NESPRIN-2G and CELL-MATRIX ADHESIONS INFLUENCE CELL MIGRATION DURING SKIN WOUND HEALING by ALEXANDRA VICTORIA WOYCHEK a Dissert

NESPRIN-2G AND CELL-MATRIX ADHESIONS INFLUENCE CELL MIGRATION

DURING SKIN WOUND HEALING

ALEXANDRA VICTORIA WOYCHEK

A dissertation submitted in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

WASHINGTON STATE UNIVERSITY School of Molecular Biosciences

MAY 2019

To the Faculty of Washington State University:

The members of the Committee appointed to examine the dissertation of ALEXANDRA

VICTORIA WOYCHEK find it satisfactory and recommend that it be accepted.

Ryan Driskell, Ph.D., Co-Chair

Jonathan Jones, Ph.D., Co-Chair

Kwanhee Kim, Ph.D.

Michael Konkel, Ph.D.

Gary Wayman, Ph.D.

ii ACKNOWLEDGMENT

I would like to thank those who have aided my research projects. My advisor Dr.

Jonathan Jones has provided invaluable feedback regarding experimental designs and composition of written documents. Dr. Susan Hopkinson has provided valuable assistance in the design and implementation of molecular biology experiments.

iii NESPRIN-2G AND CELL-MATRIX ADHESIONS INFLUENCE CELL MIGRATION

DURING SKIN WOUND HEALING

Abstract

by Alexandra Victoria Woychek, Ph.D. Washington State University May 2019

Co-Chairs: Ryan Driskell and Jonathan Jones

Human skin is essential for separating internal organs from environmental, physical, and biological insults. The skin is composed of two layers, the epidermis and dermis, which are composed of primarily keratinocytes and fibroblasts, respectively. The epidermis prevents water loss and invasion of pathogens while the dermis plays a supportive role and gives skin its stretchy property. Damage to either of these layers initiates wound healing in which the cells of the epidermis and dermis migrate into the area to restore skin function. Wound healing in both layers begins with the infiltration of migrating cells into the wound. Cell migration is mediated by a variety of proteins, including the nucleus, actin cytoskeleton, and cell-matrix adhesions. The nucleus/actin cytoskeleton interaction is important in nuclear positioning and proper orientation during cell migration. Cell-matrix adhesions allow cells to adhere to the underlying matrix and to migrate across this matrix during cell migration. Here, we explore two cell-matrix adhesions: focal adhesions (FAs) and hemidesmosomes (HDs). We show that loss of a nuclear protein, nesprin-2G, leads to an abnormal actin cytoskeleton and FA formation in primary mouse fibroblasts. This loss of FA formation leads to inability of the knockout cells to tug on their

iv underlying matrix to generate traction forces used during cell migration. Consequently, the loss of nesprin-2G results in decreased cell migration compared to wildtype fibroblasts. However, these results are not replicated in keratinocytes. The formation of these adhesions is crucial, but it is also important to understand how these protein components assemble. To better understand this mechanism, we explored the role of the α6 integrin message in α6β4 integrin localization to

HDs. We show that a sequence within the 3’ untranslated region (UTR) of the α6 integrin message is required for localization of α6β4 integrin to HDs in A549 cells. Sequences lacking this 3’UTR allow the formation of α6β4 integrin, but it is not localized to HDs. Taken together, these results imply that adhesion formation is a tightly regulated processes which is imperative for cell attachment and migration.

v TABLE OF CONTENTS

Page

ACKNOWLEDGMENT...... iii

ABSTRACT ...... iv

LIST OF TABLES ...... vii

LIST OF FIGURES ...... viii

CHAPTER ONE: INTRODUCTION ...... 1

CHAPTER TWO: NESPRIN-2G KNOCKOUT KERATINOCYTES DISPLAY NORMAL CELL MIGRATION AND PROTEIN EXPRESSION ...... 54

CHAPTER THREE: NESPRIN-2G KNOCKOUT FIBROBLASTS EXHIBIT REDUCED MIGRATION, CHANGES IN FOCAL ADHESION COMPOSITION, AND REDUCED ABILITY TO GENERATE TRACTION FORCES ...... 72

CHAPTER FOUR: THE α6 INTEGRIN 3’UTR REGULATES LOCALIZATION OF THE α6β4 INTEGRIN HETERODIMER ...... 104

CHAPTER FIVE: CONCLUSIONS AND FUTURE DIRECTIONS ...... 127

vi LIST OF TABLES

Page

CHAPTER 3

3.1: Primer sets used for genotyping...... 92

3.2: Primer set used for message presence...... 92

3.3: Quantification of FA characteristics ...... 92

3.4: Quantification of FA characteristics after MG132 incubation ...... 93

vii LIST OF FIGURES

Page

CHAPTER 1

1.1: Skin structure ...... 36

1.2: Full- and partial-thickness wounds in human skin ...... 37

1.3: Nesprins at the nuclear envelope ...... 38

1.4: Cell-matrix adhesions protein complexes ...... 39

CHAPTER 2

2.1: Confirmation of KO keratinocytes ...... 66

2.2: Cell motility of WT and KO keratinocytes ...... 67

2.3: Immunofluorescence staining of WT and KO keratinocytes ...... 69

2.4: Flow cytometric analyses of integrin surface expression of WT and KO keratinocytes ...... 71

CHAPTER 3

3.1: Nesprin-2G knockout fibroblasts exhibit abnormal FAs ...... 94

3.2: Nesprin-2G knockout fibroblasts exhibit reduced FA protein levels and altered FA message levels ...... 96

3.3: Nesprin-2G knockout fibroblasts show decreased collective cell migration in a scratch assay and single cell motility ...... 97

3.4: Nesprin-2G knockout fibroblasts generate less traction forces (TFs) than their wildtype counterparts ...... 98

S3.1: Verification of nesprin-2G knockout ...... 99

S3.2: MG132 incubation restores FA staining in knockout cells ...... 101

viii S3.3: Nesprin-2G knockout affects nuclear staining and transmembrane actin-associated nuclear (TAN) line formation ...... 103

CHAPTER 4

4.1: The α6 integrin 3’UTR contains a zipcode which localizes the α6β4 integrin heterodimer to cell-matrix adhesions ...... 120

4.2: Overexpression of the α6 integrin 3’UTR impacts α6β4 integrin adhesions in transduced A549 cells ...... 122

4.3: The α6 integrin 3’UTR is responsible for α6β4 integrin heterodimer formation in iHEKs .123

S4.1: α6-GFP protein products interact with β4 integrin and are soluble regardless of whether their mRNAs possess the α6 integrin 3’UTR ...... 125

CHAPTER 5

5.1: Representative REMSA ...... 136

CHAPTER ONE: INTRODUCTION

Overview

Skin, the outermost layer of the body, maintains a barrier between the body and environment and is thus exposed to physical, environmental, and biological insults. The two skin layers, namely the epidermis and dermis, have different functions to prevent invasion from the environment. The epidermis acts as a barrier to prevent water loss and the infiltration of pathogens, whereas the dermis has a supportive function that gives skin its “stretchy” quality.

Damage to either of these layers can disrupt normal functioning. Epithelial cells respond to such damage by infiltrating the wound to re-establish the barrier function of the skin by means of a process that involves both the proliferation and migration of living cells in the surrounding area.

Within the dermis, cells secrete factors that are responsible for filling the open wound area. In both skin layers, wound healing is mediated by cell-matrix adhesions, the nucleus, and the actin cytoskeleton. Adhesion components promote the secretion of extracellular matrix across which cells migrate. The hemidesmosome and focal adhesion are two well-studied cell-adhesion complexes, but they have different functions and localize differently. The nucleus/actin cytoskeleton interaction is crucial for maintaining nuclear positioning and promotes adhesion formation during cell migration. Therefore, cell migration is a highly complex and finely tuned process that requires the interaction of a variety of protein components.

The introduction of this dissertation describes the structure and function of skin, the process of wound healing, the effect of a nuclear protein on cell migration, adhesion formation, and messenger RNA localization. As stated above, both skin layers contribute to wound healing.

The second and third chapters of the dissertation will explore the roles of keratinocytes and fibroblasts in wound healing, respectively. The fourth chapter aims to add to the growing

literature concerning the localization of messenger RNAs and integrin heterodimer formation in epithelial cells.

Skin structure

Skin is the largest organ of the human body and is composed of two distinct layers, the epidermis and dermis, together with appendages, including glands and hair follicles (Figure 1).

The primary function of this organ is to keep internal organs separated from the exterior environment. The epidermis protects the body from water loss, environmental insult, and mechanical forces (Baroni et al., 2012), whereas the dermis provides an underlying, flexible, support structure that contains blood vessels and nerves (Thulabandu et al., 2018). These two layers are connected via the basement membrane, which forms the dermal-epidermal junction.

Both basal keratinocytes and fibroblasts produce extracellular matrix components that contribute to this region (Sorrell and Caplan, 2004).

Epidermal layer

The epidermis is composed of many cell types including melanocytes, Langerhans cells,

Merkel cells, and, most notably, keratinocytes. Whereas melanocytes, Langerhans cells, and

Merkel cells are found at the basal surface of the epidermis, keratinocytes are found throughout the epidermis in both the basal and suprabasal layers. Melanocytes are responsible for pigment production in the skin (Hirobe, 2005). Langerhans cells are dendritic leukocytes found within the epidermis and are involved in immune responses of the skin (Romani et al., 2003). Finally,

Merkel cells are the mechanoreceptors of the skin (Tai, 2013).

Multiple layers of keratinocytes create a stratified epithelium that regenerates every 40-

56 days in humans. The regeneration capacity of the epidermis is controlled by epidermal stem cells, which are interspersed between basal keratinocytes. Epidermal stem cells are responsible

for the production and rapid turnover of keratinocytes, and as these stem cells differentiate, they first become basal keratinocytes and later differentiate into suprabasal keratinocytes (Koster,

2009). Basal keratinocytes are responsible for anchoring the epidermis to the dermis at the basement membrane via cell-matrix adhesions, including hemidesmosomes and focal adhesions.

Basal keratinocytes also have cell-cell junctions that are important for the barrier function of skin. When basal keratinocytes differentiate and move upward to become the thicker spinous layer, they lose their cell-matrix adhesions but retain cell-cell contacts. Cells in the spinous layer gradually flatten during organelle degradation. The uppermost cornified keratinocytes produce lipids and proteins that protect the body from water loss, and dead keratinocytes are eventually sloughed off into the environment (Chamcheu et al., 2011).

A defining component of a keratinocyte is the overwhelming presence of keratin intermediate filaments; hence these cells are termed keratinocytes. Keratin intermediate filaments are composed of one acidic keratin (type I) and one neutral or basic keratin (type II) that dimerize. Interestingly, all the genes encoding type I keratins are found in a similar location on human chromosome 17, and the genes encoding type II keratins are found close together on human chromosome 12. The dimers come together side-by-side to form a stable tetramer that then associates end-to-end to form protofilaments. Eight protofilaments attach end-to-to-end to form a keratin filament (Chamcheu et al., 2011).

As keratinocytes differentiate, they express different keratin heterodimers that can be used as markers for different cell types. Keratins 5, 14, and 15 are markers of basal keratinocytes, and mutations in these genes can result in the skin blistering disorder, epidermolysis bullosa. Keratins 6, 16, and 17 can be found in cells of the spinous layer. Keratins

1, 2, 9, and 10 are present in the uppermost layer of keratinocytes, and mutated genes for these keratins can lead to keratinophathic ichthyosis diseases (Chamcheu et al., 2011; Pan et al., 2013).

Dermal layer

The dermis provides structural support for the epidermis and is organized into an upper papillary dermis and lower reticular dermis. The three cell types found within the dermis include fibroblasts, macrophages, and adipocytes. Additionally, the dermis is composed of extracellular matrix, blood vessels, and hair follicles. Fibroblasts are the primary support cells of the dermis and produce extracellular matrix, including collagen and fibronectin, which give the structure of the dermis (Sorrell and Caplan, 2004). More than one type of fibroblast is found within the dermis, and their genetic lineages have recently been traced (Driskell and Watt, 2015). The three types of fibroblasts include: papillary fibroblasts, which are found in the upper (papillary) dermis, reticular fibroblasts, which are found in the lower (reticular) dermis, and hair-follicle- associated fibroblasts. These fibroblasts reside in different extracellular matrix environments.

Papillary fibroblasts form thin, poorly organized, collagen fiber bundles, whereas reticular fibroblasts form thick, well-organized fibers within the extracellular matrix (Sorrell and Caplan,

2004). Another cell type of the dermis, macrophages, are immune cells that detect and kill microbes crossing the skin barrier. Moreover, adipocytes have been historically thought merely to support the basal surface of the dermis, they have recently been shown to have metabolic functions (Rivera-Gonzalez et al., 2014).

Wound healing

The process of wound healing restores function to damaged areas of the body. When organs are injured, the affected cells are removed, and immune cells infiltrate the wound to prevent infection from environmental microbes. Typically, a temporary blood clot is formed to

re-establish a barrier. Meanwhile, healthy cells begin the process of proliferation and migration into the wounded area.

Skin is active in body temperature regulation, sensation, and water retention (Hopkinson et al., 2014) and, being the outer layer of the body, is exposed to a variety of insults, including the infiltration of environmental pathogens and abrasions. After wounding, these insults can perturb the normal functions of the skin, and various degrees of wounds can form: partial- or full-thickness wounds (Figure 2). Partial-thickness wounds primarily involve the epidermis and may or may not damage the dermis. These wounds are filled by a process known as re- epithelization, which is initiated from stem cells within the sweat glands and pilosebaceous units of the epidermis (Levy et al., 2005; Rousselle et al., 2018). Full-thickness wounds involve injury to both the epidermis and dermis. In such cases, the dermal wound must first be filled by generating granulation tissue before the occurrence of re-epithelialization (Rittié, 2016), which begins at the wound margins, because glands within the epidermis are destroyed (Rousselle et al., 2018).

Human skin utilizes an intertwined cascade of mechanisms including blot clot formation, inflammation, re-epithelization by keratinocyte infiltration, granulation tissue formation, neovascularization, and tissue contraction to heal wounds in three phases: inflammation, tissue formation, and tissue remodeling (Rittié, 2016; Rousselle et al., 2018). These mechanisms involve interactions between the epidermis and dermis and primarily involve cell migration and proliferation.

Cell migration occurs when a cell responds to environmental cues and polarizes. This can be in response to a variety of factors including growth factors (chemotaxis) or substrate rigidity

(durotaxis). A migrating cell first protrudes its membrane, forming a lamellipodium and new

cell-matrix adhesions. These adhesions allow the cell to pull itself across the substrate. This is a complicated process that requires many protein components and signaling molecules.

Epidermal closure

The process of re-establishing the epidermal barrier after injury is known as re- epithelialization and begins 16-24 hours after injury (Rousselle et al., 2018). Immediately after injury, neutrophils, monocytes, and macrophages are recruited to the wound to initiate an immune response (Barrientos et al., 2008). Keratinocytes then become activated by various growth factors that induce the cells to polarize and migrate across the underlying matrix.

Activated keratinocytes undergo changes to their surface receptors and cytoskeletal elements, and tumor necrosis factor-α maintains keratinocytes in the activated state (Nickoloff and Turka,

1993; Rousselle et al., 2018). Next, lymphocyte signaling induces the expression of keratin 17, which enables keratinocytes to contract the underlying matrix (Jiang et al., 1994; Rousselle et al.,

2018). The underlying dermal fibroblasts secrete transforming growth factor-beta, which prevents keratinocytes from differentiating. As the keratinocytes migrate across the dermal- epidermal junction, they produce matrix metalloproteinase-9, which degrades parts of the matrix for more efficient migration (Michopoulou and Rousselle, 2015; Thomas et al., 2001).

Migration in epidermal closure

To begin the process of re-epithelialization, keratinocytes must completely reorient their actin cytoskeleton and dissolve anchoring protein components to prepare keratinocytes for migration. Rather than moving as individual cells across the wound bed, keratinocytes move as a sheet that contains leader and follower cells with distinct functions and characteristics. Leader cells, the first row of cells of the sheet that moves towards the wound, are responsible for controlling the directed migration of the entire sheet. The cells behind the leader cells in the

sheet are follower cells, and follower cells respond to signals and physical cues from the leader cells to determine where and how the follower cells will move (Theveneau and Linker, 2017).

When keratinocytes are induced to migrate, they elongate and flatten to produce lamellipodia, which are cytoplasmic projections formed in the direction of cell migration

(Rousselle et al., 2018). During the early phase of re-epithelialization, keratinocytes are unaffected by mitotic inhibitors indicating that keratinocytes first migrate into the wound area before proliferating; these two processes do not occur simultaneously. However, later in re- epithelialization, mitosis is induced in follower cells to help push the leader cells towards the wound bed (Rousselle et al., 2018). Recently, rapidly migrating keratinocytes have been shown to experience high tension at the rear of the cell thereby allowing the disassembly of cell-matrix adhesions as cells migrate into the wound bed (Zhao et al., 2018). Basal keratinocytes are contact-inhibited in the wound bed via actomyosin contractions which re-establishes the barrier function of the skin (Hirata et al., 2017).

A variety of models have been proposed to describe this re-epithelization because of the variable results observed in different organisms and wound systems. One model has been proposed suggesting that, as the basal keratinocytes migrate into the wound, they drag suprabasal keratinocytes with them in a confluent layer (Buck, 1979; Radice, 1980). In a second model, suprabasal keratinocytes are thought to migrate to the leading edge of the wound and then to de- differentiated into basal keratinocytes that can subsequently act as leader cells to pull the suprabasal cells into the wound bed (Krawczyk, 1971; Usui et al., 2005). A third model involves the follower basal keratinocytes having proliferative and/or migratory behaviors that pull newly formed basal keratinocytes into the wound bed, while the suprabasal keratinocytes remain together at the wound edge (Headon, 2017; Park et al., 2017; Rousselle et al., 2018).

Proliferation and differentiation in epidermal closure

Since suprabasal keratinocytes have lost the ability to proliferate (Pastar et al., 2014), it is the proliferation of basal keratinocytes within the sheet during wound closure that generates the cells needed to fill a wound. The leader cells within the sheet do not proliferate themselves.

Rather, interfollicular stem cells give rise to keratinocytes that fill the wound. (Aragona et al.,

2017).

Once the basal keratinocytes fill the wound, they experience contact inhibition that causes them to begin proliferating and differentiating upward to complete the healing process.

Keratinocytes become stratified and reform all of the suprabasal layers of the skin (Pastar et al.,

2014).

Dermal closure

A full-thickness wound is defined as a wound that damages both the dermis and epidermis. This type of wound can be detrimental to an organism if not healed as it allows the external environment to interact with the inside of the body. This broken barrier can lead to the invasion of microbes that might be hazardous for the organism. Unsurprisingly, these wounds require the orchestration of many more cells and signaling pathways than partial-thickness wounds. Such wounds require the dermis to be repaired before re-epithelialization can occur.

Cross-talk between keratinocytes and fibroblasts is crucial for wound healing initiation.

Upon wounding, keratinocytes release interleukin-1, which stimulates fibroblasts to release keratinocyte growth factor/fibroblast growth factor-7 and interleukin-6, which in turn induces the proliferation of keratinocytes after wounding (Grossman et al., 1989; Shephard et al., 2004;

Werner et al., 2007).

After wounding, the damage to capillaries induces the formation of a blood clot that primarily contains fibrin and fibrinogen (Darby et al., 2014). The larger the wound, the longer the clot takes to form, leading to increased healing times compared with smaller, less invasive wounds. This blood clot allows other immune cells to be recruited to the wound where they then release a variety of other chemokines that aid the inflammatory response and removal of environmental microbes (Barrientos et al., 2008). Additionally, fibroblasts are recruited to this provisional matrix to begin filling the wound (Darby et al., 2014). Once this provisional matrix is formed, keratinocytes dissolve their cell-cell and cell-matrix contacts, a process that allows them to polarize and migrate across the matrix to re-epithelialize the wound.

Fibroblast role in dermal wound healing

New fibroblasts must be recruited to the wound area because wounded fibroblasts are less proliferative than unwounded fibroblasts (Schäffer et al., 1997). Fibroblasts recruited to the wound area only begin proliferating once blood flow is restored; this proliferation is induced by nerve growth factor (Chen et al., 2014; Darby et al., 2014). Next, fibroblasts develop granulation tissue islands at the base of hair follicles (Bishop, 1945). Granulation tissue is a temporary matrix used to re-establish the dermis and contains fibroblasts, immune cells, and blood vessels supported by a loose matrix consisting of both collagen and fibronectin. In full-thickness wounds, both reticular and papillary fibroblasts infiltrate the wound to restore the dermis, but this migration into the wound bed is not dependent on nerve growth factor (Chen et al., 2014;

Driskell et al., 2013). Reticular fibroblasts from the wound edge are immediately recruited to the wound, whereas papillary fibroblasts are recruited to the wound only during re-epithelialization

(Driskell et al., 2013).

Fibroblasts are responsible for secreting the extracellular matrix required to form the granulation tissue. Once the preliminary granulation tissue is formed, fibroblasts differentiate into proto-myofibroblasts that form more mature adhesion complexes and can lay down the first collagen bundles (Hinz, 2010; Rittié, 2016). Proto-myofibroblasts contract the surrounding extracellular matrix by means of F-actin stress fibers pulling on the surrounding matrix. The tension thus created in the cell then recruits alpha-smooth muscle actin, which causes these cells to differentiate into myofibroblasts (Hinz et al., 2001a; Hinz et al., 2001b). A second marker of myofibroblasts is the increased responsiveness to TGF-β attributable to the loss of nuclear factor kappa-light-chain-enhancer of activated B cells (NF-κB) (Bitzer et al., 2000; Shephard et al.,

2004; Werner et al., 2007). Because the myofibroblasts tug on the extracellular matrix, the granulation tissue stiffens over time (Hinz, 2010). Contraction of the granulation tissue is only partially understood (Rittié, 2016). The contraction of collagen fibers is performed by myofibroblasts shortening a collagen fiber, cleaving fiber loops, and repetition of this process

(Castella et al., 2010).

The transition from granulation tissue to dermis requires drastic changes in extracellular matrix components and the apoptosis of excess myofibroblasts (Darby et al., 2014; Desmoulière et al., 1995). Whereas the granulation tissue primarily contains collagen I, dermis contains collagen III and elastin (Darby et al., 2014). Once this transition is complete, nerves innervate the tissue to re-form the supporting skin layer.

As described above, the skin is an essential organ that prevents the invasion of environmental microbes and protects internal organs from environmental hazards. The two layers of skin, the epidermis and dermis, each have specific functions that aid the other in both homeostatic and wounded conditions. The dermis is supportive tissue that contains fibroblasts,

nerve endings, immune cells, adipocytes, and extracellular matrix. This support structure is required for the epidermis to form properly, as keratinocytes from the epidermis cannot heal a wound without a functioning dermis. Although dermal fibroblasts and epidermal keratinocytes have different functions, they also possess similar characteristics. Both cell types contain cell- matrix adhesion protein complexes, produce extracellular matrix, and have a variety of cytoskeletal interactions.

Human vs. mouse skin wound healing

Researchers typically use animal models to study human disorders and diseases. Mice are often used in the field of wound healing. However, many characteristics that contribute to skin wound healing differ between humans and mice. Human skin is thicker than mouse skin, and mice have loose skin and the presence of a panniculus carnosus muscle found below the dermis

(Zomer and Trentin, 2018). Contraction of this muscle in mice closes wounds about 50% within a day of wounding, and 90% of full-thickness wounds heal by contraction in mice (Fang and

Mustoe, 2008; Rittié, 2016). Mice have more hair follicles than humans, and researchers have shown that human skin with more dense hair follicles heal more quickly than skin sections with fewer hair follicles. Additionally, mice can regenerate hair follicles into adulthood, whereas adult humans do not have this ability (Martinot et al., 1994; Zomer and Trentin, 2018). Although mice can be used to study wound healing in skin, the limitations of this organism must be taken into consideration.

Procedures can be performed on mice which allow researchers to mimic human wound healing in mice. One of these protocols utilizes a stint to keep the wound open by inhibiting the contraction of the panniculus carnosus muscle. Alternatively, researchers can decide to wound the scalp, ear, or tail of the mouse, i.e., in regions where the underlying cartilage or bone prevent

wound contraction. Nevertheless, wound healing studies in mouse skin are not always directly applicable to human skin wounding, although mice remain a valuable tool for studying this system (Zomer and Trentin, 2018).

Nesprin-2G

One of the many proteins found in the skin are the nesprin family members. Nesprins are anchored to the nucleus and have important roles in cell migration, proliferation, and signaling.

The precise mechanisms by which nesprins perform these functions are unknown. One possibility is that these functions are mediated by the interactions of nesprins with the actin cytoskeleton, intermediate filaments, and microtubules (Figure 3). Nesprins, together with other protein components, compose the linker of the nucleoskeleton and cytoskeleton (LINC) complex, which influences nuclear morphology, chromatin remodeling, and cell signaling.

Nesprins

Nesprins (nuclear envelope spectrin repeat proteins) were originally discovered to localize to the nuclear membrane in all tissues (Duong et al., 2014; Zhang et al., 2001). Four nesprin family members (-1G, -2G, -3, and -4) occur in vertebrates, and each of these are encoded by separate genes (SYNE1, SYNE2, SYNE3, and SYNE4, respectively). Multiple isoforms of each nesprin protein are generated by alternative splicing, alternative transcription start sites, and alternative transcription termination sites. The regulation of these transcriptional processes allows tissue-dependent isoform expression (Rajgor et al., 2012). Nesprins-1G and 2G are giant proteins with nesprin-1G having a molecular weight of ~1000 kDa and nesprin-2G having a molecular weight of ~800 kDa. Indeed, the G in their names stands for ‘giant’. In contrast, nesprin-3 is ~110 kDa, and nesprin-4 is ~42 kDa (Rajgor et al., 2012; Roux et al., 2009;

Wilhelmsen et al., 2005).

All nesprins are type II transmembrane proteins, which are single pass proteins with the carboxy terminus anchored to the nuclear membrane and the amino terminus exposed in the cytoplasm. The N-termini of nesprins-1G and -2G have two calponin homology domains that bind actin (Zhang et al., 2001), whereas neither nepsrin-3 nor -4 contains calponin homology domains (Ketema et al., 2007; Roux et al., 2009). Nesprin-3 interacts with plectin at its N- terminus, and nesprin-4 interacts with kinesin microtubule motors. All nesprins contain spectrin homology domains found between the N- and C-termini. However, the number of spectrin repeats varies among proteins. Specifically, the maximum number of repeats for nesprin-1G, -

2G, -3, and -4 are 74, 56, 8, and 1, respectively (Autore et al., 2013; Roux et al., 2009;

Wilhelmsen et al., 2005). A spectrin repeat domain is composed of three α-helices, and these repeats are used to form long, extended molecules which are important in stabilizing the actin cytoskeleton, intermediate filaments, and microtubules. In addition to structural roles, spectrin repeats can also serve signaling roles (Djinovic-Carugo et al., 2002). However, these signaling roles have not been investigated in nesprins.

Nesprin-associated human diseases

Unsurprisingly, because of the importance of the LINC complex in a variety of cell functions, the loss or mutations of these components can lead to human diseases. These diseases include muscular dystrophy, premature ageing, and cancer. Emery-Dreifus muscular dystrophy is a late-onset disease characterized by early joint contractures, muscle wasting, and heart defects; based on studies of fibroblasts isolated from these patients, multiple missense mutations in nesprin-1G and -2G are considered responsible for the nuclear morphology defects that lead to the observed phenotypes (Zhang et al., 2007b). In addition, nesprins have been implicated in aspects of premature human aging (Broers et al., 2006). Specifically, the S143F lamin A/C point

mutation causes a progeria, or premature aging, phenotype. Those progeria-like fibroblasts that have endogenous nesprin-2G expression show a normal nuclear architecture, whereas the progeria-like fibroblasts deficient in nesprin-2G exhibit an abnormal nuclear shape (Kandert et al., 2007). Therefore, the presence of nesprin-2G in these progeria-like cells can compensate for the mutated phenotype seen in progeria-like fibroblasts that lack nesprin-2G.

Evidence has been presented that nesprin-4 deficiency plays a role in hearing loss attributable to disruption of the nuclear movement of hair cells (Cartwright and Karakesisoglou,

2014; Horn et al., 2013). Interestingly, no human diseases have yet been associated with nesprin-

3 (Cartwright and Karakesisoglou, 2014).

The SYNE genes have been found to function in the development of various cancers, specifically lung, breast, ovarian, and colorectal cancers. Hypermethylation of the SYNE1 gene has been demonstrated to be associated with lung cancer (Tessema et al., 2008), and a SYNE1 polymorphism can lead to ovarian cancer (Cartwright and Karakesisoglou, 2014; Doherty et al.,

2010). Mutations within the SYNE2 gene are also associated with human breast and colorectal cancers (Sjöblom et al., 2006).

Nesprin mouse models

The expression of aberrant nesprin proteins has been associated with diseases in mice.

However, whether all the above findings in humans hold true in mice have not been determined, although strong evidence exists that human and mouse LINC complexes are associated with similar pathological conditions. For example, deficiencies in the expression of the mouse LINC complex proteins have been implicated in muscular wasting disorders.

Genetic studies of nesprin protein functions in mice have given a variety of phenotypes depending on the specific nesprin investigated. Generally, nesprin-1G mutant mice have

decreased survival rates, growth retardation, neurogenesis defects, and skeletal/cardiac muscle pathologies (Puckelwartz et al., 2009; Zhang et al., 2007a). Further, the removal of different portions of the protein to generate knockout mice have different phenotypes. However, this may be attributable to differences between mouse strains or the strategies that the researchers used to inactivate genes (Cartwright and Karakesisoglou, 2014). Double knockout mice for the nesprin-

1G/-2G nuclear-anchoring domain die within 20 minutes after birth because of respiratory failure, resulting from their inability to expand their alveoli air sacs. Therefore, nesprins-1G and

-2G have compensatory roles during development (Cartwright and Karakesisoglou, 2014; Zhang et al., 2007a).

Mice lacking only the nesprin-2G nuclear-anchoring domain exhibit severe learning and memory defects. The studies that identified this effect were performed by using rewarded T- maze alternation learning experiments. The mice were tested and then tested again 3 days later.

After the 3 days had passed, 85% of Syne-2+/- mice were able to complete the maze, whereas only 60% of Syne-2-/- mice completed the maze (Zhang et al., 2009).

Surprisingly, nesprin-2G deficient mice lacking the actin-binding domain are viable and do not exhibit any obvious morphological or behavioral abnormalities. However, the knockout mice interestingly show delayed skin wound closure compared with wildtype mice (Lüke et al.,

2008).

Nesprins in the LINC complex

Nesprins-1G and -2G, together with Sad1p-UNC-84 1/2 (SUN 1/2) proteins, make up the evolutionarily conserved linker of the cytoskeleton and nucleoskeleton (LINC) complex found at the nuclear envelope (Chang et al., 2015b). This multi-functional complex regulates numerous biological processes including nuclear morphology, cell signaling, force transduction, and cell

migration. The importance of the nesprin-nucleus association in various cellular functions is detailed below.

The nuclear envelope (NE) is a two-layered structure consisting of outer and inner nuclear membranes, termed the ONM and INM, respectively. These layers are perforated by nuclear pore complexes that allow the transport of proteins and signaling molecules into and out of the nucleus. The ONM is contiguous with the rough endoplasmic reticulum, and the INM contains membrane proteins that associate with the underlying lamina and chromatin. The lamina is the structural scaffold of the NE and is primarily composed of lamins, from which its name is derived (Broers et al., 2006).

Nesprin C-termini directly interact with the ONM. They are anchored to the INM by their

Klarsicht, ANC-1, and Syne Homology (KASH) domain, which interacts with SUN1/2 proteins within the inner membrane (Padmakumar et al., 2005; Zhou et al., 2012). SUN proteins in the complex interact with the nuclear lamina via lamins, emerin, and small INM nesprins, which are small nesprins capable of crossing the NE (Haque et al., 2006; Rajgor and Shanahan, 2013;

Zhang et al., 2005). The function of the KASH domain in mediating the association of the complex with the nuclear envelope is critical, since the loss of the nesprin KASH domain results in cytoplasmic nesprin localization (Zhang et al., 2001).

Interestingly, the predominance of nesprin proteins at the NE can change as myogenesis proceeds. In immature muscle fibers, nesprin-1G is predominantly found at the NE, but as muscle fibers mature, nesprin-2G partly replaces nesprin-1G at the nuclear envelope, and short nesprins become more apparent. However, this switch does not occur with SUN proteins (Natalie et al., 2010). This implies that nesprin proteins at the NE can change with the cell state.

The LINC complex influences cell behavior

Components of the LINC complex play important roles in a variety of cellular functions, including signaling, Golgi apparatus organization, chromatin organization, and cell migration.

Transient siRNA-mediated knockdown of nesprin-2G in HaCat cells, a type of immortalized keratinocyte cell line, induces a decrease in the protein expression levels of β-catenin within the nucleus (Neumann et al., 2010). Since normal Wnt signaling involves the interaction of α- and β- catenins with nesprin-2G at the ONM to enable the efficient transport of β-catenin into and out of the nucleus, the authors propose that the loss of nesprin-2G disrupts this complex, allowing β- catenin to less efficiently translocate into the nucleus (Neumann et al., 2010).

Nesprins-1G and -4 have been implicated in Golgi positioning. In Madin-Darby Canine

Kidney cells, nesprin-1G contains two domains that interact with the Golgi apparatus, and one of these domains acts as a dominant-negative that collapses the Golgi (Gough et al., 2003). HeLa cells do not express nesprin-4 endogenously. However, when its expression is forced by expressing a GFP tagged nesprin-4 plasmid, a loss of peri-nuclear Golgi staining is observed.

Moreover, when green fluorescent protein-tagged nesprin-4 KASH domain is expressed in these cells, no disruption of the Golgi structure occurs (Roux et al., 2009).

Chromatin organization is also regulated by the LINC complex. A histone acetyltransferase (hALP) localizes to the INM and interacts with SUN1 within the INM.

Knockdown of SUN1 in HeLa cells leads to delayed de-condensation of chromosomes and hypoacetylated histones. Therefore, the LINC complex is required for certain aspects of chromatin organization within the nucleus (Chi et al., 2007).

LINC proteins are known to influence cell migration. Loss of nesprin-2G reduces the wound healing ability of both keratinocytes and fibroblasts in 2D scratch wound assays (Lüke et al., 2008; Rashmi et al., 2012). Moreover, the motility of neuronal cells isolated from the

cerebral cortex is reduced in both SUN1/2 double knockout and nesprin-2G knockout mice, compared with controls. In such knockout cells, the nucleus moves significantly less during cell migration compared to their wildtype counterparts (Zhang et al., 2009). This is an important finding because nuclear movement is a crucial step in cell migration, according to the nuclear piston mechanism. Briefly, this mechanism explains how cells migrate in 3D environments by using the nucleus as a barrier between the forward and rear cytoplasmic compartments. Cells use the actin cytoskeleton to pull the nucleus forward, creating a high-pressure compartment in the direction of cell migration. The build-up of this pressure is used to push the cell forward. (Petrie et al., 2014). Interestingly, the loss of SUN2 expression in keratinocytes causes a deficiency in cell-cell adhesion and nuclear positioning; the authors propose that an altered microtubule organization leads to these defects (Stewart et al., 2015).

Nesprins function in traction force generation

Since nesprins affect cell migration, nesprins can be hypothesized to affect the generation of traction forces. Interestingly, both nepsrin-1G and -2G have been shown to play a role in this process. The effect of nesprin-1G in traction force generation has been better studied than the role of nesprin-2G. Nesprin-1G knockout human umbilical vein endothelial cells (HUVECs) exhibit increased traction force generation, increased nuclear height, and the misorientation of the nucleus compared with wildtype cells (Chancellor et al., 2010). Human myoblasts with a mutation in the nesprin-1G gene generate greater traction forces but show a decrease in nuclear height in opposition to the results of the previous study (Schwartz et al., 2017). These conflicting data might be attributable to the use of different cell types, because endothelial cells and muscle cells perform different functions within the body.

In mouse mammary gland carcinoma 4T1 cells, non-muscle myosin IIB has been shown to mediate nuclear translocation and traction force generation, and traction force generation has been suggested to be regulated by an interaction with nesprin-2G (Thomas et al., 2015).

Interestingly, nesprin-2G has been demonstrated to respond to myosin-dependent tension, but its influence on cell migration has not been thoroughly investigated (Arsenovic et al., 2016). One possibility is that a feedback loop exists such that the traction forces generated by myosin influence the transcription of focal adhesion proteins, but this has not been well studied.

Nesprin-2G affects nuclear morphology

A functional nucleus is required for cell viability, and many proteins, including the nesprins, are responsible for the development and maintenance of proper nuclear morphology.

Specifically, nesprins appear to assemble as a meshwork that supports the nucleus. Moreover, the interaction of nesprins with the ONM creates a stabilized nucleus that can react to environmental changes (Lu et al., 2012). In order to assemble a meshwork, nesprins have been shown to oligomerize with nesprins of the same type and with nesprins of different types. Nesprin-1G can also oligomerize with itself via the N- and C-terminal spectrin repeats, and nesprin-3 can oligomerize via the N-terminal spectrin repeats (Ketema et al., 2007, 3; Mislow et al., 2002).

Experimental evidence that nesprins are involved in nuclear organization comes from studies of cells in which various nesprins are knocked out or mutated. One group has created a nesprin-2G knockout mouse in which the actin-binding domain of nesprin-2G is missing; this leads to a complete loss of nesprin-2G in fibroblasts (Lüke et al., 2008). In these knockout mice, the nuclei of keratinocytes become enlarged (Lu et al., 2012; Lüke et al., 2008). Conversely, the overexpression of a small nesprin-2G isoform (termed nesprin-2G mini), which contains the actin- binding domain and the KASH domain but is missing most of the spectrin homology

repeat domains, results in a decreased nuclear size (Lu et al., 2012). Another group has investigated the role of nesprin-2G in mammal retinal development. Photoreceptor nuclei are mislocated in mice with a deletion mutation of nesprin-2G (Yu et al., 2011).

Nesprin-2G has also been shown to be important for transmembrane actin-associated nuclear (TAN) line formation, which helps to both stabilize and move the nucleus (Kutscheidt et al., 2014). In addition, the interaction of nesprin-2G with nesprin-3 at the nuclear envelope has been proposed to maintain nuclear shape (Lu et al., 2012). When the nesprin-2G ABD or KASH domains are overexpressed in HaCat cells with plasmids via electroporation, transfected cells show an increased nuclear size compared with non-transfected cells. In HaCats overexpressing the nesprin-2G mini construct (missing all spectrin homology repeat domains), nuclear size was reduced (Lu et al., 2012). This finding suggests that an interplay occurs between nesprin proteins, including nesprin-2G, which is responsible for maintaining nuclear morphology.

During cell migration, non-muscle myosin induces the contraction of actin arcs.

Chemical inhibition of F-actin polymerization shrinks nuclei, whereas the chemical disruption of microtubules leads to nuclei expansion (Chancellor et al., 2010; Mazumder and Shivashankar,

2010). These actomyosin contractions pull outward on the nucleus, whereas microtubules act to contract the nucleus (Chancellor et al., 2010). Additional forces are exerted on the nuclear envelope by the nuclear lamina and the genome. Specifically, one group has determined that the nucleus is much less pliable when lamins are absent, and these nuclei also exhibit increased chromatin movement (Schreiner et al., 2015). Another group has observed nuclear shrinkage following laser-ablation of the heterochromatin in mouse embryonic fibroblasts. In addition, both actin and microtubule depolymerization was observed. The authors suggest that the connections between the nucleus and cytoplasm are critical for regulating nuclear morphology (Mazumder

and Shivashankar, 2010). The cytoskeleton has also been implicated in nuclear integrity because it can regulate the deformation ability of the nucleus while allowing the nucleus to be mechanosensitive at the same time (Wang et al., 2018).

Nesprin-2G influences transcription

When forces are applied to the nucleus, both the transcription of certain genes and cellular signaling cascades can be induced because the nucleus is able to respond to mechanosensitive cues, resulting from matrix stiffness or other cellular processes (Wang et al.,

2009). Loss of LINC complex proteins has been reported to lead to the disruption of transcription of genes associated with adhesion, motility, and wound healing (Alam et al., 2015). Thus, motility-related force transduction to nesprin-2G might result in the transcription of genes that regulate motility. Changes in nuclear shape can also affect chromatin conformation, which can determine the transcriptional availability of certain genes (Osmanagic-Myers et al., 2015). If chromatin is in the open conformation, transcription factors can interact with the DNA, thus allowing transcription. In contrast, steric hindrance can prevent transcription factors from binding to DNA when chromatin is in the closed conformation (Alam et al., 2015). The finding that the loss of nesprin-2G leads to nuclear morphology defects implies that these defects lead to a reduction in the recruitment of transcription factors required for various cellular processes. For example, wounds in nesprin-2G knockout mice show abnormal levels of PPARβ/δ, E2F1, and

EGR-1 at 1- and 3-days post-wounding, but at 5 days after wounding, no defects in nesprin-2G knockout wounds are apparent (Rashmi et al., 2012).

Nesprin-2G affects cell signaling

In addition to force-mediated changes, the loss of nesprins can also lead to numerous changes in cell signaling pathways. Cells that are deficient in nesprin-2G show a loss of β-

catenin at cell-cell junctions, altered β-catenin transcription levels, and an accumulation of active

β-catenin within the nucleus. This can result in abnormal levels of transcription (Zhang et al.,

2016). Another group has found that the siRNA-mediated knockdown of nesprin-2G or the overexpression of a nesprin-2G dominant-negative construct (missing the KASH domain) causes perturbations of the MAPK1/2 signaling pathway, which can elevate cell proliferation (Warren et al., 2010).

Nesprin-2G and wound healing

Evidence has been presented that nesprin-2G is a positive regulator of wound healing.

One group has shown that deleting the actin-binding domain from nesprin-2G leads to decreased skin wound healing in mice (Lüke et al., 2008). Other work from their laboratory indicates that both isolated primary keratinocytes and fibroblasts from the knockout animals exhibit reduced wound closure in in vitro scratch assays (Lüke et al., 2008; Rashmi et al., 2012). The hypothesis that nesprin-2G affects cell migration is further supported by findings from another group who have demonstrated that fibroblasts lacking nesprin-2G display a similar decrease in cell migration and polarization compared with wildtype cells (Lombardi et al., 2011). Finally, the effects of nesprin-2G on cell migration do not appear to be limited to keratinocytes and fibroblasts. The loss of nesprin-2G in myoblasts (muscle cells) has been reported to lead to reduced directed cell migration (Chang et al., 2015a).

In summary, deficiencies in the expression or mutations of LINC complex proteins, particularly nesprins, can result in a wide range of human diseases. These include neurodegenerative diseases, musculoskeletal diseases, cancer, hearing loss, and premature aging.

The generation of LINC-deficient mouse lines can aid the better understanding of the mechanism underlying these diseases. Perhaps the most poorly understood of the mechanisms driving these

pathological conditions is the role of nesprin-2G in mediating wound healing. As chapter 3 will show, nesprin-2G can influence wound healing by affecting the formation of cell-matrix adhesions, which are discussed in further detail in the next section.

Matrix adhesions of the skin

All adherent cells use cell-matrix adhesions as an anchoring mechanism to attach to the underlying substrate, the extracellular matrix (ECM). These adhesions contain a variety of proteins, including transmembrane and cytosolic components, that interact with the cytoskeleton.

Adaptor proteins are recruited to the complex within the cytoplasm to promote cellular responses to signaling cues that can influence cell behavior and matrix deposition.

The two main types of cell-matrix adhesions are hemidesmosomes (HDs) and focal adhesions (FAs) (Figure 4). FAs have primarily been implicated in cell migration. Historically, research into keratinocyte migration has focused on FAs as the primary adhesion complex utilized in cell migration, whereas HDs have been regarded as important in stable cell adherence.

However, HDs have recently been established also to be crucial for migrating cells (Sehgal et al.,

2006). These two types of transmembrane adhesion complexes contain different protein components, perform different functions, and have different localizations within the cell. HDs have fewer protein components than FAs, as FAs can incorporate 200+ potential proteins. HDs interact with the keratin intermediate filament network, whereas FAs interact with the actin cytoskeleton. This difference allows for different responses to signaling cascades within the cell

(Hopkinson et al., 2014). FAs tend to be clustered at the leading edge of migrating cells, whereas

HDs are typically situated more rearward (Ozawa et al., 2010). However, despite several differences existing between these adhesion complexes, they both contain an integrin heterodimer at their core and interact with ECM.

Cell-matrix adhesions: hemidesmosomes

Hemidesmosomes (HDs) are found in intact human skin and maintain skin integrity under mechanical stress. They anchor basal keratinocytes of the epithelia to the underlying basement membrane. HDs were originally identified by electron microscopy, which revealed that they have a tripartite structure composed of a cytoplasmic plaque, a sub-basal dense plate, and extracellular anchoring filaments that extend into the basement membrane. They obtained their name because they resemble one half of a desmosome, which is found in cell-cell contacts (Jones et al., 1998). True HDs are found in tissue but not in cultured cells, except in 804G cells

(Hopkinson et al., 1998). Cultured cells are thought not to form true HDs, because a lower density of HD proteins occurs in cultured cells than in intact skin (Geuijen and Sonnenberg,

2002).

HDs are assemblies of proteins that contain the α6β4 integrin heterodimer, BP180

(collagen XVII, BPAG2), BPAG1e (BP230), plectin, and laminin-332 (the extracellular binding partner of integrin) (Litjens et al., 2006). The membrane-spanning proteins that anchor the cytoplasmic and extracellular proteins together include the integrin heterodimer α6β4 and

BP180. The cytoplasmic proteins (BPAG1e, plectin) are used to anchor intermediate filaments to the basal lamina, whereas extracellular matrix proteins (laminin-332) are required to anchor the two layers of epithelia together (Hopkinson et al., 2014). Keratin is the intermediate filament (IF) found in the HD and is essential for maintaining tissue integrity when bound to the integrin heterodimer and other proteins (Seltmann et al., 2013). HDs are necessary for cell adhesion but also have functions in cell motility, proliferation, differentiation, and signal transduction via both inside-out and outside-in signaling pathways (Hopkinson et al., 2014).

As cell migration is initiated, the HD is disassembled to reduce interactions with the

ECM. This disassembly is facilitated by the phosphorylation of specific residues within the β4 integrin tail (Castañón et al., 2013; Rabinovitz et al., 2004), and a variety of proteins have been implicated in this process, including epidermal growth factor and protein kinase C family members, to name just two (Rabinovitz et al., 2004). This reduction in cell-matrix interaction allows the cell to migrate.

Type I vs. type II HDs

There are two types of HDs. Type I is present in skin (discussed above), and type II is found in simple epithelia, e.g., intestinal epithelia, astrocytes, Schwann cells, and endothelial cells (Litjens et al., 2006; Sterk et al., 2000; Uematsu et al., 1994). In contrast to type I HDs, type

II HDs are comprised of only α6β4 integrin, plectin, and tetraspanin CD151 (note the lack of BP antigens). Despite type II HDs being present within cell junctions, they are not found in epidermal HDs and have thus been classified as a separate type of HD (Uematsu et al., 1994).

Type II HDs are not associated with intermediate filaments as frequently as type I HDs (Geuijen and Sonnenberg, 2002), and some researchers have hypothesized that type II HDs are a precursor complex to mature type I HDs (Raymond et al., 2007).

The β4 cytoplasmic tail can be phosphorylated in type II HDs, just as in type I HDs, but a different outcome occurs when the β4 of the type II HD is phosphorylated. Unlike type I HDs where the complex disassembles upon β4 phosphorylation in order to promote migration, the

EGF-induced phosphorylation of the type II HD complex in COS-7 cells does not result in increased migration (Rabinovitz et al., 2004). For some reason, the type II HD is more resistant to disassembly-induced migration than the type I HD; this might be related to the observation that cells that express type II HDs are continuously migrating (Rabinovitz et al., 2004).

Integrins

Integrins are transmembrane, heterodimeric proteins composed of one α and one β subunit. In total, 18 α subunits and 8 β subunits have been found that assemble into 24 distinct heterodimers in mammals (Hynes, 2002). α subunits contain a large extracellular N-terminal globular head domain (~1000 amino acids), a transmembrane segment, and C-terminal cytoplasmic tail (Hynes, 2002). These α subunits undergo post-translational cleavage of covalently linked disulfide bridges resulting in heavy and light α integrins (de Melker and

Sonnenberg, 1999). After cleavage, the heavy chain remains extracellular, whereas the light chain comprises the transmembrane region and cytoplasmic tail. β subunits contain a smaller N- terminal extracellular globular head domain (~700 amino acids), a transmembrane segment, and a C-terminal cytoplasmic tail that interacts with cytoskeletal/intermediate filament networks and acts as a signaling hub. The cytoplasmic tail is usually short (~50 amino acids), although the β4 integrin subunit has a cytoplasmic tail longer than 1000 amino acids (Hynes, 2002).

Most integrin heterodimers localize to FAs, and different heterodimers are expressed in the intact and wounded skin (Koivisto et al., 2014). Integrin heterodimers are cell-type- and tissue-specific and interact with different ligands. Interestingly, the α6β4 integrin heterodimer is exclusively found in HDs and HD-like complexes (Colburn and Jones, 2016; Hiroyasu et al.,

2013). Another unique aspect of this heterodimer is that α6 integrin is the only α subunit that can interact with β4 integrin, although α6 integrin can also bind to β1 integrin (Hemler et al., 1989;

Lee et al., 1992).

The α6β4 heterodimer is central to HD formation as other components are added after its recruitment. When either α6 or β4 integrin is mutated or missing, dysadhesion of the epidermal- dermal junction occurs, leading to skin blistering diseases, including junctional epidermolysis

bullosa with pyloric atresia (Tsuruta et al., 2011). Mutations within the β4 integrin gene (ITGB4) are more common than mutations within the α6 integrin gene (ITGA6) but are all inherited in an autosomal recessive manner (Turcan and Jonkman, 2014). Mice failing to express β4 have been shown not to form intact skin layers. Notably, in such mice, HD structures cannot be detected in affected or unaffected tissue (van der Neut et al., 1996). Moreover, young mice with the complete knockout of α6 have been demonstrated to exhibit detachment of the dermal layers

(Georges-Labouesse et al., 1996). α6β4 integrin is also used as a marker for certain cancers. For example, in the prostate, an increase in α6β1 occurs when prostate tissue becomes cancerous and more invasive (Demetriou and Cress, 2004). This increased binding to α6β1 instead of α6β4 is attributable to a post-translational cleavage event that occurs in α6 (Demetriou and Cress, 2004).

Such binding is important because β1 is usually associated with FAs that are utilized for cell motility. α6β4 integrin is also upregulated in some metastatic cancers including melanoma, breast carcinoma, and lung cancers (Hsu et al., 2013; Uematsu et al., 1994).

Cell-matrix adhesions: focal adhesions

Focal adhesions (FAs) are protein complexes found in a variety of cells and tissue types that are important for cell migration, adherence to the underlying substrate, mechanotransduction, and signaling within the cell (Leube et al., 2015). Similar to HDs, FAs contain an integrin core, e.g., α2β1, α2β5, or α3β1, that is responsible for interaction with ECM components, such as collagen, fibronectin, or laminin (Hynes, 2002). However, FAs interact with the actin cytoskeleton and can contain more than 200 types of proteins.

FA maturation is a process by which FAs become more complex and undergo functional changes. After the initial clustering of integrins and their interaction with actin, other proteins are recruited by actin tugging on the FA and thus revealing cryptic binding sites within FA proteins.

These initial complexes are termed nascent adhesions, and their fate can be either disassembly or maturation. As integrins cluster further (Maheshwari et al. 2000, Selhuber-Unkel et al. 2008) and recruit other proteins, these adhesions are termed focal contacts and then form FAs. As FAs mature, they become more stable (less likely to disassemble) and interact with the ECM (Gardel et al., 2010). Integrins can further stabilize FAs by producing more ECM (Gallant et al. 2005,

Palecek et al. 1997).

There are three regions within the FA, and their protein components are discussed in more detail below (Kanchanawong et al., 2010). The integrin signaling layer is closest to the cell membrane and contains integrins and paxillin. The force-transducing layer lies directly above the integrin layer and contains vinculin. The uppermost layer is the actin regulatory layer and contains zyxin and actin filaments (Kanchanawong et al., 2010). Tensin-1 will also be detailed below but was not included in the previous localization studies.

Paxillin

The paxillin gene (PXN) in humans is highly conserved and has 4 isoforms that differ in the N-terminal region (López-Colomé et al., 2017). Isoform 1α is the most prominent and is found throughout the body (López-Colomé et al., 2017; Salgia et al., 1995). The paxillin protein has four LIN-11, Isl1, and MEC-3 (LIM) domains at its C-terminus and five leucine-rich motifs at its N-terminus. LIM domains contain two zinc-finger domains separated by a hydrophobic linker. These domains function in protein-protein interactions and are critical in cytoskeleton organization. The third LIM domain in the C-terminus anchors paxillin to the plasma membrane

(Brown et al., 1996; Smith et al., 2014). The N-terminus interacts with FA-associated proteins including Src family members, focal adhesion kinase (FAK), vinculin, and GTPase-activating proteins (López-Colomé et al., 2017). Various serine and tyrosine residues have been identified

along the protein and promote and inhibit cell signaling for cell migration via interaction with other FA proteins (López-Colomé et al., 2017).

Paxillin is a focal adhesion structural and scaffolding protein that is important in regulating cell polarity and motility (Dubois et al., 2017; Yu et al., 2010). Interestingly, paxillin messenger RNAs are recruited to and locally translated at the leading edge in migrating cells, allowing efficient incorporation into nascent FAs (Woods et al., 2002). The early incorporation of paxillin into FAs is facilitated by FAK (Brown et al., 1996; Hu et al., 2014; Humphries et al.,

2007). Whereas paxillin is primarily found at FAs, it has been localized to the nucleus where it is believed to act as a transcription factor (Dong et al., 2009).

Vinculin

The vinculin protein is made up of a globular head, a proline-rich linker region, and a tail domain. All three domains have been shown to interact with a variety of FA proteins, but the most studied interaction is that of vinculin and talin (Ziegler et al., 2006). Interestingly, vinculin is found in two states, namely activated and inactivated, and the activated form is found in FAs

(Chen et al., 2005). The protein is considered activated when the head and tail regions are no longer in contact, revealing all its ligand binding sites (Ziegler et al., 2006). This initial dissociation of the head and tail regions has many hypothesized pathways, but clearly a substrate must first interact with the inactivated form (Janssen et al., 2006).

Vinculin is recruited to FAs by phosphorylated paxillin (Case et al., 2015). As FAs mature in response to tension, vinculin becomes activated, which further recruits proteins to the

FA (Rio et al., 2009). Vinculin has been shown to have a variety of functions within the FA including the regulation of cell migration, cell spreading, the maturation of FAs, and FA turnover

(Subauste et al., 2004; Ziegler et al., 2006). Loss of vinculin leads to increased cell migration,

decreased cell spreading, and smaller and fewer FAs compared with wildtype cells (Xu et al.,

1998).

Tensin-1

Four tensin family members are found in higher eukaryotes and have been extensively studied with regard to cell adhesion, cell migration, and cytoskeletal rearrangement (Haynie,

2014). Tensins are recruited to FAs after integrin clustering and opening of the β integrin cytoplasmic tail. Once integrated into the FA, tensins act as scaffolding proteins for docking sites of other FA components (Legate and Fässler, 2009). Specifically, tensin-1 interacts with

GTPase-activating proteins (Li et al., 2011) and with protein phosphatase 1α (Eto et al., 2007) and contains three actin-binding domains (Lo et al., 1994).

Tensin-1 knockouts and mutations provide insight into the function of this protein.

Introduction of a tensin-1 mutant that cannot interact with actin inhibits the cell migration of mouse embryonic fibroblasts (Zhao et al., 2016). Tensin-1 protein regulation is dependent on

TGF-β receptor activity. Increased TGF-β receptor signaling leads to increased myofibroblast differentiation, but loss of tensin-1 leads to the loss of FA formation and actin stress fiber formation (Bernau et al., 2016). The same group that reported the latter finding also found that fibronectin formation was dependent on tensin-1 localization to FAs (Bernau et al., 2016).

Overall, tensin-1 is important for the development and stability of FAs through the recruitment of other FA components and the regulation of ECM formation.

Zyxin

Zyxin is an FA protein with a C-terminal LIM domain and an N-terminal α-actinin- binding domain. The LIM domain is important in protein-protein or protein-DNA interactions, and these domains are required for the FA localization of zyxin (Uemura et al., 2011). Through

the α-actinin-binding domain, zyxin interacts with the actin cytoskeleton (Reinhard et al., 1999).

This zyxin/actin interaction allows the FA to participate in the regulation of the cytoskeleton during cell adhesion and migration.

Zyxin is the last protein recruited to FAs and is considered a marker of mature FAs

(Zaidel-Bar et al., 2004). The localization of zyxin to FAs is dependent upon FAK activation and contractile forces generated by the cell during cell migration. The mechanosensitivity of this protein is its most studied property (Smith et al., 2014). The loss of zyxin in fibroblasts leads to increased integrin-dependent adhesion, increased migration speed, and reduced actin filament formation (Hoffman et al., 2006). Another research group provided further support for this mechanism when they overexpressed the FA targeting domain of zyxin; these cells showed a loss of actin polymerization at FAs (Hirata et al., 2008). If cells that have formed mature FAs are placed under high tension to induce actin stress fiber formation, zyxin is reduced at FAs and enhanced at actin filaments, thereby causing the thickening and stability of actin filaments

(Yoshigi et al., 2005). Without this FA/actin linker, cells are unable to form and maintain FAs and the actin cytoskeleton.

Actin cytoskeleton

The actin cytoskeleton is one of the main frameworks within the cell and helps to maintain the shape of a cell by being both rigid and flexible, depending on cell status and signaling. Actin filaments play a role in both the stability of non-migrating cells and the migration of motile cells because of the rigidity and stability of the filament. Rac and Cdc42

GTPases promote actin filament formation, whereas the GTPase Rho aids the formation of actomyosin contractile structures (Nobes and Hall, 1995). Additionally, actin filaments are assembled at the leading edge of the lamellipodium protrusion and disassembled at the other end

of the filament, as needed, while the cell migrates (Ponti et al., 2004). This action not only pushes the migrating cell forward, but also pushes the cytoskeletal network backward, thereby generating the retrograde flow phenomena associated with actin (Forscher and Smith, 1988).

Unfortunately, the disassembly of actin filaments has been much less studied than their formation (Gardel et al., 2010).

The formation of integrin clusters induces actin filaments to interact with FAs, and this interaction allows the cell to pull on the ECM to promote cell migration (Leube et al., 2015).

Actin-binding proteins facilitate the interaction with the FA, and the act of actin pulling on the

FA causes it to recruit more proteins to the complex leading to FA maturation (Gardel et al.,

2010). Additionally, this actin/FA interaction facilitates trafficking along actin filaments, and loss of actin by blebbistatin treatment reverses this defect (Gardel et al., 2010). Therefore, the actin cytoskeleton is imperative for cell adhesion and migration processes.

In summary, a variety of proteins within HDs and FAs are responsible for maintaining skin integrity, both in a relaxed state and when placed under mechanical tension. The formation of these complexes is performed in a step-wise manner with the integrin core undergoing clustering at the membrane to facilitate HD and FA formation. However, the messages of these proteins must be targeted to the site of protein translation, and this specific targeting of messages is outlined in the following section. mRNA localization mRNA structure

The central dogma of molecular biology states that DNA is transcribed to create messenger RNAs (mRNAs), which are then translated into proteins. Other types of RNAs

(microRNAs, transfer RNAs, ribosomal RNAs, etc.) are transcribed, but only mRNAs are

translated into proteins. DNA transcription is a tightly regulated process in eukaryotes, involving transcription factors binding to enhancer and repressor regions within the promoter region of the gene. Post-transcriptional processing creates a mature mRNA transcript by adding a 5’ cap and

3’ poly-A tail and by removing introns. The mature mRNA transcript is composed of three regions: 5’UTR (untranslated region), coding sequence, and 3’UTR. The coding sequence contains the genetic information that will eventually be translated into the mature protein

(Szostak and Gebauer, 2013). The 5’ and 3’ UTRs have separate and overlapping functions depending on the gene, but, generally, these regions are AU-rich and play a role in mRNA stability, degradation, and translation. The 3’UTR is especially important for transcript localization within the cell (Mayr, 2017).

3’UTRs in mRNA localization

Eukaryotic cells are composed of a variety of compartments created from membrane- bound organelles. This separation of space creates a problem for mRNAs, which are transcribed in the nucleus and translated outside of the nucleus. These newly translated proteins are then either incorporated into a membrane or into cytosolic protein complexes. Messages are exported from the nucleus in an inactive state to the cytoplasm. Once an mRNA reaches its site of translation, it becomes activated and begins translation. Generally, proteins are translated near where they will be utilized within the cell. This localized activation allows a concentrated pool of proteins to form in certain cellular locations, and a single mRNA can be translated a multitude of times to enrich a region for certain proteins (Ryder and Lerit, 2018). The 3’UTR has been shown to be one of the main regulators for this switch between inactive to active forms because of the specific binding sequences within the 3’UTR and the 3’UTR’s interaction with other molecules that regulate localized translation.

Zipcodes

The zipcode is a sequence within the 3’UTR that is the smallest required sequence for the cellular localization of an mRNA, and these sequences vary in size from a few nucleotides to one kilobase (Weis et al., 2013). One of the first proteins to have its zipcode identified was β-actin, which was found at the leading edge of migratory cells, specifically at FAs (Katz et al., 2012;

Lawrence and Singer, 1986). These zipcode sequences have also been implicated in other cell processes, including cell polarization, which is a crucial step in cell migration. Cell polarization allows increased protein concentrations locally within cell protrusions at the leading edge of the cell (Mardakheh et al., 2015). Consequently, this zipcode sequence is essential for localized translation that leads to proper cell functioning.

RNA-binding proteins

Two primary classes of molecules directly interact with the 3’UTR: microRNAs, which inhibit translation of the transcript, and RNA-binding proteins (RBPs), which promote the translation of the transcript (Szostak and Gebauer, 2013). RBPs interact with 3’UTR zipcodes either by complementary base pairing and/or by binding the secondary or tertiary structure of the mRNA. This interaction recruits other effector proteins that direct the fate of a transcript. If an mRNA is destined to be translated into protein, RBPs direct this localized translation process by facilitating interaction with microtubule motors that move along the cytoskeleton network

(Gagnon and Mowry, 2011). Commonly, proteins with the same cellular localization are regulated by one or a few of the same RBPs (Mayr, 2017; Ryder and Lerit, 2018). In the case of

β-actin, the zipcode-binding protein-1 (ZBP1) has been identified as the responsible RBP that interacts with the β-actin zipcode to direct its localized translation. Cells lacking ZBP1 do not

localize the β-actin mRNA properly; this leads to reduced FA stability and reduced cell migration (Katz et al., 2012).

Summary

Skin is a complex organ responsible for separating the body’s internal organs from external insults. When the skin is wounded, a variety of cells and cellular components are required to re-establish the barrier function. Wound healing is facilitated by cell migration which is a complex process that involves interactions between many different proteins and signaling components. An actively migrating cell must not only maintain its cell morphology with the support of cytoskeletal networks, but also enable the formation of adhesion complexes at the anterior of the cell. In chapters 2 and 3 of this dissertation, the importance of the formation of cell-matrix adhesions is explored in relation to integrin heterodimers in the model systems of primary mouse keratinocytes and fibroblasts, respectively. In chapter 3, the role of the nucleus in fibroblast cell migration is also investigated. Finally, chapter 4 will describe a novel finding for

α6β4 integrin heterodimer localization to cell-matrix adhesions in epithelial cells.

References

Alam, S. G., Lovett, D., Kim, D. I., Roux, K. J., Dickinson, R. B. and Lele, T. P. (2015). The nucleus is an intracellular propagator of tensile forces in NIH 3T3 fibroblasts. J Cell Sci 128, 1901–1911.

Aragona, M., Dekoninck, S., Rulands, S., Lenglez, S., Mascré, G., Simons, B. D. and Blanpain, C. (2017). Defining stem cell dynamics and migration during wound healing in mouse skin epidermis. Nature Communications 8, 14684.

Arsenovic, P. T., Ramachandran, I., Bathula, K., Zhu, R., Narang, J. D., Noll, N. A., Lemmon, C. A., Gundersen, G. G. and Conway, D. E. (2016). Nesprin-2G, a component of the nuclear LINC complex, is subject to myosin-dependent tension. Biophysical Journal 110, 34–43.

Autore, F., Pfuhl, M., Quan, X., Williams, A., Roberts, R. G., Shanahan, C. M. and Fraternali, F. (2013). Large-scale modelling of the divergent spectrin repeats in nesprins: giant modular proteins. PLOS ONE 8, e63633.

Baroni, A., Buommino, E., De Gregorio, V., Ruocco, E., Ruocco, V. and Wolf, R. (2012). Structure and function of the epidermis related to barrier properties. Clin. Dermatol. 30, 257–262.

Barrientos, S., Stojadinovic, O., Brem, H. and Tomic-Canic, M. (2008). Growth factors and cytokines in wound healing. Wound Repair & Regeneration 16, 585–601.

Bernau, K., Torr, E. E., Evans, M. D., Aoki, J. K., Ngam, C. R. and Sandbo, N. (2016). Tensin 1 is essential for myofibroblast differentiation and extracellular matrix formation. Am J Respir Cell Mol Biol 56, 465–476.

Bishop, G. H. (1945). Regeneration after experimental removal of skin in man. American Journal of Anatomy 76, 153–181.

Bitzer, M., Gersdorff, G. von, Liang, D., Dominguez-Rosales, A., Beg, A. A., Rojkind, M. and Böttinger, E. P. (2000). A mechanism of suppression of TGF–β/SMAD signaling by NF- κB/RelA. Genes Dev. 14, 187–197.

Broers, J. L. V., Ramaekers, F. C. S., Bonne, G., Yaou, R. B. and Hutchison, C. J. (2006). Nuclear Lamins: Laminopathies and Their Role in Premature Ageing. Physiological Reviews 86, 967–1008.

Brown, M. C., Perrotta, J. A. and Turner, C. E. (1996). Identification of LIM3 as the principal determinant of paxillin focal adhesion localization and characterization of a novel motif on paxillin directing vinculin and focal adhesion kinase binding. The Journal of Cell Biology 135, 1109–1123.

Buck, R. C. (1979). Cell migration in repair of mouse corneal epithelium. Invest. Ophthalmol. Vis. Sci. 18, 767–784.

Cartwright, S. and Karakesisoglou, I. (2014). Nesprins in health and disease. Seminars in Cell & Developmental Biology 29, 169–179.

Case, L. B., Baird, M. A., Shtengel, G., Campbell, S. L., Hess, H. F., Davidson, M. W. and Waterman, C. M. (2015). Molecular mechanism of vinculin activation and nanoscale spatial organization in focal adhesions. Nat Cell Biol advance online publication.

Castañón, M. J., Walko, G., Winter, L. and Wiche, G. (2013). Plectin–intermediate filament partnership in skin, skeletal muscle, and peripheral nerve. Histochem Cell Biol 140, 33– 53.

Castella, L. F., Buscemi, L., Godbout, C., Meister, J.-J. and Hinz, B. (2010). A new lock-step mechanism of matrix remodelling based on subcellular contractile events. J Cell Sci 123, 1751–1760.

Chamcheu, J. C., Siddiqui, I. A., Syed, D. N., Adhami, V. M., Liovic, M. and Mukhtar, H. (2011). Keratin Gene Mutations in Disorders of Human Skin and its Appendages. Arch Biochem Biophys 508, 123–137.

Chancellor, T. J., Lee, J., Thodeti, C. K. and Lele, T. (2010). Actomyosin Tension Exerted on the Nucleus through Nesprin-1 Connections Influences Endothelial Cell Adhesion, Migration, and Cyclic Strain-Induced Reorientation. Biophys J 99, 115–123.

Chang, W., Antoku, S., Östlund, C., Worman, H. J. and Gundersen, G. G. (2015a). Linker of nucleoskeleton and cytoskeleton (LINC) complex-mediated actin-dependent nuclear positioning orients centrosomes in migrating myoblasts. Nucleus 6, 77–88.

Chang, W., Worman, H. J. and Gundersen, G. G. (2015b). Accessorizing and anchoring the LINC complex for multifunctionality. J Cell Biol 208, 11–22.

Chen, H., Cohen, D. M., Choudhury, D. M., Kioka, N. and Craig, S. W. (2005). Spatial distribution and functional significance of activated vinculin in living cells. J Cell Biol 169, 459–470.

Chen, J.-C., Lin, B.-B., Hu, H.-W., Lin, C., Jin, W.-Y., Zhang, F.-B., Zhu, Y.-A., Lu, C.-J., Wei, X.-J. and Chen, R.-J. (2014). NGF Accelerates Cutaneous Wound Healing by Promoting the Migration of Dermal Fibroblasts via the PI3K/Akt-Rac1-JNK and ERK Pathways. BioMed Research International.

Chi, Y.-H., Haller, K., Peloponese, J.-M. and Jeang, K.-T. (2007). Histone Acetyltransferase hALP and Nuclear Membrane Protein hsSUN1 Function in De-condensation of Mitotic Chromosomes. J. Biol. Chem. 282, 27447–27458.

Colburn, Z. T. and Jones, J. C. R. (2016). α6β4 Integrin Regulates the Collective Migration of Epithelial Cells. Am J Respir Cell Mol Biol 56, 443–452.

Darby, I. A., Laverdet, B., Bonté, F. and Desmoulière, A. (2014). Fibroblasts and myofibroblasts in wound healing. Clin Cosmet Investig Dermatol 7, 301–311. de Melker, A. A. and Sonnenberg, A. (1999). Integrins: alternative splicing as a mechanism to regulate ligand binding and integrin signaling events. Bioessays 21, 499–509.

Demetriou, M. C. and Cress, A. E. (2004). Integrin clipping: A novel adhesion switch? J. Cell. Biochem. 91, 26–35.

Desmoulière, A., Redard, M., Darby, I. and Gabbiani, G. (1995). Apoptosis mediates the decrease in cellularity during the transition between granulation tissue and scar. Am J Pathol 146, 56–66.

Djinovic-Carugo, K., Gautel, M., Ylänne, J. and Young, P. (2002). The spectrin repeat: a structural platform for cytoskeletal protein assemblies. FEBS Letters 513, 119–123.

Doherty, J. A., Rossing, M. A., Cushing-Haugen, K. L., Chen, C., Berg, D. J. V. D., Wu, A. H., Pike, M. C., Ness, R. B., Moysich, K., Chenevix-Trench, G., et al. (2010). ESR1/SYNE1 Polymorphism and Invasive Epithelial Ovarian Cancer Risk: An Ovarian Cancer Association Consortium Study. Cancer Epidemiol Biomarkers Prev 19, 245–250.

Dong, J.-M., Lau, L.-S., Ng, Y.-W., Lim, L. and Manser, E. (2009). Paxillin nuclear-cytoplasmic localization is regulated by phosphorylation of the LD4 motif: evidence that nuclear paxillin promotes cell proliferation. Biochemical Journal 418, 717–717.

Driskell, R. R. and Watt, F. M. (2015). Understanding fibroblast heterogeneity in the skin. Trends in Cell Biology 25, 92–99.

Driskell, R. R., Lichtenberger, B. M., Hoste, E., Kretzschmar, K., Simons, B. D., Charalambous, M., Ferron, S. R., Herault, Y., Pavlovic, G., Ferguson-Smith, A. C., et al. (2013). Distinct fibroblast lineages determine dermal architecture in skin development and repair. Nature 504, 277–281.

Dubois, F., Alpha, K. and Turner, C. E. (2017). Paxillin regulates cell polarization and anterograde vesicle trafficking during cell migration. Mol. Biol. Cell 28, 3815–3831.

Duong, N. T., Morris, G. E., Lam, L. T., Zhang, Q., Sewry, C. A., Shanahan, C. M. and Holt, I. (2014). Nesprins: Tissue-Specific Expression of Epsilon and Other Short Isoforms. PLOS ONE 9, e94380.

Eto, M., Kirkbride, J., Elliott, E., Lo, S. H. and Brautigan, D. L. (2007). Association of the Tensin N-terminal Protein-tyrosine Phosphatase Domain with the α Isoform of Protein Phosphatase-1 in Focal Adhesions. J. Biol. Chem. 282, 17806–17815.

Fang, R. C. and Mustoe, T. A. (2008). Animal models of wound healing: utility in transgenic mice. J Biomater Sci Polym Ed 19, 989–1005.

Forscher, P. and Smith, S. J. (1988). Actions of cytochalasins on the organization of actin filaments and microtubules in a neuronal growth cone. The Journal of Cell Biology 107, 1505–1516.

Gagnon, J. A. and Mowry, K. L. (2011). Molecular motors: directing traffic during RNA localization. Critical Reviews in Biochemistry and Molecular Biology 46, 229–239.

Gardel, M. L., Schneider, I. C., Aratyn-Schaus, Y. and Waterman, C. M. (2010). Mechanical Integration of Actin and Adhesion Dynamics in Cell Migration. Annual Review of Cell and Developmental Biology 26, 315–333.

Georges-Labouesse, E., Messaddeq, N., Yehia, G., Cadalbert, L., Dierich, A. and Le Meur, M. (1996). Absence of integrin α6 leads to epidermolysis bullosa and neonatal death in mice. Nat Genet 13, 370–373.

Geuijen, C. A. W. and Sonnenberg, A. (2002). Dynamics of the α6β4 Integrin in Keratinocytes. Mol. Biol. Cell 13, 3845–3858.

Gough, L. L., Fan, J., Chu, S., Winnick, S. and Beck, K. A. (2003). Golgi Localization of Syne- 1. Mol Biol Cell 14, 2410–2424.

Grossman, R. M., Krueger, J., Yourish, D., Granelli-Piperno, A., Murphy, D. P., May, L. T., Kupper, T. S., Sehgal, P. B. and Gottlieb, A. B. (1989). Interleukin 6 is expressed in high levels in psoriatic skin and stimulates proliferation of cultured human keratinocytes. PNAS 86, 6367–6371.

Haque, F., Lloyd, D. J., Smallwood, D. T., Dent, C. L., Shanahan, C. M., Fry, A. M., Trembath, R. C. and Shackleton, S. (2006). SUN1 Interacts with Nuclear Lamin A and Cytoplasmic Nesprins To Provide a Physical Connection between the Nuclear Lamina and the Cytoskeleton. Mol. Cell. Biol. 26, 3738–3751.

Haynie, D. T. (2014). Molecular physiology of the tensin brotherhood of integrin adaptor proteins. Proteins: Structure, Function, and Bioinformatics 82, 1113–1127.

Headon, D. (2017). Reversing stratification during wound healing. Nature Cell Biology 19, 595– 597.

Hemler, M. E., Crouse, C. and Sonnenberg, A. (1989). Association of the VLA alpha 6 subunit with a novel protein. A possible alternative to the common VLA beta 1 subunit on certain cell lines. J. Biol. Chem. 264, 6529–6535.

Hinz, B. (2010). The myofibroblast: Paradigm for a mechanically active cell. Journal of Biomechanics 43, 146–155.

Hinz, B., Mastrangelo, D., Iselin, C. E., Chaponnier, C. and Gabbiani, G. (2001a). Mechanical Tension Controls Granulation Tissue Contractile Activity and Myofibroblast Differentiation. Am J Pathol 159, 1009–1020.

Hinz, B., Celetta, G., Tomasek, J. J., Gabbiani, G., Chaponnier, C. and Matsudaira, P. T. (2001b). Alpha-Smooth Muscle Actin Expression Upregulates Fibroblast Contractile Activity. MBoC 12, 2730–2741.

Hirata, H., Tatsumi, H. and Sokabe, M. (2008). Mechanical forces facilitate actin polymerization at focal adhesions in a zyxin-dependent manner. Journal of Cell Science 121, 2795–2804.

Hirata, H., Samsonov, M. and Sokabe, M. (2017). Actomyosin contractility provokes contact inhibition in E-cadherin-ligated keratinocytes. Scientific Reports 7, 46326.

Hirobe, T. (2005). Role of keratinocyte-derived factors involved in regulating the proliferation and differentiation of mammalian epidermal melanocytes. Pigment Cell Research 18, 2– 12.

Hiroyasu, S., Ozawa, T., Kobayashi, H., Ishii, M., Aoyama, Y., Kitajima, Y., Hashimoto, T., Jones, J. C. R. and Tsuruta, D. (2013). Bullous Pemphigoid IgG Induces BP180 Internalization via a Macropinocytic Pathway. The American Journal of Pathology 182, 828–840.

Hoffman, L. M., Jensen, C. C., Kloeker, S., Wang, C.-L. A., Yoshigi, M. and Beckerle, M. C. (2006). Genetic ablation of zyxin causes Mena/VASP mislocalization, increased motility, and deficits in actin remodeling. The Journal of Cell Biology 172, 771–782.

Hopkinson, S. B., Findlay, K., deHart, G. W. and Jones, J. C. R. (1998). Interaction of BP180 (Type XVII Collagen) and α6 Integrin is Necessary for Stabilization of Hemidesmosome Structure. Journal of Investigative Dermatology 111, 1015–1022.

Hopkinson, S. B., Hamill, K. J., Wu, Y., Eisenberg, J. L., Hiroyasu, S. and Jones, J. C. R. (2014). Focal contact and hemidesmosomal proteins in keratinocyte migration and wound repair. Adv Wound Care (New Rochelle) 3, 247–263.

Horn, H. F., Brownstein, Z., Lenz, D. R., Shivatzki, S., Dror, A. A., Dagan-Rosenfeld, O., Friedman, L. M., Roux, K. J., Kozlov, S., Jeang, K.-T., et al. (2013). The LINC complex is essential for hearing. J Clin Invest 123, 740–750.

Hsu, Y.-L., Wu, C.-Y., Hung, J.-Y., Lin, Y.-S., Huang, M.-S. and Kuo, P.-L. (2013). Galectin-1 promotes lung cancer tumor metastasis by potentiating integrin α6β4 and Notch1/Jagged2 signaling pathway. Carcinogenesis 34, 1370–1381.

Hu, Y.-L., Lu, S., Szeto, K. W., Sun, J., Wang, Y., Lasheras, J. C. and Chien, S. (2014). FAK and paxillin dynamics at focal adhesions in the protrusions of migrating cells. Scientific Reports 4, 6024–6030.

Humphries, J. D., Wang, P., Streuli, C., Geiger, B., Humphries, M. J. and Ballestrem, C. (2007). Vinculin controls focal adhesion formation by direct interactions with talin and actin. J Cell Biol 179, 1043–1057.

Hynes, R. O. (2002). Integrins: Bidirectional, Allosteric Signaling Machines. Cell 110, 673–687.

Janssen, M. E. W., Kim, E., Liu, H., Fujimoto, L. M., Bobkov, A., Volkmann, N. and Hanein, D. (2006). Three-Dimensional Structure of Vinculin Bound to Actin Filaments. Molecular Cell 21, 271–281.

Jiang, C. K., Flanagan, S., Ohtsuki, M., Shuai, K., Freedberg, I. M. and Blumenberg, M. (1994). Disease-activated transcription factor: allergic reactions in human skin cause nuclear translocation of STAT-91 and induce synthesis of keratin K17. Molecular and Cellular Biology 14, 4759–4769.

Jones, J. C. R., Hopkinson, S. B. and Goldfinger, L. E. (1998). Structure and assembly of hemidesmosomes. Bioessays 20, 488–494.

Kanchanawong, P., Shtengel, G., Pasapera, A. M., Ramko, E. B., Davidson, M. W., Hess, H. F. and Waterman, C. M. (2010). Nanoscale architecture of integrin-based cell adhesions. Nature 468, 580–584.

Kandert, S., Lüke, Y., Kleinhenz, T., Neumann, S., Lu, W., Jaeger, V. M., Munck, M., Wehnert, M., Müller, C. R., Zhou, Z., et al. (2007). Nesprin-2 giant safeguards nuclear envelope architecture in LMNA S143F progeria cells. Hum Mol Genet 16, 2944–2959.

Katz, Z. B., Wells, A. L., Park, H. Y., Wu, B., Shenoy, S. M. and Singer, R. H. (2012). β-Actin mRNA compartmentalization enhances focal adhesion stability and directs cell migration. Genes Dev. 26, 1885–1890.

Ketema, M., Wilhelmsen, K., Kuikman, I., Janssen, H., Hodzic, D. and Sonnenberg, A. (2007). Requirements for the localization of nesprin-3 at the nuclear envelope and its interaction with plectin. Journal of Cell Science 120, 3384–3394.

Koivisto, L., Heino, J., Häkkinen, L. and Larjava, H. (2014). Integrins in Wound Healing. Adv Wound Care (New Rochelle) 3, 762–783.

Koster, M. I. (2009). Making an epidermis. Ann N Y Acad Sci 1170, 7–10.

Krawczyk, W. S. (1971). A Pattern of Epidermal Cell Migration During Wound Healing. J Cell Biol 49, 247–263.

Kutscheidt, S., Zhu, R., Antoku, S., Luxton, G. W. G., Stagljar, I., Fackler, O. T. and Gundersen, G. G. (2014). FHOD1 interaction with nesprin-2G mediates TAN line formation and nuclear movement. Nature Cell Biology 16, 708–715.

Lawrence, J. B. and Singer, R. H. (1986). Intracellular localization of messenger RNAs for cytoskeletal proteins. Cell 45, 407–415.

Lee, E. C., Lotz, M. M., Steele, G. D. and Mercurio, A. M. (1992). The integrin alpha 6 beta 4 is a laminin receptor. J Cell Biol 117, 671–678.

Legate, K. R. and Fässler, R. (2009). Mechanisms that regulate adaptor binding to β-integrin cytoplasmic tails. Journal of Cell Science 122, 187–198.

Leube, R. E., Moch, M. and Windoffer, R. (2015). Intermediate filaments and the regulation of focal adhesion. Current Opinion in Cell Biology 32, 13–20.

Levy, V., Lindon, C., Harfe, B. D. and Morgan, B. A. (2005). Distinct Stem Cell Populations Regenerate the Follicle and Interfollicular Epidermis. Developmental Cell 9, 855–861.

Li, G., Du, X., Vass, W. C., Papageorge, A. G., Lowy, D. R. and Qian, X. (2011). Full activity of the deleted in liver cancer 1 (DLC1) tumor suppressor depends on an LD-like motif that binds talin and focal adhesion kinase (FAK). PNAS 108, 17129–17134.

Litjens, S. H. M., de Pereda, J. M. and Sonnenberg, A. (2006). Current insights into the formation and breakdown of hemidesmosomes. Trends in Cell Biology 16, 376–383.

Lo, S. H., Janmey, P. A., Hartwig, J. H. and Chen, L. B. (1994). Interactions of tensin with actin and identification of its three distinct actin-binding domains. The Journal of Cell Biology 125, 1067–1075.

Lombardi, M. L., Jaalouk, D. E., Shanahan, C. M., Burke, B., Roux, K. J. and Lammerding, J. (2011). The Interaction between Nesprins and Sun Proteins at the Nuclear Envelope Is Critical for Force Transmission between the Nucleus and Cytoskeleton. J. Biol. Chem. 286, 26743–26753.

López-Colomé, A. M., Lee-Rivera, I., Benavides-Hidalgo, R. and López, E. (2017). Paxillin: a crossroad in pathological cell migration. Journal of Hematology & Oncology 10, 50.

Lu, W., Schneider, M., Neumann, S., Jaeger, V.-M., Taranum, S., Munck, M., Cartwright, S., Richardson, C., Carthew, J., Noh, K., et al. (2012). Nesprin interchain associations control nuclear size. Cell. Mol. Life Sci. 69, 3493–3509.

Lüke, Y., Zaim, H., Karakesisoglou, I., Jaeger, V. M., Sellin, L., Lu, W., Schneider, M., Neumann, S., Beijer, A., Munck, M., et al. (2008). Nesprin-2 giant (NUANCE) maintains nuclear envelope architecture and composition in skin. Journal of Cell Science 121, 1887–1898.

Mardakheh, F. K., Paul, A., Kümper, S., Sadok, A., Paterson, H., Mccarthy, A., Yuan, Y. and Marshall, C. J. (2015). Global Analysis of mRNA, Translation, and Protein Localization: Local Translation Is a Key Regulator of Cell Protrusions. Developmental Cell 35, 344– 357.

Martinot, V., Mitchell, V., Fevrier, P., Duhamel, A. and Pellerin, P. (1994). Comparative study of split thickness skin grafts taken from the scalp and thigh in children. Burns 20, 146– 150.

Mayr, C. (2017). Regulation by 3′-Untranslated Regions. Annu. Rev. Genet. 51, 171–194.

Mazumder, A. and Shivashankar, G. V. (2010). Emergence of a prestressed eukaryotic nucleus during cellular differentiation and development. Journal of The Royal Society Interface 7, S321–S330.

Michopoulou, A. and Rousselle, P. (2015). How do epidermal matrix metalloproteinases support re-epithelialization during skin healing? European Journal of Dermatology 25, 33–42.

Mislow, J. M. K., Holaska, J. M., Kim, M. S., Lee, K. K., Segura-Totten, M., Wilson, K. L. and McNally, E. M. (2002). Nesprin-1α self-associates and binds directly to emerin and lamin A in vitro. FEBS Letters 525, 135–140.

Natalie, R. K., Thanh, L. L., A, S. C., Megan, P., Denis, F., Manfred, W., M, M. E. and E, M. G. (2010). Nesprins, but not sun proteins, switch isoforms at the nuclear envelope during muscle development. Developmental Dynamics 239, 998–1009.

Neumann, S., Schneider, M., Daugherty, R. L., Gottardi, C. J., Eming, S. A., Beijer, A., Noegel, A. A. and Karakesisoglou, I. (2010). Nesprin-2 Interacts with α-Catenin and Regulates Wnt Signaling at the Nuclear Envelope. J. Biol. Chem. 285, 34932–34938.

Nickoloff, B. J. and Turka, L. A. (1993). Keratinocytes: key immunocytes of the integument. Am J Pathol 143, 325–331.

Nobes, C. D. and Hall, A. (1995). Rho, rac, and cdc42 GTPases regulate the assembly of multimolecular focal complexes associated with actin stress fibers, lamellipodia, and filopodia. Cell 81, 53–62.

Osmanagic-Myers, S., Dechat, T. and Foisner, R. (2015). Lamins at the crossroads of mechanosignaling. Genes Dev. 29, 225–237.

Ozawa, T., Tsuruta, D., Jones, J. C. R., Ishii, M., Ikeda, K., Harada, T., Aoyama, Y., Kawada, A. and Kobayashi, H. (2010). Dynamic Relationship of Focal Contacts and Hemidesmosome Protein Complexes in Live Cells. J Invest Dermatol 130, 1624–1635.

Padmakumar, V. C., Libotte, T., Lu, W., Zaim, H., Abraham, S., Noegel, A. A., Gotzmann, J., Foisner, R. and Karakesisoglou, I. (2005). The inner nuclear membrane protein Sun1 mediates the anchorage of Nesprin-2 to the nuclear envelope. J Cell Sci 118, 3419–3430.

Pan, X., Hobbs, R. P. and Coulombe, P. A. (2013). The expanding significance of keratin intermediate filaments in normal and diseased epithelia. Curr Opin Cell Biol 25, 47–56.

Park, S., Gonzalez, D. G., Guirao, B., Boucher, J. D., Cockburn, K., Marsh, E. D., Mesa, K. R., Brown, S., Rompolas, P., Haberman, A. M., et al. (2017). Tissue-scale coordination of cellular behaviour promotes epidermal wound repair in live mice. Nature Cell Biology 19, 155–163.

Pastar, I., Stojadinovic, O., Yin, N. C., Ramirez, H., Nusbaum, A. G., Sawaya, A., Patel, S. B., Khalid, L., Isseroff, R. R. and Tomic-Canic, M. (2014). Epithelialization in Wound Healing: A Comprehensive Review. Adv Wound Care (New Rochelle) 3, 445–464.

Petrie, R. J., Koo, H. and Yamada, K. M. (2014). Generation of compartmentalized pressure by a nuclear piston governs cell motility in a 3D matrix. Science 345, 1062–1065.

Ponti, A., Machacek, M., Gupton, S. L., Waterman-Storer, C. M. and Danuser, G. (2004). Two Distinct Actin Networks Drive the Protrusion of Migrating Cells. Science 305, 1782– 1786.

Puckelwartz, M. J., Kessler, E., Zhang, Y., Hodzic, D., Randles, K. N., Morris, G., Earley, J. U., Hadhazy, M., Holaska, J. M., Mewborn, S. K., et al. (2009). Disruption of nesprin-1 produces an Emery Dreifuss muscular dystrophy-like phenotype in mice. Hum Mol Genet 18, 607–620.

Rabinovitz, I., Tsomo, L. and Mercurio, A. M. (2004). Protein Kinase C-α Phosphorylation of Specific Serines in the Connecting Segment of the β4 Integrin Regulates the Dynamics of Type II Hemidesmosomes. Mol. Cell. Biol. 24, 4351–4360.

Radice, G. P. (1980). The spreading of epithelial cells during wound closure in Xenopus larvae. Developmental Biology 76, 26–46.

Rajgor, D. and Shanahan, C. M. (2013). Nesprins: from the nuclear envelope and beyond. Expert Reviews in Molecular Medicine 15, E5.

Rajgor, D., Mellad, J. A., Autore, F., Zhang, Q. and Shanahan, C. M. (2012). Multiple Novel Nesprin-1 and Nesprin-2 Variants Act as Versatile Tissue-Specific Intracellular Scaffolds. PLOS ONE 7, e40098.

Rashmi, R. N., Eckes, B., Glöckner, G., Groth, M., Neumann, S., Gloy, J., Sellin, L., Walz, G., Schneider, M., Karakesisoglou, I., et al. (2012). The nuclear envelope protein Nesprin-2 has roles in cell proliferation and differentiation during wound healing. Nucleus 3, 172– 186.

Raymond, K., Kreft, M., Song, J.-Y., Janssen, H. and Sonnenberg, A. (2007). Dual Role of α6β4 Integrin in Epidermal Tumor Growth: Tumor-suppressive Versus Tumor-promoting Function. Mol. Biol. Cell 18, 4210–4221.

Reinhard, M., Zumbrunn, J., Jaquemar, D., Kuhn, M., Walter, U. and Trueb, B. (1999). An α- Actinin Binding Site of Zyxin Is Essential for Subcellular Zyxin Localization and α- Actinin Recruitment. J. Biol. Chem. 274, 13410–13418.

Rio, A. del, Perez-Jimenez, R., Liu, R., Roca-Cusachs, P., Fernandez, J. M. and Sheetz, M. P. (2009). Stretching Single Talin Rod Molecules Activates Vinculin Binding. Science 323, 638–641.

Rittié, L. (2016). Cellular mechanisms of skin repair in humans and other mammals. J Cell Commun Signal 10, 103–120.

Rivera-Gonzalez, G., Shook, B. and Horsley, V. (2014). Adipocytes in Skin Health and Disease. Cold Spring Harb Perspect Med 4.

Romani, N., Holzmann, S., Tripp, C. H., Koch, F. and Stoitzner, P. (2003). Langerhans cells – dendritic cells of the epidermis. APMIS 111, 725–740.

Rousselle, P., Braye, F. and Dayan, G. (2018). Re-epithelialization of adult skin wounds: Cellular mechanisms and therapeutic strategies. Advanced Drug Delivery Reviews.

Roux, K. J., Crisp, M. L., Liu, Q., Kim, D., Kozlov, S., Stewart, C. L. and Burke, B. (2009). Nesprin 4 is an outer nuclear membrane protein that can induce kinesin-mediated cell polarization. Proc Natl Acad Sci U S A 106, 2194–2199.

Ryder, P. V. and Lerit, D. A. (2018). RNA localization regulates diverse and dynamic cellular processes. Traffic 19, 496–502.

Salgia, R., Li, J.-L., Lo, S. H., Brunkhorst, B., Kansas, G. S., Sobhany, E. S., Sun, Y., Pisick, E., Hallek, M., Ernst, T., et al. (1995). Molecular Cloning Of Human Paxillin, a Focal Adhesion Protein Phosphorylated by P210BCR/ABL. J. Biol. Chem. 270, 5039–5047.

Schäffer, M. R., Tantry, U., Ahrendt, G. M., Wasserkrug, H. L. and Barbul, A. (1997). Stimulation of fibroblast proliferation and matrix contraction by wound fluid. The International Journal of Biochemistry & Cell Biology 29, 231–239.

Schreiner, S. M., Koo, P. K., Zhao, Y., Mochrie, S. G. J. and King, M. C. (2015). The tethering of chromatin to the nuclear envelope supports nuclear mechanics. Nature Communications 6, 7159.

Schwartz, C., Fischer, M., Mamchaoui, K., Bigot, A., Lok, T., Verdier, C., Duperray, A., Michel, R., Holt, I., Voit, T., et al. (2017). Lamins and nesprin-1 mediate inside-out mechanical coupling in muscle cell precursors through FHOD1. Scientific Reports 7, 1253.

Sehgal, B. U., DeBiase, P. J., Matzno, S., Chew, T.-L., Claiborne, J. N., Hopkinson, S. B., Russell, A., Marinkovich, M. P. and Jones, J. C. R. (2006). Integrin β4 regulates migratory behavior of keratinocytes by determining laminin-332 organization. J. Biol. Chem. 281, 35487–35498.

Seltmann, K., Roth, W., Loschke, F., Lederer, M., Huttelmaier, S. and Magin, T. M. (2013). Keratins mediate localization of hemidesmosomes and repress cell motility. J Invest Dermatol 133, 181–190.

Shephard, P., Martin, G., Smola-Hess, S., Brunner, G., Krieg, T. and Smola, H. (2004). Myofibroblast Differentiation Is Induced in Keratinocyte-Fibroblast Co-Cultures and Is Antagonistically Regulated by Endogenous Transforming Growth Factor-β and Interleukin-1. Am J Pathol 164, 2055–2066.

Sjöblom, T., Jones, S., Wood, L. D., Parsons, D. W., Lin, J., Barber, T. D., Mandelker, D., Leary, R. J., Ptak, J., Silliman, N., et al. (2006). The Consensus Coding Sequences of Human Breast and Colorectal Cancers. Science 314, 268–274.

Smith, M. A., Hoffman, L. M. and Beckerle, M. C. (2014). LIM proteins in actin cytoskeleton mechanoresponse. Trends in Cell Biology 24, 575–583.

Sorrell, J. M. and Caplan, A. I. (2004). Fibroblast heterogeneity: more than skin deep. Journal of Cell Science 117, 667–675.

Sterk, L. M. T., Geuijen, C. A. W., Oomen, L. C. J. M., Calafat, J., Janssen, H. and Sonnenberg, A. (2000). The Tetraspan Molecule Cd151, a Novel Constituent of Hemidesmosomes, Associates with the Integrin α6β4 and May Regulate the Spatial Organization of Hemidesmosomes. J Cell Biol 149, 969–982.

Stewart, R. M., Zubek, A. E., Rosowski, K. A., Schreiner, S. M., Horsley, V. and King, M. C. (2015). Nuclear–cytoskeletal linkages facilitate cross talk between the nucleus and intercellular adhesions. J Cell Biol 209, 403–418.

Subauste, M. C., Pertz, O., Adamson, E. D., Turner, C. E., Junger, S. and Hahn, K. M. (2004). Vinculin modulation of paxillin–FAK interactions regulates ERK to control survival and motility. J Cell Biol 165, 371–381.

Szostak, E. and Gebauer, F. (2013). Translational control by 3′-UTR-binding proteins. Briefings in Functional Genomics 12, 58–65.

Tai, P. (2013). A Practical Update of Surgical Management of Merkel Cell Carcinoma of the Skin. ISRN Surg 2013.

Tessema, M., Willink, R., Do, K., Yu, Y. Y., Yu, W., Machida, E. O., Brock, M., Neste, L. V., Stidley, C. A., Baylin, S. B., et al. (2008). Promoter Methylation of Genes in and around the Candidate Lung Cancer Susceptibility Locus 6q23-25. Cancer Res 68, 1707–1714.

Theveneau, E. and Linker, C. (2017). Leaders in collective migration: are front cells really endowed with a particular set of skills? F1000Res 6.

Thomas, G. J., Poomsawat, S., Lewis, M. P., Hart, I. R., Speight, P. M. and Marshall, J. F. (2001). αvβ6 Integrin Upregulates Matrix Metalloproteinase 9 and Promotes Migration of Normal Oral Keratinocytes. Journal of Investigative Dermatology 116, 898–904.

Thomas, D. G., Yenepalli, A., Denais, C. M., Rape, A., Beach, J. R., Wang, Y., Schiemann, W. P., Baskaran, H., Lammerding, J. and Egelhoff, T. T. (2015). Non-muscle myosin IIB is critical for nuclear translocation during 3D invasion. J Cell Biol 210, 583–594.

Thulabandu, V., Chen, D. and Atit, R. P. (2018). Dermal fibroblast in cutaneous development and healing. Wiley Interdisciplinary Reviews: Developmental Biology 7, e307.

Tsuruta, D., Hashimoto, T., Hamill, K. J. and Jones, J. C. R. (2011). Hemidesmosomes and focal contact proteins: Functions and cross-talk in keratinocytes, bullous diseases, and wound healing. J Dermatol Sci 62, 1–7.

Turcan, I. and Jonkman, M. F. (2014). Blistering disease: insight from the hemidesmosome and other components of the dermal-epidermal junction. Cell Tissue Res 1–25.

Uematsu, J., Nishizawa, Y., Sonnenberg, A. and Owaribe, K. (1994). Demonstration of Type II Hemidesmosomes in a Mammary Gland Epithelial Cell Line, BMGE-H. J Biochem 115, 469–476.

Uemura, A., Nguyen, T.-N., Steele, A. N. and Yamada, S. (2011). The LIM Domain of Zyxin Is Sufficient for Force-Induced Accumulation of Zyxin During Cell Migration. Biophysical Journal 101, 1069–1075.

Usui, M. L., Underwood, R. A., Mansbridge, J. N., Muffley, L. A., Carter, W. G. and Olerud, J. E. (2005). Morphological evidence for the role of suprabasal keratinocytes in wound reepithelialization. Wound Repair and Regeneration 13, 468–479. van der Neut, R., Krimpenfort, P., Calafat, J., Niessen, C. M. and Sonnenberg, A. (1996). Epithelial detachment due to absence of hemidesmosomes in integrin β4 null mice. Nat Genet 13, 366–369.

Wang, N., Tytell, J. D. and Ingber, D. E. (2009). Mechanotransduction at a distance: mechanically coupling the extracellular matrix with the nucleus. Nat Rev Mol Cell Biol 10, 75–82.

Wang, X., Liu, H., Zhu, M., Cao, C., Xu, Z., Tsatskis, Y., Lau, K., Kuok, C., Filleter, T., McNeill, H., et al. (2018). Mechanical stability of the cell nucleus: roles played by the cytoskeleton in nuclear deformation and strain recovery. J Cell Sci jcs.209627.

Warren, D. T., Tajsic, T., Mellad, J. A., Searles, R., Zhang, Q. and Shanahan, C. M. (2010). Novel Nuclear Nesprin-2 Variants Tether Active Extracellular Signal-regulated MAPK1 and MAPK2 at Promyelocytic Leukemia Protein Nuclear Bodies and Act to Regulate Smooth Muscle Cell Proliferation. J. Biol. Chem. 285, 1311–1320.

Weis, B. L., Schleiff, E. and Zerges, W. (2013). Protein targeting to subcellular organelles via mRNA localization. Biochimica et Biophysica Acta (BBA) - Molecular Cell Research 1833, 260–273.

Werner, S., Krieg, T. and Smola, H. (2007). Keratinocyte–Fibroblast Interactions in Wound Healing. Journal of Investigative Dermatology 127, 998–1008.

Wilhelmsen, K., Litjens, S. H. M., Kuikman, I., Tshimbalanga, N., Janssen, H., Bout, I. van den, Raymond, K. and Sonnenberg, A. (2005). Nesprin-3, a novel outer nuclear membrane protein, associates with the cytoskeletal linker protein plectin. The Journal of Cell Biology 171, 799–810.

Woods, A. J., Roberts, M. S., Choudhary, J., Barry, S. T., Mazaki, Y., Sabe, H., Morley, S. J., Critchley, D. R. and Norman, J. C. (2002). Paxillin associates with poly(A)-binding protein 1 at the dense endoplasmic reticulum and the leading edge of migrating cells. J. Biol. Chem. 277, 6428–6437.

Xie, S. J., Lu, Y., Yoon, S., Yang, J. and Park, D. S. (2015). Intensity Variation Normalization for Finger Vein Recognition Using Guided Filter Based Singe Scale Retinex. Sensors 15, 17089–17105.

Xu, W., Baribault, H. and Adamson, E. D. (1998). Vinculin knockout results in heart and brain defects during embryonic development. Development 125, 327–337.

Yoshigi, M., Hoffman, L. M., Jensen, C. C., Yost, H. J. and Beckerle, M. C. (2005). Mechanical force mobilizes zyxin from focal adhesions to actin filaments and regulates cytoskeletal reinforcement. J Cell Biol 171, 209–215.

Yu, J. A., Deakin, N. O. and Turner, C. E. (2010). Emerging role of paxillin-PKL in regulation of cell adhesion, polarity and migration. Cell Adh Migr 4, 342–347.

Yu, J., Lei, K., Zhou, M., Craft, C. M., Xu, G., Xu, T., Zhuang, Y., Xu, R. and Han, M. (2011). KASH protein Syne-2/Nesprin-2 and SUN proteins SUN1/2 mediate nuclear migration during mammalian retinal development. Hum Mol Genet 20, 1061–1073.

Zaidel-Bar, R., Cohen, M., Addadi, L. and Geiger, B. (2004). Hierarchical assembly of cell– matrix adhesion complexes. Biochemical Society Transactions 32, 416–420.

Zhang, Q., Skepper, J. N., Yang, F., Davies, J. D., Hegyi, L., Roberts, R. G., Weissberg, P. L., Ellis, J. A. and Shanahan, C. M. (2001). Nesprins: a novel family of spectrin-repeat- containing proteins that localize to the nuclear membrane in multiple tissues. Journal of Cell Science 114, 4485–4498.

Zhang, Q., Ragnauth, C. D., Skepper, J. N., Worth, N. F., Warren, D. T., Roberts, R. G., Weissberg, P. L., Ellis, J. A. and Shanahan, C. M. (2005). Nesprin-2 is a multi-isomeric protein that binds lamin and emerin at the nuclear envelope and forms a subcellular network in skeletal muscle. Journal of Cell Science 118, 673–687.

Zhang, X., Xu, R., Zhu, B., Yang, X., Ding, X., Duan, S., Xu, T., Zhuang, Y. and Han, M. (2007a). Syne-1 and Syne-2 play crucial roles in myonuclear anchorage and motor neuron innervation. Development 134, 901–908.

Zhang, Q., Bethmann, C., Worth, N. F., Davies, J. D., Wasner, C., Feuer, A., Ragnauth, C. D., Yi, Q., Mellad, J. A., Warren, D. T., et al. (2007b). Nesprin-1 and -2 are involved in the pathogenesis of Emery–Dreifuss muscular dystrophy and are critical for nuclear envelope integrity. Hum Mol Genet 16, 2816–2833.

Zhang, X., Lei, K., Yuan, X., Wu, X., Zhuang, Y., Xu, T., Xu, R. and Han, M. (2009). SUN1/2 and Syne/Nesprin-1/2 Complexes Connect Centrosome to the Nucleus during Neurogenesis and Neuronal Migration in Mice. Neuron 64, 173–187.

Zhang, Q., Minaisah, R.-M., Ferraro, E., Li, C., Porter, L. J., Zhou, C., Gao, F., Zhang, J., Rajgor, D., Autore, F., et al. (2016). N-terminal nesprin-2 variants regulate β-catenin signalling. Experimental Cell Research 345, 168–179.

Zhao, Z., Tan, S. H., Machiyama, H., Kawauchi, K., Araki, K., Hirata, H. and Sawada, Y. (2016). Association between tensin 1 and p130Cas at focal adhesions links actin inward flux to cell migration. Biology Open 5, 499–506.

Zhao, Y., Wang, Y., Sarkar, A. and Wang, X. (2018). Keratocytes generate high integrin tension at the trailing edge to mediate rear de-adhesion during rapid cell migration. iScience 9, 502–512.

Zhou, Z., Du, X., Cai, Z., Song, X., Zhang, H., Mizuno, T., Suzuki, E., Yee, M. R., Berezov, A., Murali, R., et al. (2012). Structure of Sad1-UNC84 Homology (SUN) Domain Defines Features of Molecular Bridge in Nuclear Envelope. J. Biol. Chem. 287, 5317–5326.

Ziegler, W. H., Liddington, R. C. and Critchley, D. R. (2006). The structure and regulation of vinculin. Trends in Cell Biology 16, 453–460.

Zomer, H. D. and Trentin, A. G. (2018). Skin wound healing in humans and mice: Challenges in translational research. Journal of Dermatological Science 90, 3–12.

Figures

Figure 1. Skin structure reproduced from Xie et al., 2015. The skin is composed of two primary layers interspersed with hair follicles, nerves, blood vessels, and secretory glands. The epidermis is primarily composed of keratinocytes while the dermis is primarily composed of fibroblasts.

The upper layers of the epidermis are collectively referred to suprabasal layers, and the keratinocytes within these layers are less metabolically active than the basal keratinocytes.

Underlying these skin layers are a layer of adipose tissue which have recently been shown to have metabolic functions.

Figure 2. Full- and partial-thickness wounds in human skin reproduced from Rittié, 2016. a)

Intact skin. b) Formation of partial thickness wound that damages the epidermis and may or may not damage part of the dermis. c) Healing of partial thickness wound by re-epithelialization. d)

Restoration of normal skin after partial thickness wound. e) Formation of full thickness wound that damages both epidermal and dermal layers. f) The first step in healing a partial thickness wound where a blood clot is formed, and fibroblasts infiltrate the wound bed. g) The dermal layer is now primarily composed of granulation tissue which is extracellular matrix secreted by fibroblasts, and the process of re-epithelization of the epidermis begins. h) Re-epithelialization of the epidermis is complete and restores the barrier function of skin. Note that full thickness wounds impair the normal functioning of the dermal layer after healing.

Figure 3. Nesprin family members at the nuclear envelope adapted from Rajgor and Shanahan,

2013. Nesprins are anchored to the outer nuclear membrane (ONM) via the C-terminal KASH domain which interacts with SUN proteins. SUN proteins are anchored to the inner nuclear membrane (INM), where these proteins can influence nuclear shape and DNA transcription.

Each nesprin family member interacts with different cell stabilizing elements: nesprin-1G and

2G interact with the actin cytoskeleton via two actin binding domains, nesprin-2 interacts with plectin which stabilizes the keratin intermediate filament network, and nesprin-4 interacts with microtubule motors which stabilizes microtubules.

Figure 4. Cell-matrix adhesions reproduced from Hopkinson et al., 2014. The protein complex on the left represents focal adhesion (FA) components while the complex on the right represents hemidesmosome (HD) components. Both complexes contain an integrin heterodimer core which is responsible for integrating the extracellular matrix and cytoplasmic protein components. FAs can contain over 250 different proteins and interact with the actin cytoskeleton. HDs contain a standard seven proteins and interact with keratin intermediate filaments (IFs).

CHAPTER TWO: NESPRIN-2G KNOCKOUT KERATINOCYTES DISPLAY NORMAL

CELL MIGRATION AND PROTEIN EXPRESSION

Abstract

The epidermis of human skin functions as a barrier to prevent water loss and infiltration of environmental pathogens. Keratinocytes are the primary cell of the epidermis, and during wound healing, these cells move into wounds to re-establish the skin barrier. The movement of keratinocytes is facilitated by cytoskeleton-linked cell-matrix adhesion protein complexes, namely the hemidesmosome and focal adhesions. Nesprins are a family of proteins that have been suggested to have roles in cell migration by acting as linkers of the cytoskeleton and nucleoskeleton. Prior studies have shown that the loss of nesprin-2G leads to decreased skin wound healing in mice. Here, we show that nesprin-2G knockout primary mouse keratinocytes do not show morphological differences from wildtype primary mouse keratinocytes. Knockout cells exhibit no clear differences from wildtype keratinocytes with regard to their individual motility, collective cell migration, or expression of cell-matrix adhesion proteins. Therefore, we propose that keratinocytes from the epidermis do not contribute to the delayed wound healing phenotype observed in mice engineered to lack nesprin-2G.

Introduction

Human skin is a dual-layered organ composed of both epidermis and dermis. The epidermis prevents water loss and infiltration of pathogens, and it primarily contains one cell type, the keratinocyte (Baroni et al., 2012). When the skin is wounded, it is the responsibility of keratinocytes to re-establish the barrier function of skin. After wounding, keratinocytes become polarized and migrate into the wound bed across extracellular matrix via cell-matrix adhesion protein complexes termed hemidesmosomes (HDs) and focal adhesions (FAs) (Theveneau and

Linker, 2017). While FAs are prevalent in most cells, HDs are found only in certain epithelial cell types, most notably the basal keratinocytes within the epidermis, where they anchor the epidermis to the dermis. Both HDs and FAs contain an integrin heterodimer core which recruit other scaffolding and signaling proteins to form these complexes to allow for cell migration. The integrin heterodimer found in HDs is α6β4 whereas the heterodimer found in FAs can vary depending on cell type (Hynes, 2002).

The family of nesprin proteins contains 4 members (-1G, -2G, -3, and -4). These family members vary in size, but all are responsible for maintaining cell morphology through interactions with the intermediate filament and cytoskeleton networks within the cell (Rajgor and

Shanahan, 2013). Interestingly, these proteins have roles in cell migration as loss of nesprin-1G,

-2G, or -3 in endothelial cells impedes cell migration (King et al., 2014; Morgan et al., 2011).

Here, we focus on nesprin-2G because prior work indicated that loss of nesprin-2G in mice results in a wound healing defect and that this wound healing defect was likely due to changes in

FAs leading to motility abnormalities in keratinocytes (Lüke et al., 2008; Rashmi et al., 2012).

Nesprin-2G is an ~800kDa protein responsible for the nucleus-actin cytoskeleton interaction.

The C-terminal domain is anchored to the outer nuclear envelope via interactions with SUN

proteins. The N-terminal domain extends into the cytoplasm where it interacts with the actin cytoskeleton via two actin binding domains (Chang et al., 2015; Rajgor and Shanahan, 2013).

We were interested in whether nesprin-2G loss might impact HDs as well as FAs in keratinocytes. Moreover, we explored the motility defects reported in keratinocytes by the previous workers in more depth. To our great surprise, we observed that a loss of nesprin-2G in keratinocytes fails to lead to clear abnormalities in cell migration nor changes in the expression/localization of HD and FA components (Lüke et al., 2008; Rashmi et al., 2012).

Results and discussion

Motility of wildtype and nesprin-2G knockout primary mouse keratinocytes

A nesprin-2G knockout (KO) mouse line was created by another group in which the actin binding domain was deleted, resulting in a non-functional protein product (Lüke et al., 2008).

We were able to obtain this mouse line to establish a breeding colony at WSU. We assayed genomic DNA isolated from wildtype (WT) and KO mice tail snips to ensure their genotype

(Figure 1A). For the following experiments, primary keratinocytes were isolated from the epidermis of 2-day old WT and KO mice, and cultured in serum free media. To ensure our isolation protocol was effective, keratinocytes were imaged following staining with antibodies against keratin, a marker of epithelial cells, and vimentin, a marker of mesenchymal cells (Figure

1B). Our presumed keratinocytes stain positively for keratin but are not recognized by the vimentin antibody, indicating that our epithelial cell isolation protocol is effective. Isolation protocols and experimentation with dermal fibroblasts are explained in chapter 3.

Our first analysis involved comparing the motility of individual WT and KO keratinocytes. Rose plots of the movement of a typical group of 5 WT and KO cells are depicted in Figure 2A. There is no clear difference in the motility patterns of individual KO cells compared to WT cells over 3 h of imaging (Figure 2A-B). Next, we analyzed how the cells move in a scratch assay, a measure of their collective cell migration. In this assay, a monolayer of confluent cells is scratched with a pipette tip and the movement of keratinocytes into the wound is monitored for 12 h, post wounding. We observed no difference in the ability of KO cells to fill the wound compared with WT cells (Figure 2C-D), To extend and confirm our analyses, we assayed collective cell migration over 24 h, but there was still no obvious difference between

WT and KO cells (Figure 2D). In summary, the results of our single cell and scratch assays are contrary those reported in Rashmi et al., 2012.

Protein expression of wildtype and nesprin-2G KO primary mouse keratinocytes

We next assayed cell-matrix adhesions by WT versus KO cells since it has been reported that FAs are “altered” in KO keratinocytes and form more robustly upon plating compared with

WT cells (Rashmi et al., 2012). It should be noted that these same authors did not investigate the localization of HD proteins in isolated KO cells. Thus, we processed WT and KO cells for immunofluorescence analyses using antibodies against various cell-matrix adhesion proteins

(i.e., both FA and HD components). WT and KO cells were co-stained for paxillin, a marker of

FAs, and α6 integrin, a marker of HDs, and actin. Paxillin staining reveals similar sizes and numbers of FAs in WT and KO cells (Figure 3A). Likewise, α6 integrin staining is not obviously changed in KO cells (Figure 3B). We extended these observations to HDs by processing WT and

KO cells for double label immunofluorescence microscopy using antibodies against the HD integrin pair, α6 and β4 integrin. There was no clear difference in HD localization in WT compared to KO cells, with α6 integrin and β4 integrin antibody staining patterns being of similar pattern and intensity (Figure 3B). This finding is consistent with studies revealing that

α6β4 integrin localization is similar in KO versus WT skin sections prepared for immunocytochemistry (Lüke et al., 2008). Therefore, in summary, our data indicate that loss of nesprin-2G is not accompanied by any obvious changes in either FAs or HDs in individual keratinocytes. The former finding is again inconsistent with one report in the literature (Rashmi et al., 2012). Therefore, my studies indicate that there is not a difference in adhesion site formation in KO cells.

To provide additional data that there are no obvious defects in HDs in KO cells, we also compared the localization of the ligand of α6β4 integrin, laminin 332, in both WT and KO keratinocytes. To do so, cells were prepared for immunofluorescence using a polyclonal antibody against laminin 332. The antibody stains the matrix underlying both WT and KO cells (Figure

3C). Staining appears diffusely although, in some instances, appears in whirls and arcs. There is no obvious difference between WT and KO cells.

Next, we were interested in whether there were changes in integrin heterodimers, other than α6β4 integrin, due to the loss of nesprin-2G in keratinocytes. Using flow cytometry, we assayed the surface expression of α6 integrin, α5β1 integrin, and α2β1 integrin. α5β1 integrin is a receptor of fibronectin and is a component of FAs, while α2β1 integrin interacts with collagen I

(Hynes, 1992). There were no differences in α6 integrin and α2β1 integrin expression between

WT and KO cells (Figure 4A, C). However, there was an increase in α5β1 integrin surface expression in KO compared to WT cells (Figure 4B). This is consistent with the reported increased FA formation observed in KO cells compared to WT (Rashmi et al., 2012). However, as we already mentioned, our immunostaining did not provide evidence for such an increase at the immunostaining level (Figure 3).

In summary, our data do not support changes in either cell-matrix adhesion proteins nor the motility of keratinocytes as a consequence of nesprin-2G loss. In other words, we have been unable to replicate a report by Rashmi et al 2012. We have no obvious explanation for this discrepancy other than the possibility that differences in cell isolation protocols or assay conditions are responsible. With all due respect to Rashmi et al 2012, the images shown of FAs in WT and KO cells that they show do not indicate a dramatic change. Moreover, the image they present of their scratch assay is difficult to interpret since the cells are barely visible.

Nonetheless, based on our finding of no clear defects in motility of keratinocytes due to nesprin-

2G loss, we suggest that wound healing defects of nesprin-2G KO cells is not a result of keratinocyte abnormalities, at least regarding their FAs and motility behavior. Rather, the defects in wound healing are more likely to occur due to abnormalities in the fibroblasts in skin. In the next chapter we investigate this possibility.

Materials and methods

Primary keratinocyte isolation

Mice were maintained in accordance with local governmental and institutional animal care regulations and all procedures had institutional approval (Washington State University IACUC protocol numbers 04444-006 and 04824-004). Keratinocytes were isolated from 48 hour-old

C57/Bl6 wildtype and nesprin-2G knockout mice as previously published (Hamill et al., 2012).

Briefly, skins were incubated in 2.4 U/mL dispase overnight to separate epidermis from dermis.

The epidermis was placed in 0.5mL Accutase (CnT-Accutase-100) for 30 minutes followed by gentle rubbing of the epidermis to remove keratinocytes. Cells were spun down and plated for use.

Keratinocyte culture conditions

Primary keratinocytes were cultured in CnT-07 epidermal keratinocyte complete media

(CELLnTEC, Bern, Switzerland), and grew at 37° in 8% humidified CO2.

PCR

Genomic DNA was isolated from 2mm tail snips of wildtype, knockout, and heterozygote mice.

Each tail was incubated in 300μL digest buffer (5mM EDTA, pH 8; 200mM NaCl; 100mM Tris, pH 8; 0.2% SDS) supplemented with 0.4mg proteinase K/mL overnight at 55°C. Next, 1mL

100% ethanol was added to each tube, centrifuged 16,000xg for 30 minutes, supernatant removed, 1mL 70% ethanol was added to each tube, spun at 16,000xg for 20 minutes, supernatant removed, dissolved DNA pellet with 300μL TE buffer, placed tubes at 55°C for 2 h with lids open, and stored DNA at 4°C. DNA was run with either wildtype primers (forward: 5’-

CCATGAAAGCTCTAGTCAGAAAGAGC-3’, reverse: 5’-

CAGCCACACTTGTCCTGATATCC-3’) or knockout primers (forward: 5’-

TGTCTGCCTACATGTTACTATGGTC-3’, reverse: 5’-

TGCGAGGCCAGAGGCCACTTGTGTAGC-3’) and run on 1% agarose gel.

Immunostaining

Cells were plated onto coverslips, fixed with 3.7% formaldehyde, rinsed x3 with PBS, permeabilized with 0.05% triton-X in PBS, and rinsed x3 with PBS. For coverslip staining, the primary antibody was incubated for 1 hour at 37°, rinsed x3 with PBS, secondary antibodies were incubated for 1 hour at 37°, rinsed x3 with PBS, and mounted.

Cell motility assays

Single cell motility and scratch assays were performed as previously published (Sehgal et al.,

2006). In single cell motility assays, cells were plated at 5x104 cells per 35mm well 12-18 hours before imaging. Cells were imaged every 5 minutes for 3 hours. In scratch wound assays, cells were plated confluently onto 35mm dishes 12-18 hours before imaging. A scratch wound was created, and cells were imaged every 10 minutes for 12 hours. Imaging was performed with a

Leica DMi8 microscope (Leica Microsystems, Buffalo Grove, IL, USA), and cell motility was analyzed using MetaMorph software (Molecular Devices, Sunnyvale, CA, USA). For each cell type, speed and processivity were calculated. Processivity was calculated as maximum cell displacement from cell’s origin divided by path length.

Flow cytometry

Adherent cells were trypsinized and fixed with 3.7% formaldehyde for 5 minutes. Cells were then rinsed 3x with PBS, incubated with primary antibody (1:200) and 30% normal goat serum at room temperature for 1 h, rinsed 3x with PBS, incubated with secondary antibody (1:200) at room temperature for 1 h, rinsed 3x with PBS, and analyzed with a Millipore GUAVA easyCyte flow cytometer (Hayward, CA, USA).

Antibodies and immunofluorescent probes

Mouse monoclonal against pan-keratin (4545) and rabbit monoclonal against vimentin (D21H3) were purchased from Cell Signaling (Beverly, MA, USA). Rat monoclonal against α6 integrin

(J1B5) was a generous gift from Dr. Caroline Damsky (University of California, San Francisco,

USA). Mouse monoclonal against β4 integrin (CD104) was purchased from BD Biosciences

(San Jose, CA, USA). Rabbit polyclonal antibody against paxillin (Y113) was purchased from

Abcam (Cambridge, MA, USA). Rat monoclonal antibody against α6 integrin (GOH3), rat monoclonal antibody against α5β1 integrin (BMC5), and mouse monoclonal antibody against

α2β1 integrin (MAB2141z) were purchased from EMD Millipore (Billenca, MA, USA).

Fluorescein-conjugated goat anti-mouse IgG antibody (115-095-166), rhodamine-conjugated goat anti-rabbit IgG antibody (111-025-144), fluorescein-conjugated goat anti-rabbit IgG antibody (111-095-144), rhodamine-conjugated goat anti-mouse IgG antibody (109-025-003) , and fluorescein-conjugated goat anti-rat IgG antibody (112-095-167) were obtained from

Jackson ImmunoResearch Laboratories (West Grove, PA, USA). Alexa-647 conjugated phalloidin was purchased from Life Technologies (Eugene, OR, USA).

References

Baroni, A., Buommino, E., De Gregorio, V., Ruocco, E., Ruocco, V. and Wolf, R. (2012). Structure and function of the epidermis related to barrier properties. Clin. Dermatol. 30, 257–262.

Chang, W., Worman, H. J. and Gundersen, G. G. (2015). Accessorizing and anchoring the LINC complex for multifunctionality. J Cell Biol 208, 11–22.

Hamill, K. J., Hopkinson, S. B., Hoover, P., Todorović, V., Green, K. J. and Jones, J. C. R. (2012). Fibronectin expression determines skin cell motile behavior. J. Invest. Dermatol. 132, 448–457.

Hiroyasu, S., Colburn, Z. T. and Jones, J. C. R. (2016). A hemidesmosomal protein regulates actin dynamics and traction forces in motile keratinocytes. FASEB J. 30, 2298–2310.

Hynes, R. O. (1992). Integrins: Versatility, modulation, and signaling in cell adhesion. Cell 69, 11–25.

Hynes, R. O. (2002). Integrins: Bidirectional, Allosteric Signaling Machines. Cell 110, 673–687.

King, S. J., Nowak, K., Suryavanshi, N., Holt, I., Shanahan, C. M. and Ridley, A. J. (2014). Nesprin-1 and nesprin-2 regulate endothelial cell shape and migration. Cytoskeleton 71, 423–434.

Morgan, J. T., Pfeiffer, E. R., Thirkill, T. L., Kumar, P., Peng, G., Fridolfsson, H. N., Douglas, G. C., Starr, D. A. and Barakat, A. I. (2011). Nesprin-3 regulates endothelial cell morphology, perinuclear cytoskeletal architecture, and flow-induced polarization. Mol. Biol. Cell 22, 4324–4334.

Rajgor, D. and Shanahan, C. M. (2013). Nesprins: from the nuclear envelope and beyond. Expert Rev. Mol. Med. 15, E5.

Theveneau, E. and Linker, C. (2017). Leaders in collective migration: are front cells really endowed with a particular set of skills? F1000Research 6.

Figures and figure legends

Figure 1. Confirmation of nesprin-2G KO keratinocytes. A) PCR of genomic DNA isolated from mouse tails. The WT primers produce a 1.8kB band and KO primers produce a 1.2kB band. Both primers work with DNA isolated from a heterozygote mouse. B) Immunofluorescence of WT and KO keratinocytes which are keratin positive and vimentin negative, indicating no cross- contamination of fibroblasts in the keratinocyte cell cultures.

Figure 2. Cell motility of WT and KO keratinocytes. A) Rose plots indicating the movement of 5 representative cells per cell type over 3 h with the movement of each cell beginning at 0,0. B)

Analyses of individual cell speed and processivity, a measure of directed, as detailed previously

(Hiroyasu et al., 2016). N = 3 independent replicates with >100 cells per experiment C)

Representative images of a scratch wound assay at 0 h and 12 h post scratch. D) Analyses of wound closure rate shown in C. Note that the KO cells fill the wound at a similar rate to WT cells after 12 h and 24 h post-scratch. N = 3 independent replicates with observing >10 points along the wound per experiment.

Figure 3. Immunofluorescence staining of WT and KO keratinocytes using antibodies against paxillin, an FA marker, and α6β4 integrin, a marker of HDs. A) WT and KO keratinocytes show similar α6 integrin and paxillin staining patterns. Actin staining marks the cell edge. B) WT and

KO keratinocytes show similar α6 and β4 integrin localization patterns. Actin staining marks the cell edge. C) Laminin-332, the ligand of α6β4 integrin, is organized similarly in the matrix of

WT and KO keratinocytes.

Figure 4. Flow cytometric analyses of integrin surface expression in WT and KO keratinocytes.

Cells were stained for A) α6 integrin, B) α5β1 integrin, and C) α2β1 integrin. Note that KO keratinocytes have increased α5β1 integrin compared to WT cells while the other heterodimers show no change.

CHAPTER THREE: NESPRIN-2G KNOCKOUT FIBROBLASTS EXHIBIT REDUCED

MIGRATION, CHANGES IN FOCAL ADHESION COMPOSITION, AND REDUCED

ABILITY TO GENERATE TRACTION FORCES*

Author Information

Alexandra Woychek, MPH, and *Jonathan C. R. Jones, PhD

School of Molecular Biosciences

PO Box 647520

Washington State University

Pullman, WA 99164-7520

*Corresponding author

Acknowledgements: We would like to thank Drs. Angelika A. Noegel for providing the nesprin-

2G knockout mouse line and Kevin Hamill for assistance in cell motility data analyses.

Funding statement: This research did not receive any specific grant from funding agencies in the public, commercial, or not-for-profit sectors.

*Published in Cytoskeleton, 2019, pp 1-9. Reprinted here by permission of John Wiley & Sons,

Inc.

Abstract

The nuclear envelope protein nesprin-2G is a component of the LINC (linker of nucleoskeleton and cytoskeleton) complex and is responsible for mechanical and signaling crosstalk between the nucleus and cytoskeleton. A prior study has demonstrated that nesprin-2G knockout mice show delayed wound healing. Our goal was to elucidate the mechanism underlying the delayed wound closure in this mouse model. We isolated primary fibroblasts from wildtype and knockout neonatal mice. Knockout cells exhibited decreased focal adhesion (FA) size, number, and intensity. Consistent with this result, FA protein expression levels were decreased in knockout cells. Additionally, knockout fibroblasts displayed an abnormal actin cytoskeleton, as evidenced by loss of TAN line formation and both cytoplasmic and peri-nuclear actin staining. Using collective and single cell motility assays, we found that knockout cells exhibited a reduction in both speed and directed migration. Traction force microscopy revealed that knockout fibroblasts generated fewer traction forces compared to WT fibroblasts. In summary, our data indicate that changes in actin organization and defects in FAs result in a reduced ability of knockout fibroblasts to generate traction forces needed for efficient motility.

Introduction

The nesprin protein family contains four members (nesprin-1G, -2G, -3, and -4) which localize to the nuclear membrane and are components of the LINC (linker of nucleoskeleton and cytoskeleton) complex, responsible for maintaining nuclear morphology (Rajgor and Shanahan,

2013). Nesprins are anchored to the outer nuclear membrane via a Klarsicht, ANC-1, and Syne

Homology (KASH) domain at the C-terminus which interacts with Sad1p-UNC-84 1/2 (SUN

1/2) proteins, found within the inner nuclear membrane (Rajgor and Shanahan, 2013). In the cytoplasm, variable N-terminal motifs of the different nesprin family members mediate their interactions with distinct cytoskeletal systems (Chang et al., 2015). Varying numbers of spectrin homology repeat (SR) domains are found between the N and C termini, depending on the nesprin protein and whether it has undergone alternative splicing (Autore et al., 2013; Roux et al., 2009;

Wilhelmsen et al., 2005). These SR domains serve both structural and functional purposes as they allow the interaction of the cytoskeleton and nucleus while acting as docking sites for cytoskeletal and signal transduction proteins (Djinovic-Carugo et al., 2002).

In this study, we focus on nesprin-2 Giant (nesprin-2G), an approximately 800kDa protein that interacts with the filamentous actin cytoskeleton in the cytoplasm via its two N-terminal calponin homology (CH) domains (Arsenovic et al., 2016). The C-terminal KASH domain of nesprin-2G interacts with SUN 1/2 proteins which are anchored to the inner nuclear membrane by lamin and emerin (Libotte et al., 2005). In humans, loss of nesprin-2G has been reported to cause musculoskeletal and neuronal defects (Cartwright and Karakesisoglou, 2014). In contrast, nesprin-2G knockout mice do not display a dramatic phenotype as they are normal in appearance and fertile. However, these mice exhibit delayed skin wound closure compared to their wildtype

counterparts (Lüke et al., 2008). Moreover, fibroblasts isolated from the cells of nesprin-2G knockout mice heal scratch wounds in vitro more slowly than wildtype cells (Lüke et al., 2008;

Rashmi et al., 2012). The mechanism underlying the delay in wound healing both in vivo and in vitro is not entirely clear although it has been suggested that this defect relates to changes in the polarity of the knockout cells (Lüke et al., 2008).

Cell polarity and directed migration depend on numerous structural and signaling proteins, including components of a cell-matrix adhesive device termed the focal adhesion (FA). FAs are complex, multiprotein structures, containing up to 200 proteins, and are built around a core of integrin heterodimers (Hopkinson et al., 2014). Integrins anchor the protein complex to the plasma membrane, interact with the actin cytoskeleton, and mediate cell adhesion to the extracellular matrix. In this study, we tested the hypothesis that FAs and/or the actin cytoskeleton are structurally and functionally aberrant in nesprin-2G knockout fibroblasts. To test our hypotheses, we assayed the expression of four FA proteins: paxillin, vinculin, tensin-1, and zyxin. These are all common markers of FAs and are incorporated into FAs at different stages during FA maturation (Partridge and Marcantonio, 2006). Paxillin and vinculin are incorporated early during FA assembly (Hu et al., 2014; Humphries et al., 2007), tensin-1 is recruited as adhesions mature (Zamir et al., 1999), and zyxin is a marker of mature FAs (Zaidel-Bar et al.,

2004). Paxillin can act as both a structural and a scaffolding component for many signaling intermediates involved in regulating polarity and motility (Dubois et al., 2017; Yu et al., 2010).

Interestingly, paxillin messenger RNAs are recruited to and locally translated at the leading edge in migrating cells, allowing efficient incorporation into nascent FAs (Woods et al., 2002).

Paxillin recruitment to FAs is mediated by vinculin, which also functions as both a structural and

signaling component of the FA (Rahman et al., 2016; Subauste et al., 2004; Ziegler et al., 2006).

Tensin-1 is a structural protein within the FA which interacts with other FA proteins to mediate anchorage of actin stress fibers to each FA (Bernau et al., 2016; Zhao et al., 2016). Zyxin is reported to be the “last” protein incorporated into a mature FA and stabilizes FA-actin interaction

(Zaidel-Bar et al., 2004).

FAs not only regulate actin cytoskeleton organization and signaling but facilitate the generation of traction forces, which are required for cell migration (Zaidel-Bar et al., 2004). Indeed,

“tugging” on actin stress fibers attached to FAs creates the tension required to allow a cell to pull itself forward (Han et al., 2016). Interestingly, as FA size increases, the generation of traction forces also increases (Goffin et al., 2006). Thus, as part of our analyses we also assessed whether traction forces are impacted by a loss of nesprin-2G. Indeed, here we demonstrate that loss of nesprin-2G in fibroblasts leads to an altered actin cytoskeleton, aberrant FAs, and loss of traction force generation. Our data indicate that these abnormalities are the cause of the reduced migration capacity of nesprin-2G knockout cells.

Results

Expression of FA proteins and actin in nesprin-2G knockout fibroblasts

Nesprin-2G knockout mice were developed by others by deletion of the actin binding domain of nesprin-2G resulting in mice functionally null for nesprin-2G (Lüke et al., 2008). Since these knockout mice were viable and fertile, it was possible to maintain a homozygous nesprin-2G knockout mouse line. The genotype of the knockout mice was confirmed by PCR (Fig. S1A,

Table 1). We isolated primary fibroblasts from the skin of both wildtype and nesprin-2G knockout mice. Fibroblasts prepared from the latter mice exhibited lack of expression of mRNA encoding nesprin-2G (Fig S1B, Table 2). Consistent with prior studies on the same knockout animal, immunoblotting analyses revealed that nesprin-2G was absent in extracts of the knockout cells (Fig. S1C-D) (Lüke et al., 2008). In contrast, there is no change in the levels of lamin A/C in the knockout cells compared with wildtype cells and so, for this and subsequent studies, lamin

A/C was used as a loading control for immunoblots (Fig. S1C). Several nesprin-2 isoforms of molecular weight ranging from ~36 to >250 kDa, produced from the gene encoding nesprin-2G, have been reported (Lüke et al., 2008). The levels of such isoforms are either the same or reduced compared to those in wildtype fibroblasts (Fig. S1C-D). This finding was also reported by Luke and his co-workers (Lüke et al., 2008).

Wildtype and nesprin-2G knockout cells were imaged to identify potential changes in FA structure and/or cytoskeleton organization due to the absence of nesprin-2G protein. We used confocal microscopy to focus on the substratum-attached surface of the cells. In wildtype fibroblasts, FAs, stained with antibodies against four FA components (paxillin, vinculin, zyxin, and tensin-1), were arranged towards the edge of each cell. Wildtype cells contained actin-rich

fibers that traversed the cytoplasm and extended toward FAs (Fig. 1A-D). In nesprin-2G knockout fibroblasts, which spread less well than wildtype cells, FAs appeared smaller and less well stained by antibodies against the four FA components we assayed (Fig. 1A-D, Table 3). We quantified FA number, size, and staining intensity in wildtype and knockout fibroblasts (Fig.

1A’-D’; Table 3). Knockout fibroblasts had significantly fewer and smaller FAs than wildtype fibroblasts. Moreover, the FAs of knockout fibroblasts were less intensely stained by FA antibodies (Fig. 1A’-D’, Table 2). It should be noted that, whereas all wildtype and knockout cells were positively stained by antibodies against paxillin, vinculin and zyxin, only 85% of wildtype cells and 50% of knockout cells were recognized by antibodies against the FA protein tensin-1.

The intensity of actin staining at all FAs of knockout cells was reduced compared to the FAs in wildtype fibroblasts (Fig. 1A’-D’, Table 3). In this regard, 93% of knockout fibroblasts also displayed less peri-nuclear actin (Fig. S3A-B). This is consistent with studies demonstrating that a reduction in expression or inhibition of nesprin-2G interaction with other proteins resulted in loss of transmembrane actin-associated nuclear (TAN) lines (Kutscheidt et al., 2014; Luxton et al., 2011). As part of this study we used DAPI staining to visualize nuclei in both wildtype and knockout cells. Interestingly, whereas DAPI stained the nuclei of wildtype cells efficiently,

DAPI staining in the knockout fibroblasts was barely detectable (Fig. S3A-B).

Consistent with reduction in staining of FA components, knockout fibroblasts showed decreased levels of paxillin, vinculin, zyxin, and tensin-1 protein as determined by immunoblotting (Fig.

2A-B). In contrast, message levels of paxillin increased in nesprin-2G knockout fibroblasts while

message levels of vinculin, zyxin, and tensin-1 appeared unaffected, relative to GAPDH (Fig.

2C). This suggested that the knockout cells exhibit either a translational defect or an inability to incorporate translated FA proteins into FAs. To address this issue, knockout cells were incubated with MG132, a proteasome degradation inhibitor. FA proteins in the knockout cells showed significant increases in their expression following drug treatment, compared to untreated controls

(Fig. 2A-B). Moreover, when knockout cells were incubated with MG132 there was a rescue of

FA staining, number and size (Fig S2, Table 4). Together these results indicate that the loss of

FA protein is not due to a translational defect in knockout cells. Rather, our results imply that their loss is a result of accelerated protein turnover.

Nesprin-2G knockout fibroblasts exhibit migration defects

Consistent with previous studies, we observed a defect in the ability of nesprin-2G fibroblasts to close a scratch wound in vitro compared to their wildtype counterparts (Lüke et al., 2008) (Fig.

3A-B). To extend this observation and shed light into why the cells show a migration defect, we undertook single cell motility assays which are represented by rose plots (Fig. 3C). We compared the speed and processivity of individual wildtype and nesprin-2G knockout fibroblasts. Speed was calculated as the total cell movement divided by the total time of the assay. Processivity, a measure of directed cell migration, was the maximum displacement from the point of origin of the cell divided by the length of the path traveled (Hiroyasu et al., 2016). The knockout fibroblasts exhibited a 23% reduction in speed and a 16% reduction in processivity compared with their wildtype counterparts (Fig. 3D).

Nesprin-2G knockout fibroblasts exhibit decreased traction forces

Nesprin-2G has been shown to respond to myosin-dependent tension using a fluorescence resonance energy transfer-based tension biosensor (Arsenovic et al., 2016). This led us to hypothesize that nesprin-2G knockout fibroblasts would exhibit defects in their ability to generate traction forces which are regulated by the acto-myosin cytoskeleton as well as FAs

(Zaidel-Bar et al., 2004). To test our hypothesis, wildtype and knockout fibroblasts were plated onto coverslips coated with polyacrylamide gel containing fluorescent beads and coated with fibronectin. Change in bead displacement before and after cell trypsinization was measured to determine traction forces generated by an individual cell (Fig. 4A). Nesprin-2G knockout fibroblasts generated 33.9% fewer total traction forces and each traction force generated was

38.7% smaller in knockout fibroblasts (Fig. 4B). Even when controlling for wildtype and knockout fibroblast cell size, these differences were still significant. In addition, knockout fibroblasts displayed 22% reduction in total traction forces generated per square micron of cell area and 24% reduction in magnitude per square micron of cell area (Fig. 4B).

Discussion

In this study we have demonstrated that, compared to wildtype cells, nesprin-2G knockout fibroblasts exhibit dramatic aberrations in the organization of their actin cytoskeleton.

Specifically, nesprin-2G knockout fibroblasts displayed reduced actin at FAs, reduced overall actin staining, and decreased TAN lines, the latter observation being consistent with a previous report (Lüke et al., 2008). This is despite the presence of nesprin-2 isoforms in the knockout cells. In addition to changes in actin organization, we also report that the nuclei of knockout cells exhibit reduced DAPI staining compared to their wildtype counterparts. This reduction suggests that there may be a loss of heterochromatin or premature aging of knockout cells (Tsurumi and

Li, 2012). Further experimentation will be required to distinguish these possibilities. Regardless, loss of heterochromatin is indicative of global changes in transcription (Fan et al., 2005, 1).

Interestingly, we do not detect transcriptional changes in the expression of genes encoding FA proteins. Rather, as we discuss next, our data demonstrate that changes in expression of FA proteins in knockout cells is controlled post-transcriptionally.

FAs in nesprin-2G cells knockout cells were smaller, and protein expression levels of several key

FA components were significantly decreased. These deficiencies were not the result of down regulation in transcription of FA proteins since the message levels of paxillin, vinculin, tensin-1, and zyxin in knockout cells were elevated or the same as wildtype fibroblasts. Rather, the FA proteins we analyzed were translated, but were not incorporated into FAs. Indeed, our studies indicate that the loss of FA proteins in knockout cells is due to their being targeted for degradation via the proteasome pathway. This raises the question why the incorporation of FA proteins into FAs is impeded in nesprin-2G knockout cells. We hypothesize that reduced levels

of actin at assembling FAs reduces vinculin recruitment to nascent adhesions since vinculin binds actin. This reduction then will have consequences for paxillin recruitment since paxillin binds activated vinculin (Carisey and Ballestrem, 2011; Humphries et al., 2007).

We also demonstrate that FAs in nesprin-2G knockout fibroblasts are deficient in zyxin compared to wildtype cells. Indeed, zyxin is the last protein incorporated into FAs, and staining with zyxin is commonly used to quantify FA maturation (Stehbens and Wittmann, 2014; Zaidel-

Bar et al., 2004). An absence of zyxin in FAs implies that FAs of knockout cells have failed to mature. This lack of maturity is also consistent with the decreased association of actin with the

FAs of knockout cells since actin contraction and intracellular tension are needed to promote maturation of FAs (Hinz et al., 2003).

I. Finally, our data reveal that nesprin-2G knockout fibroblasts exert fewer traction forces on their substrates than their wildtype counterparts. Contraction of the actin cytoskeleton and the tension the cytoskeleton exerts on FAs are needed to produce traction forces which are needed to pull a cell forward (Gallegos et al., 2011; Han et al., 2016; Oakes and Gardel, 2014). Thus, the reduced ability of knockout cells to generate traction forces is readily explained by a decrease in expression of FA proteins, the small size of FAs and a perturbation of the actin-FA interaction we have observed in knockout cells. In addition, the decreases in traction forces induced by nesprin-2G knockout cells can account for the reduction in their speed. It should also be noted that others have reported that cells exhibiting nesprin-2G knockout have cell polarity defects evidenced by centrosome and Golgi positioning abnormalities (Lüke et al., 2008). Indeed, together, these cell polarity defects and reduced traction force issues help explain why knockout

cells show aberrant directional migration/processivity as well as collective cell migration, manifested in delayed closure of scratch wounds of monolayer cultures.

In summary, our data indicate that changes in actin organization and FA defects result in a reduced ability of nesprin-2G knockout fibroblasts to generate the traction forces on their substrate needed for efficient motility. In mice, skin wounds heal by contracture of the dermis, which is dependent on the activities and force generation capacities of stromal fibroblasts

(Zomer and Trentin, 2018). Prior reports indicate that wounds in the skin of mice lacking nesprin-2G heal more slowly than those of their wildtype counterparts. Our data provide a mechanistic explanation for this observation. In wounded skin, the reduced capacity of nesprin-

2G knockout cells to generate traction forces, due to abnormal FA and actin localization, would impede the ability of stromal fibroblasts to contract the dermis, leading to an inhibition of wound closure.

Materials and methods

Primary cell isolation

C57/Bl6 wildtype and nesprin-2G knockout mice as previously published (Hamill et al., 2012).

Briefly, skins were incubated in 2.4 U/mL dispase overnight to separate epidermis from dermis.

The dermis was chopped up and then incubated in 0.05% trypsin for 30 minutes. The addition of

Dulbecco’s modified Eagle’s medium (DMEM) (HyClone, Logan, UT, USA) supplemented with

10% fetal bovine serum (Sigma-Aldrich, St. Louis, MO, USA) was added to the trypsin solution at a 1:1 ratio, and skin debris allowed to settle for 4 minutes. Cells in the resulting supernatant were collected by centrifugation. That fibroblasts were isolated was confirmed by the presence of vimentin and absence of keratin by immunostaining.

Cell culture conditions

Primary mouse fibroblasts were cultured in Dulbecco’s modified Eagle’s medium (DMEM)

(HyClone, Logan, UT, USA) supplemented with 10% fetal bovine serum (Sigma-Aldrich, St.

Louis, MO, USA) and 1% penicillin/streptomycin mixture. Cells were maintained at 37°C in 8% humidified CO2. For some studies, cells were incubated with 10μM MG132 for 2 h, and subsequently processed for immunostaining or SDS-PAGE/western blotting as below.

Cell motility assays

Single cell motility and scratch assays were performed as previously described (Sehgal et al.,

2006). In single cell motility assays, cells were plated at 5x104 cells per 35mm well 12-18 hours before imaging. Cells were imaged every 5 minutes for 3 hours. In scratch wound assays, cells

were plated at confluence onto 35mm dishes 12-18 h before imaging. A scratch wound was created, and scratch area imaged every 10 minutes for 12 h. Imaging was performed with a Leica

DMi8 microscope (Leica Microsystems, Buffalo Grove, IL, USA), and cell motility was analyzed using MetaMorph software (Molecular Devices, Sunnyvale, CA, USA). For single cell motility assays, speed and processivity were calculated as detailed previously. Processivity was calculated as maximum cell displacement from cell’s origin divided by path length (Hiroyasu et al., 2016).

Traction force microscopy

Traction force microscopy was performed as described elsewhere (Hiroyasu et al., 2016).

Briefly, fibroblasts were plated onto coverslips coated with polyacrylamide gel containing

Alexa-647 fluorescent beads. Live cells and beads were imaged using a Leica TCS SP8 X microscope (Leica Microsystems, Buffalo Grove, IL, USA). Then, cells were trypsinized to image beads in an uncontracted state. Bead displacement was used to calculate traction forces using StackReg, Iterative Particle Image Velocimetry, and FTTC plugins for ImageJ, following protocols detailed by others (Tseng et al., 2012).

Immunostaining

Cells were plated onto fibronectin coated coverslips (10ug/mL), fixed with 3.7% formaldehyde, rinsed x3 with PBS, permeabilized with 0.05% triton-X in PBS, and rinsed x3 with PBS. For coverslip staining, the primary antibody was incubated for 1 hour at 37°C, rinsed x3 with PBS, secondary antibodies were incubated for 1 hour at 37°C, rinsed x3 with PBS, and mounted.

Quantification of FA size, number, and actin intensity

Fibroblasts were imaged with an SP5 confocal microscope (Leica Microsystems, Buffalo Grove,

IL, USA). Images were taken at the substratum-attached surface of the cells. The images of FA

proteins were thresholded in ImageJ and segmented using the R package Bioi version 0.2.9. The segmented images were used to determine the total intensity of the FA proteins and actin at FAs.

Two-sample Wilcoxon tests were performed to quantify differences between different fibroblast populations.

SDS-PAGE, immunoblot, and RT-qPCR

Cell lysates were prepared for SDS-PAGE/western blotting as detailed elsewhere (Harlow and

Lane, 2014). To assess loading, cell lysates were run on 15% polyacrylamide gels for 40 minutes, and transferred for 2 h at 110V to nitrocellulose, and probed with antibodies to GAPDH and lamin A/C. Once loading levels were determined, lysates containing comparable levels of protein were run on 6% gels for 7.5 h and transferred to nitrocellulose for 16 h at 30V to detect the large nesprin-2G protein or run on 7.5% gels for 40 minutes and transferred to nitrocellulose for 2 h at 110V to detect lower molecular weight nesprin-2 isoforms. Western blots were visualized using a myECL Imager (Thermo Fisher Scientific) and quantified using ImageJ.

Molecular weights were determined relative to the Thermo Scientific Page Ruler Plus Prestained

Protein Ladder. Total RNA was extracted from cells using a GeneJET RNA Purification Kit

(Thermo Fisher Scientific). 5 ng of extracted RNA was used to synthesize cDNA using a

Maxima H Minus First Strand cDNA Synthesis Kit with dsDNase (Thermo Fisher Scientific). mRNA expression was analyzed using a Step One Plus Real-Time PCR System (Thermo Fisher

Scientific) with TaqMan Fast Advanced Master Mix and TaqMan Gene Expression Assays

(Thermo Fisher Scientific) for GAPDH (Mm99999915_g1), paxillin (Mm00433209_m1), vinculin (Mm00447745_m1), zyxin (Mm00496120_m1), and tensin-1 (Mm00452886_m1).

Relative gene expression was quantified by ΔΔCt and normalized to GAPDH.

Antibodies and immunofluorescent probes

Rabbit polyclonal antibody against paxillin (Y113) was purchased from Abcam (Cambridge,

MA, USA). Rabbit antibody against lamin A/C (2032S) was purchased from Cell Signaling

Technology (Beverly, MA, USA). Mouse monoclonal antibody against vinculin (V9131), mouse monoclonal antibody against zyxin (Z0377), and rabbit antibody against tensin-1 (SAB4200297) were purchased from Sigma (St. Louis, MO, USA). Rabbit polyclonal antibody against zyxin

(ABC1387) and mouse polyclonal antibody against glyceraldehyde-3-phosphate dehydrogenase

(GAPDH, AB2302) were purchased from EMD Millipore (Temecula, CA, USA). 4′,6- diamidino-2-phenylindole, dilactate (DAPI, D3571) and rabbit polyclonal antibody against nesprin-2G (PA5-78438) were purchased from Invitrogen (Eugene, OR, USA). Fluorescein- conjugated goat anti-rabbit IgG antibody (111-095-144) was obtained from Jackson

ImmunoResearch Laboratories (West Grove, PA, USA). Rhodamine-conjugated phalloidin was purchased from Life Technologies. Horseradish peroxidase–conjugated goat anti-mouse IgG

(115-035-166) and anti-rabbit IgG (7074S) antibodies were purchased from Jackson

ImmunoResearch Laboratories (West Grove, PA, USA) and Cell Signaling Technology

(Beverly, MA, USA), respectively.

References

Carisey, A. and Ballestrem, C. (2011). Vinculin, an adapter protein in control of cell adhesion signalling. Eur. J. Cell Biol. 90, 157–163.

Cartwright, S. and Karakesisoglou, I. (2014). Nesprins in health and disease. Semin. Cell Dev. Biol. 29, 169–179.

Chang, W., Worman, H. J. and Gundersen, G. G. (2015). Accessorizing and anchoring the LINC complex for multifunctionality. J Cell Biol 208, 11–22.

Djinovic-Carugo, K., Gautel, M., Ylänne, J. and Young, P. (2002). The spectrin repeat: a structural platform for cytoskeletal protein assemblies. FEBS Lett. 513, 119–123.

Dubois, F., Alpha, K. and Turner, C. E. (2017). Paxillin regulates cell polarization and anterograde vesicle trafficking during cell migration. Mol. Biol. Cell 28, 3815–3831.

Fan, Y., Nikitina, T., Zhao, J., Fleury, T. J., Bhattacharyya, R., Bouhassira, E. E., Stein, A., Woodcock, C. L. and Skoultchi, A. I. (2005). Histone H1 depletion in mammals alters global chromatin structure but causes specific changes in gene regulation. Cell 123, 1199–1212.

Gallegos, L., Ng, M. R. and Brugge, J. S. (2011). The myosin-II-responsive focal adhesion proteome: a tour de force? Nat. Cell Biol. 13, 344–346.

Goffin, J. M., Pittet, P., Csucs, G., Lussi, J. W., Meister, J.-J. and Hinz, B. (2006). Focal adhesion size controls tension-dependent recruitment of α-smooth muscle actin to stress fibers. J Cell Biol 172, 259–268.

Hamill, K. J., Hopkinson, S. B., Hoover, P., Todorović, V., Green, K. J. and Jones, J. C. R. (2012). Fibronectin expression determines skin cell motile behavior. J. Invest. Dermatol. 132, 448–457.

Han, S. J., Rodriguez, M. L., Al-Rekabi, Z. and Sniadecki, N. J. (2016). Spatial and temporal coordination of traction forces in one-dimensional cell migration. Cell Adhes. Migr. 10, 529–539.

Harlow, E. and Lane, D. (2014). Antibodies a laboratory manual. 2nd ed. (ed. Greenfield, E. A.) Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press.

Hinz, B., Dugina, V., Ballestrem, C., Wehrle-Haller, B. and Chaponnier, C. (2003). α-smooth muscle actin is crucial for focal adhesion maturation in myofibroblasts. Mol. Biol. Cell 14, 2508–2519.

Hiroyasu, S., Colburn, Z. T. and Jones, J. C. R. (2016). A hemidesmosomal protein regulates actin dynamics and traction forces in motile keratinocytes. FASEB J. 30, 2298–2310.

Hu, Y.-L., Lu, S., Szeto, K. W., Sun, J., Wang, Y., Lasheras, J. C. and Chien, S. (2014). FAK and paxillin dynamics at focal adhesions in the protrusions of migrating cells. Sci. Rep. 4, 6024–6030.

Libotte, T., Zaim, H., Abraham, S., Padmakumar, V. C., Schneider, M., Lu, W., Munck, M., Hutchison, C., Wehnert, M., Fahrenkrog, B., et al. (2005). Lamin A/C–dependent localization of nesprin-2, a giant scaffolder at the nuclear envelope. Mol. Biol. Cell 16, 3411–3424.

Luxton, G. W. G., Gomes, E. R., Folker, E. S., Worman, H. J. and Gundersen, G. G. (2011). TAN lines: a novel nuclear envelope structure involved in nuclear positioning. Nucleus 2, 173–181.

Oakes, P. W. and Gardel, M. L. (2014). Stressing the limits of focal adhesion mechanosensitivity. Curr. Opin. Cell Biol. 30, 68–73.

Partridge, M. A. and Marcantonio, E. E. (2006). Initiation of attachment and generation of mature focal adhesions by integrin-containing filopodia in cell spreading. Mol. Biol. Cell 17, 4237–4248.

Rahman, A., Carey, S. P., Kraning-Rush, C. M., Goldblatt, Z. E., Bordeleau, F., Lampi, M. C., Lin, D. Y., García, A. J. and Reinhart-King, C. A. (2016). Vinculin regulates directionality and cell polarity in two- and three-dimensional matrix and three- dimensional microtrack migration. Mol. Biol. Cell 27, 1431–1441.

Rajgor, D. and Shanahan, C. M. (2013). Nesprins: from the nuclear envelope and beyond. Expert Rev. Mol. Med. 15, E5.

Stehbens, S. and Wittmann, T. (2014). Analysis of focal adhesion turnover: A quantitative live cell imaging example. Methods Cell Biol. 123, 335–346.

Tseng, Q., Duchemin-Pelletier, E., Deshiere, A., Balland, M., Guillou, H., Filhol, O. and Théry, M. (2012). Spatial organization of the extracellular matrix regulates cell–cell junction positioning. Proc. Natl. Acad. Sci. 109, 1506–1511.

Tsurumi, A. and Li, W. (2012). Global heterochromatin loss. Epigenetics 7, 680–688.

Yu, J. A., Deakin, N. O. and Turner, C. E. (2010). Emerging role of paxillin-PKL in regulation of cell adhesion, polarity and migration. Cell Adhes. Migr. 4, 342–347.

Zaidel-Bar, R., Cohen, M., Addadi, L. and Geiger, B. (2004). Hierarchical assembly of cell– matrix adhesion complexes. Biochem. Soc. Trans. 32, 416–420.

Zamir, E., Katz, B. Z., Aota, S., Yamada, K. M., Geiger, B. and Kam, Z. (1999). Molecular diversity of cell-matrix adhesions. J. Cell Sci. 112, 1655–1669.

Ziegler, W. H., Liddington, R. C. and Critchley, D. R. (2006). The structure and regulation of vinculin. Trends Cell Biol. 16, 453–460.

Zomer, H. D. and Trentin, A. G. (2018). Skin wound healing in humans and mice: Challenges in translational research. J. Dermatol. Sci. 90, 3–12.

Tables and table legends

Wildtype forward primer 5’ CCATGAAAGCTCTAGTCAGAAAGAGC 3’

Wildtype reverse primer 5’ CAGCCACACTTGTCCTGATATCC 3’

Knockout forward primer 5’ TGTCTGCCTACATGTTACTATGGTC 3’

Knockout reverse primer 5’ TGCGAGGCCAGAGGCCACTTGTGTAGC 3’

Table 1. Primer sets used for PCR analysis to identify wildtype and knockout alleles of the nesprin-2G gene (SYNE2).

Exon 1 forward primer 5’ TGCTGCCCACGGAAGATGGAGAGGG 3’

Exon 8 reverse primer 5’ GGCCAGGCGCATCCTTCGAATA 3’

Table 2. Primer sets used for PCR analysis to identify nesprin-2G message presence in wildtype and knockout cells.

Protein Reduction of Reduction of Reduction of Reduction of Reduction of FA number in FA area in KO FA intensity actin intensity total actin KO relative to relative to in KO relative at FA in KO intensity in wildtype cells wildtype cells to wildtype relative to KO relative to cells wildtype cells wildtype cells Paxillin 52% 42.3% 47.8% 40.4% 17.8%

Vinculin 31.9% 16% 19.8% 26.3% 46.1%

Tensin-1 49.5% 14.3% 21.6% 20.3% 31.8%

Zyxin 36.5% 12.5% 12.8% 53.5% 50.3%

Table 3. Quantification of the differences in the numbers of FAs, their size and staining intensity in knockout cells versus wildtype cells following staining with antibodies against paxillin, vinculin, tensin-1, and zyxin. Actin association with FAs as well as total actin staining is also reduced in knockout versus wildtype cells.

Protein FA number in FA area in KO FA intensity Actin intensity Total actin KO + MG132 + MG132 in KO + at FA in KO + intensity in relative to relative to MG132 MG132 KO + MG132 wildtype cells wildtype cells relative to relative to relative to wildtype cells wildtype cells wildtype cells Paxillin 91.3% 94.4% 102.1% 97.4% 92.4%

Table 4. Numbers of FAs, FA size, FA staining intensity (with paxillin antibody), actin association with FAs, and total actin staining in knockout cells incubated with MG132 for 2 h relative to wildtype cells.

Figures and figure legends

(Figure legend on next page)

Figure 1. Nesprin-2G knockout fibroblasts exhibit abnormal FAs. Representative images of wildtype and nesprin-2G knockout fibroblasts stained for paxillin and actin, vinculin and actin, tensin-1 and actin, and zyxin and actin are shown in A, B, C, and D, as indicated. A’, B’, C’, and

D’ present quantifications of the images presented in A-D. In A’ the average areas covered by wildtype and knockout cells are shown. In A’-D’, FA numbers per cell, FA area, intensity of staining of FAs, actin intensity at FAs and total actin intensity throughout the cell are depicted.

Scale bars in A-D = 20 μm. N = 3 independent experiments, > 50 cells per experiment. One- tailed Wilcoxon rank sum test with Benjamini-Hochberg correction. *, P<0.05; **, P<0.01; ***,

P<0.001. Error bars are standard error of the mean.

Figure 2. Nesprin-2G knockout fibroblasts exhibit reduced FA protein levels and altered FA message levels. A) Representative images of immunoblots of the extracts of the indicated cells, processed using antibodies against the FA proteins paxillin, vinculin, tensin-1, and zyxin. Lamin is the loading control. B) Quantification of A. N = 3 independent experiments. In A, immunoblots of extracts of cells treated with MG132 are also shown. C) Message levels of the indicated FA components in knockout relative to wildtype cells, normalizing to GAPDH. N = 3 independent experiments. *, P<0.05; ***, P<0.001. Error bars are standard error of the mean.

Figure 3. Nesprin-2G knockout fibroblasts show decreased collective cell migration in a scratch assay and single cell motility. In A) images of the same wildtype and knockout cells were taken at the time of scratching and then 12 h later. The dotted red lines indicate the wound margins.

Scale bar = 100nm. B) Quantification of wound closure rate of the scratches. N = 3 independent experiments. C) Representative rose plots of single cell motility with 4 cells per type shown. The movement of each cell begins at 0,0, and lines indicate motility over 3 h. D) Quantitation of speed and processivity of the cells shown in C. N = 3 independent experiments, > 100 cells per experiment. One-tailed Wilcoxon rank sum test. *, P<0.05; **, P<0.01; ***, P<0.001. Error bars are standard error of the mean.

Figure 4. Nesprin-2G knockout fibroblasts generate less traction forces (TFs) than their wildtype counterparts. A) Representative images of wildtype and knockout cells viewed by differential interference microscopy (DIC). The particle image velocimetry (PIV) image of the same cells indicates where TFs are generated by the cells. B) Quantification of the TFs generated by wildtype and knockout cells. Pa = pascals. N = 3 independent experiments, > 20 cells per experiment. One-tailed Wilcoxon rank sum test with Benjamini-Hochberg correction. *, P<0.05;

**, P<0.01; ***, P<0.001. Error bars are standard error of the mean.

Supplementary figures and supplementary figure legends

Figure S1. Verification of nesprin-2G knockout. A) Genotyping of heterozygote, wildtype, and knockout mouse lines using primer sets to identify wildtype (WT) (Table 1), resulting in a product of 1.8 kB, and knockout (KO) alleles (Table 1), resulting in a product of 1.2 kB. B) PCR of nesprin-2G message level in WT and KO cells passaged once (Table 2). The arrow indicates amplification of a nesprin-2G message in WT cells but not in KO cells. C) Western blotting of wildtype and KO cell extracts are shown. Cell lysates were first processed using antibodies against GAPDH and lamin A/C in order to establish appropriate gel loading. Subsequently, cell lysates were prepared for Western blotting to enable detection of nesprin-2 isoforms in the 60-

250kDa range (see Materials and Methods). Arrows indicate isoforms identified by previous

authors (Lüke et al., 2008). D) Lysates were also prepared for Western blot so that nesprin-2G at approximately 800kDa could be detected (see Materials and Methods). Arrow indicates absence of nesprin-2G in KO cell extracts.

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Figure S2. MG132 incubation restores FA staining in knockout cells. A) Representative images of paxillin and actin staining in wildtype fibroblasts, nesprin-2G knockout fibroblasts, and

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nesprin-2G knockout fibroblasts incubated with MG132 for 2 h. B) Quantification of FAs in cells treated as in panel A. Cell area, FA numbers per cell, FA area, intensity of staining of FAs, actin intensity at FAs and total actin intensity throughout the cell are presented. Scale bar = 20

μm. N = 3 independent experiments, > 50 cells per experiment. One-tailed Wilcoxon rank sum test with Benjamini-Hochberg correction. ***, P<0.001. Error bars are standard error of the mean.

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Figure S3. Nesprin-2G knockout affects nuclear staining and transmembrane actin-associated nuclear (TAN) line formation. In A and B, images of a wildtype and knockout cell stained with

DAPI and actin, respectively. Images were generated using the same laser settings. Scale bar =

10 μm.

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CHAPTER FOUR: THE α6 INTEGRIN 3’UTR REGULATES LOCALIZATION OF THE α6β4

INTEGRIN HETERODIMER

Abstract

The α6β4 integrin heterodimer is an essential component of hemidesmosomes (HDs) and HD- related structures, which adhere epithelial cells to the underlying extracellular matrix. In this study, we focused on the importance of the α6 integrin 3’ untranslated region (UTR) in α6β4 integrin localization. To do so, A549 cells (a type II lung alveolar cell line) and immortalized human epidermal keratinocytes (iHEK) were infected with adenovirus encoding the entire α6 integrin protein with or without portions of its 3’UTR. In infected A549 cells, we detected α6β4 integrin heterodimers containing the product of the adenovirus, regardless of whether the α6 integrin 3’UTR was present. However, only those α6 integrin proteins whose messages contained bases 4770-5633 of the α6 integrin 3’UTR were targeted to matrix adhesion sites.

Moreover, overexpression of the full length α6 integrin 3’UTR, minus the coding sequence, in

A549 cells disrupts the localization of endogenous α6β4 integrin heterodimers. In α6 integrin depleted iHEKs, restoring α6 integrin expression using the coding sequence alone resulted in α6 integrin preferentially forming α6β1 rather than α6β4 integrin heterodimers. α6β4 integrin was only observed in knocked down cells following infection of adenovirus encoding the α6 integrin coding sequence with its 3’UTR. In summary, our data indicate that the α6 integrin 3’UTR is a key regulator of α6β4 integrin heterodimer assembly and incorporation at sites of cell-matrix adhesion.

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Introduction

Cell adhesion and migration are critical processes for normal cell functioning, and these processes are aided by cell-matrix adhesion protein complexes: hemidesmosomes (HDs) and focal adhesions (FAs). The core of these complexes contains integrin heterodimers which interact with both the extracellular matrix and other cytoplasmic elements (Hynes, 1992). Each integrin heterodimer is a transmembrane, non-covalently bound complex that contains one α subunit and one β subunit. There are 18 α subunits and 8 β subunits which can form a variety of heterodimers, for example α2β1, α3β1, αvβ1, and α2β5. However, only 24 different heterodimers have been identified (de Melker and Sonnenberg, 1999). The α6β4 integrin heterodimer is unusual, not only because of the large size of the β4 integrin cytoplasmic tail, but also because it is only found in HDs and HD-related matrix adhesions, rather than in FAs (Hopkinson et al.,

2014). HDs adhere epithelial cells to the extracellular matrix in several tissues including the skin, cornea, bladder, and some glands. In contrast, HD-related adhesions are found in certain alveolar cells in the lung and gut epithelial cells where they are presumed to mediate adhesion to the underlying connective tissue (Colburn and Jones, 2016; Colburn and Jones, 2018; Jones et al.,

1991).

The α6 integrin subunit has two possible β subunit partners: β1 and β4 integrin. In cells which express both β1 and β4 integrin, α6 integrin preferentially pairs with β4, but how this is controlled is unclear (Hemler et al., 1989; Lee et al., 1992). Moreover, the targeting mechanism of α6β4 integrin to HD and HD-related matrix adhesion sites over other substrate attachment sites, including FAs, is also not understood. In this study, we have identified the 3’ untranslated region (UTR) of the α6 integrin messenger RNA (mRNA) as a key regulator of the localization

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of α6β4 integrin heterodimers as well as the formation of α6β4 versus α6β1 integrin heterodimers. In this regard, the 3’UTRs of mRNAs are known to regulate the shuttling of mRNAs to their site of translation by a sequence known as a zipcode. Although zipcode sequences can be located within coding regions of mRNA, generally, they are located within the

3’UTR (Andreassi and Riccio, 2009). For example, the zipcode in the β-actin message directs it to the site of FAs where it is translated (Katz et al., 2012). Likewise, zipcodes in vinculin, actinin-4, and α3 integrin mRNAs direct their translation to the FA (Adereth et al., 2005; Babic et al., 2009). Therefore, we tested the hypothesis that there is a zipcode within the α6 integrin

3’UTR involved in localization and/or formation of the α6β4 integrin heterodimer.

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Results and discussion

A zipcode within the α6 integrin 3’UTR is required for α6β4 heterodimer localization

Zipcodes are sequences within 3’UTRs of mRNAs that direct their localization and subsequent translation. For example, the zipcode of α3 integrin was demonstrated to be necessary for α3 integrin protein translation and incorporation into FAs (Adereth et al., 2005).

We therefore posited that the α6 integrin 3’UTR might be an important determinant of targeting

α6β4 integrin heterodimers to HDs and HD-related adhesions. To assess this possibility, we created adenoviruses encoding a GFP-tagged α6 integrin coding sequence with and without its

3’UTR and used them to infect A549, type II alveolar-like, epithelial cells (Figure 1A). In A549 cells, α6β4 integrin is distributed in puncta at the basal surface towards the edges of the cell

(Figure 1B), as we previously demonstrated (Colburn and Jones, 2016; Colburn and Jones,

2018). We first evaluated whether the 3’UTR was necessary for ensuring incorporation of the

GFP tagged α6 integrin into such puncta and co-distribution with its binding partner, β4 integrin.

Adenoviral transduced cells were processed for immunofluorescence microscopy using GFP (α6 integrin) and β4 integrin antibodies (Figure 1C-D). GFP-tagged α6 integrin protein encoded by mRNA lacking the 3’UTR fails to colocalize with β4 integrin at the basal surface of transduced

A549 cells. In contrast, cells expressing GFP α6 integrin protein encoded by mRNA containing the full 3’UTR localizes with endogenous α6β4 integrin-containing puncta along the substratum- attached surface of infected cells (Figure 1C-D).

To identify the region of the α6 integrin message containing a zipcode, we generated a series of deletions its 3’UTR and used these adenoviral constructs to transduce A549 cells. GFP- tagged α6 protein encoded by the mRNA sequences with fragments of the α6 integrin 3’UTR

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lacking bases 4133-5770 (α6-GFP+3’UTR (3233-4133), α6-GFP+3’UTR (3233-4770), and α6-

GFP+3’UTR (3233-5440)) fail to colocalize with β4 integrin at the basal surface of infected A549 cells (Figure 1 A and C-D). In sharp contrast, GFP-tagged α6 protein encoded by the mRNA sequences encompassing bases 4133-5770 (α6-GFP+3’UTR, α6-GFP+3’UTR (4133-5633) and α6-

GFP+3’UTR (4770-5770)) exhibit robust co-distribution with β4 integrin in a punctate pattern towards the edge and at the substrate-attached surface of cells, similar to the localization of endogenous α6β4 integrin protein (Figure 1A and C-D). Together these data support the notion that there is a zipcode contained within the sequence 5440-5633 of the α6 3’UTR mRNA which determines α6β4 integrin heterodimer localization to specific matrix/substrate adhesion sites.

The lack of α6β4 integrin heterodimer localization at HD-related adhesions in A549 cells may be due to an inability of its subunits to co-assemble. To test this possibility, we performed an immunoprecipitation using anti-GFP beads in cells infected with α6-GFP, α6-GFP+3’UTR

(3233-4133), or α6-GFP+3’UTR (4133-5633) constructs (Figure S1A, Figure 1A). Regardless of whether the coding sequence lacked (α6-GFP and α6-GFP+3’UTR (3233-4133)) or contained (α6-

GFP+3’UTR (4133-5633)) the fragment of the α6 integrin 3’UTR necessary for targeting α6 integrin product to specific matrix adhesion sites, the resulting α6 integrin protein was precipitated by

GFP antibodies, together with β4 integrin (Figure S1A). This indicates that α6β4 integrin heterodimers form even when the α6 integrin message lacks a functional 3’UTR. We also observed no change in solubility of the α6β4 integrin complexes in cells infected with adenovirus encoding α6 integrin with (α6-GFP+3’UTR (4133-5633)) and without (α6-GFP+3’UTR (3233-4133)) fragments of the 3’UTR that contain the sequence needed for specific matrix site targeting

(Figure S1B). These data indicate that the α6 integrin 3’UTR is not necessary for assembly or

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solubility of α6β4 integrin heterodimers in A549 cells. Rather, the α6 integrin 3’UTR is required for targeting the α6β4 heterodimer to specific adhesion sites.

Overexpression of the α6 integrin 3’UTR alters protein expression and motility of A549s

To further validate the role of the α6 integrin 3’UTR in determining localization of α6β4 integrin complexes, we next infected A549s with adenovirus encoding the full length α6 integrin

3’UTR tagged with mCherry (α6-mCherry) without the α6 integrin coding sequence.

Interestingly, α6β4 integrin heterodimer localization is dramatically disrupted in such cells compared with cells transduced with control adenovirus (Figure 2A). Despite protein mislocalization, A549 cells induced to express the α6 integrin 3’UTR adenovirus had increased levels of both α6 and β4 integrins at the surface (Figure 2B). We previously demonstrated α6β4 integrin is important for directed/processive cell migration. To assess the impact of expression of its 3’UTR alone, we performed individual cell motility assays on control and transduced cells

(Sehgal et al., 2006). A549s expressing adenovirus encoding the α6 integrin 3’UTR show increased processivity compared to control cells (Figure 2C). Therefore, these data support the notion that although overexpression of the α6 integrin 3’UTR severely inhibits α6β4 integrin localization to specific adhesion sites, an increase in surface α6β4 integrin promotes processive cell migration.

The α6 integrin 3’UTR is required for α6β4 integrin heterodimer formation in keratinocytes

So far, we have detailed studies using A549 cells, but we were interested in the function of the α6 integrin 3’UTR in other cell types. Thus, to extend our analyses, we used immortalized human epidermal keratinocytes (iHEKs) which assemble HD-like adhesions arranged in a

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rosette-like organization (Figure 3A). As with A549 cells, we infected iHEKs with adenovirus encoding the different α6 integrin constructs depicted in Figure 1A. To our great surprise, α6 integrin products of these viruses all co-localized with β4 integrin within HD-like adhesions, regardless of whether their mRNAs possessed a 3’UTR (Figure 3B). This is clearly contrary to the results we obtained with A549 cells (compare Figures 1C-D and 3B-C). We speculate that one possible reason for this contradiction may lie in the greater amount of mRNA encoding α6 integrin, as well as β4 integrin, in iHEK compared to A549 cells (Figure 3D). To test this possibility, we utilized a line of iHEKs in which we stably knocked down expression of endogenous α6 integrin (Kligys et al., 2012). Previous studies have demonstrated that these α6 integrin-depleted (α6KD) cells have a reduced expression of β4 integrin (Figure 3E-G) (Kligys et al., 2012). These α6KD cells were infected with adenovirus encoding GFP tagged α6 integrin either lacking the α6 integrin 3’UTR (α6KD+α6-GFP) or containing the α6 integrin full length

3’UTR (α6KD+α6-GFP-3’UTR). Interestingly, in α6KD+GFP-α6 cells processed for immunofluorescence, α6 integrin colocalizes with β1 integrin (Figure 3E). In contrast, in

α6KD+α6-GFP-3’UTR cells, α6 integrin is found associated with β4 integrin in HD-like arrays

(Figure 3E). These results were confirmed by western immunoblotting and immunoprecipitation

(Figure 3F). Immunoblotting and immunofluorescence reveals that β4 integrin was restored to a higher degree in α6KDs expressing the α6 integrin coding sequence with its 3’UTR compared to

α6KDs induced to express the α6 integrin coding sequence alone (Figure 3F-G). Taken together, these data imply that the 3’UTR of α6 integrin message is not only influencing α6 integrin localization but, in some way, is necessary for either promoting translation of β4 integrin message and/or stabilization of β4 integrin protein, thereby allowing for both assembly and subsequent localization of α6β4 integrin heterodimers. However, we are still left with the

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conundrum of why, in control iHEKs, the product of constructs encoding α6 integrin localize to

HD-like structures regardless of whether their mRNAs contain the α6 integrin 3’UTR but this is not the case in A549 cells. One explanation for this contradiction is due simply to cell differences in the way α6 integrin messages are translated. Alternatively, the difference may derive from the significantly larger amounts of endogenous α6 integrin mRNA in iHEKs versus

A549 cells (Figure 3D). We speculate that in iHEKs expression of α6 integrin mRNA lacking a

3’UTR associates with large quantity of endogenous α6 integrin message by some sort of piggy back mechanism, termed hitchhiking. This process was reported in Drosophilia oocytes and involves annealing of exogenous messages to endogenous messages via their 3’UTRs (Jambor et al., 2011). Regardless, in iHEKs, both messages may then be translated simultaneously, or at least in the same location, allowing for their products to be incorporated into HD-like adhesions.

In contrast, in A549s, α6 integrin messages without an appropriate 3’UTR fail to associate with the relatively small amounts of endogenous α6 integrin mRNAs and therefore do not facilitate targeting of α6 integrin protein to HD-related adhesions.

In summary, our data indicate that the α6 integrin 3’UTR is essential not only for the formation and localization of the α6β4 heterodimer to the cell-matrix adhesions but also its stability. In other words, we have uncovered a novel mechanism via which assembly of HDs and

HD-related adhesion devices is regulated. Further, we demonstrate that the localization of this heterodimer is dependent on a zipcode within the 3’UTR of the α6 integrin message, and this data parallel the study of α3 integrin whose message is targeted to FAs by its 3’UTR (Adereth et al., 2005). Our data also have implications for the “rescue” of cells and tissues of patients with skin diseases (junctional epidermolysis bullosa) in which α6 integrin protein is missing

(Schumann et al., 2013). The ability to restore α6 integrin by induction of exogenous message

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requires both the coding sequence and the α6 integrin 3’UTR to ensure both the assembly of

α6β4 integrin heterodimers and their targeting to sites of cell-matrix adhesions.

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Materials and methods

Cell culture conditions

A549s were cultured in HyClone MEM/EBSS media (HyClone, Logan, UT, USA) supplemented with 10% fetal bovine serum (Sigma-Aldrich, St. Louis, MO, USA), 2% L-glutamine, and 1% penicillin/streptomycin mixture. Immortalized human epithelial keratinocytes (iHEKs) were cultured in defined K-SFM keratinocyte media (DKSFM) (Thermo Fisher, Waltham, MA) supplemented with keratinocyte supplement and 1% penicillin/streptomycin mixture. Cells were maintained at 37°C in 8% humidified CO2.

Immunostaining

Cells were plated onto sterile coverslips. The next day, cells were fixed with 3.7% formaldehyde for 5 minutes, rinsed x3 with PBS, permeabilized with 0.05% triton-X 100 in PBS for 7 minutes, and rinsed x3 with PBS. Primary antibody was incubated on cells for 1 hour at 37°C and rinsed x3 with PBS. Secondary antibodies were incubated on cells for 1 hour at 37°C, rinsed x3 with

PBS, and mounted.

Adenoviral and lentiviral constructs

To generate the α6 integrin 3’UTR constructs tagged with GFP, the α6 integrin coding sequence,

GFP tag, and full length 3’UTR were cloned into the polylinker of the pENTR4 vector

(Clontech, Palo Alto, CA, USA). To generate the 3’UTR deletion constructs, point mutations were made at various sites in the 3’UTR to create AgeI restriction sites using the QuikChange II

XL site-directed mutagenesis kit (Agilent Technologies, Santa Clara, CA, USA). Constructs were subsequently digested with AgeI, gel purified with GeneJET Gel Extraction Kit

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(ThermoScientific, Waltham, MA, USA), and religated with Instant Stick-end Ligase Master

Mix (New England BioLabs, Ipswich, MA, USA). In addition, the α6 integrin 3’UTR was also ligated into the polylinker of the pENTR4 vector containing mCherry and deletions of the 3’UTR were carried out as described above. (Clontech, Palo Alto, CA, USA). All constructs were transformed into E. cloni 10G chemically competent cells (Lucigene, Middleton, WI, USA).

Sequences containing cDNA of α6 integrin with and without the 3’UTR were transferred from the ENTR4 vector into the pAD/CMV/V5-DEST vector using the LR recombination reaction

(Invitrogen, Carlsbad, CA, USA). The resultant adenoviral constructs were linearized with PacI enzyme and transfected into 293A cells using Lipofectamine 2000, following the manufacturer’s protocol (Invitrogen, Carlsbad, CA, USA). After 10 days, the crude viral lysate was harvested and used to amplify the adenovirus. The amplified viral stock was titered, and A549s were infected at a multiplicity of infection of 1:100 (α6 3’UTR mCherry) or 1:50 (α6 3’UTR GFP) in cell medium. The cells were used for analyses 48 h after infection.

To generate α6 knockdown keratinocytes, shRNA against α6 integrin was used as previously reported (Kligys et al., 2012) using the BLOCK-iT lentiviral RNAi expression system and individual clones isolated by serial dilution and selection in blastocidin (Invitrogen, Carlsbad,

CA, USA) (Kligys et al., 2012). α6 integrin protein was re-expressed in the knockdown keratinocytes by transduction of the cells with adenovirus expressing α6 integrin/GFP with and without the 3’UTR or GFP.

Flow cytometry

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Adherent cells were incubated with the α6 3’UTR mCherry adenovirus overnight. The next day, cells were trypsinized and fixed with 3.7% formaldehyde for 5 minutes. Cells were then rinsed

3x with PBS, incubated with primary antibody (1:200) and 30% normal goat serum at room temperature for 1 h, rinsed 3x with PBS, incubated with secondary antibody (1:200) at room temperature for 1 h, rinsed 3x with PBS, and analyzed with a Millipore GUAVA easyCyte flow cytometer (Hayward, CA, USA).

Single cell motility assays

Single cell motility was performed as previously described (Sehgal et al., 2006). Cells were plated at 5x104 cells per 35mm well 12-18 hours before imaging. Cells were observed for 3 hours and imaged every 5 minutes. Imaging was performed with a Leica DMi8 microscope

(Leica Microsystems, Buffalo Grove, IL, USA) and analyzed using MetaMorph software

(Molecular Devices, Sunnyvale, CA, USA). Processivity was calculated as detailed previously.

(Hiroyasu et al., 2016).

Immunoprecipitation and solubility assays

For immunoprecipitations of GFP constructs, adenovirus expressing GFP-tagged α6 integrin without and with portions of the 3’UTR was used to transduce 5x105 A549 cells at an MOI of

1:50 for 48 h. Cells were trypsinized, washed 3x with PBS, lysed with RIPA buffer (25mM Tris-

HCl, 150mM NaCl, 1% NP-40, 1% sodium deoxycholate, 0.1% SDS) supplemented with 1% protease inhibitor for 30 minutes at 4°C, and spun for 2 minutes to pellet cell debris. Lysate was removed and incubated with GFP beads for 4 h at 4°C. Beads were pelleted and washed x2 with

RIPA buffer, x1 with PBS, and resuspended in 1x Lamelli buffer supplemented with 10% BME.

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Samples were boiled for 4 minutes, centrifuged, and lysate was run on an SDS/PAGE gel as described below.

For solubility experiments, A549 cells were transduced as described above. After 48 h, cells were lysed in 1% Triton-X supplemented with 1% protease inhibitors and spun at 14,000 RPM to separate soluble and insoluble fractions. The soluble fraction was boiled in 1x Lamelli buffer supplemented with 10% BME. The insoluble fraction was incubated in urea/SDS sample buffer supplemented with 10% BME and sonicated. Lysates were run on SDS-PAGE gels as described below.

SDS-PAGE and immunoblot

Cell lysates were prepared for SDS-PAGE/western blotting as detailed elsewhere (Harlow and

Lane, 2014). Briefly, cells were lysed in 10% BME and 1% protease inhibitors in urea/SDS sample buffer, unless otherwise stated. Lysates containing comparable levels of protein were run on 7.5% or 15% gels and transferred to nitrocellulose for 1 h (15%) or 2 h (7.5%) at 110V.

Western blots were visualized using a myECL Imager (Thermo Fisher Scientific) and quantified using ImageJ. The Thermo Scientific Page Ruler Plus Prestained Protein Ladder was used to determine molecular weights.

Antibodies and immunofluorescent probes

Mouse monoclonal against β4 integrin (CD104) was purchased from BD Biosciences (San Jose,

CA, USA). Rat monoclonal against α6 integrin (MAB1378) and mouse monoclonal against β4 integrin (MAB1964) were purchased from EMD Millipore (Billenca, MA, USA). Rat monoclonal against α6 integrin (J1B5) was a generous gift from Dr. Caroline Damsky

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(University of California, San Francisco, USA). Rabbit monoclonal against GFP (G10362) was purchased from Life Technologies (Eugene, OR, USA). Rabbit antibody against lamin A/C

(2032S) was purchased from Cell Signaling Technology (Beverly, MA, USA). Fluorescein- conjugated goat anti-rabbit IgG antibody (111-095-144) and rhodamine-conjugated goat anti- mouse IgG antibody (109-025-003) were obtained from Jackson ImmunoResearch Laboratories

(West Grove, PA, USA). Alexa-647 conjugated phalloidin was purchased from Life

Technologies (Eugene, OR, USA). Horseradish peroxidase–conjugated goat anti-mouse IgG

(115-035-166) was purchased from Jackson ImmunoResearch Laboratories (West Grove, PA,

USA). Horseradish peroxidase–conjugated goat anti-rabbit IgG (7074S) was purchased from

Cell Signaling Technology (Beverly, MA, USA).

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References

Adereth, Y., Dammai, V., Kose, N., Li, R. and Hsu, T. (2005). RNA-dependent integrin α3 protein localization regulated by the Muscleblind-like protein MLP1. Nat. Cell Biol. 7, 1240–1247.

Andreassi, C. and Riccio, A. (2009). To localize or not to localize: mRNA fate is in 3′UTR ends. Trends Cell Biol. 19, 465–474.

Babic, I., Sharma, S. and Black, D. L. (2009). A Role for Polypyrimidine Tract Binding Protein in the Establishment of Focal Adhesions. Mol. Cell. Biol. 29, 5564–5577.

Colburn, Z. T. and Jones, J. C. R. (2016). α6β4 Integrin Regulates the Collective Migration of Epithelial Cells. Am. J. Respir. Cell Mol. Biol. 56, 443–452.

Colburn, Z. T. and Jones, J. C. R. (2018). Complexes of α6β4 integrin and vimentin act as signaling hubs to regulate epithelial cell migration. J Cell Sci 131, jcs214593. de Melker, A. A. and Sonnenberg, A. (1999). Integrins: alternative splicing as a mechanism to regulate ligand binding and integrin signaling events. BioEssays 21, 499–509.

Harlow, E. and Lane, D. (2014). Antibodies a laboratory manual. 2nd ed. (ed. Greenfield, E. A.) Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press.

Hiroyasu, S., Colburn, Z. T. and Jones, J. C. R. (2016). A hemidesmosomal protein regulates actin dynamics and traction forces in motile keratinocytes. FASEB J. 30, 2298–2310.

Hynes, R. O. (1992). Integrins: Versatility, modulation, and signaling in cell adhesion. Cell 69, 11–25.

Jambor, H., Brunel, C. and Ephrussi, A. (2011). Dimerization of oskar 3′ UTRs promotes hitchhiking for RNA localization in the Drosophila oocyte. RNA 17, 2049–2057.

Jones, J. C., Kurpakus, M. A., Cooper, H. M. and Quaranta, V. (1991). A function for the integrin alpha 6 beta 4 in the hemidesmosome. Cell Regul. 2, 427–438.

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Kligys, K. R., Wu, Y., Hopkinson, S. B., Kaur, S., Platanias, L. C. and Jones, J. C. R. (2012). α6β4 Integrin, a Master Regulator of Expression of Integrins in Human Keratinocytes. J. Biol. Chem. 287, 17975–17984.

Lee, E. C., Lotz, M. M., Steele, G. D. and Mercurio, A. M. (1992). The integrin alpha 6 beta 4 is a laminin receptor. J. Cell Biol. 117, 671–678.

Schumann, H., Kiritsi, D., Pigors, M., Hausser, I., Kohlhase, J., Peters, J., Ott, H., Hyla-Klekot, L., Gacka, E., Sieron, A. l., et al. (2013). Phenotypic spectrum of epidermolysis bullosa associated with α6β4 integrin mutations. Br. J. Dermatol. 169, 115–124.

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Figures and figure legends

Figure 1. The α6 integrin 3’UTR contains a zipcode which localizes the α6β4 integrin heterodimer to cell-matrix adhesions. A) Depiction of the α6 integrin-GFP 3’UTR constructs

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used in the studies detailed in the text. All constructs include the entire α6 integrin coding sequence tagged with GFP and various regions of the 3’UTR, indicated by grey blocks. The

3’UTR begins at base number 3233. B) Confocal immunofluorescence microscopical analyses of a control A549 cell co-stained with antibodies against α6 and β4 integrin subunits, as indicated.

Actin was visualized using Alexa-647 conjugated phalloidin. C) Confocal immunofluorescence microscopical analyses of A549 cells transduced with adenovirus encoding the constructs depicted in A. Cells were stained with antibodies against β4 integrin. D) Quantification of β4 integrin and GFP localization at the basal cell membrane of the indicated transduced cells. N = 3 independent experiments. >50 cells imaged per experiment. **, P<0.01.

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Figure 2. Overexpression of the α6 integrin 3’UTR impacts α6β4 integrin adhesions in transduced A549 cells. A) Confocal immunofluorescence microscopical analyses of A549 cells transduced with adenovirus either encoding mCherry (top panels) or α6 3’UTR-mCherry (bottom panels). The cells were stained with antibodies against α6 and β4 integrin as indicated. Note only one of the two cells in the bottom panel set expresses mCherry. This cell shows disruption in

α6β4 integrin localization. B) Flow cytometric analyses of α6 and β4 integrin in A549 cells transfected with adenovirus encoding α6 3’UTR-mCherry. C) Quantitation of single cell processivity of control A549 cells or A549 cells expressing adenovirus either encoding mCherry or α6 3’UTR-mCherry. N = 3 independent experiments. >50 cells imaged per experiment. ***,

P<0.001.

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Figure 3. The α6 integrin 3’UTR is responsible for α6β4 integrin heterodimer formation in

HEKs. A) Confocal immunofluorescence microscopical analyses of iHEKs stained with antibodies against α6 integrin and β4 integrin. Actin was visualized using Alexa-647 conjugated phalloidin. B) iHEKs transduced with adenovirus encoding a GFP-tagged α6 integrin construct, with and without a 3’UTR, as indicated, prepared for confocal immunofluorescence microscopical analyses using antibodies against β4 integrin. C) Quantification of GFP and β4 integrin co-localization at HD cell adhesion sites in iHEKs transduced with the indicated adenovirus. These are shown in Figure 1A. N = 3 independent experiments. 100 cells counted per experiment. D) Comparison of α6 integrin and β4 integrin mRNA levels in iHEKs and A549 cells by qPCR. N = 3 independent experiments. ***, P<0.001 E) Confocal immunofluorescence microscopical analyses of iHEKs, in which α6 integrin has been knocked down, transduced with virus encoding just the α6 coding sequence tagged with GFP (α6KD+α6-GFP). Cells were co- stained with either antibodies against β4 integrin (top panels) or β1 integrin (bottom panels). F)

Extracts of iHEKs, α6KD cells or α6KD cells transduced with either adenovirus encoding the α6 integrin coding sequence (α6KD+α6-GFP) or the α6 integrin coding sequence with its 3’UTR

(α6KD+α6-GFP+3’UTR) were prepared for immunoblotting using antibodies against α6 and β4 integrin as well as lamin A, for a loading control. G) Immunofluorescence of iHEKs, α6KD, and

α6KD+α6-GFP+3’UTR co-stained for α6 and β4 integrins. Panel G is reproduced from Kligys et al., 2012.

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Supplementary figure and supplementary figure legend

Figure S1. α6-GFP protein products interact with β4 integrin and are soluble regardless of whether their mRNAs possess the α6 integrin 3’UTR. A) A549 cells were infected with adenovirus constructs encoding GFP alone, GFP-tagged α6 integrin coding sequence, GFP- tagged α6 integrin coding sequence together with bases 3233 to 4133 of its 3’UTR and GFP-α6 integrin coding sequence together with bases 3233-5633 of its 3’UTR, as indicated. Extracts of the cells were prepared for immunoprecipitation using GFP beads and the resulting immunoprecipitates processed for immunoblotting using β4 integrin and GFP antibodies as indicated. Immunoprecipitated protein extracts are shown on the right and initial extracts (inputs) used for immunoprecipitation are shown on the left. β4 integrin is pulled down by GFP beads from all the extracts except cells infected with virus encoding GFP alone. In B) extracts of the

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same populations of viral infected A549 cells as in A were divided into soluble (S) and insoluble

(I) pools and both sets of pools were prepared for immunoblotting using antibodies against β4 integrin and lamin A. β4 integrin is found in the soluble pool while lamin is found in the insoluble fraction of the cells.

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CHAPTER FIVE: CONCLUSIONS AND FUTURE DIRECTIONS

Nesprin-2G studies

Conclusions

Skin wound healing is a complex process that requires cell migration of both epidermal and mesenchymal (fibroblast) cells which are found in the epidermis and dermis, respectively. In a full-thickness wound, both the dermis and epidermis are wounded. In these wounds, the dermis must be re-established before the epidermis. In the dermis, a blood clot forms and immune cells invade the region. Next, fibroblasts secrete extracellular matrix to form granulation tissue which eventually becomes the new dermis. Next, the process of re-epithelialization occurs where keratinocytes from the wound edge migrate across this provisional matrix to re-establish the barrier function of skin (Rittié, 2016). This process of cell migration is crucial post-wounding, and it requires a variety of cellular components including the nucleus and cell-matrix adhesions.

Nuclear positioning has become recognized as an important aspect of cell migration in both 2D and 3D culture systems. This nuclear positioning is dependent upon both the cytoskeletal network and transmission of traction forces. The cytoskeleton prevents abnormal deformations of the nucleus due to an indirect actin/nuclear interaction via nesprin proteins. The inability of the nucleus and cytoskeleton to interact leads to abnormal cell motility. Specifically, loss of nesprin-2G and their interacting partners at the inner nuclear membrane, SUN proteins, leads to reduced cell migration (Lüke et al., 2008; Rashmi et al., 2012; Zhang et al., 2009).

Traction forces are generated by a cell when a matrix adhesion tugs on the underlying matrix to promote cell migration. Interestingly, the nucleus resides where the traction forces cancel out, termed the point of maximum tension (PMT). In actively migrating cells, the PMT changes

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frequently, but the cell only takes minutes to move the nucleus to the new PMT location (Alam et al., 2015).

Cell-matrix adhesions are important in anchoring stable cells to their underlying matrix and aiding migrating cells by generating traction forces from tugging on the underlying substrate.

Two matrix adhesions were explored in this dissertation, namely hemidesmosomes (HDs) and focal adhesions (FAs). Only epithelial cells contain HDs, but most cells types have FAs, including epithelial cells. There is a vast amount of literature indicating how loss or over expression of many of these protein components can influence cell migration. The HD component the α6β4 integrin is important in the migration of keratinocytes (Jones et al., 1991).

Loss of paxillin, a component of FAs, leads to increased migration of endothelial cells (German et al., 2014). FAs promote cell migration by interacting with both the extracellular matrix and the nucleus, via actin, by generating traction forces. Actin is a known regulator of cell migration and

FA formation, and loss of actin leads to reduced migration and increased FA formation in mouse embryonic fibroblasts (Bunnell et al., 2011; Gardel et al., 2010).

Chapters 2 and 3 of this dissertation explored the role of nesprin-2G as a linker between the nucleus and cell-matrix adhesions in actively migrating primary mouse keratinocytes and fibroblasts. Previously, a research group published findings that keratinocytes isolated from nesprin-2G knockout (KO) cells showed reduced migration compared to wildtype (WT) cells

(Rashmi et al., 2012). However, we were unable to replicate these results in chapter 2 even though we used the same cell model. We found no difference between the migration capacity of

WT and KO keratinocytes. Moreover, we also found no change in HD or FA protein components in the same cells. In sharp contrast, we found considerable differences in the ability of nesprin-

2G KO fibroblasts to move compared with their WT counterparts. Our results are consistent with

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studies indicating that KO fibroblasts have a decreased ability to fill a wound created in vitro compared to WT fibroblasts (Lüke et al., 2008). In addition, unlike KO keratinocytes, KO fibroblasts possessed abnormal FAs. We are unable to explain the discrepancy of our and the previous study regarding KO keratinocyte motility (Rashmi et al., 2012). One possible explanation for the discrepancy may lie in the “keratinocyte” population assayed by Rashmi et al. (2012). This population may not have been was pure and may have contained fibroblasts, a common contaminant in primary epithelial cell cultures. In addition, we have no explanation for our finding that FAs are normal appearing in nesprin-2G KO keratinocytes but abnormal in KO fibroblasts. This requires a more detailed examination of FAs in KO cells and may be indicative of differences in FAs in keratinocytes versus fibroblasts.

Chapter 3 of this dissertation aimed to explore the underlying mechanism responsible for the migration defect in nesprin-2G KO fibroblasts. We observed that KO fibroblasts displayed reduced FA characteristics and an abnormal actin cytoskeleton. Specifically, FAs do not form as efficiently or mature properly. Next, we showed that KO fibroblasts had a reduced ability to generate traction forces compared to WT fibroblasts. We hypothesize that loss of nesprin-2G, which directly binds actin, causes a collapse of the actin cytoskeleton because it is no longer able to communicate with the nucleus via nesprin-2G force transduction. Overall, the results from chapters 2 and 3 provide a mechanistic insight into the wound healing deficit seen in these nesprin-2G KO mice (Lüke et al., 2008). We propose that the fibroblasts from the dermis, rather than keratinocytes from the epidermis, are responsible for this delayed wound closure in mice.

Future directions

While our work allowed us to derive a better understanding of the mechanism by which nesprin-2G influences motility, there are still many unexplored topics related to nesprin-2G. For

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example, why do we see no change in the motility of nesprin-2G KO keratinocytes compared to

WT cells while KO fibroblasts clearly exhibit migration defects? This may be due to a compensation mechanism regulated by keratinocyte HDs or their protein components, since these are not found in fibroblasts. Possibly this relates to the ability of HD proteins to regulate directed migration of keratinocytes (Jones et al., 1991). Another possibility is that the high amount of keratin in keratinocytes stabilizes cell shape, leading to stabilization of other cytoskeletal elements such as actin, thereby compensating for the loss of nesprin-2G.

That nesprin-2G KO leads to defective FA maturation suggests that a better understanding of signaling associated with the nesprin/actin interaction and how this signaling cascade leads to FA maturation is needed. We know that trans-factors, including FHOD1, are required to facilitate nesprin-2G/actin interaction, but there has been no study indicating how these factors are recruited (Kutscheidt et al., 2014). Preliminary studies indicate that the tugging of nesprin-2G on the nuclear membrane can increase transcription of certain genes (Luxton et al.,

2011; Neumann et al., 2010), but how this may influence the transcription of needs to be exploration.

Lastly, it will be important to determine how nesprin-2G interacts with the Golgi to orient cells during collective cell migration (Rashmi et al., 2012). Nesprin-1G has 2 sites which interact with the Golgi and are important in cell polarization (Gough et al., 2003). Because nesprins-1G and -2G have similar structures and interact with similar cytoskeletal elements, nesprin-2G may also interact with the Golgi. This interaction may, in turn, influence cell polarity and how FAs assemble. In this regard, although it is known that the Golgi apparatus determines cell polarity, how it does so remains unknown. Our results indicate that a thorough analysis of nesprin-2G-

Golgi interaction might lead to valuable new insights into how cell polarity is established.

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α6 integrin

Conclusions

As mentioned in chapters 1 and 5 of this dissertation, α6 integrin, together with its  integrin binding partner, is an important regulator of directed cell migration in keratinocytes

(Jones et al., 1991). In part, it does so by nucleating assembly of HDs (Hopkinson et al., 1998).

Yet, little is known about how α6β4 integrin heterodimers form and how assembled heterodimers are directed to their site of insertion into the plasma membrane. Our studies begin to answer these questions since they indicate that the 3’ untranslated region (UTR) of the α6 integrin mRNA contains a so-called zipcode which directs the localized incorporation of the α6β4 integrin heterodimer to HD-related matrix adhesion sites in A549 cells (Szostak and Gebauer,

2013). Specifically, in chapter 4 of this dissertation, we identified a zipcode located within bases

4770 to 5633 of the α6 integrin 3’UTR. This zipcode sequence is required for localized heterodimer incorporation into HD-related adhesions in A549s. Interestingly, the α6 integrin product of adenoviruses encoding α6 integrin message that does not include the zipcode still form heterodimers with β4 integrin in infected A549 cells, but the heterodimers fail to localize to

HD-related adhesions at the cell membrane.

We were interested in whether the importance of the α6 integrin 3’UTR was replicated in other cell types. Thus, we began experimentation with cultured immortalized keratinocytes

(iHEKs). Unlike the case of A549 cells, the α6 integrin product of adenovirus encoding α6 integrin message with no 3’UTR was incorporated into HD-like adhesions in infected iHEKs.

This was an unexpected result. However, we also noted that iHEKs express much more α6 integrin message than A549 cells and wondered whether this could explain the discrepancy in our results. To test this possibility, we used α6 integrin knockdown iHEKs (α6KD). These cells

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have little if any α6 and β4 integrin proteins (Kligys et al., 2012). Interestingly, when only the α6 coding sequence alone is re-expressed in α6KD cells by adenoviral means, α6β1, rather than

α6β4, integrin heterodimers form. α6β4 integrin heterodimers are only formed in α6KD cells and assemble into HD-like adhesions following α6KD cell infection with adenovirus encoding the α6 integrin coding sequence and its 3’UTR. This implies that the 3’UTR of α6 integrin is important in promoting β4 integrin protein expression as well as targeting the formed heterodimers to HDs.

Moreover, to explain why the protein product of α6 integrin message without a 3’UTR in iHEKs localizes normally, we speculate that the adenoviral-derived α6 integrin message piggy backs onto the large amount of endogenous α6 integrin message to facilitate not only α6β4 integrin heterodimer formation but also the incorporation of such heterodimers at HD-like sites of cell- matrix adhesion.

Future directions

Although we have identified a putative zipcode within the α6 integrin 3’UTR, we have yet to uncover how precisely this zipcode may function. Based on a prior report on targeting of

α3 integrin by protein interaction with tis 3’UTR, we suggest that the 3’UTR of α6 integrin may also possess protein binding sites. These proteins may be crucial for targeting message to sites of translation. We have begun preliminary studies into identifying proteins that bind the 3’UTR of

α6 integrin mRNA by running an RNA electromobility shift assay (REMSA). The α6 integrin

3’UTR is 2.5kb, but this assay requires RNA ~300 bases long so we generated ~250-300 base portions of the α6 integrin 3’UTR that were biotinylated, incubated with whole cell lysate, and run on a non-denaturing 6% polyacrylamide gel (Figure 1). The α6 integrin 3’UTR pieces that contain the zipcode sequence were the only sequences that specifically interacted with protein, indicated by an upshift in lanes 9 and 10. The nature of the protein or proteins that interact with

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the 3’UTR have not yet been determined. However, once candidates have been identified then it would be possible to assess their role in targeting message and α6β4 integrin proteins using genetic manipulations, including knockout via CRISPR or knock down approaches.

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References

Bunnell, T. M., Burbach, B. J., Shimizu, Y. and Ervasti, J. M. (2011). β-Actin specifically controls cell growth, migration, and the G-actin pool. Mol Biol Cell 22, 4047–4058.

German, A. E., Mammoto, T., Jiang, E., Ingber, D. E. and Mammoto, A. (2014). Paxillin controls endothelial cell migration and tumor angiogenesis by altering neuropilin 2 expression. J Cell Sci 127, 1672–1683.

Gough, L. L., Fan, J., Chu, S., Winnick, S. and Beck, K. A. (2003). Golgi Localization of Syne- 1. Mol Biol Cell 14, 2410–2424.

Jones, J. C., Kurpakus, M. A., Cooper, H. M. and Quaranta, V. (1991). A function for the integrin alpha 6 beta 4 in the hemidesmosome. Cell Regul 2, 427–438.

Luxton, G. W. G., Gomes, E. R., Folker, E. S., Worman, H. J. and Gundersen, G. G. (2011). TAN lines: a novel nuclear envelope structure involved in nuclear positioning. Nucleus 2, 173–181.

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Rittié, L. (2016). Cellular mechanisms of skin repair in humans and other mammals. J Cell Commun Signal 10, 103–120.

Szostak, E. and Gebauer, F. (2013). Translational control by 3′-UTR-binding proteins. Briefings in Functional Genomics 12, 58–65.

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Figure and figure legend

Figure 1. Representative REMSA. REMSA showing a transcript not binding (3947) and a transcript that specifically binds (5262) a protein in the whole cell lysate. Biotinylated RNA transcripts from α6 integrin 3’UTR are run alone (lanes 1, 6); with transcript and lysate (2, 7); transcript, lysate, and unbiotinylated RNA (lanes 3, 8); antisense biotinylated RNA, lysate, unbiotinylated RNA (4, 9); and antisense biotinylated RNA, lysate, unbiotinylated RNA x2 (5,

10).

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