPG and lipid only is shown in Figure 3.13 (A). A distinctive drop in the Tm of Tβ4 in lipid indicates the insertion of the peptide into lipid bilayers.

In Figure 3.13 (B), the relative transition enthalpy (∆H/∆H0) as a function of increasing peptide concentration is plotted for DMPC/DMPG lipid vesicles with molar ratios of 70:30 and 30:70 to vary the strength of the negative charge of the lipid vesicles. For both lipid types, a drop in the relative transition enthalpy was measured with increasing peptide concentration. Raising the negative charge of the lipids caused a slight decrease in peptide insertion into the membrane. As expected, no changes in relative transition enthalpy were recorded with increasing concentrations of insulin.

These results are a clear indication of the membrane binding properties of Tβ4.

Results 3

3.5.2 Fluorescence correlation spectroscopy (FCS) of membrane-bound fluorescent Tβ4 Although DSC is a well established method for examining peptide-membrane insertion, it cannot accurately measure binding kinetics. Moreover, this method is not suitable for binding measurements for nanomolar concentrations, as loss of peptide and lipid are bound to occur as they adsorb onto the walls of measurement containers and pipettes. The fluorescence correlation spectroscopy (FCS) method, however, allows for a direct measurement of single-molecules in solution.

Originally, FCS had been utilized to study molecular diffusion and reaction dynamics [79, 110-111] . This method involves the measurement of fluorescence intensity

fluctuations caused by the movement of fluorescently labeled Tβ4 moving in and out of a detection volume (1 µm3) defined by the focus of an excitation laser beam. Unbound fluorescent peptide diffusing rapidly through the volume causes a highly fluctuating fluorescence signal. Peptides bound to the much larger SUVs move at much slower rates and result in a longer correlated pattern of bursts in the fluorescence signal. Auto-correlation analysis of the fluorescence fluctuations yields information on

the diffusion kinetics of bound and unbound fluorescent Tβ4. Using this method, we

are able to resolve the single-molecule diffusion of the fluorescently-labeled Tβ4 on lipid membranes.

The binding of Tβ4 to synthetic lipid membranes was examined with fluorescence

correlation spectroscopy (FCS) using Alexa488-labeled Tβ4 and SUVs. The SUVs had PS/PC ratios of 1:5 and 1:6 to mimic cellular membranes, which have concentrations between 10-30% of monovalent acidic lipids (mainly DMPS) [107]. While measurements were made with increasing concentrations of lipid, the concentration

of Tβ4 was kept constant at a minimum of 50 ng Tβ4-Alexa488 /300 µl lipid buffer to ensure that a minimum number of fluorophores on average (1..10) will be in the detection volume at a time during the measurement. Control measurements were

done on Tβ4- Alexa488 in lipid buffer only. Single photon counts were registered at a rate of 250 kHz for 10 s to maximize the resolution of yielded information. Autocorrelation plots ( ) obtained from each measurement were fitted according to the normalized autocorrelation function (see Materials and Methods 2.2.8). 퐺 휏

3 Results

Figure 3.14 Lipid membrane binding of Tβ4 measured through fluorescence correlation spectroscopy. (A) Autocorrelation curves of Tβ4-Alexa488 in increasing concentrations of 1:6 PS/PC lipids. There is no significant difference in lag times of the measured fluorescent signal. (B) An increase in the amount of acidic lipids (1:5 PS /PC) results in a significant right shift of autocorrelation curves with increasing lipid concentrations. These results suggest longer lag times (τ) of the fluorescent peptide in the detection volume, consistent with the binding of fluorescently labeled Tβ4 on much larger SUVs. The concentration of Tβ4-Alexa488 was kept to a minimum (50 ng Tβ4/300 µl lipid buffer) to limit the particles in the detection volume for measurement efficiency. Autocorrelation curves are taken as an average of 10 measurements. (C) A typical pattern of photon count rate as a function of time for Tβ4-Alexa488 in lipid buffer only. (D) Fluorescent signal pattern of Alexa488- Tβ4 in 5 µM 1:5 PS/PC. Labeled SUVs result in a photon count rate bursts and higher signals.

Results 3

Figure 3.14 (A) shows the autocorrelation plot of Tβ4-Alexa488 in increasing lipid concentrations (PS/PC of 1:6) and the control (no lipids). Each curve is obtained from an average of 10 measurements. As seen in Figure 3.14 (A) The addition of a 1:6 DMPS/DMPC lipid mixture (concentration ranging from 1 – 5 µM) to an Alexa488- labeled Tβ4 solution caused only a marginal change in the correlation time (τD) when compared to control (no lipids added), indicating that the peptide did not bind to or insert into the SUV lipid membrane. At a DMPS/DMPC ratio of 1:5, however, the correlation time increased dramatically with increasing lipid concentrations, indicating that Tβ4 bound to the lipid membrane as seen in Figure 3.14 (B) and Figure

3.15, confirming that Tβ4 and not the fluorophore bound to the SUVs. Taken together, these results show that Alexa488-Tβ4 is able to bind to lipid membranes, and furthermore indicate that basic residues in the Tβ4 peptide are mainly responsible for lipid binding through their interaction with acidic lipid head groups. A similar mechanism of binding was reported previously for the effector domain of MARCKS (residues 151-175) [107].

The insertion and binding of extracellular Tβ4 to the cell plasma membrane offers a mechanism of how the local concentration of the peptide can be greatly increased while at the same time its diffusion is greatly decreased so as to increase the reaction probability with other membrane-associated signaling proteins such as ILK and

PINCH [112-113]. Previous studies have shown the co-localization of actin-free Tβ4 and ILK around cell edges in migratory cells [55]. In addition, the binding affinity of

Tβ4 with G-actin is much higher (dissociation constant, Kd = 0.5 – 2.5 µM, [114]) compared to binding with the lipid membrane (Kd = 72 µM as measured in this study using FCS and lipid vesicles with a DMPS/DMPC ratio of 1:5). Therefore, intracellular

Tβ4 is not expected to display a pronounced association with the cell membrane in the presence of G-actin. This explains our finding that the binding of small concentrations of extracellular Tβ4 to the lipid membrane can induce dramatic cellular responses even in the presence of large intracellular concentrations of Tβ4.

3 Results

Figure 3.15 The effect of acidic lipid head groups on the binding of Tβ4 -Alexa488 on membranes. Correlation times of Tβ4-Alexa488 in the presence of SUVs for varying lipid concentrations with a PS/PC ratio of 1:5 (squares) or 1:6 (triangles). As control, Alexa488 fluorophore without Tβ4 was added to lipids with a PS/PC ratio of 1:5 (circles). Error bars (mean ± SE, n=10) are not shown when smaller than data symbols.

4 Discussion

4.1 The effects of Tβ4 on Cellular Mechanics and Motility

The results obtained from the course of this work have shown that Tβ4, when applied exogenously to SW480 cells, does not act classically as an actin sequestering protein. Past studies give evidence of Tβ4 having multiple functions [34]. The following section discusses in depth how Tβ4 can affect the mechanics of the cytoskeleton and the regulation of SW480 cell motility.

4.1.1 Tβ4 regulates the stiffness and contractility of SW480 cells The stiffness of a cell is a measure of the number of the interactions and the elasticity of bonds in the cytoskeletal network which transfers mechanical forces from the cell to the fibronectin-coated bead. The degree of internal cellular prestress, cell geometry, the thickness of the cell below the fibronectin-coated bead, the number of cell surface integrins bound to the bead, and the number of focal adhesions and associated proteins involved in binding the bead are all factors that influence the stiffness of a cell. It has been shown so far that up to a concentration of 0.2 µM, that increasing

amounts of exogenously applied Tβ4 results in the increase in stiffness of SW480 cells.

The forces during the tweezing experiments that are needed to detach the fibronectin coated bead from the cell characterize the bead binding strength. As with cell stiffness, the bead binding strength is related directly to the number of molecular interactions that relay mechanical forces between the integrin-bound bead and the cytoskeleton [18-19]. These include the connections in the cell involved in the out-in force transmission, such as the fibronectin-coated bead with the integrins, the integrins with the focal adhesion proteins, and the focal adhesion proteins with the cytoskeleton. Central in this network are mechano-transducers in the focal adhesion complex. Bead binding strength also depends on how these structures yield to the forces. In experiments involving externally induced forces of up to 10 nN, no

difference in bead detachment was seen across all concentrations of Tβ4 used and the

control. This leads to the conclusion that Tβ4 does not compromise the effect of force transmitting elements between integrins, focal adhesions and the cytoskeleton.

The yielding of the cell to pulling forces exerted onto the fibronectin-coated bead can also be measured by recording the differential cell stiffness at increasing forces. Irreversible yielding can occur through induced stress and will decrease the stiffness of the cell. The stretching of the cytoskeleton under stress results in the non-linear response of cellular mechanics. According to most models, this phenomenon has its origins from the prestress or contractile tension of the filamentous cytoskeletal network. Cellular prestress is a mechanism responsible for the stability of cell shape [93, 115] and is caused by acto-myosin contraction. The prestress can be estimated from the tractions generated through the adhesion of cells onto the extracellular matrix. In tweezing experiments, stepwise increasing external force from 0.5 nN to

Discussion 4

10 nN in 0.2 µM Tβ4 treated cells produced less increase in stiffness (one and a half fold increase) when compared with the control and 1 µM Tβ4 treated cells (both over twofold increase in stiffness). Taken the observations in bead detachment and cellular prestress together, it can be concluded that although Tβ4 may not alter passive force transmission through the focal-adhesion complex, up to a maximum working dose of 0.2 µM, it does, however, influence the active acto-myosin contraction regulating cytoskeletal pre-stress in SW480 cells.

In each individual creep measurement across all concentrations of Tβ4, the recorded response of bead displacement over time followed a power law, in agreement with past literature [89-90, 116-119]. A higher power-law exponent value, b, would indicate a more viscous system and, in sum, a higher turnover of interactions that transmit the externally exerted stress intracellularly. When comparing the power-law exponent, b, of SW480 cells, no statistically significant difference was recorded at all force steps throughout all Tβ4 concentrations and the control. Even at forces of 10 nN, no change of the power-law exponent was observed for all concentrations. Thus, Tβ4 does not result in a change of yielding of the cell to increasing induced stress, nor does it contribute to the fluidization of the force transmitting structures within the cell. As in the observations in stress stiffening, these results confirm that Tβ4 does not affect the transmission of externally induced forces from the extracellular environment, through the integrins and focal adhesion to the cytoskeleton. Taken together, all the results so far have shown that Tβ4 does not cause fluidization of the cytoskeleton as would be the case if the peptide would act predominantly as a classical actin- sequestering protein.

While magnetic tweezing assays record the active cellular forces resisting the externally induced deformation of the cytoskeleton, cellular traction microscopy measures the contractile prestress, or the intrinsic strain energy stored within the matrix. Although SW480 cells do not exhibit a pronounced development of stress fibers, they generate substantial traction forces when compared to other cell types

[75, 120]. SW480 cells treated with 0.2 µM Tβ4 showed more than a fourfold increase in strain energy generated per cell when compared with the negative control. A tight correlation between cell stiffness and contractility has been shown in previous studies [19, 121]. Extensive experimental evidence have shown that cell traction forces are generated through acto-myosin contraction induced by the phosphorylation of the myosin light chain (MLC) by myosin light chain kinase (MLCK) and Rho-kinase [3, 88,

122]. In line with the effect of Tβ4 on cellular stiffness and prestress discussed previously, these results show that up to a concentration of 0.2 µM, Tβ4 plays a role in actively influencing the acto-myosin contraction in SW480 cells.

4 Discussion

4.1.2 Tβ4 regulation of acto-myosin contractility is involved in the biphasic response of SW480 cell morphology and motility Maximum levels of cellular stiffness and contractility of SW480 cells were reached

using a Tβ4 concentration of 0.2 µM. The use of Tβ4 concentrations higher than 0.2 µM

causes a reduction in cell stiffness; SW480 cells treated overnight with 1 µM Tβ4 did not show a significant difference in stiffness or cytoskeletal prestress to those treated with the control. The same phenomenon was observed in the traction microscopy

assays using increasing amounts of Tβ4 for the stimulation of SW480 cells. Cells

treated with 1.0 µM Tβ4 showed a reduction in strain energy per cell when compared

with cells treated with 0.2 µM Tβ4. As discussed previously, no difference in bead detachment and cellular yielding to externally induced distortion was recorded across

all force steps at all concentrations of Tβ4. These results indicate that the biphasic

response to Tβ4 seen in SW480 cell stiffness and tractions is due to its active biphasic regulation of acto-myosin contractility. At this point so far, the assays indicate that the

biphasic cellular response of Tβ4 stimulation does not affect the mechano-coupling processes of force transmission mediated by the focal adhesions.

The cell interacts with its environment by applying tractions through the integrins to adhere to the extracellular substrate or matrix [123]. In a feedback loop, the tension from the cell is resisted by the extracellular matrix and results in an out-in signaling, which triggers cellular responses that include cell motility, differentiation or growth. The cycle of acto-myosin contractility is one of the major processes governing the tension within the cell. As discussed previously, acto-myosin interaction is promoted through the phosphorylation of the regulatory myosin light chain (MLC) [124-125] by MLCK (myosin light chain kinase). The activity of MLCK is controlled by the activity of ROCK and RhoA. Through the action of ROCK, MLC is not dephosphorylated and, thereby, remains in an active state.

The extracellularly applied Tβ4 forms a complex with PINCH to activate ILK, a focal adhesion scaffolding protein that interacts directly with cell surface integrins and that

is known to be activated by Tβ4 [40]. Here, ILK plays a dual role; it acts as a Ca2+ independent myosin light chain kinase [103] as well as a RhoA regulator [102].

Initially, at lower levels of Tβ4, ILK promotes the phosphorylation of MLC, resulting in increased acto-myosin contractility as confirmed in the magnetic tweezer and

tractions microscopy experiments; up to a working concentration of 0.2 µM Tβ4 cells

become stiffer and more contractile. At concentrations of Tβ4 exceeding 0.2 µM, however, the action of ILK as a RhoA regulator may inhibit the action of ROCK. Through the dephosphorylation and inactivation of MLC, acto-myosin contraction

decreases, as seen in assays with 1.0 µM Tβ4 induction of SW480 cells. In this effect,

Discussion 4

the interaction of Tβ4 and ILK becomes essentially a cellular sensor for outside-in signaling governing cell contractility.

The consequences of the biphasic regulation of acto-myosin contractility are manifested directly in the morphology and motility of SW480 cells. On 2D surfaces, cells treated with 0.2 µM Tβ4 were observed to adopt a more polarized form with a distinct formation of a leading edge and trailing edge. Past literature gives evidence that point to a pivotal role for myosin II activity in maintaining cell polarity [87]. The ability of cells to move on 2D surfaces [27, 99] is influenced by the adhesion strength, adhesion dynamics and cytoskeletal remodeling. In this effect, a cycle of detachment and adhesion to the extracellular substrate occurs in a migrating cell [87]. The migration speed is inversely correlated with the adhesion strength of the cell to the extracellular substrate. Cells which are highly adherent have typically large integrin clusters and low migration speeds. This phenomenon was observed in 2D cell motility assays of SW480 cells. Although cells treated with 0.2 µM Tβ4 were seen to move slower (lower apparent diffusivity values, D, in µm2 per second), they showed a high persistence in movement (power-law coefficient value). With an extracellular induction of 1.0 µM Tβ4, lower levels of tractions and cell migration persistence were recorded in SW480 cells. Thus, the effect of Tβ4 at a maximum working concentration of 0.2 µM on 2D cell migration is the combination of increased acto-myosin induced contractility and the persistence of cell movement.

Compared to movement on 2D substrates, a three dimensional extracellular matrix provides a physical obstruction to cell movement. According to Zaman et al. [126- 127], four biophysical processes influence the speed of cell invasion in 3D environments: 1) contractile forces are needed to overcome the steric hindrance of the extracellular matrix, 2) these forces are transmitted to the surrounding matrix through cell surface adhesions (integrins) and must be in a balance with detachment from the matrix to allow the cell to move itself forward. 3) As the cell moves through the matrix, it undergoes massive changes in cell morphology; the cell’s intrinsic force resisting the change in shape must be small otherwise a dynamic remodeling of the cytoskeleton must accommodate for this change, 4) The physical obstruction in form of elastic forces provided by the matrix must be low enough for the cell to squeeze through otherwise the cell must degrade the ECM through the production of matrix proteinases [27].

Again, a distinctive biphasic response in three dimensional cell morphology and migration of SW480 cells in collagen gels was also observed with increasing concentrations of Tβ4 induction. Previous studies have shown than cells overexpressing Tβ4 have increased levels of matrix metalloproteinase-7 (MMP-7) [48] coupled with lower levels of E-cadherin causing the disruption of cell-cell contacts. 79

4 Discussion

This phenomenon has been correlated with the increase in ILK activity [62]. The combination of all these biochemical responses leads to the degradation of the ECM and the epithelial-mesenchymal transition of the originally non-malignant SW480 cells, enabling a single cell to break away from a non-motile cell colony and invade through the ECM.

Although levels of MMP-7 and E-cadherin were not measured in SW480 cells induced

with extracellular Tβ4 in the assays performed, changes in cell shape as seen in the

overexpression of Tβ4 in SW480 cells was observed in 3D collagen gels; cells treated

with 0.2 µM Tβ4 developed a mesenchymal morphology with a fibroblastic appearance and long extended filapodia. Here, it can also be assumed that up to a concentration of 0.2 µM, the effect of acto-myosin contractility increases, as does the decrease in E-cadherin associated with the altered shape of the cells to a more fibroblastic morphology. These effects combined with the production matrix proteinases have enabled cells to move furthest within a 3D collagen matrix as shown in the invasion assays with an induction of 0.2 µM.

With a treatment of 1.0 µM Tβ4, however, cells show a mixed morphology of round cells in clusters and differentiated fibroblastic-like cells. The reduction in invasivity of the SW480 cells is in part due to the lower levels contractility of these cells as a result of RhoA inhibition. At this point, it is not known whether the levels of MMP-7 or E-

cadherin expression due to Tβ4 induction have contributed to this effect. For more conclusive evidence, this must be looked into further.

The inhibition of cell contractility at higher levels of Tβ4 induction (at 1.0 µM working concentration) was bypassed using calyculin-A in 3D cell invasion assays. As a MLC phosphatase inhibitor, calyculin-A sustained phosphorylation of MLC and, thereby, maintained the contractility of SW480 cells. Use of calyculin-A (10 nM and 50 nM) and

Tβ4 of up to 1 µM together produced a progressive increase in 3D cell invasivity. No

biphasic response with the increase in Tβ4 working concentration in cell treatment

was recorded. In 3D invasion assays with calyculin-A and Tβ4 induction of 0.2 µM, the

recorded SW480 cell invasivity was lower than that of with using 0.2 µM Tβ4 alone. Taking these results together, it is shown that sustaining overall cell contractility alone will not account for the increase in cell invasion. It can be concluded here that the balance in cell adhesion, contractility and dynamic remodeling of the cytoskeleton is essential for the motility and, finally, the invasivity of a cell. Shifts in this systematic balance will have a dramatic effect on cell migration. In sum, in addition to the

regulation of MLC phosphorylation through a putative Tβ4-PINCH-ILK pathway, it may be also the influence of that pathway on RhoA activity and the associated effects on

Discussion 4

cell polarity, adhesion behaviour and cytoskeletal dynamics that contribute to the pronounced biphasic response in cell invasiveness.

4 Discussion

4.2 Tβ4 and the cell lipid membrane: a novel mechanism for outside-in signaling The cellular plasma membrane is not only a biological barrier between the cell interior and the extracellular environment; it is involved in a myriad of cellular process including cell adhesion, biochemical signal transduction [128] and molecular transport. The cellular lipid membrane provides an anchor to the cytoskeleton, a scaffold for cytoskeletal and membrane associated proteins involved in outside-in and inside-out signal transduction. This section discusses the novel mechanism of Tβ4 lipid binding and its consequences in the signaling cascade and cellular responses.

Firstly, the binding of Tβ4 to synthetic lipid membranes in the form of small unilamellar lipid vesicles (SUVs) was recorded in DSC measurements using a mixture of DMPC/DMPG lipid vesicles. Increasing the overall negative charge of the lipids by increasing the DMPG lipid portion resulted in a slight increase in binding capacity of the peptide only at very high concentrations (175 µM). As a slightly negatively

charged protein with an isoelectric point of 5.1, Tβ4 will only bind weakly to neutral lipids. Consequently, by increasing the negative charge of the lipids, electrostatic

binding of Tβ4 to the membrane was not enhanced significantly. In DSC measurements, the change in specific heat is recorded over a large time span and

gives only a general indication of the binding capacity of Tβ4 to SUVs. Due to this, the kinetics of peptide-lipid binding cannot be resolved with this method. Furthermore, measurement inaccuracy due to the loss of peptide and lipids, which usually adsorb to the walls of assay containers and pipettes during the course of the experiment, cannot be avoided.

The binding of Tβ4 to synthetic lipid membranes was examined further and confirmed

with the FCS method using Alexa488 labeled Tβ4 and a PS/PC lipid vesicle mixture. As expected, increasing the mole fraction of acidic lipids (PS) and, thereby, allowing more

electrostatic interaction between the basic residues on the Tβ4 peptide and the lipid

vesicles resulted in the dramatic increase in Tβ4-lipid binding. The FCS technique has been used extensively to study intermolecular interactions of complimentary DNA oiligomers [78], protein-protein interactions between membranes, and the binding of myristoylated alanine-rich C kinase substrate (MARCKS) [107] to synthetic lipid

vesicles. Not only does FCS allow for the use of only minute quantities of Tβ4 peptide, single-molecule events of protein-lipid binding can be recorded. The main advantage of using this method is that it allows for the measurement at very short time scales, which enables the quantification of the protein-lipid binding kinetics.

While the basic structure of the cellular membrane is given by the lipid bilayers, its functions are carried out by membrane proteins. The number and types of proteins can differ vastly according to the specific function of the membrane and type of cell [85]. Membrane proteins are known to span or insert into a lipid bilayers through a

Discussion 4

single or multiple α-helices. Many of these proteins are amphipathic; meaning some regions of the protein are hydrophobic and some hydrophilic. Hydrophobic residues of the peptide can react with the hydrophobic tails of the phospholipid molecules located inside the lipid bilayers, whereas the hydrophilic residues lie in water either on the cytoplasmic side or the extracellular side of the membrane [85].

Figure 4.1 3D stucture of Tβ4. Shown is the structure of the Tβ4 peptide as generated by the molecular visualization software, PyMOL. In this conformation, the peptide is made up of two -helices, the first from residues 4 – 16 and the second from 30 – 40. 60 % of the peptide is made up of basic and hydrophobic residues; basic residues seen in blue, hydrophobic in red. α

NMR spectroscopic analysis of Tβ4 in solution have shown that the wild-type Tβ4 peptide is largely unstructured in water and are made up of two -helices in fluorinated alcohols [30]; the first helix extending from residues 4 – 16 and a second α helix from residues 30-40. Amino acid analysis of Tβ4 showed that clusters of basic and hydrophobic residues make up about 60% of the entire peptide (Figure 4.1). Among the 43 amino-acids making up the entire peptide, 9 residues are basic and 1 is aromatic [108]. These characteristics combined, make Tβ4 a good candidate for membrane binding. Similar modes of membrane association have been recorded for other proteins such as MARCKS [107] and HIV-1 [109]. The insertion or translocation of proteins, in this case Tβ4, either from the extracellular environment to the cell plasma membrane will not only increase the concentration of the peptide at this location in the membrane up to a thousand fold [107], it will reduce significantly the diffusion time so as to increase the reaction probability with other membrane- associated signalling proteins, such as ILK and PINCH. Thus, the mechanism of Tβ4 binding to the lipid membrane would explain the phenomenon of how induction with minute concentrations of Tβ4 can elicit a dramatic cellular response.

4 Discussion

4.3 The Tβ4-ILK Molecular Switch

In this section, a putative model for the biphasic response of Tβ4 on cellular response on a global scale is elucidated. A direct correlation will be made from the biophysical results measured and observed in the course of this work with past biochemical experiments done in vitro and in vivo. Finally, future implications of this work in clinical therapeutic application are discussed.

Previous studies have associated Tβ4 with increased cellular motility and invasivity.

The genetic over-expression of Tβ4 caused the epithelial-mesenchymal transition and the subsequent increase in invasivity of SW480 colorectal carcinoma cells, an otherwise non-malignant cell line [47-48, 62]. In embryonal cardiomyocytes,

extracellular treatment with Tβ4 induced increased migration in cardiac outflow tract explants [40, 129]. Both phenomena have been associated with the activity of ILK. In previous work, however, no distinction has been made between the effects of

extracellular induction and genetic overexpression of Tβ4 cellular responses. The

dose-dependent effect of Tβ4 application and overexpression on cell motility has yet to be quantified.

The results of this work have demonstrated the direct biphasic cellular response of SW480 extracellular induction in cell stiffness, contractility and invasivity. A novel

mechanism of the Tβ4 binding to the cell membrane has also been shown. Close

analysis of the recorded cellular biophysical responses and Tβ4 –membrane binding in this work combined with extensive knowledge of the biochemical pathways involved

with Tβ4 expression, extracellular stimulation and its interaction with ILK retrieved from past literature have contributed to the following model.

Discussion 4

4 Discussion

Figure 4.2 The regulation of SW480 cellular mechanics in response to extracellular Tβ4 concentration – a biphasic molecular switch mechanism. A schematic diagram of the biphasic cellular responses in stiffness, spreading, tractions and motility due to increasing levels of exogenous Tβ4 stimulation; at low extracellular concentrations, Tβ4 activation of ILK results in the phosphorylation of MLC, acto-myosin contractility and consequent increase in cell stiffness and contractility leading to increased motility. At concentrations above 0.2 µM, Tβ4 and ILK activity leads to the inhibition of RhoA and a decrease in MLC phosphorylation.

It is known that Tβ4 circulates in serum and wound fluid; its accumulation is assumed to be a product of cellular secretion or its release from damaged cells [34]. Near the

wound, a concentration gradient is present with the highest concentrations of Tβ4 directly at the wound site (see Figure 4.2 (B)). As passively circulating cells approach

the wound, the cells responsive to Tβ4 stimulation such as SW480 cells, ‘sense’ the Tβ4

concentration gradient. Tβ4 starts to bind to the cell membrane and then interacts

with ILK, a membrane associated protein. Local Tβ4 concentrations at the membrane

are amplified, thus, even minute amounts of extracellular Tβ4 can trigger a response.

At lower Tβ4 concentrations, Tβ4 activates ILK which acts as a Ca2+ independent MLC kinase, increasing MLC phosphorylation and, consequently, causing acto-myosin mediated cell contractility. As a result, the cell moves persistently in the direction of

the concentration gradient. Above a threshold level of Tβ4 in the extracellular environment, the further activation of ILK results in the negative regulation of RhoA (see Figure 4.2 (C)), leading gradually to the ablation of cell contractility and invasivity as it reaches the site of the wound. In the wound environment, the cell generates enough tractions to adhere at the site [87] without further movement.

The role of Tβ4 as a molecular switch in cell motility lies, firstly, in its role as a membrane-binding peptide and, secondly, in its interaction with ILK as a molecular

switch. The extent of the cellular response to Tβ4 concentrations in the extracellular environment may depend on the individual characteristics of the membranes in different cell lines such as the fraction of acidic phospholipids and the overall electrostatic charge of the membrane. Thus, it may be assumed that not all cells will respond in the concentration gradient as shown in SW480 colorectal carcinoma cells. The complicated dual role of ILK in regulating cell contractility by, on one hand, negatively regulating RhoA [102, 130] and, on the other hand, phosphorylating MLC as a Ca2+ independent MLC kinase has already been documented extensively in the past

[131]. However, by elucidating the biphasic cellular response of Tβ4, the two opposing

pathways are reconciled in the function of Tβ4 as a molecular switch in regulating SW480 cell motility.

Discussion 4

As demonstrated in this work, it is clear, that several aspects must still be considered in developing Tβ4 as a potential therapeutic agent. Firstly, studies involving the topical or extracellular application of Tβ4 should be separated from those involving the overexpression of the peptide. Secondly, the overall consequences of Tβ4 application must be considered in the mitigation of each pathological condition. For example, in the course of accelerating wound healing, concerns must be raised with regard to the role Tβ4 plays in tumor metastases and invasivity. Until now, no in vitro or in situ experimental data has been recorded on the effect of Tβ4 application on existing tumors. Finally, the sensitivity of individual cell types to Tβ4 concentrations may vary greatly due to the unique membrane characteristics distinguishing one cell type from the other.

Bibliography

1. Li, S., J.-L. Guan, and S. Chien, Biochemistry and biomechanics of cell motility. Annu Rev Biomed Eng, 2005. 7: p. 105-150. 2. Chen, H., B.W. Bernstein, and J.R. Bamburg, Regulating actin-filament dynamics in vivo. Trends Biochem Sci, 2000. 25(1): p. 19-23. 3. Ridley, A.J., Rho GTPases and cell migration. J Cell Sci, 2001. 114(Pt 15): p. 2713-22. 4. Raftopoulou, M. and A. Hall, Cell migration: Rho GTPases lead the way. Dev Biol, 2004. 265(1): p. 23-32. 5. Machesky, L.M. and K.L. Gould, The Arp2/3 complex: a multifunctional actin organizer. Curr Opin Cell Biol, 1999. 11(1): p. 117-21. 6. Ressad, F., et al., Control of actin filament length and turnover by actin depolymerizing factor (ADF/cofilin) in the presence of capping proteins and ARP2/3 complex. J Biol Chem, 1999. 274(30): p. 20970-6. 7. Nishida, E., Opposite effects of cofilin and profilin from porcine brain on rate of exchange of actin-bound adenosine 5'-triphosphate. Biochemistry, 1985. 24(5): p. 1160-4. 8. Didry, D., M.F. Carlier, and D. Pantaloni, Synergy between actin depolymerizing factor/cofilin and profilin in increasing actin filament turnover. J Biol Chem, 1998. 273(40): p. 25602-11. 9. Sun, H.Q., K. Kwiatkowska, and H.L. Yin, Actin monomer binding proteins. Curr Opin Cell Biol, 1995. 7(1): p. 102-10. 10. Sun, H.Q. and H.L. Yin, The beta-thymosin enigma. Ann N Y Acad Sci, 2007. 1112: p. 45-55. 11. Carlier, M.F., et al., Modulation of the interaction between G-actin and thymosin beta 4 by the ATP/ADP ratio: possible implication in the regulation of actin dynamics. Proc Natl Acad Sci U S A, 1993. 90(11): p. 5034-8. 12. Keely, P., L. Parise, and R. Juliano, Integrins and GTPases in tumour cell growth, motility and invasion. Trends Cell Biol, 1998. 8(3): p. 101-6. 13. Schwartz, M.A. and S.J. Shattil, Signaling networks linking integrins and rho family GTPases. Trends Biochem Sci, 2000. 25(8): p. 388-91. 14. Cary, L.A., D.C. Han, and J.L. Guan, Integrin-mediated signal transduction pathways. Histol Histopathol, 1999. 14(3): p. 1001-9. 15. Wu, C. and S. Dedhar, Integrin-linked kinase (ILK) and its interactors: a new paradigm for the coupling of extracellular matrix to actin cytoskeleton and signaling complexes. J Cell Biol, 2001. 155(4): p. 505-10. 16. Alenghat, F.J., et al., Analysis of cell mechanics in single vinculin-deficient cells using a magnetic tweezer. Biochem Biophys Res Commun, 2000. 277(1): p. 93-9. 17. Kollmannsberger, P. and B. Fabry, High-force magnetic tweezers with force feedback for biological applications. Rev Sci Instrum, 2007. 78(11). 18. Mierke, C.T., The role of vinculin in the regulation of the mechanical properties of cells. Cell Biochem Biophys, 2009. 53(3): p. 115-126. 88

Bibliography

19. Mierke, C.T., et al., Mechano-coupling and regulation of contractility by the vinculin tail domain. Biophys J, 2008. 94(2): p. 661-670. 20. Fabry, B., et al., Focal adhesion kinase stabilizes the cytoskeleton. Biophys J, 2011. 101(9): p. 2131-2138. 21. Kasza, K.E., et al., Filamin A is essential for active cell stiffening but not passive stiffening under external force. Biophys J, 2009. 96(10): p. 4326-4335. 22. Pelham, R.J., Jr. and Y. Wang, High resolution detection of mechanical forces exerted by locomoting fibroblasts on the substrate. Mol Biol Cell, 1999. 10(4): p. 935-45. 23. Mierke, C.T., et al., Vinculin facilitates cell invasion into three-dimensional collagen matrices. J Biol Chem, 2010. 285(17): p. 13121-13130. 24. Legant, W.R., et al., Measurement of mechanical tractions exerted by cells in three- dimensional matrices. Nat Methods, 2010. 7(12): p. 969-71. 25. Koch, T.M., et al., 3D Traction forces in cancer cell invasion. PLoS One, 2012. 7(3): p. e33476. 26. Raupach, C., et al., Stress fluctuations and motion of cytoskeletal-bound markers. Phys Rev E Stat Nonlin Soft Matter Phys, 2007. 76(1 Pt 1). 27. Mierke, C.T., et al., Contractile forces in tumor cell migration. Eur J Cell Biol, 2008. 87(8-9): p. 669-676. 28. Huff, T., et al., beta-Thymosins, small acidic peptides with multiple functions. Int J Biochem Cell Biol, 2001. 33(3): p. 205-220. 29. Czisch, M., et al., Conformation of thymosin beta 4 in water determined by NMR spectroscopy. Eur J Biochem, 1993. 218(2): p. 335-44. 30. Zarbock, J., et al., Solution conformation of thymosin beta 4: a nuclear magnetic resonance and simulated annealing study. Biochemistry, 1990. 29(34): p. 7814-21. 31. Hannappel, E., et al., Thymosin beta 4: a ubiquitous peptide in rat and mouse tissues. Proc Natl Acad Sci U S A, 1982. 79(7): p. 2172-5. 32. Xu, G.J., et al., Synthesis of thymosin beta 4 by peritoneal macrophages and adherent spleen cells. Proc Natl Acad Sci U S A, 1982. 79(13): p. 4006-9. 33. Hannappel, E. and W. Leibold, Biosynthesis rates and content of thymosin beta 4 in cell lines. Arch Biochem Biophys, 1985. 240(1): p. 236-41. 34. Goldstein, A.L., E. Hannappel, and H.K. Kleinman, Thymosin beta4: actin-sequestering protein moonlights to repair injured tissues. Trends Mol Med, 2005. 11(9): p. 421-429. 35. Safer, D., R. Golla, and V.T. Nachmias, Isolation of a 5-kilodalton actin-sequestering peptide from human blood platelets. Proc Natl Acad Sci U S A, 1990. 87(7): p. 2536-40. 36. Safer, D., M. Elzinga, and V.T. Nachmias, Thymosin beta 4 and Fx, an actin-sequestering peptide, are indistinguishable. J Biol Chem, 1991. 266(7): p. 4029-32. 37. Safer, D., T.R. Sosnick, and M. Elzinga, Thymosin beta 4 binds actin in an extended conformation and contacts both the barbed and pointed ends. Biochemistry, 1997. 36(19): p. 5806-16. 38. Kabsch, W., et al., Atomic structure of the actin:DNase I complex. Nature, 1990. 347(6288): p. 37-44.

Bibliography

39. Frohm, M., et al., Biochemical and antibacterial analysis of human wound and blister fluid. Eur J Biochem, 1996. 237(1): p. 86-92. 40. Bock-Marquette, I., et al., Thymosin beta4 activates integrin-linked kinase and promotes cardiac cell migration, survival and cardiac repair. Nature, 2004. 432(7016): p. 466-472. 41. Malinda, K.M., et al., Thymosin beta4 accelerates wound healing. J Invest Dermatol, 1999. 113(3): p. 364-8. 42. Philp, D., et al., Thymosin beta 4 and a synthetic peptide containing its actin-binding domain promote dermal wound repair in db/db diabetic mice and in aged mice. Wound Repair Regen, 2003. 11(1): p. 19-24. 43. Philp, D., et al., Thymosin beta4 promotes matrix metalloproteinase expression during wound repair. J Cell Physiol, 2006. 208(1): p. 195-200. 44. Philp, D. and H.K. Kleinman, Animal studies with thymosin beta, a multifunctional tissue repair and regeneration peptide. Ann N Y Acad Sci, 2010. 1194: p. 81-86. 45. Sosne, G., et al., Thymosin beta 4 promotes corneal wound healing and modulates inflammatory mediators in vivo. Exp Eye Res, 2001. 72(5): p. 605-8. 46. Sosne, G., et al., Thymosin beta 4 promotes corneal wound healing and decreases inflammation in vivo following alkali injury. Exp Eye Res, 2002. 74(2): p. 293-9. 47. Wang, W.-S., et al., Overexpression of the thymosin beta-4 gene is associated with malignant progression of SW480 colon cancer cells. Oncogene, 2003. 22(21): p. 3297-3306. 48. Wang, W.-S., et al., Overexpression of the thymosin beta-4 gene is associated with increased invasion of SW480 colon carcinoma cells and the distant metastasis of human colorectal carcinoma. Oncogene, 2004. 23(39): p. 6666-6671. 49. Malinda, K.M., A.L. Goldstein, and H.K. Kleinman, Thymosin beta 4 stimulates directional migration of human umbilical vein endothelial cells. FASEB J, 1997. 11(6): p. 474-481. 50. Philp, D., et al., The actin binding site on thymosin beta4 promotes angiogenesis. FASEB J, 2003. 17(14): p. 2103-2105. 51. Parks, W.C., Matrix metalloproteinases in repair. Wound Repair Regen, 1999. 7(6): p. 423- 32. 52. Blain, E.J., D.J. Mason, and V.C. Duance, The effect of thymosin beta4 on articular cartilage chondrocyte matrix metalloproteinase expression. Biochem Soc Trans, 2002. 30(Pt 6): p. 879-82. 53. Shrivastava, S., et al., Thymosin beta4 and cardiac repair. Ann N Y Acad Sci, 2010. 1194: p. 87-96. 54. Srivastava, D., et al., Thymosin beta4 is cardioprotective after myocardial infarction. Ann N Y Acad Sci, 2007. 1112: p. 161-70. 55. Fan, Y., et al., Spatial coordination of actin polymerization and ILK-Akt2 activity during endothelial cell migration. Dev Cell, 2009. 16(5): p. 661-674. 56. Sosne, G., et al., Thymosin-beta4 modulates corneal matrix metalloproteinase levels and polymorphonuclear cell infiltration after alkali injury. Invest Ophthalmol Vis Sci, 2005. 46(7): p. 2388-95. 57. Woodhouse, E.C., R.F. Chuaqui, and L.A. Liotta, General mechanisms of metastasis. Cancer, 1997. 80(8 Suppl): p. 1529-37.

Bibliography

58. Liotta, L.A., P.S. Steeg, and W.G. Stetler-Stevenson, Cancer metastasis and angiogenesis: an imbalance of positive and negative regulation. Cell, 1991. 64(2): p. 327-36. 59. Clark, E.A., et al., Genomic analysis of metastasis reveals an essential role for RhoC. Nature, 2000. 406(6795): p. 532-5. 60. Kobayashi, T., et al., Thymosin-beta4 regulates motility and metastasis of malignant mouse fibrosarcoma cells. Am J Pathol, 2002. 160(3): p. 869-882. 61. Cha, H.-J., M.-J. Jeong, and H.K. Kleinman, Role of thymosin beta4 in tumor metastasis and angiogenesis. J Natl Cancer Inst, 2003. 95(22): p. 1674-1680. 62. Huang, H.C., et al., Thymosin beta4 triggers an epithelial-mesenchymal transition in colorectal carcinoma by upregulating integrin-linked kinase. Oncogene, 2006. 63. Hannappel, E. and M. van Kampen, Determination of thymosin beta 4 in human blood cells and serum. J Chromatogr, 1987. 397: p. 279-85. 64. Mannherz, H.G. and E. Hannappel, The beta-thymosins: intracellular and extracellular activities of a versatile actin binding protein family. Cell Motil Cytoskeleton, 2009. 66(10): p. 839- 851. 65. Huff, T., et al., Thymosin beta4 is released from human blood platelets and attached by factor XIIIa (transglutaminase) to fibrin and collagen. FASEB J, 2002. 16(7): p. 691-696. 66. Guarnera, G., A. DeRosa, and R. Camerini, The effect of thymosin treatment of venous ulcers. Ann N Y Acad Sci, 2010. 1194: p. 207-12. 67. Morris, D.C., et al., Thymosin beta4: a candidate for treatment of stroke? Ann N Y Acad Sci, 2010. 1194: p. 112-7. 68. Dunn, S.P., et al., Treatment of chronic nonhealing neurotrophic corneal epithelial defects with thymosin beta4. Ann N Y Acad Sci, 2010. 1194: p. 199-206. 69. Thymosins in Health and Disease. Proceedings of the 2nd International Symposium. September 30-October 2, 2009. Catania, Italy. Ann N Y Acad Sci, 2010. 1194: p. ix-xi, 1-229. 70. Guarnera, G., DeRosa A., Camerini, R. The Effect of Thymosin Beta 4 (TB4) Treatment of Venous Stasis Ulcers - Answers from a Well-Controlled European Clinical Trial. in Thymosins in Health and Disease. 2009 Catania, Italy. 71. Sambrook, J.a.R., D.W., ed. Molecular Cloning: A Laboratory Manual. 2001, Cold Spring Harbor Laboratory. 72. Huff, T., et al., Thymosin beta(4) serves as a glutaminyl substrate of transglutaminase. Labeling with fluorescent dansylcadaverine does not abolish interaction with G-actin. FEBS Lett, 1999. 464(1-2): p. 14-20. 73. Raupach, C., et al., Stress fluctuations and motion of cytoskeletal-bound markers. Phys Rev E Stat Nonlin Soft Matter Phys, 2007. 76(1 Pt 1): p. 011918. 74. Raupach, C., et al., Stress fluctuations and motion of cytoskeletal-bound markers. Phys Rev E Stat Nonlin Soft Matter Phys, 2007. 76 (1 Pt 1): p. 011918. 75. Butler, J.P., et al., Traction fields, moments, and strain energy that cells exert on their surroundings. Am J Physiol Cell Physiol, 2002. 282(3): p. C595-C605. 76. Kollmannsberger, P., Nonlinear microrheology of living cell, in Biophysics Group. 2009, FAU Erlangen-Nuremberg.

Bibliography

77. W.H. Goldmann, B.B., and T.P. Lele, Cytoskeletal proteins at the lipid membrane. in: T.H. T., and O.-L. A., in Planar lipid bilayers (BLM's) and their applications. 2007, Elsevier. p. 227-255. 78. Rieger, R., Röcker, C., Nienhaus, G., Fluctuation correlation spectroscopy for the advanced physics laboratory. American Journal of Physics, 2005. 73(12): p. 1129-1134. 79. Petrásek, Z. and P. Schwille, Precise measurement of diffusion coefficients using scanning fluorescence correlation spectroscopy. Biophys J, 2008. 94(4): p. 1437-1448. 80. Maulucci, G., et al., Particle size distribution in DMPC vesicles solutions undergoing different sonication times. Biophys J, 2005. 88(5): p. 3545-50. 81. Qiu, F.-Y., et al., Thymosin beta4 induces endothelial progenitor cell migration via PI3K/Akt/eNOS signal transduction pathway. J Cardiovasc Pharmacol, 2009. 53(3): p. 209-214. 82. Metzner, C., et al., Simple model of cytoskeletal fluctuations. Phys Rev E Stat Nonlin Soft Matter Phys, 2007. 76(2 Pt 1): p. 021925. 83. Thiery, J.P., Epithelial-mesenchymal transitions in tumour progression. Nat Rev Cancer, 2002. 2(6): p. 442-454. 84. Savagner, P., Leaving the neighborhood: molecular mechanisms involved during epithelial- mesenchymal transition. Bioessays, 2001. 23(10): p. 912-923. 85. Alberts, Molecular Biology of the Cell, 4th Edition. 4th Edition ed. 2002, New York: Garland Science. 86. Pollard, T.D. and G.G. Borisy, Cellular motility driven by assembly and disassembly of actin filaments. Cell, 2003. 112(4): p. 453-465. 87. Ridley, A.J., et al., Cell migration: integrating signals from front to back. Science, 2003. 302(5651): p. 1704-1709. 88. Parsons, J.T., A.R. Horwitz, and M.A. Schwartz, Cell adhesion: integrating cytoskeletal dynamics and cellular tension. Nat Rev Mol Cell Biol, 2010. 11(9): p. 633-643. 89. Fabry, B., et al., Scaling the microrheology of living cells. Phys Rev Lett, 2001. 87(14). 90. Lenormand, G., et al., Linearity and time-scale invariance of the creep function in living cells. J R Soc Interface, 2004. 1(1): p. 91-97. 91. Fernández, P. and A. Ott, Single cell mechanics: stress stiffening and kinematic hardening. Phys Rev Lett, 2008. 100(23). 92. Gardel, M.L., et al., Stress-dependent elasticity of composite actin networks as a model for cell behavior. Phys Rev Lett, 2006. 96(8). 93. Wang, N., et al., Cell prestress. I. Stiffness and prestress are closely associated in adherent contractile cells. Am J Physiol Cell Physiol, 2002. 282(3): p. C606-C616. 94. Vicente-Manzanares, M., et al., Non-muscle myosin II takes centre stage in cell adhesion and migration. Nat Rev Mol Cell Biol, 2009. 10(11): p. 778-790. 95. Burack, W.R. and T.W. Sturgill, The activating dual phosphorylation of MAPK by MEK is nonprocessive. Biochemistry, 1997. 36(20): p. 5929-33. 96. Sugiura, R., et al., The MAPK kinase Pek1 acts as a phosphorylation-dependent molecular switch. Nature, 1999. 399(6735): p. 479-83. 97. Kieran, M.W., et al., Concentration-dependent positive and negative regulation of a MAP kinase by a MAP kinase kinase. Oncogene, 1999. 18(48): p. 6647-57. 92

Bibliography

98. Zhang, S.Q., et al., Recruitment of the IKK signalosome to the p55 TNF receptor: RIP and A20 bind to NEMO (IKKgamma) upon receptor stimulation. Immunity, 2000. 12(3): p. 301-11. 99. Horwitz, A.R. and J.T. Parsons, Cell migration--movin' on. Science, 1999. 286(5442): p. 1102-3. 100. Huang, J., et al., Gi-coupled receptors mediate phosphorylation of CPI-17 and MLC20 via preferential activation of the PI3K/ILK pathway. Biochem J, 2006. 396(1): p. 193-200. 101. Khyrul, W.A.K.M., et al., The integrin-linked kinase regulates cell morphology and motility in a rho-associated kinase-dependent manner. J Biol Chem, 2004. 279(52): p. 54131-54139. 102. Kogata, N., et al., Integrin-linked kinase controls vascular wall formation by negatively regulating Rho/ROCK-mediated vascular smooth muscle cell contraction. Genes Dev, 2009. 23(19): p. 2278-2283. 103. Wilson, D.P., et al., Integrin-linked kinase is responsible for Ca2+-independent myosin diphosphorylation and contraction of vascular smooth muscle. Biochem J, 2005. 392(Pt 3): p. 641- 648. 104. Henson, J.H., et al., Actin-based centripetal flow: phosphatase inhibition by calyculin-A alters flow pattern, actin organization, and actomyosin distribution. Cell Motil Cytoskeleton, 2003. 56(4): p. 252-266. 105. Leopoldt, D., H.F. Yee, and E. Rozengurt, Calyculin-A induces focal adhesion assembly and tyrosine phosphorylation of p125(Fak), p130(Cas), and paxillin in Swiss 3T3 cells. J Cell Physiol, 2001. 188(1): p. 106-119. 106. Chartier, L., et al., Calyculin-A increases the level of protein phosphorylation and changes the shape of 3T3 fibroblasts. Cell Motil Cytoskeleton, 1991. 18(1): p. 26-40. 107. Rusu, L., et al., Fluorescence correlation spectroscopy studies of Peptide and protein binding to phospholipid vesicles. Biophys J, 2004. 87(2): p. 1044-1053. 108. Czisch, M., et al., Conformation of thymosin beta 4 in water determined by NMR spectroscopy. Eur J Biochem, 1993. 218(2): p. 335-344. 109. Gerlach, H., et al., HIV-1 Nef membrane association depends on charge, curvature, composition and sequence. Nat Chem Biol, 2010. 6(1): p. 46-53. 110. García-Sáez, A.J. and P. Schwille, Fluorescence correlation spectroscopy for the study of membrane dynamics and protein/lipid interactions. Methods, 2008. 111. Ries, J. and P. Schwille, New concepts for fluorescence correlation spectroscopy on membranes. Phys Chem Chem Phys, 2008. 10(24): p. 3487-3497. 112. Kholodenko, B.N., J.B. Hoek, and H.V. Westerhoff, Why cytoplasmic signalling proteins should be recruited to cell membranes. Trends Cell Biol, 2000. 10(5): p. 173-8. 113. McCloskey, M.A. and M.M. Poo, Rates of membrane-associated reactions: reduction of dimensionality revisited. J Cell Biol, 1986. 102(1): p. 88-96. 114. Huff, T., et al., beta-Thymosins, small acidic peptides with multiple functions. Int J Biochem Cell Biol, 2001. 33(3): p. 205-20. 115. Wang, N., et al., Mechanical behavior in living cells consistent with the tensegrity model. Proc Natl Acad Sci U S A, 2001. 98(14): p. 7765-7770.

Bibliography

116. Puig-de-Morales, M., et al., Cytoskeletal mechanics in adherent human airway smooth muscle cells: probe specificity and scaling of protein-protein dynamics. Am J Physiol Cell Physiol, 2004. 287(3): p. C643-54. 117. Laudadio, R.E., et al., Rat airway smooth muscle cell during actin modulation: rheology and glassy dynamics. Am J Physiol Cell Physiol, 2005. 289(6): p. C1388-95. 118. Bursac, P., et al., Cytoskeletal remodelling and slow dynamics in the living cell. Nat Mater, 2005. 4(7): p. 557-61. 119. Fabry, B., et al., Selected contribution: time course and heterogeneity of contractile responses in cultured human airway smooth muscle cells. J Appl Physiol, 2001. 91(2): p. 986-94. 120. Stamenovic, D., et al., Rheology of airway smooth muscle cells is associated with cytoskeletal contractile stress. J Appl Physiol, 2004. 96(5): p. 1600-5. 121. Nagayama, M., et al., Contribution of cellular contractility to spatial and temporal variations in cellular stiffness. Exp Cell Res, 2004. 300(2): p. 396-405. 122. Ridley, A.J., Life at the leading edge. Cell, 2011. 145(7): p. 1012-1022. 123. Aikawa, R., et al., Rho family small G proteins play critical roles in mechanical stress- induced hypertrophic responses in cardiac myocytes. Circ Res, 1999. 84(4): p. 458-66. 124. Somlyo, A.P. and A.V. Somlyo, Signal transduction and regulation in smooth muscle. Nature, 1994. 372(6503): p. 231-6. 125. Gallagher, P.J., B.P. Herring, and J.T. Stull, Myosin light chain kinases. J Muscle Res Cell Motil, 1997. 18(1): p. 1-16. 126. Zaman, M.H., et al., Computational model for cell migration in three-dimensional matrices. Biophys J, 2005. 89(2): p. 1389-97. 127. Zaman, M.H., et al., Migration of tumor cells in 3D matrices is governed by matrix stiffness along with cell-matrix adhesion and proteolysis. Proc Natl Acad Sci U S A, 2006. 103(29): p. 10889- 94. 128. DiNitto, J.P., T.C. Cronin, and D.G. Lambright, Membrane recognition and targeting by lipid- binding domains. Sci STKE, 2003. 2003(213): p. re16. 129. Bock-Marquette, I., et al., Thymosin beta4 mediated PKC activation is essential to initiate the embryonic coronary developmental program and epicardial progenitor cell activation in adult mice in vivo. J Mol Cell Cardiol, 2009. 46(5): p. 728-38. 130. Yamazaki, T., et al., EphA1 interacts with integrin-linked kinase and regulates cell morphology and motility. J Cell Sci, 2009. 122(Pt 2): p. 243-255. 131. Montanez, E., et al., Alpha-parvin controls vascular mural cell recruitment to vessel wall by regulating RhoA/ROCK signalling. EMBO J, 2009. 28(20): p. 3132-3144.

Acknowledgements

First and foremost, I would like to thank my supervisor, Prof. Ben Fabry, for his guidance and support. Through Prof. Fabry, I was able to build a network of contacts and participate in numerous international conferences to pursue further my scientific endeavors. Many thanks also to Prof. Wolfgang Goldmann for the insightful comments and constructive criticism.

I would like to express my gratitude to Prof. Ewald Hannappel, who introduced me to the world of Thymosins and, in particular, Thymosin beta 4. His profound knowledge in this field had singularly inspired me to begin my work with Tβ4. The motivation and feedback he provided as well as the constant supply of the potent peptide were all key ingredients to the success of this study. A special acknowledgement here to Christine App, Prof. Hannappel’s PhD student, for producing the much coveted fluorescently labeled Tβ4 and all her help throughout my dissertation.

For funding my research and making this study possible, I would like to thank the International Max-Planck Research School Physics of Light and their staff.

Thanks to all my colleagues during my stay at the institute: Carina, Phillip, Daniel, Claudia, Martina, Navid, Nadine, Johannes, Anna and Thorsten not only for motivating me but also for supporting my strengths and enduring my weaknesses. I would like to give special credit to Nadine for her direct contribution to the FCS setup and measurements. Very special thanks to my roommates, Anna and Thorsten, for their constant and everlasting friendship and good conversation, scientific and otherwise, and to Johannes, our auxiliary member, for showing me the lighter side of physics. Thank you to Prof. Ana Smith for her inexhaustible enthusiasm, wisdom and optimism.

Last but never least; I would like to express my deepest gratitude to all my family members for their unwavering support, encouragement and patience throughout the years leading to the completion of this dissertation. This work is dedicated to my daughters, Sanna and Ellen, my constant source of strength and inspiration.

Publications/Conferences

Lorenz C, Brunner JG, Kollmannsberger P, Jaafar L, Fabry B, Virtanen S. Effect of surface pre-treatments on biocompatibility of magnesium. Acta Biomater. 2009 Sep;5(7):2783-9. doi: 10.1016/j.actbio.2009.04.018. Epub 2009 May 4

Jaafar L, App C, Lang N, Hannappel E, and Fabry B The Effect of Extracellular Thymosin β4 on Cell Mechanics and Motility PLoS One, 2014 Jan, submitted

Cell Biomechanics Meeting, Paris (Sept. 2005) Invited talk Title : The Role of Vinculin in Cellular Mechanotransduction

ASCB Annual Meeting, San Francisco (Dec. 2005) Poster Session Title : Role of Vinculin in Cytoskeletal Dynamics and Regulation

Cell Biomechanics Meeting, Barcelona (Sept. 2007) Invited talk Title :

BiophysicalEffects Society of Thymosin 50th Annual β4 on Meeting, Cytoskeletal Salt LakeOrganization City (Feb. 2006) Poster Session Title : Vinculin Regulates Cytoskeletal Dynamics and Prestress in the Cell

Thymosins in Health and Disease (Oct.. 2009) Poster Session Title :

Effects of Thymosin β4 on Cytoskeletal Organization

Curriculum Vitae

Personal information

Birth place and date: Kuala Lumpur, Malaysia, 09.07.1975

Citizenship: Malaysisch

Address: Max-Busch-Str. 22 91054 Erlangen [email protected]; [email protected]

Tertiary Education

2006- present Doctoral studies in Biophysics Biophysics Group (Prof. B. Fabry) FAU Erlangen-Nuremberg, Germany

2005 – 2006 Diploma thesis in Molecular Biology Department of Microbiology (Prof. M. Niederweiß) FAU Erlangen- Nuremberg, Germany

2000 –2005 Undergraduate studies in Biology (Diploma) FAU Erlangen- Nuremberg, Germany

1999 –2000 German language course for the Deutschen Sprachprüfung für den Hochschulzugang (DSH)

1994 –1997 Undergraduate studies in Medicine (MBBS) University of New South Wales, Sydney, Australia

Primary/Secondary Education

1993 –1994 UNSW Foundation Certificate University of New South Wales Foundation Course Kuala Lumpur, Malaysia

1987 –1992 Malaysian High School Certificate Sri Aman Secondary School Kuala Lumpur, Malaysia (1987-1992)

1982 –1986 Elementary school Louise Archer Elementary School Vienna, Virginia, USA

Awards

2006 – 2010 Stipend for doctoral studies International Max-Planck Research School for Physics of Light

Job Experience

2013 - present Project manager R&D/Product Development, Containment Systems AREVA GmbH

2010 - 2013 Computational Engineer Radiation Protection & Neutron Fluence AREVA GmbH

1999 - 2000 Software User-Interface and Testing for Syngo Siemens Medical Solutions Siemens AG