role of flightless i in cell migration · monomers and mitochondria, but partially with paxillin....
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ROLE OF FLIGHTLESS I IN CELL MIGRATION
by
Ibrahim Mohammad
A thesis submitted in conformity with the requirements for the degree of Master of Science Graduate Department of Dentistry
University of Toronto
© Copyright by Ibrahim Mohammad 2010
ii
Role of Flightless I in Cell Migration
Ibrahim Mohammad
Master of Science
Graduate Department of Dentistry University of Toronto
2010
Abstract
A central process in connective tissue homeostasis is cell migration, which involves dynamic
interactions between focal adhesions, the actin cytoskeleton and mitochondria, but the role of
focal adhesion proteins in cell migration is not wholly defined. We examined focal adhesion-
associated proteins from mouse fibroblasts and identified Flightless I (FliI) as a potential focal
adhesion protein. We determined that FliI is distributed in the cytosol and co-localizes with actin
monomers and mitochondria, but partially with paxillin. Biochemical assays showed that FliI
associates with both actin monomers and short oligomers/filaments. Migration assay determined
that cells with reduced FliI expression migrated more quickly and that FliI knockdown inhibited
activation of β1 integrins. Consistent with these data, cell adhesion assay demonstrated that FliI
knockdown cells were less adherent than wildtype cells. Our findings indicate that FliI may
regulate cell migration by interacting with the actin monomers and the mitochondria to affect cell
adhesion.
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Acknowledgments I have been blessed for having an opportunity to complete a master’s degree with the Matrix
Dynamics Group at the Faculty of Dentistry in University of Toronto. There are many people
who I would like to thank because of their dedication and assistance towards the completion of
my degree.
I would like to thank my supervisor Dr. C.A. McCulloch for giving me the opportunity to create,
develop, and embellish my scientific skills in his laboratory. He has provided extraordinary
amount of guidance throughout my research. I have found my weekly meetings with him very
insightful. His office door has always been open for me and my colleagues to approach him
openly about any subject. Dr. McCulloch sincerely cares about others people’s career goals. He
has been very supportive of my desire wanting to become a dentist in the future.
I also would like to thank my committee members Dr. M. Manolson and Dr. C. Simmons for
taking the time out of their busy schedules to sit on the advisory committee. They have provided
valuable advice and direction towards my project.
I would like to thank Pam Arora for teaching me the fundamentals of scientific experiments.
Over the last two years Pam has on more than several occasions taken time out of her personal
schedule to assist me with the experiments. She has encouraged me to think outside the box, and
has always reminded me to keep a positive attitude no matter what the outcome of the
experiments is. Her guidance towards my education and life has been invaluable.
I would like to thank my colleagues in our laboratory who have made my experience in the
laboratory exhilarating. I would like to thank Carol Laschinger, Wilson Lee, Dhaarmini
Rajshankar, Reza Termei, Hugh Kim, Mathew Chan, Anne Koehler, and Cheung Lo.
Last but not least, I would like to thank Jeff Li, a summer student from University of
Pennsylvania School of Dental Medicine, and Rhodaba Ebaday, a volunteer student from
University of Toronto, both of whom who provided valuable assistance towards my research.
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Table of Contents Abstract ii
Acknowledgements iii
Table of Contents iv
List of Tables vii
List of Figures viii
Chapter 1 - Literature Review 1
1. Connective Tissue Homeostasis 1
1.1. What are Connective Tissues? 1
1.2. Homeostatic Systems in Matrix Biology 2
1.3. Pathologies associated with connective tissue homeostasis 4
1.4. Cell function in matrix remodeling 4
2. Cell Migration 5
2.1. Mechanics of cell migration 5
2.2. Cell Adhesions 7
2.2.1. Families of matrix receptors 7
2.2.2. Integrins 8
2.2.3. Focal adhesions 10
2.3. Actin 12
2.3.1. Control of actin assembly 12
2.3.2. Actin binding proteins 14
v
2.3.3. Role in leading edge protrusion 15
2.4. Endoplasmic Reticulum 16
2.4.1. Structure and function 16
2.4.2. Ca2+ homeostasis and cell signaling 17
2.4.3. ER-focal adhesion linkage 18
2.5. Mitochondria 19
2.5.1. Role of actin in mitochondrial function 19
2.5.2. Functions in cell motility and actin assembly 21
3. Flightless I 23
3.1. Discovery 23
3.2. Structure and Function of Gelsolin Family proteins 25
3.3. Structure and Function of Flightless I 25
3.4. Flightless I in Cell Migration 27
Statement of the Problem 28
Introduction 30
Materials and Methods 33
Results 39
Discussion 45
Figure Legends 50
Figures 54
Tables 62
Future Directions 66
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Conclusion 69
References 70
vii
List of Tables Table 1: Proteins identified by mass spectrometry from bead-associated fractions 62
Table 2: Proteins identified by mass spectrometry from FliI immunoprecipitates 62
viii
List of Figures Figure 1A: Immunoblot of Bead-Associated Proteins 54
Figure 1B: TIRF Images of GFP-Paxillin and Flightless I 54
Figure 1C: Confocal Images of GFP-Paxillin and Flightless I 54
Future 1D: Sectional Images of Flightless I 54
Figure 2A: Knockdown of Flightless I with siRNA 55
Figure 2B: In-Vitro Scratch Assay 55
Figure 2C: Graph of Migration Rate 55
Figure 2D: Quantification of Flightless I fluorescence from migratory and non-
migratory cells 55
Figure 3A: Graph of Jet Wash Assay 56
Figure 3B: Quantification of 9EG7 fluorescence from migratory and non-
migratory cells 56
Figure 3C: FRAP comparisons of wildtype and Flightless I knockdown cells 56
Figure 4A: Flightless I immunoprecipitates 57
Figure 4B: Differential centrifugation of actin 57
Figure 4C: Confocal images of actin and Flightless I 57
Figure 4D: Quantification of phalloidin fluorescence 58
Figure 4E: Treatment of cells with jasplakinolide and latrunculin 58
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Figure 5A: Mitochondrial and endoplasmic reticulum fractions 59
Figure 5B: Confocal images of Flightless I and KDEL 59
Figure 5C: Confocal images of Flightless I and Mito-tracker Red 59
Figure 5D: TIRF images of GFP-Paxillin and Mito-tracker Red 59
Figure 5E: Confocal images of actin monomer and Mito-tracker Red 60
Figure 6: Proposed mechanism of Flightless I in the organization of actin filaments 61
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Chapter 1 Literature Review
1 Connective Tissue Homeostasis
1.1 What are Connective Tissues?
Mammalian connective tissues provide structural support and metabolic needs for other tissues
and organs throughout the body (Heath and Young, 2000). Soft connective tissues are comprised
of the extracellular matrix, which forms a supporting structural and informational framework
(Alberts et al, 2002), and cells, which include fibroblasts, myofibroblasts, adipocytes, mast cells,
macrophages, and leukocytes (Alberts et al, 2002). In mineralizing connective tissues,
specialized cell types including odontoblasts, chondrocytes, osteoblasts and osteoclasts,
contribute to the formation and remodeling of these tissues.
The extracellular matrix, which is the predominant component of connective tissues, plays a
wide variety of important roles such as providing structural support and attachment for tissues
and organs, and regulating cell survival, development, migration, proliferation, shape and
function of cells (Alberts et al, 2002). The matrix is composed of a large array of proteins
(collagen, elastin, fibronectin, and laminin), glycosaminoglycans and proteoglycans. Variations
in the relative amounts and composition of matrix macromolecules contribute to the diversity of
connective tissue form and function. For example, bones and teeth are comprised of calcified
matrices, which exhibit high levels of compressive and tensile strength whereas the transparent
matrix of the cornea enables vision (Alberts et al, 2002).
2
Collagens are the major proteins of the extracellular matrix and are the most abundant proteins of
mammals (Perez-Tamayo, 1978), comprising 25% of total protein mass. There are 20 types of
collagen molecules. The major collagen types in mammalian connective tissues include types I,
II, III, V, and XI. Another important matrix glycoprotein is fibronectin, which plays a central
role in cell attachment to the matrix and in guiding cell migration. Connective tissues serve many
biological roles, and as a result undergo extensive remodeling in development, wound healing
and aging. Remodeling of healthy tissues is tightly regulated by a large array of homeostatic
mechanisms that ensure proper form and function.
1.2 Homeostatic Systems in Matrix Biology
Homeostasis of connective tissues in mammals is determined by the balanced synthesis,
degradation and remodeling of extracellular matrices. Several different types of mesenchymal
cells synthesize and degrade matrix components. Matrix synthesis is performed by resident cells,
which include fibroblasts, osteoblasts, chondrocytes and odontoblasts. These specialized cells
produce different components of the matrix that are required for growth, wound healing and the
development of fibrotic lesions (Grant and Prockop, 1972; Ross, 1975). The amount and type of
matrix proteins vary between tissues, in different stages of development, and in different
pathological circumstances (Grant and Prockop, 1972). In gingiva and periodontal ligament, the
synthesis and degradation of collagen are extraordinarily rapid compared to other connective
tissues (Sodek, 1977) and are carefully balanced to maintain oral mucosal health (McCulloch,
2004). The appropriate and timely breakdown of the extracellular matrix is essential for
appropriate tissue remodeling (Nagase et al, 1997).
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There are several matrix degradation pathways, which include the matrix metalloproteinase
(MMP), plasmin-dependent, polymorphonuclear serine proteinase, phagocytic and osteoclastic
pathways (Hansen et al, 1993). The MMP pathway involves zinc-dependent endopeptidases,
which exhibit catalytic activity against most extracellular matrix macromolecules (Woessner,
1991). Cells use these proteases to degrade matrix molecules and certain types of signaling
molecules to effect matrix remodeling. Most connective tissues normally exhibit low levels of
matrix protease activity, which is controlled in part by inhibitors such as the tissue inhibitors of
metalloproteinases (Woessner, 1991; Nagase and Woessner 1999; Berrier and Yamada, 2007).
The plasmin-dependent pathway plays an important role in the remodeling of the extracellular
matrix wherein activated plasmin cleaves Lys-Arg peptide bonds exposed on the surface of a
wide range of native proteins, such as fibrin and fibronectin (Hansen et al, 1993). The neutrophil
pathway mediates degradation of extracellular matrix macromolecules by release of serine
proteinases, elastase and cathepsin G (Hansen et al, 1993). These proteinases can cleave a variety
of extracellular matrix proteins such as collagen (type IV), laminin, fibronectin, and
proteoglycans (Hansen et al, 1993). The fibroblast phagocytic pathway is involved in
degradation of extracellular matrix by the process of internalizing collagen fibrils and degrading
them in lysosomes (Cate and Syrbu, 1974; Arora et al, 2001). Collagen degradation by the
phagocytic pathway is prominent in tissues with rapid collagen turnover such as the periodontal
ligament and the uterus (Harkness and Moralee, 1956; Parakkal; 1969; Melcher and Chan, 1981).
Fine balancing of synthesis and degradation pathways of extracellular matrix is essential for
maintenance of normal tissue structure; small shifts that favor synthesis or degradation can lead
to several, clinically important and high prevalence diseases.
4
1.3 Pathologies associated with connective tissue homeostasis
In many inflammatory diseases there is a shift towards increased matrix protease activity (e.g.
collagenase), which is seen for example in patients with rheumatoid arthritis (Cawsten et al,
1984). Elevated collagenase activity is also a predominant feature of periodontitis (Overall et al,
1991). Collagenase disrupts the fibrous meshwork of the extracellular matrix and increases
matrix porosity (Berrier and Yamada, 2007), which enables more rapid migration of leukocytes
through the tissue, thereby enhancing host defense in response to infecting microorganisms. In
high prevalence disorders such as osteoporosis, abnormal bone tissue homeostasis results from
an imbalance between bone formation by osteoblasts and bone resorption by osteoclasts, thereby
leading to net bone resorption and loss of bone mass.
Analogous to diseases arising from imbalances of degradative pathways, excessive production of
matrix proteins is also associated with several diseases (McCulloch and Knowles, 1993). For
example, excessive production of fibronectin, collagen, and glycosaminoglycans lead to fibrosis
(Korn et al, 1992). In hepatic fibrosis, there is a net accumulation of collagen types I and III in
liver. Excessive production and deposition of matrix proteins in the heart by cardiac fibroblasts
can lead to cardiomyopathy (Eghbali et al, 1988; Porter and Turner, 2009). The regulation of
synthesis and degradation of extracellular matrices by the activities of resident cells is central to
our understanding of pathogenic mechanisms in connective tissues.
1.4 Cell function in matrix remodeling
Cells interact with the extracellular matrix mechanically and chemically, often with dramatic
effects on the architecture of connective tissues (Alberts et al, 2002). The macromolecules that
constitute the matrix are mainly produced by local cells that reside in the matrix. In addition to
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matrix synthesis, cells remodel the matrix in various ways. As noted above, cells can release
enzymes such as MMPs that degrade matrix proteins. In addition, cells such as macrophages or
fibroblasts can phagocytose matrix components and degrade these proteins in lysosomes (Cate
and Syrbu, 1974; Arora et al, 2001). In healing wounds, connective tissue cells can migrate into
the blood clot of a wound and deposit matrix proteins to enable apposition of the cut edges, and
ultimately, wound closure. The aforementioned processes of matrix secretion, deposition and
phagocytosis, as well as the migration of cells, are critical for the remodeling of extracellular
matrices.
2 Cell Migration
A central process for achieving and maintaining connective tissue homeostasis is cell migration,
which involves a tightly regulated sequence of mechanical and biochemical events that enable
cells to explore and remodel connective tissue matrices in order to maintain normal tissue
structure. Cell migration also plays important roles in embryonic development, the inflammatory
response, wound repair, and tumor formation and metastasis (Lauffenburger and Horwitz, 1996).
2.1 Mechanics of Cell Migration
Cell migration involves coordinated regulation of the formation and function of cell adhesions at
the leading and trailing edges of the cell, and the assembly and disassembly of actin filaments.
Migrating cells undergo extensive and sequential changes of cell shape and structure
(Lauffenburger and Horwitz, 1996). Cell migration is initiated by extension of the leading edge,
a process which relies on actin polymerization (Stossel, 1993), followed by attachment of the
leading edge to the substratum (Izzard and Lochner, 1976), disassembly of adhesions at the
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trailing edge (Chen, 1981; Regen and Horwitz: 1992), and net forward translocation of the cell
(Jay et al, 1995). The morphologies of the protrusions in the leading edge include large, broad
lamellipodia or spike-like filopodia, which are stabilized by adherence to the extracellular matrix
or to adjacent cells via transmembrane receptors linked to the actin cytoskeleton (Ridley et al,
2003).
Cell adhesions serve as traction sites for migrating cells. Similar to a caterpillar tractor, cells
move over adhesions and then the adhesions are disassembled at the trailing edge of the cell,
allowing the cell to detach (Ridley et al, 2003). The rate of forward displacement is regulated in
part by the net adhesive strength of the cell to the matrix, which in some cell types is biphasic
over time as the cell moves forward (Palecek et al, 1997: Diagram 1). Cells are thought to
migrate most quickly when adhesive strength is intermediate (Palecek et al, 1997). Cell
migration rate is lowest when adhesive strength is very low; in these circumstances cells cannot
generate the traction forces that are needed for net forward displacement. When adhesive
strength is high, cells become so tightly anchored to the substrate that they cannot detach from
the matrix and translocate (Palecek et al, 1997). A comprehensive understanding of cell
migration requires identification of the factors controlling the assembly and disassembly of cell
adhesions, the organization and behaviour of the actin filament network, and the factors that
govern the spatial relationships of organelles such as the endoplasmic reticulum and
mitochondria in the translocating cell.
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Diagram 1 – Biphasic curve of migration rate
2.2 Cell Adhesions
2.2.1 Families of Matrix Receptors
Cell adhesion is a complex process involving several classes of cell adhesion molecules
including integrins, immunoglobulin-like domain containing cell adhesion molecules (IgCAMs),
selectins, cadherins, cell surface heparin sulfate proteoglycans (HSPGs), and ADAMs (A
Disintegrin and A Metalloprotease) (Dzamba et al, 2001; Volkmer, 2001; Mcever, 2001; Radice
and Takeichi, 2001; Reizes et al, 2001; White et al, 2001). Controlled regulation of cell adhesion
is necessary for normal development of tissues and for wound healing (Sjaastad and Nelson,
1997).
IgCAMs are part of the immunoglobulin superfamily, a widespread and complex protein family
of more than 100 members that is associated with many different functions (Volkmer, 2001).
8
IgCAMs function as cell adhesion and signaling receptors that transduce extracellular signals
from neighbouring cells or from the extracellular matrix to the intracellular signaling machinery
(Volkmer, 2001). Selectins are a family of trans-membrane molecules, expressed on the surface
of leukocytes and activated endothelial cells (Mcever, 2001) that are important in the
inflammatory response and for the migration of leukocytes to infected or injured sites. Cadherins
participate in the maintenance of proper cell-to-cell contacts (Radice and Takeichi, 2001) and are
particularly important for the maintenance of epithelial integrity and in creating syncytia in
fibroblast populations of soft connective tissues (Ko and McCulloch, 2001). HSPGs function as
important cell-cell and cell-matrix adhesion receptors (Reizes et al, 2001). ADAMs are a family
of trans-membrane and secreted proteins that function in cell adhesions (White et al, 2001). One
of the major groups of matrix receptors that play a particularly pivotal role in cell adhesion is the
integrin family.
2.2.2 Integrins
Integrins are trans-membrane glycoproteins that mediate adhesion of cells to the extracellular
matrix (Dzamba et al, 2001). They are composed of two different subunits (α and β) that are non-
covalently linked (Hynes et al, 1989; Calderwood et al, 2004). Various combinations of 18
different α subunits and 8 different β subunits combine to produce a large group of matrix and
intercellular adhesion molecules that exhibit marked ligand specificity. Most cells express more
than one integrin (Diaz-Gonzales et al, 1996; Porter and Hogg, 1997). In particular, β1and β3
integrins regulate cell adhesion, cell spreading, and cell migration in connective tissue cells
(Calderwood et al, 2004). Integrins provide an important functional connection between
extracellular matrix proteins and the cytoskeleton. Matrix proteins such as collagen, fibronectin,
laminin and tenascin bind to specific types of integrins (Hynes et al, 1989; Horwitz et al, 1985;
9
Giancotti et al, 1999), indicating a measure of ligand specificity. For example fibronectin binds
to β1 and β3 integrins on the basis of the arginine-glycine-aspartic acid (RGD) sequence in
fibronectin (Alberts et al, 2002).
In addition to their role in cell adhesion, integrins can transduce signals either from the inside of
the cell to the outside (inside-outside signaling) or from the outside to the inside (Liddingtion
and Ginsberg, 2002; Hynes, 2002). Associated with these signaling processes, integrins exhibit
the property of being able to switch from a low to a high affinity state, which is accompanied by
activation of adaptor and signaling molecules that induce changes in the conformation of the
integrin (Carman and Springer, 2003). In cell protrusions, many integrins are in a high-affinity
state (Schwartz and Ginsberg, 2002), which enables strong binding to the underlying substrate.
Activation of integrins is dependent in part on binding of talin, a focal adhesion protein, to the
cytoplasmic tail of β1 integrins and β3 integrins (Calderwood et al, 1999; Calderwood et al,
2004). Talin binding alters allosteric conformational states of integrins and increases their
affinity to matrix ligands. Disruption of talin-integrin interactions prevents integrin activation
(Tadokoro et al, 2003). Signaling through integrins is important for cell migration, especially in
terms of regulating cell adhesion. The net strength of cellular adherence to the substrate is
influenced by several integrin-related variables including the types of integrins that are
expressed, their density on the plasma membrane, and the affinity state of individual receptors
(Dzamba et al, 2001). Highly adherent cells have increased density of activated integrins on the
plasma membrane while in contrast, motile cells have fewer activated integrins and are not as
tightly adherent to the substrate. As the cytoplasmic tail of integrins is short and lacks enzymatic
activity (Giancotti et al, 1999), integrins are thought to transduce adhesion-related signals by
associating with adaptor proteins that also provide a mechanical connection between integrins
10
and the cytoskeleton. The adaptor proteins include a large group of focal adhesion proteins,
many of which bind actin filaments.
2.2.3 Focal Adhesions
Focal adhesions are specialized organelles of cultured cells that contact the substrate (Izzard and
Lochner, 1976) and are believed to play a critical role in cell migration in vitro and possibly in
vivo. During the process of cell migration, focal adhesions undergo assembly at the leading edge
of the cell and disassembly at the trailing edge (Burridge et al, 1988). In motile cells focal
adhesions are often small and transient while they are typically large and stable in stationary
cells (Couchman and Rees, 1979; Kolega et al, 1982). The constituents of focal adhesions
include the matrix proteins to which cells are attached, integrins, cytoskeletal-associated or focal
adhesion proteins (such as talin, paxillin, vinculin, α-actinin), and actin filaments. Accordingly,
focal adhesions provide sites of mechanical continuity between the extracellular matrix and the
actin cytoskeleton. The function of focal adhesions is very dependent on protein-protein
interactions involving integrins and focal adhesion proteins. Importantly, the linkage between
integrins and the actin cytoskeleton is critical for the integrity, stability and remodeling of
adhesions (Vicente-Manzanares et al, 2009), which in turn are required for cell migration.
Talin and α-actinin bind to integrin cytoplasmic domains, which provide a connection to the
actin cytoskeleton (Horwitz et al, 1986). The actin binding protein vinculin also binds to talin
and α-actinin (Burridge and Mangeat, 1984) and is an important determinant of adhesion. Talin
provides a mechanical link between integrins and the actin cytoskeleton, which stabilizes
adhesions (Zhang et al, 2008). Disruption of focal adhesion components (i.e. linkages between
the substratum and actin) compromises the integrity of adhesions and can lead to their
disassembly. For example, loss of talin expression leads to impaired cell adhesion, spreading,
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and migration (Calderwood et al, 1999; Zhang et al, 2008). Further, calpain, which is a calcium-
dependent protease, helps to disassemble adhesions by its catalytic action on talin (Chan et al,
2010). Disruption of other components of focal adhesions also affects adhesion dynamics
(Vicente-Manzanares et al, 2009). For example, genetic deletion of vinculin increases cell
migration (Xu et al, 1998) while knockdown of α-actinin leads to the formation of small
adhesions (Choi et al, 2008). Notably, cells that cannot form activated integrins generate weak
adhesions (Zhang et al, 2008).
The formation of focal adhesions is regulated by members of the rho subfamily of the ras family
of GTP-binding proteins (cdc 42, rac, rho) (Yamada and Geiger, 1997). Initially, when focal
adhesions are formed, focal adhesion and cytoskeletal-related proteins, such as the focal
adhesion kinase (FAK), paxillin, and tensin, are phosphorylated (Schaller and Parsons, 1994).
These post-translational modifications alter the binding affinity of these proteins with each other
and with other focal adhesion-associated proteins, thereby regulating adhesive processes. Once
focal adhesions are formed at the leading edge of the cell, they increase in size and adhesive
strength as the cell migrates forward (Ridley et al, 2003). Adhesions remain fixed to the
substratum until they reach the rear or an edge of the cell as the cell migrates over them (Ridley
et al, 2003).
For a cell to migrate rapidly, adhesions at the trailing edge of the cell need to be disassembled
efficiently (Chen, 1981). The release of adhesions in the rear part of the cell results from a
combination of several mechanisms that include cytoskeletal contraction and contributions from
signaling pathways involving rho, tyrosine kinases and calpain (Chen, 1981; Regen and Horwitz:
1992; Jay et al, 1995; Chan et al, 2010). As the cell moves forward due to the release of
adhesions, portions of the cell-associated integrins remain attached to the substratum (Regen and
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Horwitz, 1992). Interestingly, no focal adhesion proteins (such as talin and paxillin) are left
behind in these integrin-membrane remnants (Regen and Horwitz, 1992). Integrins that remain
on the cell surface undergo two possible fates: 1) regulated release, in which they disperse on the
cell surface or 2) endocytosis into vesicles that accumulate in the cell body (Regen and Horwitz,
1992). Assembly and disassembly of focal adhesions is a critical process for cell migration,
which also involves the tightly regulated remodeling of the actin cytoskeleton.
2.3 Actin
2.3.1 Control of Actin Assembly
The cytoskeleton is comprised of actin filaments, microtubules, and intermediate filaments,
which are responsible for the mechanical properties and shapes of cells (Pollard and Cooper,
2009). More specifically, actin filaments provide mechanical strength and structure, and are
required for the motility of animal cells. Microtubules are responsible for separating
chromosomes, for the long-range transport of large particles in all eukaryotic cells and for the
formation of cell adhesions. Intermediate filaments in vertebrate cells function as intracellular
ligaments and struts that resist mechanical forces (Pollard and Copper, 2009).
Actin is the most abundant protein in most mesenchymal cells and is essential for cell survival
(Pollard and Cooper, 2009). Actin exists as monomers and filaments (Lee and Dominguez,
2010). Actin filaments provide internal mechanical support, tracks for movements of
intracellular materials, and help to generate the forces that enable cell movement (Janmey, 1994;
Mitchison and Cramer, 1996; Pollard and Cooper, 2009). Under physiological conditions, actin
monomers spontaneously polymerize into long, stable filaments (Pollard and Cooper, 2009).
Actin filaments are polar in that the subunits in the filament point in the same direction. The
13
barbed ends of actin filament grow much faster than the pointed ends (Schafer and Copper,
1995) as actin monomers are preferentially added on to barbed ends.
Actin cytoskeletal dynamics are regulated by controlling the balance between pools of actin
monomers and filaments. The molarity of monomeric actin in solution under steady state
conditions is defined as the critical concentration (Theriot, 1994). Monomeric actin in vitro
polymerizes in a head-to-tail fashion to form filaments with a distinct structural polarity (Theriot,
1994). The critical concentration is different at the two ends of the actin filament, and also varies
depending on the concentration of salts, the concentration of magnesium and calcium, and the
relative concentrations of ATP and ADP (Theriot, 1994; Schafer and Cooper, 1995).
Actin is assembled into filaments initially by nucleation (Winder and Ayscough, 2005).
Polymerization of actin is energetically unfavourable unless there is a nucleus of three
associating monomers (Pollard, 1986; Winder and Ayscough, 2005; Pollard, 2007). Rapid
nucleation of filaments can be mediated by various actin binding proteins (ABPs). Notably, new
filaments can also be formed from the side of existing filaments or by severing an existing
filament (Winder and Ayscough, 2005; Pollard, 2007). Different proteins function to promote
each of these modes of filament elongation. The Arp2/3 complex can nucleate filaments from the
sides of existing filaments (Winder and Ayscough, 2005; Pollard, 2007). This is important in
motile cells because polymerization of actin drives cell migration at the leading edge of the cell
(Pollard and Cooper, 2009) by a so-called Brownian ratchet mechanism. Interactions of actin
filaments with myosin also produce movement. Myosin generates forces that act on adjacent
actin filaments, thereby producing contractions that pull the rear of the moving cell forward
(Pollard and Cooper, 2009). Actin polymerization can also be nucleated by formin proteins.
14
Once nucleated, actin filaments are able to grow rapidly by addition of monomers at their barbed
ends (Winder and Ayscough, 2005; Pollard, 2007).
Actin monomers bind ATP and extend filaments as they are incorporated at the barbed end
(Winder and Ayscough, 2005; Pollard and Cooper, 2009). As the filament matures, ATP bound
in the central cleft of actin is hydrolysed, phosphate is released and the resulting ADP-actin
filament is disassembled by loss of monomers from the pointed end. The release of ADP-actin
monomers then undergoes nucleotide exchange to generate ATP-actin monomers that can be
used for new rounds of polymerization (Winder and Ayscough, 2005; Pollard and Cooper, 2009).
Eukaryotic cells use more than 100 actin binding proteins (ABPs) to shape the actin
cytoskeleton. By their actions, ABPs maintain a pool of actin monomers, initiate actin
polymerization, restrict the length of actin filaments, regulate assembly and turnover of actin
filaments, and cross-link filaments into networks or bundles (Pollard and Cooper, 2009).
2.3.2 Actin binding proteins
ABPs determine the organization and the behaviour of the actin cytoskeleton (Hartwig and Yin,
1988). ABPs are classified into seven groups (Dos Remedios et al, 2003): 1) monomer-binding
proteins sequester G-actin and prevent polymerization (e.g. thymosin β4, DNase I); 2) filament-
depolymerizing proteins induce the conversion of F-actin to G-actin (e.g. CapZ and cofilin); 3)
filament end-binding proteins cap the ends of actin filaments, thereby preventing the exchange of
monomers at the pointed end (e.g. tropomodulin) and at the barbed end (e.g. CapZ); 4) filament
severing proteins shorten the average length of filaments by binding to the side of F-actin and
cutting it into two pieces (e.g. gelsolin); 5) cross-linking proteins contain at least two binding
sites for F-actin, thus facilitating the formation of filament bundles, branching filaments, and
15
three-dimensional networks (e.g. filamin A). 6) Stabilizing proteins bind to the sides of actin
filaments and prevent depolymerization (e.g. tropomyosin). 7) Motor proteins that use F-actin as
a track upon which to move (e.g. myosin family of motors) (Dos Remedios et al, 2003). The
functions of several ABPs are not limited to one class. For example, gelsolin can sever actin
filaments and cap barbed ends while the Arp2/3 complex can nucleate filament formation,
elongate filaments, and establish branch points in actin networks (Dos Remedios et al, 2003).
2.3.3 Role in leading edge protrusions
Actin filaments play an important role in the protrusion of the leading edge, which enables the
cell to move forward. Migrating cells move forward by extension of lamellipodia and filopodia,
which are controlled by GTPases such as rac, cdc42, and rhoG (Hall, 1994; Ridley et al, 2003).
At the cell edge, an increase in the number of sites for actin polymerization is a first step,
followed by the net addition of actin monomers to actin filament growth sites (Condeelis, 1993;
Hall, 1994). New barbed ends that are available for actin monomer addition arise by a
combination of mechanisms including uncapping of already-existing filaments, their severing, or
both, as well as by the formation of new actin trimer nucleation sites (Lauffenburger and
Horwitz, 1996). The gelsolin family of proteins is important for the generation of actin
nucleation sites because these proteins regulate both severing and uncapping of actin filaments
(Hartwig and Yin, 1988).
The regulation of free monomeric actin levels may be a primary effector for membrane extension
(Lauffenburger and Horwitz, 1996). The numbers of actin filaments that can be generated at the
cell cortex can be increased by raising the concentration of actin monomers, which exists in two
pools: free monomeric actin and actin monomers bound to monomer-binding proteins (e.g. β-
thymosins, profilins, and ADFs/cofilins) (Sun et al, 1995; Theriot, 1994). These proteins serve as
16
a potential source of actin monomers that are necessary for filament extension at the leading cell
edge.
Protrusion of the membrane is tightly coupled to polymerization of actin filaments at the leading
edge (Mitchison and Cramer, 1996). Actin polymerization in lamellipodia is mediated in part by
Arp2/3 complexes, which bind to the sides or tip of a pre-existing actin filament and induce the
formation of a new daughter filament that branches off the mother filament (Ridley et al, 2003).
Formin proteins such as mDia1 and mDia2 are also involved in promoting actin growth in a
linear fashion by binding to the barbed end of actin filaments and there promoting filament
extension by nucleating filament growth (Vicente-Manzanares et al, 2005).
2.4 Endoplasmic reticulum
2.4.1 Structure and function
The endoplasmic reticulum (ER) exhibits a heterogeneous structural organization in the cell
(Berridge, 2002) and is highly plastic, which enables it to assume many configurations in order
to perform its many functions (Berridge, 2002). The ER can appear as flattened sacs, which are
important for its functions in protein synthesis, or it can exist as an interconnected meshwork of
tubules (Baumann and Walz, 2001). The tubular network is constantly remodeled through
processes such as tubule sliding, tubule branching and ring closure (Baumann and Walz, 2001).
The ER can be divided morphologically into rough and smooth ER. The rough ER, which
contains ribosomes, is actively involved in protein synthesis. The smooth ER is responsible in
part for Ca2+ signaling (Berridge, 2002). In addition to protein synthesis and Ca2+ signaling, the
ER modifies and transports proteins to various parts of the cell (Petersen and Verkhratsky, 2007).
17
The ER is also capable of generating a variety of specific output signals, which control gene
transcription at different levels.
As noted above, the ER plays a central role in the generation of cytosolic Ca2+ signals (Petersen
and Verkhratsky, 2007) where it acts both as a sink and source of Ca2+. The ER intraluminal
[Ca2+] is approximately 10,000 times higher than the cytosol (Petersen and Verkhratsky, 2007).
Release of Ca2+ in response to input signals is under the control of InsP3 receptors (InsP3Rs) and
ryanodine receptors (RYRs) (Berridge, 2002).
2.4.2 Ca 2+ homeostasis and cell signaling
Cell migration and changes in cell morphology, which are regulated in part by the activities of
actin binding proteins, are accompanied by transient increases in cytoplasmic Ca2+ concentration
(Marks and Maxfield, 1990; Janmey, 1994). Ca2+ alters the functions of many proteins that
affect the organization of the actin cytoskeleton (Elson, 1988; Stossel, 1989). The effects of Ca2+
depend on the kinetics of elevation of [Ca2+] as well as the actual [Ca2+] in the cytoplasm that are
generated (Janmey, 1994). In migratory cells such as leukocytes, there are transient increases in
intracellular [Ca2+]. Ca2+ is released from internal stores such as the ER, and this process can
increase cytoplasmic [Ca2+] from 100 nM to µM levels (Marks and Maxfield, 1990). The
increase in cytoplasmic [Ca2+] activates many calcium-sensitive proteins, including the gelsolin
family of proteins, which then can sever actin filaments (Yin, 1987) and enable actin remodeling
for cell migration.
As mentioned above, calcium release from the ER is involved in the regulation of a wide range
of cellular functions, such as actin binding proteins. In some cases, actin binding is activated by
Ca2+, and in other cases, it is inhibited or reversed. Gelsolin for example, which disrupts the actin
18
networks by filament severing, is activated by Ca2+, whereas α-actinin, which cross-links actin
filaments, is inhibited by Ca2+ (Janmey, 1994).
In migrating cells Ca2+ plays many important roles. It influences directional sensing, changes the
actin cytoskeleton organization, affects traction force generation, and changes position of focal
adhesions (Ridley et al, 2003; Van Haastert and Devreotes, 2004; Pettit and Fay, 1998; Brundage
et al, 1991; Lee et al, 1999). Previous studies showed that intracellular Ca2+ exhibits a rear-to-
front gradient, in which the lowest [Ca2+] is at the front of a migrating cell (Brundage et al,
1991). However, more recent studies revealed that the leading edge of a migrating cell contain
numerous proteins that require high levels of [Ca2+] for activation (Wei et al, 2009). For
example, migrating WI-38 fibroblasts exhibit high amplitude, transient increases of [Ca2+] at the
leading edge (Wei, 2009).
The intracellular [Ca2+] in cells is tightly regulated and in this context, it is notable that the ER
forms a highly dynamic interconnected network with mitochondria with which they cooperate to
maintain or generate Ca2+ signals. The ER releases Ca2+ for a specific cell function after which
mitochondria sequester Ca2+ and then return it to the ER (Berridge, 2002). This role of
mitochondria in shaping the Ca2+ signal released from the ER is important for Ca2+ homeostasis.
2.4.3 ER-focal adhesion linkage
As described above, focal adhesions are dynamic structures and other cellular organelles may be
sequestered at adhesion sites for the conduct of essential cellular processes. For example, focal
adhesions contain large numbers of proteins that are involved in apoptotic signaling pathways.
They may also provide a physical and functional connection to organelles such as the ER (Tran
et al, 2002; Wang et al, 2006; Mak et al, 2008). Evidence for an ER-focal adhesion connection
19
has been provided in studies that examined focal adhesion-associated proteins and found ER-
resident proteins such as calnexin and BiP, which are ER-resident proteins (Mak et al, 2008).
Conceivably, the integrity of the focal adhesion-ER connection is dependent on the actin
cytoskeleton. In cells treated with swinholide A, which inhibits actin filament assembly, focal
adhesion formation and co-accumulation of calnexin and BiP are blocked (Mak et al, 2008).
These data indicate that interactions between focal adhesions and ER-resident proteins depend on
the integrity of actin filaments and the formation of focal adhesions (Mak et al, 2008). In a
separate study, ER-resident proteins such as Bip, calnexin, calreticulin, and IP3 receptor (type 1)
were linked to focal adhesions using immunoprecipitation and TIRF microscopy methods (Wang
et al, 2006). Further, kinectin, which is an integral membrane protein in the ER that is involved
in vesicle transport, localizes to sites of cell adhesions (Tran et al, 2002). This finding suggests
that adhesion formation can trigger differential recruitment of cytoplasmic components involving
localization of a portion of the ER. Currently, the connection of focal adhesions to the ER with
respect to cell migration is not defined. It is plausible that the ER may be recruited to sites of cell
adhesions, where local provision of Ca2+ influences the assembly and disassembly of adhesions
during cell migration. This is similar to phagocytosis where the ER is redistributed to the site of
the forming phagosome, where it provides Ca2+ that is required for the phagocytic process to
occur (Stendhal et al, 1994).
2.5 Mitochondria
2.5.1 Role of actin in mitochondrial function
Mitochondria are dynamic organelles with important functions including energy production,
regulation of cytosolic calcium levels and modulation of apoptosis (Cavanagh et al, 2009).
20
Mitochondria are heterogeneously dispersed in cells (Anesti and Scorrano, 2006) and assume
different morphologies depending on the cell type. For example, mitochondria exhibit a range of
shapes that may include small spheres or short rods or long tubules (Detmer and Chan, 2007).
Mitochondria are often enriched at sites where energy demand is substantial or where metabolic
functions are sequestered (Anesti and Scorrano, 2006).
Actin cables are used as scaffolds for the attachment and movement of mitochondria in yeast.
Mitochondria in S. cerevisiae bind to the actin cytoskeleton in order to position themselves and
move (Anesti and Scorrano, 2006). In yeast, proteins such as Mmm1p (maintenance of
mitochondrial morphology 1 protein), Mdm10p (mitochondrial distribution and morphology 10
protein) and Mdm12p provide a connection between mitochondria and the actin cytoskeleton
(Burgess et al, 1994; Sogo and Yaffe, 1994; Berger et al, 1997). These proteins comprise an
integral mitochondrial membrane protein complex that spans the mitochondrial outer and inner
membranes and provide linkages to the actin cytoskeleton. Movement of mitochondria is driven
by actin polymerization via the Arp2/3 complex. Arp2/3 complex co-localizes with mitochondria
by its interaction with the outer membrane (Boldogh et al, 2001). While actin cables represent
the main tracks for mitochondrial movement in S. cerevisiae, in higher eukaryotes this
relationship is not as well-defined. In mammalian cells the distribution of mitochondria is largely
controlled by their movement along microtubules, which is mediated by kinesin and dynein
motors (Campello et al, 2006). However, some studies have shown that actin filaments also play
an important role in mitochondrial transport (Morris and Hollenbeck, 1995; Ligon and Steward,
2000).
The shape and size of mitochondria are highly variable and are controlled by fusion and fission
processes, which are regulated by a family of mitochondria-shaping proteins that include specific
21
mitochondrial, dynamin-related proteins (Anesti and Scorrano, 2006). Mitofusins (MFN1 and
MFN2), which are located on the mitochondrial outer membrane, are essential for fusion of the
outer mitochondrial membranes (Detmer and Chan, 2007). Mitochondrial fission requires
recruitment of a dynamin-related protein (DRP1) from the cytosol. DRP1 assembles into
punctuate spots on mitochondrial tubules, and a subset of these complexes leads to productive
fission events (Detmer and Chan, 2007). At steady state, the frequencies of fusion and fission
events are balanced to maintain the overall morphology of the mitochondrial population
(Nunnari, 1997). However, cells with a high fusion-to-fission ratio generate fewer mitochondria,
and these mitochondria are typically elongated and highly interconnected (Detmer and Chan,
2007). Conversely, cells with low fusion-to-fission ratios contain numerous mitochondria that
are small spheres or short rods, which are referred to as fragmented mitochondria (Detmer and
Chan, 2007). The coordination between movement and changes in mitochondrial shape is
important as mitochondria must divide into smaller units so they can be readily transported
(Anesti and Scorrano, 2006) to areas of the cell with high energetic demands. For example, in
neurons, mitochondria are enriched at sites of high ATP consumption at cortical regions within
the pre-synaptic zone, dendritic spines, growth cone and nodes of Ranvier (Boldogh and Pon,
2007).
2.5.2 Functions in cell motility and actin assembly
The role of mitochondria in actin assembly and ultimately, their impact on cell migration, is just
starting to be discovered. Some proteins that regulate the fusion-fission of mitochondria
orchestrate lymphocyte chemotaxis (Campello et al, 2006). MFN1, which promotes fusion,
prevents migration of lymphocytes toward a chemical gradient. Conversely, pro-fission
molecules such as DRP1 stimulate chemotaxis (Campello et al, 2006). Modulations of
22
mitochondrial fusion or fission processes are of considerable interest for defining the role of
mitochondria in cell migration. The notion that fragmentation enhances mitochondrial
redistribution and cell migration, whereas conditions that promote fusion have the opposite
effect, is intriguing.
ATP production by mitochondria plays an important role in cell migration. ATP in cells is
generated by both oxidative phosphorylation in mitochondria and by glycolysis in the cytosol.
Migrating cells require ATP production by mitochondria (Campello et al, 2006). When
lymphocytes are treated with oligomycin, a specific inhibitor of the mitochondrial F1F0-ATPase,
cell migration is impaired (Campello et al, 2006). In neurons mitochondria are present
throughout the axon, but they accumulate where the need for ATP production is especially high
(Morris and Hollenbeck, 1995; Verstreken et al, 2005).
Changes in cytoskeletal structure require ATP and ATP-producing mitochondria; accordingly
these organelles are of central importance for cytoskeletal structure dynamics (Cavanagh et al,
2009). Conversely, mitochondria need to interact with cytoskeletal elements to develop normal
motility, morphology, localization, and function (Boldogh and Pon, 2006). During cell
migration, actin undergoes assembly and disassembly. ATP is necessary for stabilizing actin
filaments (Pollard and Borsiy, 2003). Hydrolysis of ATP and dissociation of the γ-phosphate
triggers actin filament turnover by regulatory proteins in cell migration (Blanchoin and Pollard,
2002). The Arp2/3 complex initiates filament branching that grows in the direction of the barbed
end by addition of Mg-ATP actin. New branches grow rapidly and push the membrane forward
(Pollard and Borisy, 2003; Pollard and Cooper, 2009). Each filament grows transiently since
capping proteins terminate growth. Actin subunits in this branched network hydrolyze their
bound ATP and dissociate the γ-phosphate (Pollard and Borisy, 2003; Pollard and Cooper,
23
2007). Dissociation of the γ-phosphate initiates disassembly by inducing de-branching and
binding of ADF/cofilin, which in turn, promotes severing and dissociation of ADP-subunits from
actin filament ends (Pollard and Borisy, 2003; Pollard and Cooper, 2007). Profilin is an
important nucleotide exchange factor for actin, catalyzing exchange of ADP for ATP and
returning subunits to the ATP-actin-profilin pool, ready for another cycle of assembly (Pollard
and Borisy, 2003; Blanchoin and Pollard, 2002). In the steady state, ATP-actin associates at the
barbed end and ADP-actin dissociates from the pointed end, leading to a treadmilling of subunits
from the barbed end to the pointed end. ATP hydrolysis in the filament is essential for
maintaining treadmilling, a process that is vital for cell migration (Pollard and Borisy, 2003).
3 Flightless I
3.1 Discovery
Flightless I was originally discovered in studies of Drosophila melanogaster which were
investigating genes for development and differentiation that impacted fly behavior (Koana and
Hotta, 1978). Flies were initially treated with ethyl methanesulphonate (EMS), which causes
mutagenesis, and then screened for X-linked flightless mutants of Drosophila melanogaster by
using a column-type flight tester (Koana and Hotta, 1978). A number of mutations were isolated.
Chromosomal mapping and complementation experiments defined the loci of interest. One of the
loci identified was Flightless I (FliI). Mutation of the gene leads to loss of flight due to
myofibrillar abnormalities in the indirect flight muscles (Koana and Hotta, 1978). The mutants
exhibit normal ability in walking, jumping, and holding up their wings before starting flight;
these processes are governed by tubular muscles. In contrast, the Flightless mutants lack the
ability to beat their wings when tethered, suggesting a functional defect in the indirect flight
24
muscle system (Koanna and Hotta, 1978). Comparative electron microscopic studies of indirect
flight muscles found a deficiency in the Z bands and a disorganized arrangement of myofibrils
(Koanna and Hotta, 1978; Miklos and De Couet, 1990). More severe allelic mutations caused
lethality in early embryogenesis and affect the processes of cellularization and gastrulation
(Campbell, 1993).
FliI cDNAs have been isolated and characterized. These analyses predicted a protein of 1256
amino acids (Campbell et al, 1993). Data base searches revealed homologous genes in
Caenorhabditis elegans and humans. The predicted C. elegans and human proteins are 49% and
58% identical to the D. melanogaster protein (Campbell et al, 1993). The predicted proteins
contain gelsolin-like domains (GLD) as well as a leucine rich repeat (LRR) region (Diagram 3).
The similarity of the FliI sequences in different species suggest conservation of the function of
this protein over an extended evolutionary period (Campbell et al, 1997). In mice FliI is
necessary for development because FliI knockout mice are embryonic lethal (Campbell et al,
2002).
Disruption of expression or mutation of regulatory cytoskeletal genes can result in
developmental abnormalities and tumorigenesis (Claudianos and Campbell, 1995). The human
flightless gene has been mapped to chromosome 17p11.2, which is the critical region deleted in
the Smith-Magenis Syndrome (SMS) (Chen et al, 1995). The SMS is a gene micro-deletion
syndrome in which affected patients suffer from mental retardation, short stature, facial
dysmorphology, developmental delay, and self-destructive behavior (Chen et al, 1995). As
indicated above, FliI was identified as a member of the gelsolin family of proteins, based on its
gelsolin-like domains. Alignment of human FliI protein and human gelsolin protein reveals 52%
identity.
25
3.2 Structure and Function of Gelsolin Family Proteins
The gelsolin super-family of proteins consists of seven members: gelsolin, adseverin, villin,
capG, advillin, supervillin and flightless I (Silacci et al, 2004). All of these proteins contain three
or six homologous repeats of a domain that is termed the gelsolin-like domain. Villin, advillin,
supervillin and flightless I have additional domains in addition to the six-fold repeats. Gelsolin is
the founding member of this family; its actin severing activity is regulated by changes of Ca2+,
intracellular pH, phosphoinositides and tyrosine phosphorylation. Gelsolin can bind, sever, and
cap actin filaments (Janmey and Stossel, 1987; Yin, 1987; Hartwig et al, 1995). Domain 4 of
gelsolin binds to actin in a Ca2+-dependent manner (Sun et al, 1999). Domains 1-2 also bind
actin, but independently of Ca2+ (Sun et al, 1999). These domains are also responsible for
severing and capping activities.
The gelsolin family of proteins plays an important role in the organization of the actin
cytoskeleton by influencing filament length, flexibility, and concentration (Silacci et al, 2004).
These proteins, on account of their capacity to alter the structural properties of actin filaments,
are important for regulating cellular morphology and function, including cell motility (Silacci et
al, 2004). Over-expression of gelsolin (Cunningham et al, 1991) or capG (which lacks actin
severing activity) (Sun et al, 1995) increases the motility of fibroblasts in vitro.
3.3 Structure and Function of Flightless I
Flightless I (FliI) contains two segments. Segment 1 of the FliI protein comprises a 400 amino
acid leucine-rich repeat (LRR) domain at the N-terminal (Campbell et al, 1993). Segment 2 of
the FliI protein comprises the six gelsolin-like domains (GLD) at the C-terminal (Campbell et a,
1993). FliI has an estimated size of 143,672 Da (Campbell et al, 1993). The LRR of FliI may
26
play a role in several cellular processes. Proteins containing LRRs have diverse cellular
localizations (extracellular, cytoplasmic, transmembrane, and nuclear) and functions (receptor-
ligand binding, signal transduction, cell adhesion, development, bacterial virulence, DNA repair,
and RNA processing) (Liu and Yin, 1998). The LRR motif contributes to protein-protein
interactions, either directly as the ligand binding molecule, or as a regulator to enhance affinity
and/or specificity of binding to a separate ligand-binding site (Liu and Yin, 1998). Previous
studies showed several interacting proteins with the human LRR of FliI that included LRR-FLI
interacting protein 1 (LRRFIP1), LRR-FLI interacting protein 2 (LRRFIP2) (Fong et al, 1999)
and FLAP-1 (Liu and Yin, 1998). LRRFIP1 and LRRFIP2 are predominantly expressed in
cardiac tissues and skeletal muscle.
LRR GLD
6 5 4 3 2 1
Diagram 2 – Structure of Flightless I
The presence of gelsolin-like domains in FliI has led to speculation that FliI protein is involved
in reorganizing the actin cytoskeleton. The gelsolin-like domains of FliI have been implicated in
interactions with actin (Liu and Yin, 1998). However, its actin binding activity has not been
measured. Further, the regulatory pathways that control the synthesis and degradation of FliI are
not defined (Kopecki and Cowin, 2007).
FliI binds to Daam1 and mDia1 (Higashi et al, 2010). It enhances the intrinsic actin assembly of
Daam1 and mDia1 in vitro and is required for Daam1-induced actin assembly in living cells
27
(Higashi et al, 2010). Daam1 and mDia1 proteins play central roles in actin dynamics as a result
of their rates in assembling actin filaments (Higashi et al, 2010).
3.4 Flightless I in Cell Migration
Reorganization and remodeling of the actin cytoskeleton is fundamental to all aspects of tissue
repair, including the crawling of keratinocytes during wound re-epithelialisation, infiltration of
inflammatory cells, and migration of fibroblasts required for deposition and remodeling of the
extracellular matrix and wound contraction (Adams et al, 2007). FliI co-localizes with actin-
based structures (Davy et al, 2000), influences cell motility, regulates wound repair, and affects
cell proliferation (Cowin et al, 2007). In mice that are FliI +/-, skin wound healing is enhanced
compared to wild-type mice. In contrast, FliI-over-expressing mice exhibit significantly impaired
healing, with reduced cell proliferation and delayed epithelial migration. These studies reveal
that the effect of FliI on wound repair is mediated by its inhibitory effect on cell proliferation and
migration (Cowin et al, 2007; Adams et al, 2007). When skin fibroblasts or keratinocytes are
grown to confluence and scratch-wounded in vitro, the wounded area closes significantly faster
when FliI levels are reduced. In contrast, scratch wounds close significantly more slowly in FliI-
over-expressing fibroblasts (Cowin et al, 2007). Currently, the mechanisms by which FliI
impacts cell migration are not defined. Conceivably, FliI may regulate focal adhesions as FliI
interacts with proteins directly linked to the cytoplasmic domain of integrins (i.e. talin, vinculin,
paxillin) (Kopecki et al, 2009).
28
Statement of the Problem
Connective tissue homeostasis in heart, muscles and joints is an important process for the
maintenance of health. Homeostatic processes that regulate connective tissue turnover are
disrupted in various diseases including heart failure and arthritis with important consequences for
human health. Notably, the actin cytoskeleton of connective tissue cells plays a central role in
tissue development and regeneration, which are important processes for homeostasis. In
connective tissues, a wide variety of regulatory signals including mechanical forces are
transmitted through focal adhesions, which connect the actin cytoskeleton to the extracellular
matrix via integrins and other matrix receptors. My preliminary data indicated that focal
adhesions contain a protein known as Flightless I (FliI), a member of the gelsolin family of actin
severing proteins, which may provide a connection between focal adhesions and other organelles
such as the endoplasmic reticulum. The endoplasmic reticulum is a major storage site of calcium
ions, which together with focal adhesion proteins, may help to regulate important aspects of cell
metabolism that are critical for tissue development and regeneration. One of the central processes
in determining tissue homeostasis and regeneration is cell migration. Based on my data,
knockdown of Flightless I enhances cell migration but currently the mechanism by which
Flightless I is involved in regulating cell migration is not defined.
Hypothesis
There are focal adhesion-associated proteins, such as Flightless I, that provide a functional
connection to the endoplasmic reticulum, thereby regulating cell migration.
29
Objectives
1) Localize Flightless I in migrating fibroblasts.
2) Examine the role of Flightless I as a potential regulator of cell adhesions.
3) Determine the role of Flightless I in mediating activation of β1 integrins.
4) Determine the role of Flightless I in the organization of the actin cytoskeleton.
5) Determine the potential relationship of Flightless I with the endoplasmic reticulum.
30
Chapter 2
Introduction
Imbalances of connective tissue homeostasis can manifest as fibrosis or net destruction of normal
matrix structure. These perturbations are commonly seen in high prevalence diseases of humans
including heart failure, osteoporosis, arthritis and periodontitis (Porter and Turner, 2009; Berrier
and Yamada, 2007). A critical, defining element for achieving and maintaining connective tissue
homeostasis is cell migration, which involves a tightly regulated sequence of mechanical and
biochemical processes that enable cells to explore and remodel connective tissue matrices in
order to maintain normal tissue structure. Migrating cells undergo extensive and sequential
changes of cell shape and structure, which are visible microscopically as protrusions of the
leading edge of the cell, followed by adhesion of the leading edge to the underlying substrate,
disassembly of adhesions at the trailing edge and finally, net forward translocation of the cell
(Lauffenburger and Horwitz, 1996). The rate of forward displacement is regulated in part by cell
adhesion strength to the matrix, which in some cell types is biphasic over time as the cell moves
forward (Palecek et al, 1997). When adhesive strength is very low, cells cannot generate the
traction forces needed for forward translocation (Palecek et al, 1997). In contrast, cells that are
very tightly attached to the underlying matrix may be unable to efficiently detach the trailing
edge and can migrate only slowly or not at all (Palecek et al, 1997). Consequently, net forward
translocation rates are often optimized when cells exhibit intermediate levels of cell adhesive
strength, but the determinants of adhesive strength are not well-defined.
Some of the potential loci for regulation of adhesive strength in migrating cells involve the
formation and remodeling of adhesions to matrix proteins. In cultured cells, focal adhesions are
31
intensively studied and specialized cellular domains that via integrins, provide mechanical
continuity between the extracellular matrix and actin filaments (Izzard and Lochner, 1976; Heath
and Dunn, 1978; Burridge et al, 1988; Yamada and Geiger, 1997). Continuous remodeling of
actin filaments is required for cell migration (Pollard and Cooper, 2009). At the leading edge of
the cell, actin filaments undergo cycles of assembly and disassembly, enabling forward extension
of the leading lamellipodia (Pollard and Borisy, 2003; Weaver et al, 2003; Vicente-Manzanares
et al, 2005; Pollard, 2007) and the formation and reorganization of cell adhesions (Webb et al,
2004; Vicente-Manzanares et al, 2009). The structure, function and remodelling of actin
filaments are regulated by actin binding proteins, a large array of actin monomer sequestering
and actin filament severing, bundling, cross-linking and nucleating proteins. More than 100 actin
binding proteins have been identified, and a subset of these proteins has been identified in focal
adhesions (Dos Remedios et al, 2003). For example, filamin A is an actin binding protein that
cross-links actin cytoskeleton, is enriched in focal adhesions and regulates cell adhesions by
controlling cell surface expression and activation of β1 integrins (Kim et al, 2008; Meyers et al,
1998). Importantly, a subset of actin binding proteins is regulated by [Ca2+], most notably the
gelsolin superfamily of proteins.
Ca2+, which is released from the endoplasmic reticulum and plays an important role in cell
migration, is required for the regulation of the structure and dynamics of the actin cytoskeleton
(Hartwig and Yin, 1988) and the formation and disassembly of cell-substratum adhesions
(Sjaastad and Nelson, 1997). Tight regulation of Ca2+ is particularly important for actin filament
assembly and disassembly in adhesions. In general, cytosolic [Ca2+] is ~100 nM (Petersen and
Verkhratsky, 2007) but transient increases of [Ca2+] (up to 1-2 μM) can lead to activation of
actin severing proteins such as gelsolin (Janmey and Stossel, 1987), which alter the actin
32
filament network within the cell and facilitate cell motility (Cunningham 1991; Arora 1996).
Further, previous data indicate that endoplasmic reticulum-resident proteins such as kinectin,
calreticulin (Tran et al. 2002) and calnexin (Mak et al. 2008) are recruited to nascent focal
adhesions, suggesting potential functional linkages between the endoplasmic reticulum and
developing focal adhesions.
Based on the dynamic relationship between focal adhesions, the actin cytoskeleton and the
endoplasmic reticulum in migration, we used mass spectrometry to screen for proteins that may
provide a functional connectivity between these three structures to thereby regulate cell
migration. Mass spectrometry of focal adhesion fractions identified Flightless I (FliI), a member
of the gelsolin family that was originally identified in Drosophila melanogaster (Koana and
Hotta, 1978; Campbell et al, 1993). Previous studies have indicated that FliI may play a role in
remodeling the actin cytoskeleton (Campbell et al, 1993; Davy et al, 2000; Goshima et al, 1999).
Accordingly, we investigated the localization, role and function of FliI in cell migration and the
organization of actin filaments.
33
Materials and Methods
Reagents
Mouse monoclonal anti-FliI (sc-21716) and rabbit polyclonal anti-FliI (sc-30046) antibodies
were obtained from Santa Cruz Biotechnology (CA, USA). Mouse monoclonal antibody against
calnexin (610523) and rat anti-mouse CD29 (9EG7) antibody were purchased from BD
Biosciences. Mouse monoclonal anti-paxillin antibody (clone 5H11) was obtained from
Millipore. Mouse monoclonal anti-KDEL antibody (10C3) was obtained from Assay Designs
(Ann Arbor, Michigan). Mouse monoclonal anti-β-actin (clone AC-15), protease inhibitor
cocktail, and carbonyl cyanide m-chlorophenylhydrazone (CCCP) were obtained from Sigma-
Aldrich (St. Louis, s). Latrunculin B was purchased from Calbiochem. Jasplakinolide (J7473)
and Mito-tracker red (M7513) were purchased from Molecular Probes. Mouse monoclonal anti-
HA antibody (MMS-101P), biotin-SP-conjugated AffiniPure goat anti-rat IgG (H+L), biotin-SP-
conjugated AffiniPure goat anti-rabbit IgG (H+L), biotin-SP-conjugated AffiniPure goat anti-
mouse IgG, FITC-conjugated AffiniPure F(ab’)2 fragment goat anti-rat IgG (H+L), Cy 2-
conjugated streptavidin and Cy3-conjugated streptavidin were purchased from Cedarlane
Laboratories. Rabbit polyclonal anti-BiP antibody was purchased from Cell Signaling
Technology. Rabbit polyclonal anti-prohibitin (ab28172) was purchased from Abcam
(Cambridge, Massachusetts).
Cell Culture
NIH 3T3 cells obtained from the American Type Culture Collection were cultured at 37oC in
complete DME medium containing 10% fetal calf serum and antibiotics (Penicillin G
34
124units/mL, Gentamicin SO4 50µg/mL, Fugizone 0.25µg/mL). Cells were maintained in a
humidified incubator gassed with 95% air and 5% CO2, and were passaged with 0.01% trypsin
(Gibco, Burlington, ON).
Collagen bead binding
Ferric oxide beads (<5 µm) or carboxylate- or sulphate-coated polystyrene microbeads (<2
microns) were coated with collagen or BSA as described (Lee et al, 1996). Briefly, beads were
incubated with 1 mL of an acidic bovine collagen solution (InVitrogen) at a concentration of 3
mg/mL to produce collagen-coated beads. The collagen solution was neutralized to pH=7.4 with
100 µL of 1N NaOH added to 1 mL of the collagen solution to induce fibril assembly on beads.
The beads were incubated with cells at 37oC for 30 min. BSA-coated beads were produced by
incubating the beads with 1 mL of 0.15% BSA at 37oC for 30 min.
Isolation of Focal Adhesions
After addition of collagen or BSA-coated magnetite beads, bead-associated proteins were
isolated as described (Plopper and Ingber, 1993). Briefly, cells were washed with cold PBS to
remove unbound beads, scraped into cold cytoskeleton buffer (0.5% Triton X-100, 50 mM NaCl,
300 mM sucrose, 3 mM MgCl2, 10 mM PIPES pH 6.8, 1 mM phenylmethylsulfonyl fluoride,
1:50 dilution Protease Inhibitor Cocktail) and vortexed for 30 s. The beads were isolated from
the lysate with a magnet, re-suspended in cold cytoskeleton buffer, homogenized, and re-isolated
magnetically. Bead-associated proteins were removed by boiling in Laemmli sample buffer.
35
Immunoblotting
Whole cell extracts were prepared by rinsing cells with cold Ca2+-Mg2+-free PBS, scraped into
cold lysis buffer (50 mM Tris pH 7.5, 150 mM NaCl, 1% Triton X-100, 0.1% SDS, 1% sodium
deoxycholate, 1:50 dilution Protease Inhibitor Cocktail), and passed through a 23G needle 5
times. The homogenate was centrifuged at 18,000 g at 4oC for 10 min, and the supernatant was
retained for biochemical analysis. Equivalent amounts of protein (BCA assay) were separated on
SDS-PAGE gels. Immunoblotted samples were probed with appropriate antibodies and
quantified by scanning densitometry. Other cellular fractions that were prepared by differential
centrifugation were immunoblotted in a similar manner and equivalent amounts of protein were
separated on SDS-PAGE gels and probed with appropriate antibodies.
Mass spectrometry
Bead-associated proteins were isolated from cells and eluted with 50 mM glycine buffer (pH 2.3-
2.5). The eluted proteins were dialyzed for 36 hours in carbonate buffer (25 mM NH4HCO3;
pH=7.5). Trypsin (1 µg; Roche, Indianapolis, IN) was added to the sample and rotated overnight
at 37oC. Subsequently, 0.1% acetic acid was added to the sample followed by air drying with a
speed vac. Lyophilized samples were analyzed by LC-MS/MS with a QStar XL, ES1-Qq-TOF,
AB1/MDS Sciex mass spectometer (Concord, Ontario; Mass Spectrometry Facility, Hospital for
Sick Children, Toronto). Data from the LCMS run were analyzed using the National Center for
Biotechnology Information (NCBI) database and the MASCOT search engine. Scaffold 2.0
proteome software was used for analyzing search results.
36
Transfections
NIH3T3 fibroblasts were seeded on to 6-well plates and cultured until 30% confluent at the time
of transfection. FliI siRNA ON-TARGETplus SMARTpool (J-050081-09 Target Sequence:
CGGAGUUUACGGAGGA, J-050081-10 Target sequence: ACAUUGACUUCUCGCUACA, J-
050082-22 target sequence: GGAGAUGGGUGACGAGAGU, J-050081-12 target sequence:
GGUCCUGGAUGUUCGAGAA- Dharmacon Thermo Scientific) were transfected into cells
using Oligofectamine (InVitrogen) in which 175 µL of FliI siRNA (20 µM) in DME was
incubated with 15 µL of Oligofectamine for 20 min at room temperature. Then, 200 µL of
siRNA:Oligofectamine complex was added to each well. After 6 hours of incubation 1 mL of
growth medium (DME, 10% FCS, Antibiotics) was added to each well. Cells were incubated for
48 hours prior to analysis by western blot.
Cell migration
Cells in monolayers were grown on fibronectin-coated plates (10 µg/mL fibronectin in
incubation medium). A scratch in the monolayer was created with a 200 µL pipette tip to model
a wound. Images were captured after scratching and at regular intervals until the scratch was
closed by cell migration. Identical image fields were compared to quantify cell migration rate.
Immunocytochemistry
Cells were fixed with 3.7% formaldehyde in PBS, permeabilized with 0.2% Triton X-100,
blocked with 0.2% BSA, and stained with appropriate antibodies or with specific fluorescent
dyes. Confocal microscopy (Leica, Heidelberg, Germany; 40 x oil immersion lens) was used to
determine the spatial distribution of proteins of interest or of organelles. Transverse optical
sections were obtained at a nominal thickness of 1 µm. Total internal reflection fluorescence
37
(TIRF) microscopy (Leica, Heidelberg, Germany; 100 x oil immersion lens) was used to study
subcellular regions of the cell in contact with the substrate (optical penetration depth <110 nm).
Jet Wash Assay
For estimation of cell adhesion (Chong et al, 2007), cells were grown in monolayer culture (24-
well plate) and subjected to shear forces with increasing numbers of PBS washes (1-16 washes
with a repeating Eppendorf pipettor). Cells were fixed with 3.7% formaldehyde, permeabilized
with Triton X-100 and stained with DAPI (5 µg/mL). In 5 fields of view in each well, DAPI-
stained nuclei were counted to provide estimates of the number of adherent cells. Four wells
were evaluated for each wash in 3 different repeats.
Differential Centrifugation
Cell lysates were fractionated by differential centrifugation of extracts prepared in Triton X-100
(Yamamoto et al, 2001). Briefly, cells were collected in CSKB buffer containing 1 µM
phalloidin (Invitrogen) and centrifuged at 15,900 g for 2 min and then at 366,000 g for 20 min.
The pellets from each step were resuspended in the same volume of buffer as the original lysate.
Samples were boiled in SDS sample buffer and equal fractions of protein from each pool were
separated by SDS-PAGE and immunoblotted.
Subcellular fractionation
Endoplasmic reticulum and mitochondria were isolated as described (endoplasmic reticulum
Isolation Kit, Sigma). Briefly, cells (109) were harvested and resuspended in cold hypotonic
isolation buffer (100 mM HEPES, pH 7.8, 10 mM EGTA, 250 mM KCl) for 20 min, which
allowed the cells to swell. Cells were centrifuged at 600 g and re-suspended in isotonic isolation
38
buffer (50 mM HEPES, pH 7.8, 1.25 mM sucrose, 5 mM EGTA, 125 mM KCl), and disrupted
with a Dounce homogenizer (Dounce tissue grinder set, Sigma Aldrich) with 20 strokes,
followed by passing through a 30 gauge needle (20 strokes). The homogenate was centrifuged at
1000 g for 10 min. The supernatant was recovered and centrifuged at 12,000 g for 15 min. The
resulting pellet was the microsomal fraction which contained the mitochondrial fraction and
endoplasmic reticulum fraction. The specificity of the mitochondrial fraction and the ER
fractions were confirmed by immunoblotting with the mitochondrial-specific protein prohibitin
and the ER-specific protein calnexin and BiP.
Fluorescence recovery after photobleaching
Cells were seeded on to 35 mm glass bottom microwell dishes (MatTek Corporation, Ashland,
MA) and transfected with GFP-Paxillin. Samples were placed on to a heated microscope stage
and imaged with a confocal microscope (scan speed-800 Hz). Circular regions of interest (4 µm2
bleach spots) were illuminated and photobleached with an argon laser at full power (488 nm for
4 seconds). Fluorescence in the bleached spot was measured before bleaching and for 30 seconds
after bleaching. The diffusion coefficient (D, X 10-10 cm2/s) and the mobile fraction (%R) were
calculated as described (Axelrod et al, 1976; Jacobson and Wojcieszyn, 1984).
Statistical Analysis
For all continuous variable data, means and standard errors of means were computed. When
appropriate, comparisons between two samples were made by Student’s t-test with statistical
significance set at p<0.05. All experiments were performed at least three times in triplicate.
39
Results
Collagen- or BSA-coated magnetite beads were attached to the dorsal surface of cultured mouse
fibroblasts. After cell lysis, beads were purified magnetically (Plopper and Ingber, 1993) and the
bead-associated proteins were analyzed by tandem mass spectrometry (Arora et al, 2008). Of the
more than 100 different proteins identified in bead-associated fractions, Flightless I (FliI) was
notable because it was found in fractions prepared from collagen-coated beads, which mimic the
ligand-bound state of integrins, but not in BSA-coated beads, which bind non-specifically to
cells (Table 1). The presence of FliI in the collagen bead-associated fractions was confirmed by
immunoblotting (Fig. 1A). As a negative control, collagen bead-associated fractions were
immunoblotted for GAPDH. For a positive control, collagen bead-associated fractions were
immunoblotted for the focal adhesion protein paxillin.
We examined the possible relationship between FliI and focal adhesions in imaging studies using
total internal reflection fluorescence (TIRF) microscopy. Mouse fibroblasts were transfected
with GFP-paxillin, allowed to spread and migrate on fibronectin-coated cover slips, fixed and
immunostained for FliI. TIRF microscopy was used to assess adhesive cellular domains
immediately adjacent to the substrate (<150 nm above the superior surface of the cover slip).
There were both short and long streaks of GFP-paxillin near the cell periphery (Fig. 1B). The
immunostaining pattern for FliI showed a disperse distribution of dots throughout the cytoplasm.
In the merged TIRF image of GFP-paxillin and FliI immunostaining, there were small, discrete
areas of co-localization of GFP-paxillin and FliI (Pearson’s coefficient = 0.46±0.04), indicating
that only a fraction of FliI is present in focal adhesions adjacent to the substrate. Confocal
microscopy, which enables examination of a thicker focal plane (~1500 nm), showed that FliI is
40
located closer to the cytoplasmic face of focal adhesions (Fig. 1C). Importantly, when examining
three Z-section regions of fibroblasts (dorsal, middle, and ventral) using confocal microscopy,
FliI was distributed predominantly in the middle section (Fig. 1D).
We determined whether FliI impacts cell migration with an in vitro scratch assay. Fibroblasts
were grown on a fibronectin-coated substrate. Fibronectin is one of the major ECM proteins and
binds β1 integrins (Alberts et al, 2002). Confluent cultures were scratch-wounded and phase
contrast images were obtained at several time intervals thereafter to measure cell migration into
the denuded area. FliI expression was reduced by ~70% with FliI siRNA compared to irrelevant
siRNA (Fig. 2A). Cells with reduced FliI expression migrated more quickly into the denuded
area than did cells treated with irrelevant siRNA (Fig. 2B, C; p<0.05), consistent with previous
data (Cowin et al, 2007; Adams et al, 2007). As siRNA knockdown does not reduce protein
content uniformly in all cells (Ge et al, 2005), we examined the expression of FliI in siRNA-
treated cultures by immunostaining to determine whether migrating cells exhibited similar levels
of FliI as non-migrant cells. Cellular fluorescence above background was used to estimate FliI
protein expression. Migratory cells in the scratch wound exhibited reduced FliI expression (Fig.
2D; p<0.03), consistent with the finding that knockdown of FliI enhances cell migration.
As FliI evidently affected migration, we determined its potential importance in cell adhesion.
Cells were plated on fibronectin-coated substrates overnight and then subjected to repeated
washes with PBS (Chong et al, 2007). Under these conditions, cells experience shear forces (35
dynes/cm2) due to the ejected fluid from the pipette tip. For both cell types, with increased
numbers of washes, the percentage of cells that remained adherent to the plate decreased.
Notably, FliI knockdown cells were washed off more readily than cells treated with irrelevant
41
siRNA (Fig. 3A; p<0.01 for 1st and 16th washes, p<0.05 for 2nd and 4th washes, p<0.02 for 8th
wash).
As FliI is found in the vicinity of focal adhesions and since knockdown of FliI enhances cell
migration and attenuates cell adhesion, we explored the effect of FliI on integrin function. Cells
in scratch assays were stained with 9EG7, a neo-epitope antibody that binds to activated β1-
integrins (Lenter et al, 1993). Bound 9EG7 was quantified by counter-staining with biotin and
Cy3-conjugated streptavidin and TIRF microscopy-based photometry. Cells labeled with 9EG7
antibody exhibited stained streaks of variable length, indicating the presence of activated β1-
integrins in these adhesive zones. In migrating cells with reduced expression of FliI, there were
fewer cells with activated β1 integrins (Fig. 3B; p<0.03), indicating that FliI may regulate focal
adhesion function by increasing activation of β1 integrins.
We examined the effect of FliI depletion on the mobility of GFP-paxillin in focal adhesions by
fluorescence recovery after photobleaching. Cells treated with irrelevant or FliI siRNA were
transfected with GFP-paxillin and circular areas (4 µm in size) in focal adhesions were bleached
with a laser. During the recovery period, unbleached GFP-paxillin moved into the bleached area
and the fluorescence was measured. The % mobile fraction for control (irrelevant siRNA) cells
was 54±9% and for FliI knockdown cells was 52±7% (p>0.2; Fig. 3C). Similarly, the diffusion
coefficient was not significantly different between controls and FliI knockdown (control:
1.34±0.5 x 10-9 cm2/s; FliI knockdown: 1.98±0.45 x 10-9 cm2/s; p>0.2). These data indicate that
FliI does not affect the mobility of paxillin into focal adhesions and therefore does not likely
affect focal adhesion turnover.
We determined whether FliI interacts with the actin cytoskeleton, possibly regulating the
organization and behaviour of actin filaments in adhesions. FliI immunoprecipitates were
42
obtained from cell lysates and FliI-associated proteins were analyzed by tandem mass
spectrometry. Several proteins were identified by mass spectrometry, which included β-actin,
non-muscle myosin IIA (NMMIIA), and PAK1 (Table 2). We confirmed these findings by
immunoblotting FliI immunoprecipitates for β-actin, NMMIIA, and PAK1 (Fig. 4A). These
proteins were abundant in the FliI immunoprecipitates, indicating a potential functional
association between these proteins and FliI. Because of the presence of gelsolin-like domains in
FliI (Campbell et al, 1993), and because the functional relationship between FliI and actin is not
defined, we focused subsequent studies on the potential association between FliI and β-actin.
Notably, the immunoprecipitation data did not indicate whether FliI associates with actin
monomers or actin filaments. We addressed this problem by separating actin monomer and
filament pools in cells by differential centrifugation. Under these conditions, the high-speed
supernatant was enriched for actin monomers while the high-speed pellet was enriched for short
actin oligomers and actin filaments (Yamamoto et al, 2001). When equal amounts of proteins
from these fractions were immunoblotted, FliI was present in both the high speed pellet and
supernatant fractions, indicating that FliI may associate with actin monomers, oligomers and
filaments. With this approach there was no conclusive evidence to indicate that FliI preferentially
associates with one form of the actin more than the other.
We explored the spatial association of FliI with actin monomers and actin filaments using in-situ
affinity labeling. Fibroblasts were fixed and stained for actin monomers (FITC-DNAse I;
Blikstad et al, 1978) or in separate cultures for actin filaments (FITC-phalloidin; Wulf et al,
1979). In both cultures, FliI was immunostained and detected with an anti-FliI antibody and Cy3-
conjugated streptavidin reagents (Fig. 4C). Confocal microscopy showed co-localization
between FliI and actin monomers (Pearson’s coefficient = 0.52 ±0.04). This is consistent with
the biochemical findings above showing that FliI associates with actin monomers. However,
43
there was less co-localization between FliI and actin filaments (Pearson’s coefficient = 0.20
±0.03). We also examined the relative abundance of actin filaments in control and FliI
knockdown cultures. Cells were stained with FITC-phalloidin. The fluorescence attributable to
the actin filament network was lower in FliI knockdown cells compared to control cells (Fig. 4D;
p<0.02).
We investigated the association of FliI with the actin cytoskeleton by manipulating the relative
abundance of actin pools and measuring the amount of FliI in the high speed pellet and
supernatant. Cells were treated with either jasplakinolide (1 µM), which stabilizes and promotes
actin polymerization (Arora et al, 2004), or with latrunculin B (1 µM), which sequesters actin
monomers (Segal et al, 2000). The pools were isolated and immunoblotted for FliI (Fig. 4E).
When cells were treated with jasplakinolide there was more FliI in pellets than in supernatants
but latrunculin B treatment made no difference (Fig. 4E-i). In cells treated with jasplakinolide, β-
actin immunoprecipitates were prepared from pellets or supernatants and then immunoblotted for
FliI (Fig. 4E-ii). In these samples there was more FliI in the pellets than supernatants.
As we found FliI in the vicinity of focal adhesions, and as FliI evidently associated with actin,
which together play an important role in cell migration (Vicente-Manzanares et al, 2009), we
investigated the potential relationship between FliI and the endoplasmic reticulum. Notably, the
ER is also an important regulatory organelle for cell migration (Wei et al, 2009). Microsomal
fractions were prepared by differential centrifugation, and immunoblotted for calnexin and BiP
(ER resident proteins) to confirm the authenticity of ER enrichment (Fig. 5A). ER-enriched
microsomal fractions were also immunoblotted for FliI. There was abundant FliI in the ER
fractions, suggesting that FliI may be associated with the endoplasmic reticulum. However, when
cells were immunostained for FliI and for KDEL, a marker of the ER (Fig. 5B), confocal
44
microscopy showed very little or no co-localization between FliI and KDEL staining (Pearson’s
coefficient = 0.15±0.03), indicating that FliI probably does not associate with the ER. Notably,
the distribution of FliI throughout the cytoplasm suggested the possibility that it may associate
with other organelles.
We examined the microsomal fractions prepared earlier for other organelle markers. The
fractions that were separated by differential centrifugation were immunoblotted for prohibitin, an
inner mitochondrial membrane protein (Artal-Sanz and Tavernarakis, 2009). The microsomal
fraction was enriched with prohibitin, indicating the presence of mitochondrial proteins (Fig.
5A). Since FliI was also abundant in the microsomal fraction, we considered a possible spatial
association between FliI and mitochondria. Fibroblasts were loaded with Mito-tracker red, which
diffuses across the plasma membrane and concentrates in mitochondria (Minamikawa et al,
1999). Cells were fixed and immunostained for FliI (Fig. 5C). Confocal microscopy showed co-
localization between FliI and mitochondria (Pearson’s coefficient = 0.57±0.02). We next used
TIRF microscopy to analyze the distribution and localization of mitochondria adjacent to cell
adhesions. While mitochondria were not present at the most ventral aspect (i.e. the substrate
side) of focal adhesions, they were present in the vicinity of focal adhesions (Fig. 5D).
As FliI was found to associate and co-localize with both actin monomers and mitochondria, we
considered the possibility that FliI acts as a linker between actin monomers and mitochondria. In
a preliminary study, cells were loaded with Mito-tracker red, fixed and stained for actin
monomers (Fig. 5E). Confocal microcopy showed marked co-localization (Pearson’s coefficient
= 0.62±0.03) between actin monomers and mitochondria.
45
Discussion
The principal findings of this study are that FliI is present at focal adhesions and associates with
actin monomers; knockdown of FliI enhances cell migration, attenuates cell adhesion and
reduces activation of β1 integrins. Contrary to our a priori hypothesis, FliI does not associate
with the endoplasmic reticulum. Instead, FliI apparently co-localizes with mitochondria. These
findings may offer novel insights into the potential roles of Flightless I in cell migration.
Focal adhesions are specialized organelles that provide structural and functional continuity
between the cytoskeleton and the underlying extracellular matrix (Izzard and Lochner, 1976;
Burridge et al, 1988). We isolated focal adhesion-associated proteins with collagen-coated beads
and used mass spectrometry to identify FliI in the focal adhesion fractions. Our imaging studies
with TIRF microscopy showed that only a fraction of FliI was present in focal adhesions. In
addition to our findings of an association of FliI with cell adhesions, previous
immunoprecipitation studies showed that FliI associates with focal adhesion proteins including
talin, paxillin and vinculin (Kopecki et al, 2009). The possible physical association between FliI
and focal adhesions is significant as FliI is evidently one of >50 proteins that are found in these
organelles where it may contribute to cell adhesion. Because FliI is predominantly cytosolic, it is
possible that its presence in focal adhesions arises from its association with actin monomer
binding proteins.
Since we found FliI in focal adhesions, we explored its role in cell migration and adhesion
processes which are pivotal in connective tissue homeostasis and tissue regeneration (Ridley et
al, 2003; Abreu et al, 2008). Our data showed that knockdown of FliI enhances cell migration
and attenuates cell adhesion, indicating a novel role of FliI in stabilizing focal adhesions. When
46
cells expressed FliI, cells adhered to the substrate strongly. Conversely, when FliI expression
was reduced, adherence to the substrate was diminished and cell motility was enhanced. Our
studies with fluorescence recovery after photobleaching of GFP-paxillin indicate that FliI
probably does not regulate adhesion strength by affecting the recruitment of actin binding
proteins into focal adhesions. While the actual mechanism by which FliI enhances cell migration
is not defined (Kligys and Jones CR, 2009), FliI may affect the formation of cell attachments at
the leading cell edge and the release of attachments at the trailing edge, thereby regulating
migration rates.
The potential regulatory role of FliI on focal adhesion function was investigated by measurement
of activated β1 integrins in migration assays. β1 integrins enable cell migration by physically
linking actin filaments to extracellular matrix proteins including collagen and fibronectin
(Dzamba, 2001; Calderwood et al, 2004). Our studies showed that after siRNA knockdown,
migrating cells with reduced expression of FliI also exhibited lower levels of activated β1
integrins. These data suggest that integrin activation may be under the control of FliI. Since FliI
associates with talin (Kopecki et al, 2009) and talin binding to integrin is the final step in β1
integrin activation (Calderwood et al, 2004), FliI could mediate binding of talin to β1 integrins.
Increased levels of FliI may lead to higher talin-integrin binding interaction, further enhancing
β1 integrin activation and strengthening cell adhesion to the substrate. Conversely, when FliI
expression is reduced, the activation of β1 integrins is reduced, cells are less tightly anchored to
the substrate and they can migrate more quickly.
Cell migration is a highly complex and regulated process that not only involves the remodeling
of focal adhesions but also the actin cytoskeleton. Actin is a key component in the molecular
machinery of cell migration (Vicente-Manzanares et al, 2009). Because FliI plays an important
47
role in cell migration, we considered its potential interactions with the actin cytoskeleton. Our
mass spectrometry data of FliI immunoprecipitates identified β-actin as a protein that may
associate with FliI. Although an interaction of FliI with actin has not been established, previous
studies have suggested an association based on co-localization of FliI and actin (Davy et al,
2000; Li et al, 2008). Conceivably, FliI may play an important role in the organization and
behaviour of the actin cytoskeleton as a result of its structural similarities to other actin binding
proteins. FliI has 6 domains in its C-terminal that are similar to gelsolin, which is an actin
binding protein that remodels actin filaments by severing actin filaments, nucleating actin
filament assembly, and blocking the fast exchanging end of actin filaments (Yin, 1987).
Currently, the role of FliI in regulating actin filament structure has not been defined.
We found that there is a strong association between FliI and monomeric actin, which is
supported by the increased abundance of FliI in actin monomer-rich supernatants after
centrifugation. Actin assembly and the proteins that regulate actin filament organization are
essential regulators of cell migration (Berrier and Yamada, 2007). FliI may be involved in the
process of contributing to actin filament growth, thereby facilitating incorporation of actin
filaments into focal adhesions. This conjecture is supported by our imaging studies that showed a
diminished actin filament network when FliI expression was reduced. Possibly, FliI may promote
the assembly of highly organized branched actin networks by providing nucleotide-charged actin
monomers to growing actin filaments.
In addition to cellular adhesion to substrates and the remodeling of the actin cytoskeleton, cell
migration depends on temporally and spatially discrete release of Ca2+ from intracellular stores
(Wei et al, 2009). A major intracellular storage site of Ca2+ is the endoplasmic reticulum from
which Ca2+ is released during cell migration (Lee et al, 1999). Initially, our immunoblot analysis
48
of microsomal fractions indicated that FliI may associate with the ER. These data lead us to
explore a potential association of FliI with the ER, which was not confirmed in co-localization
studies. Accordingly, we re-examined the microsomal fractions, which are enriched with ER and
other organelle proteins. We found abundant amounts of prohibitin, a mitochondrial matrix
protein (Artal-Sanz and Tavernarakis, 2009). Further imaging studies showed that FliI co-
localizes with mitochondria. Collectively these findings indicate a potentially important
connection between FliI and mitochondria. Since FliI is an actin binding protein, it is possible
that FliI regulates mitochondrial movement and morphology within fibroblasts in a similar
manner as yeast. In yeast, mitochondria bind to actin filaments, which regulate both the
positioning and transport of mitochondria (Anesti and Scorrano, 2006). Conceivably, FliI
regulates the shape and distribution of mitochondria during cell migration by mediating fusion-
fission equilibrium events. When the equilibrium is shifted towards fusion, there are fewer
mitochondria and these mitochondria form tubular networks with limited capacity for movement
within the cell; under these conditions cell migration is inhibited (Campello et al, 2006; Detmer
and Chan, 2007). Conversely, when the equilibrium is shifted towards fission, there are more
numerous mitochondria that are fragmented. These mitochondria are able to move more freely
within the cell to sites of high energy dependence, therefore facilitating cell migration (Campello
et al, 2006; Detmer and Chan, 2007). FliI may bind to mitochondria to stabilize branched
networks similar to actin filaments and therefore inhibit migration. Conversely, knockdown of
FliI may lead to mitochondrial fission, resulting in enhanced cell migration. The importance of
this connection needs to be further investigated.
The spatial distribution of mitochondria throughout the cytoplasm is normally quite disperse
(Anesti and Scorrano, 2006). In addition to their cytosolic distribution, the relative numbers of
mitochondria are often concentrated at sites where energy demand is high or where sequestered
49
metabolic function is required (Anesti and Scorrano, 2006). Our data indicate that mitochondria
are enriched in the vicinity of focal adhesions where energy is required for organelle remodeling
and conceivably for adhesion. Since FliI associates with mitochondria, it is possible that FliI may
bind to mitochondria for recruitment in the vicinity of cell adhesions. At these sites mitochondria
are needed to provide ATP for generating ATP-actin, which is required for actin filament
assembly (Pollard, 2007; Blanchoin and Pollard, 2002; Pollard, 1986).
In summary, our findings show that knockdown of FliI enhances cell migration and attenuates
cell adhesion. FliI may impact cell migration by regulating adhesion function through activation
of β1 integrins, organizing the actin cytoskeleton into a more highly branched network, and in
the neighbourhoods of mitochondria, facilitating exchange of ATP-actin monomers on existing
filaments, thereby promoting filament remodeling (Fig. 6).
50
Figure Legends
Figure 1 – A) Immunoblots of collagen bead-associated proteins. Whole cell lysates and bead-
associated fractions were prepared and immunoblotted for FliI (mouse monoclonal anti-FliI),
paxillin (mouse monoclonal anti-paxillin) and GAPDH (mouse monoclonal anti-GAPDH). FliI is
present in the bead-associated fraction. Paxillin serves as a positive control and GAPDH as a
negative control for focal adhesion-associated proteins. B) 3T3 fibroblasts were transfected with
GFP-paxillin and stained with FliI antibody [mouse monoclonal anti-FliI (1:500), Biotin goat
anti-mouse (1:2000), Cy3-conjugated streptavidin (1:4000)]. TIRF microscopy was used to
observe cell adhesions adjacent to the substrate (<150 nm). In the merged image, small discrete
areas of co-localization can be observed (Pearson’s coefficient = 0.46 ±0.04). C) Similar
preparations as B but confocal microscopy was used to observe a thicker focal plane. D)
Fibroblasts stained with FliI antibody and imaged at three different regions of the cell (dorsal,
middle, and ventral) with confocal microscopy show predominantly cytosolic (middle)
distribution of FliI.
Figure 2 A) Fibroblasts were treated with FliI siRNA to knockdown expression of FliI and the
lysates were collected for immunoblotting. FliI expression was reduced by 70%. B) Differential
interference contrast images of cells treated with irrelevant siRNA or with siRNA for FliI. Cells
were examined in in vitro scratch assays at 0 hour and 26 hours post-wounding. C) Comparison
of migration rates between cells treated with irrevelant siRNA or with siRNA to knockdown FliI.
More cells migrated into the scratched area when FliI expression was knocked down (p<0.05).
D) Cells from in vitro scratch assay were stained for FliI. Immunofluorescence due to FliI was
quantified with Image J software. Migratory cells exhibited reduced expression for FliI (p<0.03).
51
Figure 3 A) Cell adhesion of wild-type and FliI knockdown cells were measured by in a shear
force assay. FliI knockdown cells were washed off more readily than cells treated with irrelevant
siRNA (p<0.05). B) Fibroblasts from in vitro scratch assay were stained for activated β1-
integrins using 9EG7 antibody [rat anti-mouse CD29 (1:100), biotin goat anti-rat (1:200), Cy3-
conjugated streptavidin (1:400)]. TIRF microscopy was used to analyze the expression of
activated β1-integrins adjacent to the cell membrane in migratory and non-migratory cells.
Migratory cells in FliI knockdown cultures exhibited reduced staining for activated β1-integrin
(p<0.03). C) Fluorescence recovery after photobleaching was used to measure mobility of GFP-
paxillin in cells treated with irrelevant siRNA or with siRNA to knock down FliI. The percent
recovery of GFP-paxillin in wild-type cells is 54 ± 9% and in FliI knockdown cells is 52 ± 7%
(p>0.2). The diffusion coefficient for wild-type cells is 1.34± 0.5 x 10-9 cm2/s and for FliI
knockdown cells is 1.98 ± 0.45 x 10-9 cm2/s (p>0.2).
Figure 4 A) Immunoprecipitation was performed with a FliI antibody (rabbit polyclonal anti-
FliI) and FliI immmuoprecipitates were immunoblotted for β-actin, NMMIIA, and PAK1. FliI
immunoprecipitates show abundant presence of β-actin, NMMIIA, and PAK1. Nebulin was used
as an irrelevant antibody for control. B) Actin monomers enriched in high speed supernatant
(HSS) and actin filaments enriched in high speed pellet (HSP) were isolated by differential
centrifugation. Fractions were immunoblotted for FliI, β-actin, and GAPDH. FliI associates with
both monomeric actin and filamentous actin. C) 3T3 fibroblasts were stained for actin monomers
(i- FITC-DNAse I) and actin filaments (ii -FITC-phalloidin), and subsequently stained with FliI
antibody. In the merged image there is co-localization between FliI and actin monomers
(Pearson’s coefficient = 0.52 ±0.04); but very little co-localization can be observed between FliI
and actin filaments (Pearson’s coefficient = 0.20 ±0.03). D) Wild-type and FliI knockdown cells
52
were labeled with FITC-phalloidin (1:50, 20 min). Immunofluorescence microscopy was used to
quantify phalloidin fluorescence. FliI knockdown cells exhibited reduced phalloidin fluorescence
(p<0.02). E) i- The relative portions of the actin monomer and filament pools in cells were
manipulated by treatment with jasplakinolide (1 μM, 1 hour), which stabilizes actin filaments
and latrunculin B (1 μM, 10 min), which sequesters actin monomers and enhances actin
depolymerization. By differential centrifugation, actin filaments (P- pellets) and actin monomers
(S: supernatants) were isolated and immunoblotted for FliI and β-actin. There is slightly higher
FliI content in the pellet than in the supernatant when cells were treated with Jasplakinolide. ii –
Similarly, fractions were prepared with the same treatments. The fractions were
immunoprecipitated with β-actin antibody and the immunoprecipitates were immunoblotted for
FliI. There is a more FliI in the pellet when cells were treated with Jasplakinolide.
Figure 5 A) Microsomal fractions were isolated by subcellular fractionation. The fractions were
immunoblotted for prohibitin (a marker for mitochondria), calnexin and BiP (markers for
endoplasmic reticulum), FliI, and β-actin. The 2nd pellet contains the mitochondria and the
endoplasmic reticulum, which was confirmed by enrichment of the expected resident proteins.
FliI is abundant in all of the fractions. B) Cells were labeled with FliI antibody and with KDEL
antibody (marker for ER). Confocal microscopy was used to assess co-localization. In the
merged image there is very little co-localization (Pearson’s coefficient = 0.15 ±0.03). C) Cells
were loaded with Mitotracker red (500 nM for 30 min) and immunostained for FliI. Confocal
microscopy was used to determine co-localization. In the merged image there is a moderate
degree of co-localization (Pearson’s coefficient =0.57 ±0.02). D) Cells were transfected with
GFP-paxillin and immunostained for mitochondria using Mitotracker red. TIRF microscopy was
used to assess cell adhesions for presence of mitochondria. Presence of mitochondria in the
53
vicinity of cellular adhesions can be observed. E) Cells were stained for actin monomers (FITC-
DNAse I) and subsequently for mitochondria (Mito-tracker red). Confocal microscopy was used
to determine co-localization. In the merged image, a moderately high degree of co-localization
can be observed (Pearson’s coefficient = 0.62 ±0.03).
Figure 6 – Proposed mechanism of Flightless I in the organization of actin filaments. FliI acts as
a monomer binding protein that may facilitate exchange of ADP for ATP on actin monomers in
the vicinity of mitochondria. FliI may then transport ATP-actin monomers to barbed end (+) of
actin filaments for assembly before dissociating from the filaments.
Figure 1
A
B
C
D
Flightless I
GAPDHWhole
Cell Lysate
Bead-associated fraction
Bead-Associated Proteins
Paxillin
GFP-Paxillin MergeFlightless I
GFP-Paxillin MergeFlightless I
Flightless I
MiddleDorsal Ventral
Flightless IFlightless I
Figure 2
A
0
20
40
60
80
100
120
Wildtype FliI KND
Rel
ativ
e %
of F
liI e
xpre
ssio
n
0 HR 26 HR Post Wounding
Wildtype
FliI KND
In-Vitro Scratch AssayFlightless IGAPDH
Wildtype FliI KND
B
C
D
Non-Migratory Cells
Non-Migratory Cells
Migratory Cells
0
20
40
60
80
100
120
Non-Migratory Cells
Migratory Cells
% o
f Pix
els a
bove
thre
shol
d in
cell
s st
ained
with
FliI
ant
ibod
y
*
Time Post-Wounding (Hours)
0.0
5.0
10.0
15.0
20.0
25.0
30.0
35.0
0 3 6 9 12 15 18 21 24 27Nu
mb
er o
f ce
lls m
igra
tin
g in
to w
oun
ded
are
a
Wildtype
FliI KND
**
**
**
Figure 3
A
B
C Wildtype
0
10
20
30
40
50
60
0 1.6 3.3 9.1 10.7 12.4 14 15.6 17.3 18.9 20.6 22.2 23.9
Time (s)
Fluo
resc
ence
(A.U
)
FLI KND
0
10
20
30
40
50
60
0 1.6 3.3 9.1 10.7 12.4 14 15.6 17.3 18.9 20.6 22.2 23.9Time (s)
Fluo
resc
ence
(A.U
)
0
20
40
60
80
100
120
0 1 2 4 8 16
Number of Washes
Perc
enta
ge o
f Adh
ered
Cell
s
Wildtype Flii KND
*
* * **
TIRF
DIC
Non-Migratory Cells
Non-Migratory
Cells
In-Vitro Scratch Assay
Migratory Cells
TIRF
DIC
Perc
enta
ge e
xpre
ssio
n of
act
ivat
edβ1
-inte
grin
s (9E
G7)
0
20
40
60
80
100
120
Non-MigratoryCells
Migratory Cells
*
Figure 4A
B
FliI
LSP HSP HSS
β-actin
GAPDH
0
0.2
0.4
0.6
0.8
1
1.2
1.4
Rel
ativ
e Ex
pres
sion
of F
liIw
ith b
eta-
actin
LSP HSP HSS
I.B. NMMIIA
I.B. PAK1
WCL FliI IP
I.B: β-actin
WCL Nebulin IP
IP: NebulinIP: FliI
Flightless I Merge
Actin Filaments Flightless I Merge
Actin Monomers
C i
ii
0
20
40
60
80
100
120
Per
cen
tage
of
pix
els
abov
e b
ackg
rou
nd
leve
ls t
oin
dic
ate
ph
allo
idin
bin
din
g
Wildtype Flii Knockdown
*
D
E
P S P S P S
Control JAS LAT
I.B: FliI
I.B: β-actin
P SP SP S
Control JAS LATIP: β-actin
I.B: FliI Light Chain
i ii
Wildtype Flii KnockdownActin Filaments Actin Filaments
Figure 5
Flii MergeER - KDEL
A
Flii MergeMitotracker
B
C
D
GFP-Paxillin Mitotracker Merge
Prohibitin
β-Actin
FliI
1st P 1st S 2nd P 2nd S
BiP
Calnexin
Actin Monomer Mitochondria Merge
E
Figure 6
Plasma membrane
ADP-Actin
ATP-Actin
FliI MitochondriaMitochondria
+-Intracellular
ExtracellularIntegrin
VinculinTalin
Filamin A
Pi
62
Tables
Table 1: Proteins identified by mass spectrometry from bead-associated fractions
# Identified Protein M. W. (kDa)
1 vimentin 46 2 unnamed protein product 22 3 Histone H1.3 (H1 VAR.4) (H1d) 22 4 heterogeneous nuclear ribonucleoprotein A2/B1 isoform 1 36 5 histone cluster 1, H2bd 14 6 lamin A isoform A 74 7 high mobility group AT-hook 2 12
8 PREDICTED: similar to heterogeneous nuclear ribonucleoprotein A3, isoform 13 30
9 Beta actin 42 10 Hist1h4c protein 11 11 H2A histone family, member X 15 12 Y box protein 1 25 13 histone cluster 1, H1e 22 14 GPI-anchored membrane protein 1, isoform CRA_a 84 15 Plectin-1 (Plectin-6) (PLTN) (PCN) 534 16 Mybbp1a protein 94 17 histone cluster 1, H1a 22 18 interleukin enhancer binding factor 3 isoform 1 98 19 mCG1128, isoform CRA_a 26 20 mCG10912, isoform CRA_a 72 21 Eukaryotic translation elongation factor 1 alpha 1 50 22 flightless I homolog 145 23 unnamed protein product 16 24 Hist1h3e protein 15 25 histone cluster 1, H1c 21 26 AHNAK nucleoprotein isoform 1 604 27 DEAD box polypeptide 17 isoform 3 29 28 retinoblastoma-binding protein mRbAp48 52 29 Nucleolin 77 30 polypyrimidine tract binding protein 1 isoform 1 59 31 topoisomerase (DNA) II alpha 173 32 ribosome binding protein 1 isoform a 158 33 mCG15678, isoform CRA_a 39 34 myeloid-associated differentiation marker 35
63
35 kappa-B motif-binding phosphoprotein 51 36 collagen, type I, alpha 1 138 37 mCG118787 24 38 unnamed protein product 31 39 mCG120696 169 40 unnamed protein product 24 41 integrin, alpha 11 131 42 Nucleolar RNA helicase 2 94 43 unnamed protein product 32 44 unnamed protein product 27 45 Yip1 domain family, member 6 6 46 vacuolar protein sorting 13C 420 47 mitochondrial ribosomal protein L39 39 48 Olfactomedin-like 3 46 49 olfactory receptor 1373 35 50 unnamed protein product 71
51 ELAV-like protein 1 (Hu-antigen R) (HuR) (Elav-like generic protein) (MelG) 36
52 unnamed protein product 17 53 caldesmon 1, isoform CRA_a 48 54 novel protein similar to Atpase class I type 8B member 2 135 55 Pde4dip protein 106 56 lamin B1 67 57 dicer1 216 58 U1 small nuclear ribonucleoprotein polypeptide A 20 59 E4f1 protein 74 60 leiomodin 2 (cardiac) 62 61 UDP-N-acetylglucosaminyltransferase 117 62 cDNA sequence BC048502 18
63 malic enzyme 3, NADP(+)-dependent, mitochondrial, isoform CRA_b 28
64 Chd4 protein 94 65 protein tyrosine phosphatase-like protein PTPLB 29 66 mCG12252, isoform CRA_a 31 67 PREDICTED: similar to Rpl23a protein 26 68 mCG23000, isoform CRA_a 12 69 N-glycan processing alpha-mannosidase IIx 131 70 amyotrophic lateral sclerosis 2 (juvenile) chromosome region 42 71 PREDICTED: hypothetical protein 198 72 p100 co-activator 99 73 mCG51561 26 74 FYVE, RhoGEF and PH domain containing 5, isoform CRA_b 65 75 erythrocyte protein band 4.1-like 2 110 76 titin 3766 77 procollagen type V alpha 2 145
64
78 PREDICTED: similar to 4930422G04Rik protein 214 79 nectin-like protein 2 33 80 unnamed protein product 39 81 mCG59859 104 82 nucleoporin 153 152 83 Lamin B2 [Mus musculus] 69 84 unnamed protein product 51 85 PREDICTED: similar to Sm D2 isoform 1 13 86 jumonji protein 137 87 RRS1 ribosome biogenesis regulator homolog 42 88 PREDICTED: hypothetical protein 6 89 mCG121979, isoform CRA_b 95 90 Septin-1 42 91 envelope polyprotein 70 92 mCG131128, isoform CRA_c 19 93 Serine/arginine repetitive matrix protein 2 295 94 programmed cell death 4 52 95 unnamed protein product 41 96 cytidine and dCMP deaminase domain containing 1 59 97 hypothetical protein LOC320135 28 98 partner of NOB1 27 99 unnamed protein product 33
100 fibronectin 161 101 PREDICTED: hypothetical protein LOC72785 12 102 catenin, delta 1 isoform 2 103 103 zinc finger RNA binding protein 114 104 ribosomal protein L22 15 105 unnamed protein product 22 106 mCG120690 104 107 small GTPase Rah 29 108 mCG11166 18 109 cytoskeleton-associated protein 4 55 110 RanBP2 protein 140 111 tankyrase 1 binding protein 1 113 112 unnamed protein product 22 113 Eif3b protein 109 114 unnamed protein product 25 115 high mobility group nucleosomal binding domain 1, isoform CRA_c 10 116 ribosomal protein L10 25 117 proteasome (prosome, macropain) subunit, alpha type 7 28 118 unnamed protein product 19 119 PAK/PLC-interacting protein 1 42 120 PREDICTED: similar to Transcription factor COE1 64 121 mCG5639 54 122 kelch-like 34 71
65
123 mCG1044286 5 124 Rps13 protein 16
Table 2: Proteins identified by mass spectrometry from FliI immunoprecipitates
# Identified Protein M. W. (kDa)
1 Non-muscle myosin IIA 226 2 Heat shock protein 8 71 3 Phosphoglycerate mutase 1 29 4 mCG144006 55 5 Beta actin 42 6 mCG20427 320 7 Unnamed protein product 39 8 Unnamed protein product 50 9 Unnamed protein product 42
10 Heat shock protein HSP 90-beta 83 11 PAK1 61 12 Immunoglobulin heavy chain variable region 13
66
Future Directions
Our data provide new insights into the role of FliI in cell migration. However, there are several
unresolved issues. The current results have focused on the role of FliI in cell migration and cell
adhesion using siRNA knockdown. A useful, complementary approach would analyze cell
migration and adhesion in cells with higher levels of FliI expression. This approach could be
achieved by transfecting cells with a FliI expression plasmid and then conducting the migration
and adhesion assays described above. We expect that over-expression of FliI would inhibit cell
migration and enhance adhesion.
We found that FliI impacts focal adhesion function by changing the levels of activated β1
integrins. To further understand the role of FliI in focal adhesion function, immunostaining of
control and FliI knockdown cells could be used to assess the effect on other focal adhesion
proteins such as talin, vinculin, and paxillin. TIRF microscopy would then be used to assess the
levels of each protein at focal adhesions. It is expected that FliI knockdown cells will exhibit
reduced talin, vinculin, and paxillin abundance in focal adhesions.
So far the actin binding functions of FliI have not been determined. These functions could be
established by the conduct of several biochemical assays. In the first assay, actin assembly would
be studied using incorporation of rhodamine actin monomers. The increase of fluorescence will
be measured microphotometrically (Arora et al, 2004). If there is enhanced monomer
incorporation into actin filaments by FliI, this would support a role for FliI in actin assembly.
Second, actin severing assays could be used to determine whether FliI has severing activity in
vitro. Briefly, rhodamine phalloidin is added to actin filaments and the rate of fluorescence loss
is measured at 570 nm fluorimetrically (Allen and Janmey, 1994). We will determine if reduction
67
of fluorescence is caused by ability of FliI to sever actin and displace phalloidin from labeled
filaments.
We found that knockdown of FliI reduces the fluorescence attributable to phalloidin-labeled
actin filaments in microscope analyses. We can more accurately quantify actin filament content
using a phalloidin extraction assay (Howard and Oresajo, 1985). This assay is based upon the
specificity of phalloidin binding to actin filaments and the solubility of phalloidin in methanol.
Control and FliI knockdown cells will be stained with FITC-phalloidin. The bound FITC-
phalloidin will be extracted in methanol and the relative fluorescence intensity of the solution
will be determined. Further, we will determine whether FliI impacts the mobility of actin
filaments. Control and FliI knockdown cells will be transfected with GFP-Lifeact (Riedl et al,
2008) and FRAP will be used to determine the fluorescence recovery of GFP-Lifeact. If FliI
stabilizes actin filaments, the % recovery of GFP-Lifeact will be higher in FliI knockdown cells
compared to cells treated with an irrelevant siRNA.
Finally, we could establish more fully the functional association of FliI with mitochondria. We
will determine the role of FliI on the mitochondrial fusion-fission equilibrium by manipulating
the levels of FliI in the cell with FliI siRNA or an expression plasmid. The cells will be stained
with mitotracker red and mitochondrial morphology will be assessed by confocal microscopy.
We will also identify the possible relationship between mitochondria and actin cytoskeleton
mediated by FliI. The levels of FliI in the cell will be manipulated and mitochondrial fractions
will be isolated by differential centrifugation. We will immunoblot those fractions for prohibitin
and β-actin. If FliI acts as a scaffold to anchor mitochondria to actin filaments, then we expect
that the mitochondrial fractions from FliI knockdown cells will exhibit reduced actin content.
68
Similarly, mitochondrial fractions from cells that over-express FliI should exhibit more actin
because FliI may provide a linkage between mitochondria and actin filaments.
The impact of mitochondria on cell migration could also be determined. Cells will be treated
with CCCP (carbonyl cyanide m-chlorophenyl hydrazone), which dissipates the mitochondrial
membrane potential. When combined with the in vitro scratch assay, we should be able to
determine the effect of mitochondrial dysfunction on cell migration.
69
Conclusion
Flightless I may regulate cell migration by interacting with actin monomers and mitochondria,
thereby affecting cell adhesion. An in-depth understanding of cell migration mechanisms could
suggest novel strategies for treatment of diseases such as fibrosis, arthritis, and vascular disease.
70
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